pH-responsive bilayer hydrogel with sequential release of morin-based nanoparticles and bFGF for the treatment of the “ice and fire” wounds

Abstract

When the skin suffers from burns or frostbite, its normal structure and function are severely damaged, leading to problems such as inflammation, oxidative stress and difficulties in vascularization, which result in difficult wound healing. Currently, the existing treatment options have limitations in efficacy. The pH value of the burns and frostbite (B/F) wound microenvironment shows a dynamic change: it is acidic in the initial stage, gradually alkalizes as the repair process progresses, and then returns to a slightly acidic state in the later stage. Based on this, we designed a pH-responsive bilayer hydrogel with sequential release of morin-based nanoparticles and basic fibroblast growth factor (bFGF) for the treatment of B/F wounds. 4-hydroxyphenylboronic acid pinacol ester (PAPE) modified fucoidan (Fu), chitosan (Cs) and morin self-assembled to form nanoparticles (CFMNPs), which exhibited excellent pH sensitivity. Then, using methacrylated hyaluronic acid (MeHA) as the main component, taking advantage of the concentration difference and the property that MeHA is easily degraded in an acidic environment, a 0.8 % MeHA lower layer hydrogel loaded with CFMNPs and a 2 % MeHA upper layer hydrogel loaded with bFGF were prepared. In vitro experiments confirmed the pH responsiveness and cytocompatibility of the bilayer hydrogel. In vivo experiments on mouse B/F wounds verified that the bilayer hydrogel can achieve sequential release of active ingredients according to the pH changes of the B/F wound, effectively inhibit inflammation, resist oxidative stress, promote new blood vessel formation, and ultimately accelerate wound healing. This provides a highly potential and efficient new strategy for the treatment of B/F wounds.

Graphical abstract

In this study, a pH-responsive bilayer hydrogel was designed and fabricated by integrating an antioxidant flavonoid with a pro-angiogenic growth factor for the treatment of burn and frostbite (B/F) wounds. The bilayer hydrogel (bFGF-CFMN) comprises a lower methacrylated hyaluronic acid (MeHA, 0.8 %) layer loaded with morin self-assembled nanoparticles (CFMNPs) and an upper MeHA (2 %) layer loaded with basic fibroblast growth factor (bFGF). By leveraging the concentration dependent, pH-responsive degradation of MeHA and the pH-triggered release characteristics of CFMNPs, bFGF-CFMN bilayer hydrogel enables staged, precise modulation of the wound microenvironment. During the early acidic phase of wound healing, the lower layer degrades rapidly to release CFMNPs, which subsequently release morin as pH shifts toward alkaline, thereby suppressing inflammation and mitigating oxidative stress. As the wound progresses into the proliferative and remodeling phases, the upper layer gradually degrades under alkaline conditions, sustaining bFGF release to promote endothelial cell proliferation and migration, enhance angiogenesis, and support collagen deposition and re-epithelialization. Animal experiments have demonstrated that the bFGF-CFMN bilayer hydrogel can effectively suppress inflammation, resist oxidative stress, and facilitate the formation of new blood vessels, ultimately accelerating wound healing. In summary, the bFGF-CFMN bilayer hydrogel holds great potential for the treatment of B/F wounds.

1. Introduction

The skin, as the largest organ of the human body, plays a crucial role as a physical, chemical and microbial barrier. It possesses unique biological functions and wholeheartedly protects the host from external threats. However, it is extremely fragile. When the skin is subjected to physical injuries, such as burns and frostbite (B/F), the normal structure and function of the skin can be severely damaged [1,2]. In the case of burns, high temperatures can cause the denaturation of proteins and necrosis of cells in the skin tissue, thereby disrupting the skin's barrier function. Simultaneously, this increases the risk of infection [3]. In severe cases, it may lead to sepsis and endanger life [4]. Frostbite, on the other hand, occurs when low temperatures cause the skin blood vessels to constrict, slowing down blood flow and leading to tissue ischemia and hypoxia. Moreover, severe frostbite can lead to skin tissue necrosis, causing symptoms such as redness, blisters, and ulcers in the local skin [5,6]. Meanwhile, it also increases the risk of infectious diseases [7]. B/F not only cause direct damage to the skin but also significantly contribute to poor wound healing through factors such as inflammation, oxidative stress, and difficulties in vascularization [[8], [9], [10]]. After B/F occur, the damaged tissue initiates an inflammatory response. The persistent inflammatory response causes tissue edema and pain. Immune cells release a large amount of reactive oxygen species (ROS) [11], which exacerbates inflammation and hinders wound healing. At the same time, B/F damage the vascular endothelium, leading to vascular spasm and thrombus formation [12]. As a result, the damaged tissue lacks nutrients, has a reduced anti-infection ability, and has difficulty in excreting metabolites, causing difficulties in vascularization, delaying wound healing, and triggering complications [13]. Currently, comprehensive treatments are mainly adopted for B/F wounds, including early debridement, the use of topical medications containing antibiotics and growth factors, and skin grafting surgery [14,15]. However, the overall effectiveness of the existing treatment regimens still has certain limitations. Therefore, there is an urgent need for effective strategies to regulate the inflammatory response, alleviate oxidative stress, promote early endothelial cell regeneration, and reconstruct vasculature to solve the clinical problem of poor wound healing of B/F.

Morin, with its excellent anti-inflammatory and antioxidant properties, is demonstrating broad application prospects in the medical field [16,17]. Research has shown that morin exhibits significant anti-inflammatory and antioxidant effects in various disease models, such as ulcerative colitis, neurodegenerative diseases and diabetes [[18], [19], [20]]. For example, in cardiovascular diseases, morin can alleviate oxidative stress and inflammatory responses in vascular endothelial cells, improve vascular endothelial function [21]. However, the hydrophobicity and low bioavailability of morin limit its application [22]. Self-assembled nanoparticles represent a promising solution to overcome this limitation [23]. These nanoparticles are formed through non-covalent interactions, such as hydrogen bonds, van der Waals forces, and electrostatic interactions, which endow them with unique properties. Chitosan (Cs), a typical natural cationic polysaccharide prepared by the deacetylation of chitin, is an excellent material for nanoparticle carriers [24]. It has characteristics such as good biocompatibility, biodegradability, and low toxicity [25]. Cs can form self-assembled nanoparticles with morin through non-covalent interactions, reducing its degradation and inactivation in the body, prolonging its circulation time, and improving the bioavailability of morin [26]. However, the poor solubility of Cs at physiological pH limits its application as a nanoparticle carrier in the physiological environment to some extent [27]. Finding suitable auxiliary materials to cooperate with Cs is an effective solution. Fucoidan (Fu) is an excellent auxiliary material. It can interact with Cs through electrostatic adsorption and hydrogen bonds to form a more stable composite carrier system [28,29]. This not only enhances the drug encapsulation ability of Cs but may also further improve the sustained release performance of morin nanoparticles [30,31].

Growth factors play a crucial role in the process of wound healing. They belong to the class of polypeptides and possess the activity to stimulate cellular growth. When exogenous growth factors are added, they can activate the body's own “active” repair mechanism, facilitate the formation of granulation tissue, and thereby accelerate the rate of wound re-epithelialization [32]. Basic fibroblast growth factor (bFGF) is an important member of the growth factor family. It can promote the proliferation and migration of endothelial cells and stimulate the synthesis and secretion of the extracellular matrix, providing a material basis for the construction of new tissues [33,34]. Meanwhile, bFGF can promote angiogenesis and improve blood circulation in the wound area [34], showing great potential in the repair of B/F wounds.

Wound repair, especially the repair of complex wounds such as B/F, is a multi-stage dynamic process involving inflammatory responses, tissue proliferation, and remodeling [35]. The pH changes in the wound microenvironment are closely related to each repair stage, exhibiting specific patterns [36]. In the case of B/F wounds, local tissue cells are damaged, resulting in ischemia and hypoxia. The intracellular metabolic process shifts from aerobic metabolism to anaerobic metabolism, generating a large amount of acidic metabolites, such as lactic acid, which leads to a decrease in the wound pH (pH 5.5–6.5) [37]. As the repair response to the injury is initiated, inflammatory cells infiltrate, and neutrophils and other cells release alkaline substances, causing an increase in the pH of the wound surface (pH 7.4–8.0) [38]. As the wound remodels and heals, the wound pH will further approach that of normal skin (pH 6.5–7.0) [39,40]. Therefore, how to adapt to the dynamic changes in the wound microenvironment through a pH-responsive degradation and release strategy to achieve precise temporal drug delivery and thus regulate the biological processes at different repair stages is a key challenge and research hotspot in the current field of wound repair.

In recent years, hydrogels have shown great application potential and unique advantages as wound dressings in the field of wound repair [41,42]. Hydrogels are highly hydrophilic, capable of absorbing a large amount of fluid exuded from the wound to maintain a moist wound environment [43]. Meanwhile, they can also carry various therapeutic components, such as nanoparticles and growth factors. Hyaluronic acid (HA), as a natural polysaccharide biological macromolecule, has significant advantages in constructing hydrogel wound dressings [44]. On the one hand, HA itself has good biocompatibility and biodegradability. On the other hand, HA can serve as an excellent carrier for therapeutic components. While ensuring the activity of the loaded drugs, it can improve their dispersibility and stability in the hydrogel and achieve the ordered release of therapeutic components as HA degrades [45]. In addition, an acidic environment promotes the hydrolysis reaction of HA and accelerates its degradation process [46,47]. However, most existing HA-based hydrogel wound dressings lack the ability to precisely respond to the dynamic changes in the microenvironment during the wound healing process. They are unable to achieve spatiotemporal controlled release of different therapeutic components according to different stages of wound healing. Moreover, drug delivery often shows randomness, making it difficult to precisely exert therapeutic effects on the wound microenvironment at specific stages, thus limiting the overall therapeutic efficacy. Therefore, our research innovatively designed a pH-responsive bilayer hydrogel. This hydrogel solves the key problems in the treatment of B/F wounds through three key innovations: First, pH-responsive morin-loaded nanoparticles (CFMNPs) can specifically release morin in the acidic microenvironment of early wounds. Second, a bilayer hydrogel composed of different concentrations of methacrylated hyaluronic acid (MeHA) can sequentially release CFMNPs in an acidic environment and release bFGF in an alkaline environment. Third, a synergistic treatment strategy combines morin (with anti-inflammatory/antioxidant effects in the early stage) and bFGF (promoting angiogenesis in the later stage) to address the challenges in the early and later stages of B/F wound healing. By integrating pH-responsive nanotechnology, bilayer hydrogel design and multi-drug therapy, the hydrogel we developed achieves stage-based and spatiotemporal controlled treatment. This represents a significant advancement compared to previous methods that lacked pH sensitivity, sequential release ability, and comprehensive treatment coordination.

Based on this, considering the dynamic changes in the pH of B/F wounds, we developed a pH-responsive bilayer hydrogel with sequential release based on morin nanoparticles-bFGF for the treatment of B/F. First, 4-hydroxyphenylboronic acid pinacol ester (PAPE) was used to modify Fu to form a pH-responsive material [48]. Then, through self-assembly, it was combined with Cs to form CFMNPs. HA was first methacrylated to obtain MeHA. Taking advantage of the fact that MeHA is more easily degraded in an acidic environment and the concentration difference, a 0.8 % MeHA was used as the lower layer hydrogel to load CFMNPs (CFMN) and a 2 % MeHA was used as the upper layer hydrogel to load bFGF (bFGF). The two layers together formed a bilayer hydrogel (bFGF-CFMN), which was designed to adapt to the dynamic changes in wound pH and achieve sequential release of effective therapeutic components (Fig. 1A and C). In short, in the early acidic stage of B/F wounds, the low-concentration MeHA in the lower layer degrades and releases CFMNPs. As the wound healing progresses and the wound gradually becomes alkaline, the CFMNPs further release anti-inflammatory and antioxidant morin. Meanwhile, the high-concentration MeHA in the upper layer slowly releases bFGF in the later alkaline wound stage, continuously promoting wound healing (Fig. 1B). We conducted in vitro and in vivo experiments, which verified that the bFGF-CFMN bilayer hydrogel can achieve sequential release of effective components according to the pH changes in B/F wounds. It effectively inhibits inflammation, resists oxidative stress, promotes the proliferation and migration of endothelial cells, promotes the formation of new blood vessels, and ultimately accelerates wound healing. In summary, the pH-responsive bilayer hydrogel based on morin nanoparticles-bFGF, with its ingenious design and excellent performance, provides a highly potential and efficient new strategy for the treatment of B/F wounds.

Fig. 1.

Schematic diagram of a bFGF-CFMN bilayer hydrogel for the treatment of B/F wounds. (A). Schematic illustration of the synthesis process of bFGF-CFMN bilayer hydrogel. (B). Mechanistic process of bFGF-CFMN bilayer hydrogel in the treatment of B/F wounds. (C). The chemical structures of the raw materials for synthesizing the bFGF-CFMN bilayer hydrogel and the relevant synthetic routes.

2. Materials and methods

2.1. Materials

Hyaluronic acid (HA, Mw: 150–250 kDa, purity: 97 %), N,N-dimethylformamide (DMF, purity: 99 %), 4-hydroxyphenylboronic acid pinacol ester (PAPE, purity: 97 %), 1-(3-dimethylaminopropyl)-3-ethylcarbodiimide (EDC, purity: 99 %), 4-dimethylaminopyridine (DMAP, purity: 99 %) and anhydrous formamide were obtained from Macklin (Shanghai, China). Fucoidan (Fu, Mw: 242 Da, purity: 98 %), chloroacetic acid (purity: 99 %), dihydroethidium (DHE, 10 mM in DMSO) and NaOH (purity: 95 %) were purchased from Aladdin (Shanghai, China). Morin (purity: 95 %), chitosan, methacrylic acid (purity: 99 %), lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP, purity: 95 %), fluorescein isothiocyanate (FITC, purity: 90 %) and Rhodamine B (purity: 95 %) were purchased from Sigma-Aldrich (St. Louis, USA). Basic fibroblast growth factor (bFGF), dimethyl sulfoxide (DMSO, purity: 99 %), live/dead viability assay kit, ROS assay kit, EdU cell proliferation kit, 4’,6-diamidino-2-phenylindole (DAPI) staining solution, cell counting kit-8 (CCK-8) and lactate assay kit were purchased from Beyotime (Shanghai, China). Lipopolysaccharide (LPS, purity: 98 %) and phosphate buffered saline (PBS) were obtained from Solarbio (Beijing, China). Mouse TNF-α enzyme-linked immunosorbent assay (ELISA) kit, mouse IL-1β ELISA kit, mouse IL-6 ELISA kit, mouse IL-10 ELISA kit, mouse VEGF ELISA kit, mouse PDGF ELISA kit and mouse TGF-β ELISA kit were all purchased from MultiSciences Biotech Co., Ltd (Hangzhou, China). Mouse Ki67 ELISA kit was obtained from Jianglai Biotechnology Co., Ltd (Shanghai, China). CD31 rabbit antibody (ab9498) and VEGF rabbit antibody (ab32152) were purchased from Abcam (Cambridge, UK). Superoxide dismutase (SOD) assay kit (WST-1 method), malondialdehyde (MDA) assay kit (TBA method) and reduced glutathione (GSH) assay kit were purchased from Nanjing Jiancheng Bioengineering Institute (Nanjing, China). Human umbilical vein endothelial cells (HUVECs) and RAW 264.7 macrophages were purchased from Pricella Biotechnology Co., Ltd (Wuhan, China).

2.2. Preparation of CFMNPs

First, 1.7 g Fu was dissolved in 20 mL of ultrapure water. Subsequently, 3.2 g NaOH and 3.7 g chloroacetic acid were added sequentially. After stirring at room temperature for 12 h, the reaction mixture was neutralized with 2 M HCl solution. Then, the mixture was dialyzed using a dialysis bag (MWCO 1000) for 3 days, with the external water being changed 6 times a day. Finally, the dialyzed product was collected and freeze-dried for 3 days to obtain the freeze-dried product Fu-COOH [49].

100 mg Fu-COOH was added to a 50 mL round bottom flask and 20 mL anhydrous formamide was added. The mixture was stirred for about 30 min until Fu-COOH was completely dissolved. Then, 30 mg DMAP was added to the above solution and the solution was stirred until fully dissolved. Subsequently, 50 mg EDC was added and stirred for 15 min to ensure its full dissolution and carboxyl group activation. Finally, 50 mg of PAPE was added to the above mixture. The reaction flask was sealed and the reaction was carried out under stirring in the dark at room temperature for 24 h. After the reaction was completed, the reaction mixture was transferred to a pre-prepared dialysis bag (MWCO 1000) and dialyzed for 3 days, with the external water being changed 8 times a day. After dialysis, the final mixture was collected and freeze-dried to obtain Fu -PAPE [48].

Equal volumes of a 5 mg/mL Cs solution (prepared in a 2 % acetic acid solution) and a 5 mg/mL Fu-PAPE aqueous solution were mixed. Then, a 5 mg/mL morin solution (pre-dissolved in DMSO) was added. The mixture was sonicated (power: 100 W, frequency: 40 kHz) for 1 h to promote the self-assembly of morin with Cs and Fu-PAPE, followed by continuous stirring for another 1 h. Subsequently, the mixture was dialyzed using a dialysis bag (MWCO 3000) for 6 h. After dialysis, the solution was collected and filtered through a 0.45 μm filter. After filtration, the filtrate was centrifuged at 15000 rpm for 15 min to collect the CFMNPs precipitate. Finally, the precipitate was freeze-dried to obtain CFMNPs.

2.3. Characterization of CFMNPs

The morphology and structure of CFMNPs were observed using a transmission electron microscope (TEM, Hitachi HT 7800, Japan). The particle sizes of CFMNPs were measured using a Malvern particle size analyzer (Malvern Zetasizer Nano ZS 90, UK). The infrared spectrum of CFMNPs was measured using a Fourier transform infrared reflection (FTIR) spectrometer (Nicolet iS50, Thermo Fisher).

Next, high performance liquid chromatography (HPLC, Agilent HPLC 1260, USA) was used to determine the encapsulation efficiency (EE) and loading efficiency (LE) of morin in CFMNPs. Before HPLC analysis, methanol was used to disrupt the structure of CFMNPs and dissolve morin. Specifically, 10 μL the sample was injected into a C18 chromatographic column (250 × 4.6 mm, 5 μm) and heated to 37 °C. Then, the sample was analyzed at a wavelength of 370 nm using a mobile phase consisting of 50:50 v/v methanol purified water at a flow rate of 1.0 mL/min [19].

The EE and LE were calculated using the following formulas:

$$\text{EE}(\%)=\left(\text{Weight}\text{of}\text{loaded}\text{morin}/\text{Total}\text{weight}\text{of}\text{added}\text{morin}\right)\times 100\%$$

$$\text{LE}(\%)=\left(\text{Weight}\text{of}\text{loaded}\text{morin}/\text{Total}\text{weight}\text{of}\text{added}\text{CFMNPs}\right)\times 100\%$$

2.4. pH-responsive release experiment of CFMNPs

CFMNPs were dispersed in PBS with different pH values, namely PBS solutions at pH 6, pH 7 and pH 8. A 5 mg/mL CFMNPs solution was added into a dialysis bag (MWCO 3000, Millipore). The solution was stirred at room temperature, and then the supernatant was taken at predetermined time points for HPLC analysis to determine the concentration of morin.

2.5. Synthesis of bFGF-CFMN bilayer hydrogel

1 g HA was dissolved in a mixed solvent of 30 mL DMF and 60 mL pure water, and the mixture was stirred until HA was completely dissolved. Then, under ice-bath conditions, 3 mL methacrylic acid was slowly added dropwise, and the reaction was continued for 24 h. After the reaction is completed, adjust the pH of the solution to 8–9 with NaOH. Subsequently, three times the volume of pre-cooled absolute ethanol was added to precipitate the product. After precipitation, the supernatant was removed, the precipitate was centrifuged at 5000 rpm for 10 min, the supernatant was removed again, an appropriate amount of pure water was added to dissolve the precipitate, and it was placed in dialysis at 4 °C for 3 days, with the water being changed 3 times a day. Then, the dialyzed aqueous solution was collected and freeze-dried to finally obtain MeHA solid.

MeHA was dissolved in pure water to obtain 0.8 % (w/v) and 2 % (w/v) MeHA solutions. After it was completely dissolved, the LAP photoinitiator (0.25 %, w/v) was added and the solutions were exposed to ultraviolet light for 30 s to form hydrogels. The drug loading capacities of morin and bFGF were determined with reference to previous studies [16,50]. The bilayer hydrogel was prepared as follows: first, 2 % MeHA was added into the mold, then bFGF (100 μg per milliliter of the hydrogel system) was added, and it was irradiated with an ultraviolet light source for 30 s. Subsequently, 0.8 % MeHA and CFMNPs (10 mg per milliliter of the hydrogel system) were added, and it was irradiated with the ultraviolet light source again for 45 s to obtain the bFGF-CFMN bilayer hydrogel.

A blank bilayer hydrogel without bFGF and CFMNPs (Blank) was prepared following similar steps as described above.

2.6. Characterization of bFGF-CFMN bilayer hydrogel

The structural characteristics of the bFGF-CFMN bilayer hydrogel were observed using a scanning electron microscope (SEM, ZEISS GeminiSEM 360, Germany). Briefly, the hydrogel was first fractured in liquid nitrogen and then freeze-dried. Subsequently, the cross-section was fixed on the sample stage. After sputtering a thin layer of gold on its surface, SEM imaging was performed. The bFGF-CFMN bilayer hydrogel, after thorough freeze-drying, was characterized using FTIR (Thermo Fisher Scientific Nicolet iS 20, USA).

2.7. Rheological properties

The rheological behavior of the bFGF-CFMN bilayer hydrogel was tested using a rheometer (Discovery HR 30, USA) equipped with a parallel-plate geometry with a diameter of 25 mm. To determine the elasticity of the hydrogel, the storage modulus (G′) and loss modulus (G″) of the bFGF-CFMN bilayer hydrogel were recorded during an angular frequency sweep from 0.1 to 100 rad/s at room temperature with a constant strain of 1 %. To determine the viscosity of the bFGF-CFMN bilayer hydrogel, the viscosity values were recorded during a shear-rate sweep from 0.1 to 100 s⁻¹ at room temperature with a constant strain of 1 %.

2.8. pH-responsive degradation experiment of bFGF-CFMN bilayer hydrogel

The bFGF-CFMN bilayer hydrogels were immersed in PBS with different pH values, namely PBS solutions with pH 6, pH 7, and pH 8. The hydrogels were incubated at 37 °C. At predetermined time points, the hydrogels were taken out, freeze-dried, and weighed. The degradation rate of the hydrogel was calculated according to the following formula.

Degradation rate (%) = [(The initial weight of the hydrogel − The weight of the hydrogel at the specified time) / The initial weight of the hydrogel] × 100 %

2.9. pH-responsive release experiment of bFGF-CFMN bilayer hydrogel

The bFGF-CFMN hydrogels were immersed in PBS with different pH values, namely PBS solutions with pH 6, pH 7, and pH 8. The samples were incubated at 37 °C. At predetermined time points, the supernatants after hydrogel immersion were taken out. The contents of morin and bFGF were detected by HPLC and a bFGF ELISA kit, respectively. The release rates of morin and bFGF were calculated according to the following formulas.

$$\text{Morin}\text{release}\text{rates}(\%)=\left[\left(\text{The}\text{initial}\text{concentration}\text{of}\text{morin}-\text{The}\text{concentration}\text{of}\text{morin}\text{in}\text{the}\text{supernatant}\text{at}\text{the}\text{specified}\text{time}\right)/\text{The}\text{initial}\text{concentration}\text{of}\text{morin}\right]\times 100\%$$

$$\text{bFGF}\text{release}\text{rates}(\%)=\left[\left(\text{The}\text{initial}\text{concentration}\text{of}\text{bFGF}-\text{The}\text{concentration}\text{of}\text{bFGF}\text{in}\text{the}\text{supernatant}\text{at}\text{the}\text{specified}\text{time}\right)/\text{The}\text{initial}\text{concentration}\text{of}\text{bFGF}\right]\times 100\%$$

2.10. Cellular cytotoxicity assessment

The CCK-8 assay was employed to evaluate the impact of bFGF-CFMN bilayer hydrogel on the viability of HUVECs [51]. Briefly, the cells were seeded into 96-well plates at a density of 3.0 × 10³ cells per well. In the experimental group, 10 μL hydrogel (prepared from 10 μL hydrogel solution; subsequent steps are similar) was added to each well, while the control group received an equal volume of PBS. At 24 h and 72 h after treatment, 10 μL CCK-8 solution was added to each well, and the cells were incubated for 1 h. The absorbance at 450 nm was measured using a microplate reader (Thermo Fisher Scientific Varioskan LUX, USA) to assess cell viability. Meanwhile, the live/dead staining method was used to evaluate the cytotoxicity of HUVECs at 24 h and 72 h. In brief, HUVECs were stained with calcein-AM and propidium iodide (PI), and then observed under a fluorescence microscope. The fluorescence intensity was detected to determine cell viability.

2.11. Scratch assay

The scratch assay was employed to evaluate the migratory effect of bFGF-CFMN bilayer hydrogel on HUVECs to simulate the wound healing process. HUVECs were cultured in 6-well plates until they reached approximately 100 % confluence, and a 200 μL pipette tip was used to scratch the cell monolayer. In the experimental group, 200 μL of the corresponding hydrogel was added to each well, while the control group received an equal volume of PBS. Subsequently, the cells were incubated in serum-free Dulbecco's Modified Eagle Medium (DMEM) for 6 and 12 h. Cell migration was captured using a microscope, and the wound width at each time point was analyzed using ImageJ to calculate the migration rate.

$$\text{Migration}\text{rate}(\%)=\left[\left(\text{Initial}\text{scratch}\text{width}-\text{Scratch}\text{width}\text{at}\text{the}\text{specified}\text{time}\right)/\text{Initial}\text{scratch}\text{width}\right]\times 100\%$$

2.12. Tube formation assay

Fifty microliters of Matrigel was evenly coated on a 24-well plate and incubated at 37 °C for 30 min. HUVECs were seeded at a density of 1 × 10⁵ cells per well. In the experimental group, 200 μL of the corresponding hydrogel was added to each well, while the control group received an equal volume of PBS. After 8 h incubation, HUVECs were stained with calcein-AM, and tube formation was observed using a fluorescence microscope.

2.13. In vitro antioxidant evaluation

RAW 264.7 cells were seeded in 24-well plates at a density of 1 × 10⁵ cells per well. Serum-free DMEM containing 50 μM 2′,7′-dichlorodihydrofluorescein diacetate (DCFH-DA) was added, and the cells were incubated in the dark for 30 min. Then, the cells were washed three times with PBS. Subsequently, each well was treated with 500 μL of a 50 μg/mL ROS inducer (Rosup) for 1 h. In the experimental group, 200 μL of the corresponding hydrogel was added, while the control group received an equal volume of PBS. After 2 h incubation in the dark, the cells were washed three times with PBS, then stained with DAPI for 5 min. Finally, the fluorescence intensity of cells in each group was observed using a confocal fluorescence microscope.

2.14. In vitro anti-inflammatory evaluation

RAW264.7 cells were seeded in 24-well plates at a density of 1 × 10⁵ cells per well and treated with 1 μg/mL LPS for 24 h to induce an inflammatory environment. In the experimental group, 200 μL of the corresponding hydrogel was added to each well, while the control group received an equal volume of PBS. After 8 h of incubation, the cell supernatants were collected, centrifuged at 1000 rpm for 5 min, and the supernatants were then taken. The concentrations of pro-inflammatory cytokines (including TNF-α, IL-1β and IL-6) were measured using an ELISA.

2.15. Cell proliferation assay

HUVECs were seeded in 6-well plates at a density of 3 × 10⁵ cells per well. In the experimental group, 200 μL of the corresponding hydrogel was added to each well, while the control group received an equal volume of PBS. After 8 h of incubation, the old medium was aspirated, and the cells were washed three times with PBS. Then, 2 mL medium containing 10 μM EdU working solution was added to each well, and the cells were further incubated for 2 h. Subsequently, the old medium was removed. The cells were fixed with 4 % paraformaldehyde for 15 min, permeabilized with a permeabilization solution for 15 min, and then incubated with an endogenous peroxidase blocking solution at room temperature for 20 min. Next, 0.5 mL Click reaction solution (provided in the kit) was added to each well. The culture plate was gently shaken to ensure that the reaction mixture evenly covered the samples, followed by incubation at room temperature in the dark for 30 min. After the incubation, the Click reaction solution was aspirated, and the cells were washed three times with the washing solution. Then, 0.2 mL DAB chromogenic solution was added to each well and incubated at room temperature for 15 min. After washing with PBS, the cells were stained with DAPI dye for 5 min. Finally, the fluorescence of cells in each group was observed using a fluorescence microscope.

2.16. Animal experiments

In this study, 30 healthy male C57BL/6 mice aged 6–8 weeks were selected. The mice were numbered and a random number table was used for random grouping to divide the mice into five groups: (1) Control group, (2) Blank group (treated with blank hydrogel), (3) CFMN group (treated with CFMN hydrogel), (4) bFGF group (treated with bFGF hydrogel) and (5) bFGF-CFMN group (treated with bFGF-CFMN bilayer hydrogel). There were six mice in each group. Blinding was implemented in this study. The researchers who evaluated the results were unaware of the grouping information of the mice. After anesthesia with 3 % sodium pentobarbital (50 mg/kg), all mice were shaved and depilated and their skin was disinfected with povidone-iodine. A 10 mm skin punch was used to create a full-thickness circular skin wound with a diameter of 10 mm on the dorsal surface of the mice. Burns and frostbite wounds were created using a heated metal rod and liquid nitrogen, respectively. Briefly, for burn wounds [52], a metal rod with a flat tip and a diameter of 10 mm was used. It was heated to 100 °C by a heating device with precise temperature control. Then, the tip of the metal rod was brought into contact with the wound surface for 5 s (depth: 2.4 mm); for frostbite wounds [53,54], a 10 mm stainless steel coin was placed in liquid nitrogen at a temperature of - 196 °C for 2 min to ensure it was fully cooled. After removal, it was pressed onto the wound surface for 5 s (depth: 2.4 mm). The day when the wounds were created was recorded as day 1. A circular hydrogel with a diameter of 10 mm was applied to the dorsal wound each mouse. On days 1, 3, 5, 10, and 14, wound images were taken using a camera and the wound area was measured using ImageJ software. The wound healing rate was calculated using the following formula:

$$\text{Wound}\text{healing}\text{rate}(\%)=\left[\left(\text{the}\text{initial}\text{wound}\text{area}-\text{the}\text{actual}\text{wound}\text{area}\text{at}\text{the}\text{specified}\text{time}\right)/\text{the}\text{initial}\text{wound}\text{area}\right]\times 100\%$$

2.17. Detection of the pH value of wound tissues

A needle-type pH microsensor (PreSens Precision Sensing GmbH, NTH-HP5, Germany) was used by inserting its needle into the wound tissues to measure the pH values of the wounds in each group at different time points.

2.18. Detection of lactic acid content in wound tissues

Wound tissue samples were collected. According to the ratio of adding 100 μL of lactate assay buffer (provided by the kit) for every 10 of mg tissue, the tissues were minced and homogenized at 4 °C. The homogenate was centrifuged at approximately 12,000×g for 5 min at 4 °C, and the supernatant was collected for subsequent detection. A lactic acid detection kit (Beyotime, China) was used to measure the lactic acid content in the supernatant according to the manufacturer's instructions.

2.19. In vivo degradation experiment

Rhodamine B (0.1 mg/mL) and FITC (0.5 mg/mL) were mixed into the pre-solutions of the lower and upper layer hydrogels, respectively, and the dual-fluorescently labeled bFGF-CFMN bilayer hydrogel was prepared using the aforementioned method. Subsequently, the hydrogel was applied to the B/F wounds on the backs of mice, respectively. On days 1, 3, 5, and 7, the bFGF-CFMN bilayer hydrogel at the wound sites was detected using an in vivo imaging system (IVIS Spectrum, PerkinElmer, USA), and the fluorescence intensity was quantitatively analyzed.

2.20. In vivo drug release experiment

The bFGF-CFMN bilayer hydrogel was applied to the B/F wounds on the backs of mice, respectively. Then, on days 1, 3, 5, and 7, the undegraded hydrogel was removed from the wounds and immersed in PBS. The pH of the solution was adjusted to completely degrade the hydrogel. The supernatant of the above mentioned mixture was collected, and the contents of morin and bFGF were measured to evaluate the drug release.

2.21. Histological staining of wound tissues

On days 5 and 15, some mice were euthanized, and the wound tissue samples from the dorsal side of the mice were collected and fixed in 4 % paraformaldehyde. After routine histological processing, the tissues were embedded in paraffin and sectioned into 4 μm slices. The slices were routinely stained with hematoxylin and eosin (HE) for histological examination, and Masson staining was used to evaluate the collagen deposition in the wounds.

For immunohistochemistry (IHC), the dewaxed and rehydrated slices were subjected to antigen retrieval, and then non - specific binding sites were blocked. Subsequently, the slices were incubated with anti-VEGF primary antibody (1:100) overnight at 4 °C. After washing, the slices were incubated with the corresponding secondary antibody for 1 h at 25 °C. Finally, the slices were observed under an optical microscope.

For tissue immunofluorescence staining (IF), the dewaxed and rehydrated slices were subjected to antigen retrieval, and then non - specific binding sites were blocked with a blocking solution. After that, the slices were incubated with anti-CD31 primary antibody (1:200) overnight at 4 °C. After washing, the slices were incubated with a fluorescence-labeled secondary antibody for 1 h at 25 °C in the dark. After further washing, the cell nuclei were counterstained with DAPI. After mounting, the slices were observed under a fluorescence microscope.

2.22. Oxidative stress level in wound tissues

The wound tissues were embedded in OCT compound and frozen in liquid nitrogen. Then, they were cut into frozen sections with a thickness of 8 μm. The sections were placed at room temperature and fixed in 4 % paraformaldehyde for 1 h. Subsequently, in the dark, the sections were first stained with 20 μM DHE for 1 h, and then stained with DAPI for 5 min. Finally, images were captured using a fluorescence microscope.

2.23. Detection of oxidative stress substance concentrations in wound tissues

Twenty mg of wound tissue was accurately weighed. Mince the tissue and grind it on ice to prepare a homogenate. Centrifuge the homogenate at 4000 rpm for 10 min and collect the supernatant for subsequent analysis. According to the manufacturer's instructions, use the corresponding kits to measure the activity of SOD, and the contents of GSH and MDA in the supernatant.

2.24. Cytokine concentrations in wound tissues

Ten mg of wound tissue was collected and weighed. Subsequently, the wound tissue samples were cut into small pieces and ground on ice, followed by ultrasonic disruption of the ground tissue. Finally, the homogenate was centrifuged at 8000 rpm for 15 min at 4 °C, and the supernatant was collected for analysis. The levels of TNF-α, IL-1β, IL-6, IL-10, VEGF, PDGF, TGF-β and Ki67 in the supernatant were quantitatively determined using the corresponding ELISA kits. All procedures were carried out in accordance with the manufacturer's instructions.

2.25. Statistical analysis

Experimental data are presented as the mean ± standard deviation (SD). The normal distribution of all data was assessed using the Shapiro-Wilk test (significance value of 0.05). If the data passed the normality test, one-way analysis of variance (ANOVA) followed by Tukey's post hoc test was used to analyze the differences between groups; if not, Kruskal-Wallis test followed by Dunn's post hoc test was used to analyze the differences between groups. The graphs and statistical analysis were carried out using GraphPad Prism software, version 8, SPSS 25.0 software and OriginPro 2019. Statistical significance is indicated as follows: ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001, ∗∗∗∗p < 0.0001; “ns” indicates no significant difference. Each experiment was repeated at least three times (n ≥ 3).

3. Results and discussion

3.1. Preparation and characterization of CFMNPs

CFMNPs were prepared using a self-assembly method. Briefly, Fu was modified with PAPE to endow it with pH-responsive ability. Then, Cs, Fu-PAPE and morin self-assembled through stirring and sonication to form nanoparticles, namely CFMNPs (Fig. 2A). Due to intermolecular hydrogen bonds and π-π stacking interactions, morin has poor solubility in water, often presenting as a turbid suspension and is prone to precipitation [55], which greatly limits its application in the biomedical field. However, when morin participates in the self-assembly process, Cs and Fu-PAPE bind to morin through hydrogen bonds and electrostatic interactions, breaking its aggregation force and effectively improving its solubility in water [56,57]. It can be observed that the CFMNPs solution is more homogeneous and transparent than the morin solution (Fig. 2B). Subsequently, the obtained CFMNPs were further characterized to comprehensively explore their structural features and potential application performance. TEM was used to observe the micro-morphology of CFMNPs, revealing a relatively uniform spherical structure with a diameter of approximately 123.94 ± 5.63 nm (Fig. 2C). Then, a nanoparticle size analyzer revealed that the diameter of CFMNPs was approximately 154.63 ± 24.20 nm, and the polydispersity index (PDI) was 0.25 ± 0.09, indicating a uniform size distribution (Fig. 2D). Infrared spectroscopy analysis was performed to verify the successful preparation of CFMNPs. FTIR spectroscopy showed that Fu had a OH stretching vibration peak at approximately 3400 cm⁻¹. After the introduction of carboxyl groups to form Fu-COOH [58], while retaining the OH stretching vibration peak at the same position, a new C=O stretching vibration peak appeared near 1700 cm⁻¹, indicating the successful introduction of carboxyl groups into the fucoidan molecule (Fig. S1A). Cs exhibited NH₂ and OH stretching vibration peaks at the same wavenumbers [59]. Morin had a C=C stretching vibration peak at 1450-1600 cm⁻¹ and obvious absorption at 1000-1300 cm⁻¹ [60]. The spectrum of the CFMN composite material showed characteristic peaks of all three components, proving that the three components were successfully combined and interacted to form stable nanoparticles (Fig. 2E). In addition, the encapsulation and loading efficiencies of CFMNPs for morin were determined by HPLC to be approximately 87.23 % and 8.31 %, respectively. Finally, the stability of CFMNPs at different temperatures was evaluated. The aqueous solutions of CFMNPs were stored at 4 °C, room temperature and 45 °C respectively, and then the changes in their particle sizes were detected. The results showed that CFMNPs could be stably preserved under low temperature and room temperature conditions. Even in an environment of 45 °C, they remained stable within 3 days (Fig. S1B). This indicates that CFMNPs have good stability in conventional environments.

Fig. 2.

CFMNPs characterization. (A). Photographs of the CFMNPs powder. (B). Photographs of the CFMNPs solution (left) and the morin solution (right). (C). TEM image of CFMNPs. Scale bar is 200 nm. (D). Size distribution of CFMNPs. (E). FTIR spectra of CFMNPs. (F). Particle size changes of CFMNPs in different pH environments in vitro. (G). The particle size of CFMNPs at 24 h in different pH environments in vitro. (H). Release profiles of CFMNPs in different pH environments in vitro.

To further verify the pH-responsive ability of CFMNPs endowed with Fu-COOH, degradation experiments of CFMNPs under different pH conditions were first carried out. The experimental results showed that in an alkaline environment (pH 8), the diameter of CFMNPs gradually decreased over time, reaching 14.77 ± 5.47 nm at 24 h. In a neutral environment (pH 7), the diameter of CFMNPs decreased, but the decrease was relatively small. In an acidic environment (pH 6), the diameter of CFMNPs hardly changed significantly, and it was still 142.79 ± 20.90 nm at 24 h (Fig. 2F and G). This indicates that CFMNPs undergo significant degradation in an alkaline environment, slightly degrade in a neutral environment (pH 7.0), and hardly degrade in an acidic environment. Moreover, under alkaline conditions (pH 8), the particle size of CFMNPs continuously and gradually decreases. The particle size distribution at 24 h exhibits a bimodal pattern, and the vast majority of the particles are concentrated in the peak with a smaller particle size. This further confirms that CFMNPs undergo a step-by-step degradation process rather than simply breaking into irregular fragments in an alkaline environment. Meanwhile, it can be seen that the supernatant of CFMNPs in an alkaline environment gradually turned from colorless to yellow over time, while there was no significant color change in the supernatant of CFMNPs in neutral and acidic environments (Fig. S1C). In an alkaline environment, the spherical morphology of CFMNPs was significantly damaged, while the morphology of CFMNPs in an acidic environment remained relatively intact (Fig. S1D). This fully demonstrates that CFMNPs can be rapidly degraded in an alkaline environment and have a certain stability in an acidic environment. The pH-responsiveness of CFMNPs is attributed to multiple factors. Firstly, in an alkaline environment, the boronic acid pinacol ester bond in PAPE is prone to nucleophilic attack by hydroxide ions. This attack significantly undermines the structural stability of the nanoparticles. Secondly, under alkaline conditions, the amino groups of Cs undergo proton dissociation. This dissociation leads to a reduction in solubility and alters the intermolecular interactions between molecular chains, which in turn has a substantial impact on the stability of the nanoparticles. Moreover, in an alkaline environment, the intermolecular forces between Fu and Cs are disrupted. All these factors collectively contribute to the pH-responsiveness of CFMNPs.

Furthermore, the ability of CFMNPs to release morin in response to pH was verified. Quantitative analysis of the morin released from CFMNPs at different time points by HPLC showed that the cumulative release of morin exceeded 80 % at only 12 h. This rapid initial release stage was attributed to the rapid destruction of the nanoparticle structure in an alkaline environment, allowing morin to quickly pass through the loosened nanoparticle shell into the surrounding environment. By 24 h, the cumulative release of morin was as high as approximately 95 %, meaning that most of the morin had been released from CFMNPs. In stark contrast, in a neutral environment, the release rate of morin from CFMNPs was significantly slower, with the cumulative release of morin being only approximately 20 % at 12 h and approximately 24 % at 24 h. This phenomenon might be due to the degradation rate of nanoparticles in a neutral environment is relatively slow, and the degree of structural damage is limited, which hinders the diffusion of morin from the inside of the nanoparticles to the surrounding environment. In an acidic environment, the release of morin from CFMNPs was extremely low, with a release rate of only approximately 11 % at 24 h, fully demonstrating the stability of CFMNPs in an acidic environment. This may be because the structure of CFMNPs is hardly damaged under acidic conditions, and morin is firmly encapsulated inside and difficult to release (Fig. 2H).

In conclusion, CFMNPs have excellent pH-responsive ability to release morin and can achieve precise regulation of morin release according to changes in the environmental pH. This feature enhances their potential as drug-delivery vehicles, especially in therapies that demand pH-triggered release.

3.2. Preparation and characterization of bFGF-CFMN bilayer hydrogel

To accommodate dynamic pH fluctuations during B/F wound healing, a pH-responsive bilayer hydrogel was developed. This innovative hydrogel utilizes MeHA as the primary material, leveraging the concentration-dependent properties of MeHA and its sensitivity to acidic environments to construct a unique bilayer structure. Based on previous experiments, we investigated the gelation and degradation of MeHA at different concentrations. The results showed that when the MeHA concentration was below 0.5 %, gelation could not occur (Fig. S2A). When the MeHA concentration was above 3 %, the hydrogel hardly degraded within 3 days, which was unfavorable for drug release (Fig. S2B). The hydrogels formed at 0.8 % and 2 % concentrations exhibit distinct degradation rates, meeting the demand for differential degradation responses. Therefore, MeHA at these two specific concentrations (0.8 % and 2 %) was selected to prepare the upper and lower layers of the bilayer hydrogel. Specifically, the upper layer hydrogel, composed of 2 % MeHA, was loaded with bFGF (bFGF), while the lower layer hydrogel, composed of 0.8 % MeHA, was loaded with CFMNPs (CFMN). Together, these layers formed the pH-responsive bilayer hydrogel (bFGF-CFMN) (Fig. 3A). The microstructure of the bFGF-CFMN bilayer hydrogel was further characterized using SEM (Fig. 3B). The results revealed that the CFMN hydrogel exhibited a relatively loose structure, with CFMNPs uniformly distributed on the surface, facilitating rapid degradation and release of CFMNPs. In contrast, the bFGF hydrogel displayed a denser network structure, effectively protecting bFGF and enabling its sustained release. Porosity, which reflects the crosslinking density of the hydrogel, was significantly lower in the upper layer hydrogel compared to the lower layer hydrogel (Fig. S3A), indicating a more compact and stable network structure in bFGF hydrogel. FTIR analysis showed no significant differences in the spectral peaks between the upper hydrogel loaded with bFGF and the unloaded upper layer hydrogel. Similarly, the lower layer hydrogel exhibited no significant changes in its main peaks regardless of CFMNPs loading (Fig. 3C). These results indicate that incorporating bFGF and CFMNPs did not alter the chemical structure of MeHA and that both agents were noncovalently incorporated (physically entrapped) within the hydrogel matrix.

Fig. 3.

bFGF-CFMN bilayer hydrogel characterization. (A). Physical appearance of the hydrogel. I represents the upper layer bFGF hydrogel, II represents the lower layer CFMN hydrogel and III represents the bFGF-CFMN bilayer hydrogel. (B). SEM image of the bFGF-CFMN bilayer hydrogel. I represents the lower layer CFMN hydrogel, Ia is a magnified view of a local area, and II represents the upper layer bFGF hydrogel. The red triangles point to the CFMNPs. The scale bars for I and II are 200 μm, and the scale bar for Ia is 20 μm. (C). FTIR spectra of the hydrogels. (D). Storage (G′) and loss (G″) modulus of the hydrogels. (E). Shear viscosity of the hydrogels. (F). In vitro degradation curves of the hydrogels under different pH environments (n = 3). (G). In vitro release curves of the hydrogels under different pH environments. (n = 3).

Rheological tests further elucidated the viscoelastic properties of the bFGF-CFMN bilayer hydrogel. The storage/loss modulus of the bFGF hydrogel was greater than that of the CFMN hydrogel (Fig. 3D), indicating higher stiffness in the bFGF hydrogel. Similarly, the viscosity of the bFGF hydrogel was also greater than that of the CFMN hydrogel (Fig. 3E). This rheological contrast enables the CFMN hydrogel to conform more effectively to the wound surface and undergo controlled degradation, while the stiffer bFGF hydrogel preserves structural integrity and provides protection against external mechanical perturbations.

Finally, the pH-responsive behavior of the bFGF-CFMN bilayer hydrogel was evaluated through degradation and drug release assays. The degradation behavior of the hydrogel was assessed by immersing it in PBS solutions at different pH values (6, 7, and 8). The results showed significant degradation of both the bFGF hydrogel and the CFMN hydrogel in acidic conditions (pH 6), while the hydrogels remained stable in neutral (pH 7) and alkaline (pH 8) environments. Interestingly, the bFGF hydrogel exhibited a gradual darkening in color under alkaline conditions, likely due to the pH-responsive degradation of CFMNPs (Fig. S3B). Further degradation experiments confirmed that both the upper and lower layer hydrogels degraded more rapidly in acidic environments, with the lower layer hydrogel exhibiting a higher degradation rate than the upper layer hydrogel (Fig. 3F). In addition, we immersed the lower layer of the bilayer hydrogel in an acidic solution (pH 6) to investigate the influence of its degradation on the stability of the upper layer. The results showed that the upper layer remained globally stable after the degradation of the lower layer in the acidic environment (Fig. S3C). Drug release experiments revealed that the release rate of bFGF from the upper layer hydrogel correlated with its degradation rate, while the release rate of morin from the lower layer hydrogel did not align with that of its degradation rate. Specifically, morin was released more rapidly in alkaline environments than in neutral or acidic conditions (Fig. 3G), likely due to the rapid degradation of CFMNPs in alkaline environments. To validate this hypothesis, the release of CFMNPs from the CFMN hydrogel was further investigated. The results showed that CFMNPs were released most rapidly in acidic environments and least in alkaline conditions, consistent with the degradation behavior of the CFMN hydrogel (Fig. S4A). This suggests that the release of morin from the CFMN hydrogel is influenced by both the hydrogel itself and the CFMNPs. In summary, the bilayer hydrogel exhibits unique pH-responsive properties, with the release rate of bFGF from the bFGF hydrogel matching its degradation rate, while the release of morin from the CFMN hydrogel is jointly regulated by the hydrogel and CFMNPs, thereby, highlighting the complexity and specificity of the drug release mechanism. The degradation rate of MeHA, the main component of the bFGF-CFMN bilayer hydrogel, is influenced by pH. In an acidic environment (pH 6), hydrogen ions protonate the glycosidic bonds and loosen the carboxyl groups, accelerating the degradation of MeHA. Conversely, in an alkaline environment (pH 8), the glycosidic bonds and carboxyl groups remain stable, leading to a slower degradation rate. These pH-dependent degradation behaviors affect the release of bFGF and CFMNPs. Moreover, CFMNPs exhibit an accelerated release of morin under alkaline conditions. This pH-responsive release characteristic is beneficial for the healing of B/F wounds. In the initial stage of wound healing, the acidic microenvironment triggers the rapid degradation of the CFMN hydrogel, resulting in the release of CFMNPs. As the wound repair process commences and the pH increases, CFMNPs rapidly release morin, which exerts anti-inflammatory effects and creates a favorable microenvironment for the subsequent action of bFGF. In the later stage of wound healing, when the microenvironment becomes more alkaline, the bFGF hydrogel is slowly and continuously released, which ultimately improves the quality and efficiency of wound healing.

3.3. Cellular experiments of bFGF-CFMN bilayer hydrogel

Cellular biocompatibility is a critical indicator for the application of hydrogels in biomedical fields, directly determining their safety and efficacy in vivo [61]. To evaluate the biocompatibility of the bFGF-CFMN bilayer hydrogel, HUVECs, which are relevant to wound repair, were used for subsequent experiments. First, the effect of the bFGF-CFMN bilayer hydrogel on cell viability was assessed using live/dead staining (Fig. 4A and C). The results showed that at 24 and 72 h of culture, cells in the CFMN, bFGF, and bFGF-CFMN groups exhibited higher green fluorescence intensity than the control group, indicating high cell viability. PI staining further confirmed low levels of red fluorescence in all treatment groups, suggesting minimal cell death. This indicates that the bFGF-CFMN bilayer hydrogel is biocompatible and does not elicit significant cytotoxicity. To further corroborate these findings, a CCK-8 cell proliferation assay was conducted (Fig. 4B and D). The results showed no significant differences in cell viability among the CFMN, bFGF, and bFGF-CFMN groups and the control group at 24 and 72 h of culture, further confirming the biocompatibility of the bFGF-CFMN bilayer hydrogel.

Fig. 4.

In vitro cell experiments of bFGF-CFMN bilayer hydrogel. Live-dead staining of HUVECs after the hydrogels treatment at 24 h (A) and 72 h (C). Cell viability of HUVECs after the hydrogels treatment at 24 h (B) and 72 h (D). (E). Cell scratch assay of HUVECs treated with the hydrogels. Scale bar is 100 μm. (F). Closure rate of HUVECs treated with the hydrogels (n = 3). (G). Tube formation assay of HUVECs treated with the hydrogels. Scale bar is 50 μm. (H). Quantitative analysis of tube formation in the tube formation assay (n = 3). All data are shown as the mean ± SD. Compared with control group, ns ≥ 0.05, ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001.

Scratch and tube formation assays are standard methods for evaluating wound repair processes at the cellular level, providing insights into cell migration and angiogenesis [62]. The scratch wound assay results showed that the wound closure rate was significantly higher in the CFMN, bFGF and bFGF-CFMN groups compared to the control group (Fig. 4E), with the highest closure rate observed in the bFGF-CFMN group (Fig. 4F). This suggests that the bFGF-CFMN hydrogel effectively promotes HUVECs migration in vitro, accelerating the wound healing process. Additionally, the wound closure rates in the bFGF and bFGF-CFMN groups were similar and higher than those in the CFMN group, suggesting that bFGF is the primary driver promoting HUVECs migration, while CFMN may provide a synergistic contribution. Next, the tube formation assay further validated the pro-angiogenic effects of the bFGF-CFMN hydrogel. Compared to the control group, the CFMN, bFGF and bFGF-CFMN groups induced the formation of more tubular structures (Fig. 4G), with the highest number observed in the bFGF-CFMN group (Fig. 4H), closely followed by the bFGF group. This indicates that bFGF predominantly drives angiogenesis, while CFMN provides ancillary support. We infer that as CFMN degrades, the released morin eliminates the adverse effects of inflammation and other factors on HUVECs. The released chitosan and fucoidan can interact with the cell surface receptors on HUVECs, activating the intracellular signaling pathways involved in angiogenesis. Consequently, they synergistically enhance the effect of bFGF. In summary, these results demonstrate that the bFGF-CFMN bilayer hydrogel not only exhibits excellent biocompatibility but also significantly enhances cell migration and angiogenesis, supporting its potential to promote wound repair.

3.4. In vitro antioxidant and anti-inflammatory effects of bFGF-CFMN bilayer hydrogel

Subsequently, to further confirm the antioxidant and anti-inflammatory capabilities of the bFGF-CFMN hydrogel, RAW 264.7 cells, a commonly used macrophage cell line for studying inflammatory responses and immune regulation, were used [63]. First, an oxidative stress environment was induced using a ROS inducer, and the hydrogel was co-incubated with RAW 264.7 cells for a period. Intracellular ROS levels were detected using the DCFH-DA fluorescent probe. The results showed that compared to the negative control group, the positive control group exhibited pronounced green fluorescence, indicating elevated ROS levels and successful induction of oxidative stress. Further observation revealed that the green fluorescence intensity was significantly reduced in the CFMN, bFGF and bFGF-CFMN treatment groups, suggesting reduced intracellular ROS generation and antioxidant activity (Fig. 5A). Additionally, the blank hydrogel also showed modest antioxidant activity. Quantitative fluorescence analysis indicated that the bFGF-CFMN group had the lowest average fluorescence intensity, with the CFMN group being slightly lower than the bFGF group (Fig. 5B). This suggests that the bFGF-CFMN bilayer hydrogel exhibits the greatest antioxidant effect, likely attributable to CFMNPs. The strong antioxidant capacity likely stems from morin released by CFMNPs in the CFMN hydrogel. In terms of anti-inflammatory effects, LPS was introduced to model an inflammatory environment, and the hydrogel was co-incubated with cells for a defined period. The cell supernatant was then collected, and the levels of inflammatory cytokines (TNF-α, IL-1β and IL-6) were measured using ELISA to assess the anti-inflammatory capacity of the bFGF-CFMN bilayer hydrogel. The results showed that compared to the control group, the CFMN, bFGF, and bFGF-CFMN groups significantly reduced the concentrations of these inflammatory cytokines, with the bFGF-CFMN group exhibiting the most pronounced inhibitory effect. Similarly, the CFMN group showed comparable anti-inflammatory effects to the bFGF-CFMN group (Fig. 5C–E). This demonstrates the superior anti-inflammatory efficacy of the bFGF-CFMN hydrogel, with CFMN likely playing a key role in inflammation suppression. Meanwhile, bFGF may synergistically promote cell repair and regeneration, creating a microenvironment conducive to inflammation resolution and tissue repair. In addition, as crucial immune cells, the polarization state of macrophages plays a vital role in inflammation. Therefore, we further investigated the effect of the bFGF-CFMN bilayer hydrogel on macrophage polarization in vitro. LPS was used to stimulate RAW 264.7 cells to mimic cellular inflammation, and then the cells were co-incubated with the hydrogel. IF was employed to detect iNOS (a characteristic protein of M1 macrophages) and CD206 (a characteristic protein of M2 macrophages). The results showed that compared with the control group, the fluorescence level of iNOS decreased, while that of CD206 increased (Fig. 5F). Quantitative analysis further confirmed these observations (Fig. 5G and H). This indicates that the bFGF-CFMN bilayer hydrogel can promote the polarization of macrophages towards the anti-inflammatory M2 phenotype and inhibit their polarization towards the pro-inflammatory M1 phenotype, thereby exerting an anti-inflammatory effect. In conclusion, the bFGF-CFMN bilayer hydrogel not only reduces oxidative stress but also effectively inhibits inflammatory responses, providing a stable and favorable microenvironment for wound repair.

Fig. 5.

In vitro anti-inflammatory, antioxidant and pro-proliferative effects of bFGF-CFMN bilayer hydrogel. (A). After RAW 264.7 were loaded with fluorescent probes and treated with Rosup inducing drugs, they were further treated with the hydrogels. Then, the fluorescence was observed under a fluorescence microscope. Scale bar is 100 μm. (B). Quantitatively analyze the fluorescence intensity of RAW 264.7 (n = 3). After LPS stimulation and hydrogel treatment, the levels of (C) TNF-α, (D) IL-1β and (E) IL-6 in RAW 264.7 were measured using ELISA (n = 3). (F). Fluorescent staining of iNOS and CD206 in RAW cells after LPS stimulation and hydrogel treatment. Scale bar is 100 μm. Quantitative analysis of the immunofluorescence intensity of (G) iNOS and (H) CD206 (n = 3). (I). EdU staining of HUVECs after the hydrogels treatment. Scale bar is 100 μm. (J). Quantitative analysis of the results of EdU staining (n = 3). All data are shown as the mean ± SD. Compared with control group, ns ≥ 0.05, ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001.

Furthermore, to assess the proliferation-promoting effects of the bFGF-CFMN bilayer hydrogel on HUVECs, an EdU cell proliferation assay was performed. Fluorescence results showed that the red fluorescence intensity in the CFMN, bFGF and bFGF-CFMN groups was higher than in the control group, indicating enhanced proliferation activity (Fig. 5I). Quantitative fluorescence analysis further revealed that the bFGF-CFMN group had the highest average fluorescence intensity, followed by the bFGF group (Fig. 5J). These findings suggest that while CFMN plays a dominant role in antioxidant and anti-inflammatory activities, bFGF exhibits a more significant advantage in promoting cell proliferation. On the other hand, bFGF typically exhibits more significant effects in a bacteria-free environment. In an environment with bacterial infection, bacteria release various toxins and enzymes that can damage the structure and activity of bFGF, making it difficult for them to function effectively. Staphylococcus aureus is one of the most common pathogens in burn wound infections. Therefore, we selected Staphylococcus aureus as the research subject and evaluated the antibacterial performance of the hydrogel using the plate colony counting method. The experimental results showed that the number of Staphylococcus aureus colonies in the CFMN and the bFGF-CFMN group decreased significantly (Fig. S4B). Further quantitative analysis of the antibacterial rate of the hydrogel revealed that the antibacterial effect of the hydrogel mainly originated from the CFMNPs within it, and pure MeHA also had a weak antibacterial effect (Fig. S4C). The above results indicate that CFMNPs composed of morin, Cs and Fu possess certain antibacterial properties. This is beneficial for reducing the risk of wound bacterial infection and creating a relatively sterile condition for bFGF to function.

In summary, the bFGF-CFMN bilayer hydrogel not only exhibits excellent antioxidant and anti-inflammatory capabilities but also significantly promotes endothelial cell proliferation. This multifunctionality supports its potential for biomedical applications, particularly in wound repair.

3.5. In vivo degradation and drug release of bFGF-CFMN bilayer hydrogel

The pH-sensitivity of the bFGF-CFMN bilayer hydrogel was initially confirmed in vitro. To further investigate its pH-responsive degradation and drug release in vivo, B/F wound models were established in mice, and the pH of the wound surface was monitored using a needle-type pH microsensor throughout the healing process. Meanwhile, we compared the HE staining of wound tissues before and after inserting the needle-type pH microsensor, which indicated that the sensor did not cause additional damage such as inflammatory reactions (Fig. S4D). We also compared the pH values measured by the sensor directly in the wound and indirectly in the tissue homogenate. The results showed no significant difference between the two detection methods (Fig. S4E), further validating the accuracy and reliability of the method for directly detecting the wound pH using the needle-type pH microsensor. The results showed that in both B/F wound models, the pH gradually acidified in the early stages of wound healing, became alkaline during the mid-stages, and returned to a slightly acidic state in the later stages (Fig. 6A). The initial acidification may be due to the release of acidic substances from damaged cells, metabolic acid production by inflammatory cells, and plasma protein hydrolysis. As healing progressed, fibroblast proliferation, angiogenesis, and macrophage phenotype switching were associated with an increase in pH toward alkalinity. The return to a slightly acidic state in the later stages may be related to metabolic acid production by keratinocytes and the maintenance of skin barrier function. The dynamic changes in wound pH are likely to affect the in vivo degradation and drug release of the bFGF-CFMN bilayer hydrogel. Bacterial infection is also an important factor influencing the pH of wounds. Therefore, we selected Staphylococcus aureus-infected wounds to further investigate the impact of infection on the wound pH. Experimental results show that the overall pH value of infected burn wounds is higher than that of non-infected ones (Fig. S4F), and a similar pattern is observed in frostbite wounds (Fig. S4G). However, the pH values of both non-infected and infected burn wounds follow the same changing trend during the healing process, and the pH-responsive properties of the bFGF-CFMN bilayer hydrogel can still play their roles. Lactate, a key factor affecting wound pH, was also measured. In both burns and frostbite models, lactate levels rapidly increased in the early stages of wound healing and gradually decreased to a stable level in the mid to late stages (Fig. 6B). The initial increase in lactate may result from enhanced anaerobic glycolysis due to tissue hypoxia, active inflammatory cell metabolism, and accelerated glycogen breakdown, which is consistent with the observed decrease in pH. The subsequent decrease in lactate levels is likely due to improved blood circulation, changes in cellular metabolism, and systemic pH regulation, resulting in increased wound pH.

Fig. 6.

In vivo degradation and release of bFGF-CFMN bilayer hydrogel. (A). Changes in the pH values of B/F wounds over the healing time (n = 5). (B). Changes in lactic acid concentration in B/F wounds over the healing time (n = 3). (C). In vivo imaging was performed on days 1, 3, 5 and 14 after application to monitor the degradation of Rhodamine B-labeled CFMN hydrogel and FITC-labeled bFGF at the B/F wounds. (D). Quantitative analysis of in vivo fluorescence imaging of burn wounds (n = 3). (E). Quantitative analysis of in vivo fluorescence imaging of frostbite wounds (n = 3). (F). In vivo drug release profile of the bFGF-CFMN bilayer hydrogel at burn wounds (n = 3). (G). In vivo drug release profile of the bFGF-CFMN bilayer hydrogel at frostbite wounds (n = 3).

The pH-responsive behavior of the bFGF-CFMN bilayer hydrogel in vivo was further investigated. The upper bFGF hydrogel was fluorescently labeled with FITC, and the lower CFMN hydrogel was fluorescently labeled with Rhodamine B. The fluorescence signal intensity of both layers was monitored over time using small animal in vivo imaging in B/F wound models. The results showed that in both models, the fluorescence intensity of the bFGF hydrogel decreased minimally by day 5, while the fluorescence intensity of the CFMN hydrogel decreased substantially. By day 14, the CFMN hydrogel had nearly completely degraded, while the bFGF hydrogel retained measurable fluorescence intensity (Fig. 6C). Quantitative analysis of fluorescence intensity revealed that the CFMN hydrogel rapidly degraded in the early stages of wound healing, while the bFGF hydrogel degraded more slowly (Fig. 6D and E). This suggests that the CFMN hydrogel responds rapidly to the early acidic environment, while the bFGF hydrogel degrades slowly in the later stages. Taken together with the dynamic changes in wound pH and lactate levels, it can be inferred that the bFGF-CFMN bilayer hydrogel modulates its degradation rate in response to pH changes in B/F wounds, thereby controlling drug release to better adapt to the different stages of wound healing.

To further assess the in vivo drug release of the bFGF-CFMN bilayer hydrogel, the local tissue levels of morin and bFGF were monitored using HPLC and ELISA, respectively. The results showed that by day 5, the release rates of morin in B/F wounds reached 80.33 ± 6.22 % and 74.27 ± 4.74 % respectively, while the release rates of bFGF did not exceed 25 % and 30 % respectively. (Fig. 6F and G). The in vivo drug release profile was consistent with the degradation behavior. In the early inflammatory stage, the acidic environment of the wound triggered rapid degradation of the CFMN hydrogel, releasing CFMNPs to address inflammation. As the wound progressed to the proliferative and remodeling stages, the alkaline environment slowed the degradation of the bFGF hydrogel, allowing for the sustained release of bFGF to promote tissue repair. In brief, the bFGF-CFMN bilayer hydrogel precisely modulates its degradation rate and drug release in response to the dynamic pH changes in B/F wounds in vivo, supporting stage-appropriate delivery and underscoring its potential as a novel wound treatment material.

3.6. Therapeutic efficacy of bFGF-CFMN bilayer hydrogel in B/F wounds

To comprehensively evaluate the potential of the bFGF-CFMN bilayer hydrogel in promoting the healing of B/F wounds in vivo, we established B/F wound models in mice for in vivo experiments (Fig. 7). The efficacy of the bFGF-CFMN bilayer hydrogel on B/F wounds was investigated. The bFGF-CFMN bilayer hydrogel was applied to B/F wounds separately, and its efficacy was compared with that of the control group. The residual wound areas were recorded and measured on days 1, 3, 5, 10 and 14 (Fig. 8, Fig. 9A). Compared with the control and blank groups, CFMN, bFGF and bFGF-CFMN groups all exhibited accelerated wound closure. Among these, the bFGF-CFMN group achieved near-complete closure on day 14, indicating its optimal ability to promote wound healing. Upon further analysis of the wound healing rates of each group, the wound healing rates of the bFGF-CFMN group reached as high as 95.92 ± 1.22 % and 94.47 ± 3.63 % for B/F wounds respectively, with the fastest healing speed (Fig. S5A and S5B). The wound healing rate of the CFMN group was slightly higher than that of the bFGF group (Fig. 8, Fig. 9D). This might be because the morin released from CFMNPs within the CFMN hydrogel has strong antioxidant and anti-inflammatory properties. In the early phases of B/F wounds, morin scavenges free radicals and inhibits proinflammatory factors, creating a favorable wound microenvironment. Meanwhile, bFGF mainly promotes cell proliferation and migration in the middle and late stages. The CFMN group controls inflammatory stress in the early stage, laying a foundation for repair and thus exhibiting a slightly better healing process.

Fig. 7.

Timeline diagram of the animal experiment on treating B/F wounds with bFGF-CFMN bilayer hydrogel.

Fig. 8.

Treatment of burn wounds with bFGF-CFMN bilayer hydrogel. (A). Representative photographs of wound healing on days 1, 3, 5, 10 and 14 after the corresponding treatments were administered to each group. (B). After different treatments, HE staining was performed on burn wound tissues to observe the tissue changes on days 5 and 14. Scale bar is 100 μm. (C). After different treatments, masson staining was performed on burn wound tissues to observe the tissue changes on days 5 and 14. Scale bar is 100 μm. (D). The wound healing rates on days 5 and 14 following the respective treatments (n = 3). (E). Quantitatively analyze the inflammatory infiltration in burn wound tissues of each group (n = 3). (F). Quantitative analysis of collagen deposition in burn wound tissues of each group (n = 3). The levels of inflammatory cytokines in burn wound tissue at different time points following various treatments: TNF-α (G), IL-1β (H) and IL-6 (I) (n = 3). All data are shown as the mean ± SD. Compared with control group, ns ≥ 0.05, ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001.

Fig. 9.

Treatment of frostbite wounds with bFGF-CFMN bilayer hydrogel. (A). Representative photographs of wound healing on days 1, 3, 5, 10 and 14 after the corresponding treatments were administered to each group. (B). After different treatments, HE staining was performed on frostbite wound tissues to observe the tissue changes on days 5 and 14. Scale bar is 100 μm. (C). After different treatments, masson staining was performed on frostbite wound tissues to observe the tissue changes on days 5 and 14. Scale bar is 100 μm. (D). The wound healing rates on days 5 and 14 following the respective treatments (n = 3). (E). Quantitatively analyze the inflammatory infiltration in frostbite wound tissues of each group (n = 3). (F). Quantitative analysis of collagen deposition in frostbite wound tissues of each group (n = 3). The levels of inflammatory cytokines in frostbite wound tissue at different time points following various treatments: TNF-α (G), IL-1β (H) and IL-6 (I) (n = 3). All data are shown as the mean ± SD. Compared with control group, ns ≥ 0.05, ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001.

Subsequently, we performed histological analysis using HE and Masson staining to further confirm the therapeutic effect of the bFGF-CFMN group. Whether it was burns or frostbite, on day 5, compared with the control group, the infiltration of inflammatory cells in the bFGF-CFMN group was significantly reduced. On day 14, the tissue structure of the bFGF-CFMN group was relatively more regular overall, with orderly cell arrangement, reducing cell damage caused by excessive inflammatory stimulation (Fig. 8, Fig. 9B). Further quantitative analysis of inflammatory cell infiltration showed that the bFGF-CFMN group had the lowest degree of inflammatory infiltration, followed by the CFMN group. The degree of inflammatory infiltration in the bFGF group was lower than that in the control group and the blank group but slightly higher than that in the CFMN group (Fig. 8, Fig. 9E). Moreover, it was found that the inflammatory damage in burn wounds was more severe than that in frostbite wounds. This verifies the remarkable efficacy of morin released from CFMNPs in the CFMN hydrogel in controlling inflammation and the synergistic advantage of the bFGF-CFMN bilayer hydrogel in regulating the inflammatory microenvironment of the wound. Masson staining showed that in both B/F wounds, there was greater collagen deposition in the bFGF-CFMN group. The collagen fibers in the skin tissue were denser, thicker, and better arranged, and the epithelial coverage was more complete. In contrast, the control group and the blank group only showed a small amount of loose and disordered collagen fibers (Fig. 8, Fig. 9C). Quantification of the collagen area indicated that the bFGF-CFMN group had the highest collagen deposition. The bFGF group was similar to the bFGF-CFMN group and was slightly greater than the CFMN group (Fig. 8, Fig. 9F). This may be due to the direct and significant role of bFGF in promoting collagen synthesis. Although the CFMN group laid a foundation for wound repair through the anti - inflammatory effect of morin in the early stage, due to the lack of direct stimulation of cell proliferation and collagen synthesis by bFGF, its collagen deposition was slightly lower than that of the bFGF group and the bFGF-CFMN group. However, the collagen deposition in the CFMN group was still significantly greater than that in the control group and the blank group, which further demonstrates the positive role of morin in regulating the wound microenvironment and indirectly promoting collagen synthesis.

These results indicate that the bFGF-CFMN bilayer hydrogel shows excellent potential in promoting the healing of B/F wounds by regulating the inflammatory response, enhancing collagen synthesis, and epithelial coverage.

3.7. Anti-inflammatory effects of bFGF-CFMN bilayer hydrogel

Excessive and persistent inflammation is a key factor hindering wound repair in B/F wounds, making the anti-inflammatory effects of the bFGF-CFMN bilayer hydrogel particularly important. Upon injury, the wound rapidly enters a state of stress, activating inflammatory signaling pathways and triggering the release of inflammatory cytokines. The excessive secretion of these cytokines not only directly damages surrounding cells but also exacerbates inflammation, creating a vicious cycle that hinders normal wound healing. Therefore, the anti-inflammatory effects of the bFGF-CFMN bilayer hydrogel in B/F wounds were further investigated. The levels of key inflammatory cytokines (TNF-α, IL-1β, and IL-6) in wound tissues were measured using ELISA to reflect the inflammatory status (Fig. 8, Fig. 9). In both B/F models, the levels of TNF-α, IL-1β, and IL-6 were higher in the control group on day 5, consistent with an early inflammatory response to injury. In contrast, the bFGF-CFMN group exhibited lower levels of these cytokines, suggesting potential anti-inflammatory effects. Additionally, the CFMN group showed slightly lower cytokine levels than the bFGF group, which is consistent with morin contributing more prominently to anti-inflammatory activity. Meanwhile, the levels of inflammatory factors in the bFGF group were also significantly lower than those in the control group, indicating that bFGF also has certain anti-inflammatory effects. This may be attributed to two aspects. On one hand, bFGF can directly interact with specific receptors on immune cells, regulating their activation and function and enhancing their ability to resist inflammation. On the other hand, as hypoxia is a major driver of inflammation, by promoting vascularization of the wound surface and alleviating hypoxia, bFGF can indirectly reduce inflammation in the wound.

The levels of the anti-inflammatory cytokine IL-10 were also measured to further evaluate the anti-inflammatory capacity of the bFGF-CFMN bilayer hydrogel. IL-10 plays a crucial role in inhibiting macrophage and T cell activation and reducing the release of inflammatory mediators. The results showed that IL-10 levels in the bFGF-CFMN group were higher than those in the control, CFMN, and bFGF groups. Although IL-10 levels decreased by day 14, they remained the highest in the bFGF-CFMN group (Fig. S6A and S6B). This suggests that the bFGF-CFMN bilayer hydrogel may promote increased IL-10 expression, which may help limit the overactivation of inflammatory cells and reduce the release of pro-inflammatory cytokines, thereby creating a stable microenvironment for wound healing.

In the early stages of wound healing, the CFMN hydrogel rapidly degrades in response to the acidic environment, releasing CFMNPs. As the wound progresses, the pH becomes alkaline, triggering the degradation of CFMNPs and the release of morin, which plays a key anti-inflammatory role. In the later stages, the alkaline environment inhibits the degradation of the bFGF hydrogel, enabling sustained release of bFGF, which may further modulate inflammation. In summary, the bFGF-CFMN bilayer hydrogel achieves potent anti-inflammatory effects through its pH-responsive degradation and drug release mechanisms, providing effective and sustained anti-inflammatory support for B/F wound healing.

3.8. Antioxidant effects of bFGF-CFMN bilayer hydrogel

During the healing process of B/F wounds, oxidative stress, alongside excessive inflammatory responses, significantly impedes wound repair. Numerous studies have shown that B/F injuries lead to the overproduction of ROS. The accumulation of ROS causes cellular and tissue damage, thereby delaying wound healing. Therefore, evaluating the antioxidant effects of the bFGF-CFMN bilayer hydrogel is crucial for a comprehensive understanding of its mechanism in promoting wound healing. To investigate the antioxidant capacity of the bFGF-CFMN bilayer hydrogel, dihydroethidium (DHE) staining was performed on B/F wound tissues in mice. The intensity of red fluorescence was used to assess the extent of oxidative stress damage (Fig. 10A and C). The fluorescence results revealed that, on days 5 and 14, the wound tissues in the control group exhibited strong red fluorescence, indicating high levels of oxidative stress. The fluorescence intensity was higher in burn wounds compared to frostbite wounds, suggesting that oxidative stress was more severe in burn wounds. This may be due to the more extensive and intense tissue damage caused by burns, which can trigger a stronger inflammatory response and greater ROS production. In contrast, the CFMN and bFGF groups showed reduced red fluorescence intensity, though less significantly than the bFGF-CFMN group. This indicates that both CFMN and bFGF hydrogels alleviated oxidative damage to some extent, while the bFGF-CFMN bilayer hydrogel more effectively suppressed oxidative stress. Quantitative analysis showed that the red fluorescence intensity in the bFGF-CFMN group was significantly lower than in the other groups on days 5 and 14 (Fig. 10B and D). These results indicate that the bFGF-CFMN bilayer hydrogel possesses strong antioxidant properties in B/F wounds, effectively reducing oxidative stress levels and promoting wound healing. Additionally, the red fluorescence intensity in the CFMN group was lower than in the bFGF group, indicating that morin in the CFMN hydrogel made a more prominent contribution in the antioxidant process. Although bFGF also reduced oxidative stress to some extent, its effect was limited.

Fig. 10.

Antioxidant effects of bFGF-CFMN bilayer hydrogel. (A). DHE fluorescence staining of burn wounds on days 7 and 14 following the hydrogels treatment. Scale bar is 100 μm. (B). Quantitative analysis of DHE fluorescence (A) intensity in each group (n = 3). (C). DHE fluorescence staining of frostbite wounds on days 7 and 14 following the hydrogels treatment. Scale bar is 100 μm. (D). Quantitative analysis of DHE fluorescence (C) intensity in each group (n = 3). Detection of antioxidant indicators in burn wound tissues on days 5 and 14 following the corresponding treatments: SOD (E), GSH (F) and MDA (G) (n = 3). Detection of antioxidant indicators in frostbite wound tissues on days 5 and 14 following the corresponding treatments: SOD (H), GSH (I) and MDA (J) (n = 3). All data are shown as the mean ± SD. Compared with control group, ns ≥ 0.05, ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001.

Antioxidant and oxidative stress indicators, including the enzyme SOD, the antioxidant GSH, and the lipid peroxidation end product MDA, play pivotal roles in the oxidative stress process during wound healing. Their levels are key biomarkers reflecting the oxidative stress status of wound tissues. Therefore, we further measured SOD activity, GSH levels and MDA levels in B/F wound tissues. Compared to the control group, the CFMN and bFGF groups showed increased SOD activity (Fig. 10E and H), elevated GSH levels (Fig. 10F and I), and reduced MDA levels (Fig. 10G and J) in wound tissues. However, these changes were less pronounced than those observed in the bFGF-CFMN group. These results are consistent with the bFGF-CFMN bilayer hydrogel being associated with higher SOD activity and GSH levels and lower MDA levels in wound tissues, consistent with enhanced tissue antioxidant defenses and reduced lipid peroxidation, thereby supporting antioxidant effects. In summary, the bFGF-CFMN bilayer hydrogel enhances the antioxidant defense capacity of B/F wound tissues.

3.9. Pro-angiogenic effects of bFGF-CFMN bilayer hydrogel

Angiogenesis is crucial for B/F wound healing by delivering essential nutrients and oxygen to the wound site, promoting cell proliferation and tissue repair. Therefore, studying the pro-angiogenic effects of the bFGF-CFMN bilayer hydrogel is highly significant. To evaluate the pro-angiogenic capacity of the bFGF-CFMN hydrogel, the expression of CD31 in B/F wound tissues was detected using IF. The bFGF-CFMN group exhibited higher CD31 fluorescence intensity compared to the control group (Fig. 11, Fig. 12A). Further quantitative analysis showed that on day 5, the bFGF-CFMN group had the highest average CD31 fluorescence intensity, followed by the bFGF group and then the CFMN group. By day 14, the difference in average fluorescence intensity between the bFGF-CFMN group and the control group further increased (Fig. 11, Fig. 12C). These results support the pro-angiogenic effects of the bFGF-CFMN hydrogel. Vascular endothelial growth factor (VEGF) is a key regulator of angiogenesis, stimulating endothelial cell proliferation, migration, and vessel formation. Therefore, we used IHC to detect VEGF expression in wound tissues. The results showed that in B/F wound models, the bFGF-CFMN group had higher VEGF expression levels than the control group (Fig. 11, Fig. 12B). This indicates that the bFGF-CFMN hydrogel more effectively promotes VEGF expression, which may support angiogenesis. Quantitative analysis of VEGF IHC revealed that the bFGF-CFMN group had the highest VEGF positivity rate, with the bFGF group slightly lower but still higher than other groups. Although the VEGF positivity rate in the bFGF-CFMN group slightly decreased by day 14, it remained the highest (Fig. 11, Fig. 12D). These results suggest that the bFGF hydrogel has a pro-angiogenic ability, while the CFMN hydrogel may synergistically enhances this effect, collectively promoting angiogenesis and tissue repair in B/F wounds. In response to the complex microenvironment of B/F wounds, the bFGF-CFMN bilayer hydrogel, compared to single-layer hydrogels, creates a more favorable intracellular and extracellular environment, potentially promoting angiogenesis more effectively. Additionally, the slight decrease in VEGF positivity rate in the bFGF-CFMN group by day 14 likely reflects physiological feedback mechanisms that temper VEGF overexpression. Furthermore, we measured VEGF content in B/F wounds (Fig. S7A and S7B), and the results were consistent with the IHC, with the bFGF-CFMN group showing the highest VEGF levels. This further confirms the exceptional ability of the bilayer hydrogel to promote VEGF synthesis and secretion, thereby supporting wound vascularization.

Fig. 11.

Angiogenic effect of bFGF-CFMN bilayer hydrogel on burn wounds. (A). CD31 IF of burn wound tissues on days 5 and 14 after the corresponding treatments. Scale bar is 50 μm. (B). VEGF IHC was performed on burn wound tissues on days 5 and 14 after the corresponding treatments. Scale bar is 50 μm. (C). Quantitative analysis of CD31 fluorescence intensity in each group (n = 3). (D). Quantitative analysis of the positive expression levels of VEGF in each group (n = 3). The contents of PDGF (E) and Ki67 (F) in burn wound tissues were detected by ELISA on days 5 and 14 after the corresponding treatments (n = 3). All data are shown as the mean ± SD. Compared with control group, ns ≥ 0.05, ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001.

Fig. 12.

Angiogenic effect of bFGF-CFMN bilayer hydrogel on frostbite wounds. (A). CD31 IF of frostbite wound tissues on days 5 and 14 after the corresponding treatments. Scale bar is 50 μm. (B). VEGF IHC was performed on frostbite wound tissues on days 5 and 14 after the corresponding treatments. Scale bar is 50 μm. (C). Quantitative analysis of CD31 fluorescence intensity in each group (n = 3). (D). Quantitative analysis of the positive expression levels of VEGF in each group (n = 3). The contents of PDGF (E) and Ki67 (F) in frostbite wound tissues were detected by ELISA on days 5 and 14 after the corresponding treatments (n = 3). All data are shown as the mean ± SD. Compared with control group, ns ≥ 0.05, ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001.

Platelet-derived growth factor (PDGF) and transforming growth factor-β1 (TGF-β1) also play important roles in angiogenesis, synergizing with VEGF to promote B/F wound healing. PDGF stimulates the proliferation and migration of fibroblasts, smooth muscle cells, and endothelial cells, participating in vessel wall construction and stabilization. TGF-β1 regulates extracellular matrix synthesis and degradation, thereby influencing vessel remodeling and maturation. To further investigate the effects of the bFGF-CFMN bilayer hydrogel on these key growth factors, we measured PDGF and TGF-β1 levels in B/F wounds. On day 5, the bFGF-CFMN group had higher PDGF content than the control group, with levels similar to those in the bFGF group. By day 14, although PDGF content began to decline and stabilize, the bFGF-CFMN group still had higher PDGF content than the control group (Fig. 11, Fig. 12E). Similarly, the differences and changes in TGF-β1 content across groups were consistent with those of PDGF (Fig. S8A and S8B). Overall, the bFGF-CFMN hydrogel positively regulates multiple key growth factors throughout the B/F wound healing process. The strong pro-angiogenic ability of bFGF provides a foundation, while the CFMN hydrogel synergistically enhances this regulatory effect, collectively promoting angiogenesis and tissue repair in B/F wounds. Finally, to further validate the proliferation-promoting effects of the bFGF-CFMN bilayer hydrogel, we detected Ki67 expression in B/F wound tissues. Ki67 is a nuclear protein closely associated with cell proliferation, and its expression level reflects cell proliferative activity. The results showed that on days 5 and 14, the bFGF-CFMN group had higher Ki67 levels than the control group, with the highest Ki67 levels observed on days 5 (Fig. 11, Fig. 12F). This suggests that the bFGF-CFMN bilayer hydrogel not only effectively promotes angiogenesis but also significantly enhances cell proliferative capacity, ensuring continuous growth and repair of new tissue. This sustained proliferation-promoting effect may be important for the complete healing of B/F wounds.

In the early acidic environment of the wound, the CFMN hydrogel degrades rapidly, releasing CFMNPs. As the wound progresses and the pH rises toward alkalinity, CFMNPs degrade and release morin, which may help create a favorable microenvironment for later-stage wound healing and vascularization. As the wound healing process progresses and the pH further increases, the bFGF hydrogel degrades slowly in the alkaline environment, enabling sustained release of bFGF, which upregulates the expression of angiogenesis-related factors such as CD31, VEGF, PDGF, and TGF-β1, and promotes wound cell proliferation, ultimately facilitating the formation and maturation of new blood vessels. Based on the above results, the bFGF-CFMN bilayer hydrogel, through the potent pro-angiogenic capability of bFGF and the synergistic effects of CFMN, collectively establishes a microenvironment conducive to vascularization, cell proliferation and tissue regeneration, thereby accelerating the healing process of B/F wounds.

3.10. Biocompatibility of bFGF-CFMN bilayer hydrogel

The in vivo biocompatibility of the bFGF-CFMN bilayer hydrogel is a critical for evaluating its clinical potential. We conducted a series of in vivo experiments to comprehensively assess the biocompatibility of the bFGF-CFMN hydrogel. To evaluate the hemocompatibility of the bFGF-CFMN bilayer hydrogel, we first performed a hemolysis test. This test directly assesses whether the hydrogel induces red blood cell hemolysis. The hemolysis test results showed that the bFGF-CFMN bilayer hydrogel did not cause significant red blood cell lysis upon contact with blood (Fig. S9A), with a hemolysis rate of well below 5 % (Fig. S9B), demonstrating its good hemocompatibility. Additionally, the bFGF-CFMN bilayer hydrogel was applied to the wounds of healthy mice, and after 14 days, their major organs (liver, kidneys, heart, lungs, and spleen) were harvested for HE staining. The staining results showed that, compared to healthy mice, the major organs of mice treated with the bFGF-CFMN hydrogel exhibited intact tissue structures and normal cell morphology, with no significant pathological damage or inflammatory infiltration (Fig. S10). These results collectively demonstrate the excellent in vivo biocompatibility of the bFGF-CFMN bilayer hydrogel.

4. Conclusion

To address the clinical challenge of B/F wounds, we developed a novel bFGF-CFMN bilayer hydrogel that leverages its unique pH-responsive degradation to regulate drug release, adapting to the dynamic physiological microenvironment at different stages of wound healing, and demonstrating exceptional therapeutic efficacy. The bFGF-CFMN bilayer hydrogel, composed of MeHA, harnesses the concentration-dependent and pH-responsive properties of MeHA, together with pH-responsive CFMNPs and the pro-healing factor bFGF, to form a bilayer structure with distinct functions. The lower layer consists of MeHA (0.8 %) loaded with CFMNPs, while the upper layer is composed of MeHA (2 %) loaded with bFGF. Together, they form a bilayer hydrogel that responds to dynamic pH changes in the wound, regulating degradation and releasing morin and bFGF in a time-dependent manner for precise therapeutic delivery. Experimental results confirmed that the bFGF-CFMN bilayer hydrogel exhibits excellent pH responsiveness and mechanical properties. In vitro cell studies revealed that the bFGF-CFMN bilayer hydrogel demonstrated superior biocompatibility, promoting HUVECs migration, tube formation and proliferation, and showed preliminary good antioxidant and anti-inflammatory capabilities. In animal B/F wound models, the bFGF-CFMN bilayer hydrogel promoted wound healing by inhibiting inflammation, counteracting oxidative stress, and enhancing angiogenesis. Additionally, the bFGF-CFMN bilayer hydrogel exhibited good biocompatibility. In conclusion, the bFGF-CFMN bilayer hydrogel, through its unique pH-responsive degradation and time-dependent release mechanism, achieves precise delivery of antioxidant, anti-inflammatory, and pro-angiogenic therapeutic components, providing an efficient and intelligent solution for the treatment of B/F wounds, with broad clinical application prospects.

CRediT authorship contribution statement

Meilin Yi: Writing – original draft, Investigation. Wenzhang Jin: Writing – review & editing, Visualization, Methodology, Data curation. Haobing Li: Validation, Formal analysis. Xiaoying Niu: Validation. Junru Wang: Formal analysis. Shunfu Wang: Validation. Wa Zhang: Formal analysis. Mengxuan Zhou: Validation. Zhe Wang: Formal analysis. Yutong Zhou: Validation. Xuchen Deng: Validation. Jingyong Huang: Resources, Funding acquisition. Xiang Su: Resources, Funding acquisition, Conceptualization.

Ethics approval statement

All experimental procedures were strictly carried out in accordance with the animal ethics guidelines approved by the Ethics Committee of the First Affiliated Hospital of Wenzhou Medical University and the Animal Experimentation Ethics Committee of Zhejiang Province (Approval No.: WYYY-IACUC-AEC-2025-027).

Declaration of competing interest

The authors declare no potential conflicts of interest with respect to the research, authorship, and publication of this article.

Appendix A. Supplementary data

The following is the Supplementary data to this article.

Data availability

Data will be made available on request.