Introduction

The largest organ of the human body, the skin, plays a primary defensive role against external environmental threats [1]. Infections that permeate this barrier through wounds may cause severe inflammation, adversely affecting healing and potentially leading to life-threatening conditions [2]. Thus, it is crucial to develop innovative, multifunctional bioactive materials that can eradicate infections and enhance wound healing and tissue regeneration. Various types of dressing materials, such as nanofiber films and hydrogels, have been designed for this purpose [3, 4]. Hydrogels are known for their high permeability, biocompatibility and moisturizing qualities, positioning them as promising materials for treating infectious wounds [5, 6]. However, traditional hydrogel dressings often lack inherent antibacterial properties, limiting their use to conjunction with antibiotics [7, 8]. Regrettably, long-term use of antibiotics may yield bacterial resistance and provoke adverse hepatic or renal side effects, impeding the wound healing process [9, 10]. Therefore, there is an urgent need to develop new types of hydrogel dressings capable of taking effect without relying on antibiotics.

Infectious wounds render a host of complications owed to particular inflammatory conditions which obstruct the physiological sequence of wound healing and considerably challenge macrophages’ polarization from pro-inflammatory (M1) to anti-inflammatory (M2) phenotypes [11]. A surfeit of M1 macrophages accumulates within the wound, excreting pro-inflammatory cytokines like tumor necrosis factor-alpha (TNF-α) and producing reactive oxygen species [12]. This action consecutively induces oxidative stress and a cellular damage microenvironment within the wound, resulting persistent inflammation and vascular alterations. This sequence culminates in impeded wound healing, amplified inflammatory mediators and a perpetuated detrimental cycle [13,14,15]. Consequently, manipulating the immune microenvironment and fostering angiogenesis are crucial for healing infectious wounds. Immune modulation triggers the expression of transforming and proliferative cytokines, renowned for their angiogenesis promoting attributes [6d). In a similar vein, the mean nodule formation experienced a surge of 7.72, 7.46, and 7.28 times relative to the control, PADM, and PADM@Mg aggregates, accordingly (Fig. 6d).

Studies have indicated that administering curcumin by gavage can enhance neovascularization and wound healing in mice with diabetic foot ulcers. Additionally, the local application of 1% turmeric extract gel has been found to have a beneficial impact on angiogenesis [48]. Furthermore, there is no substantial variance in the effects of various administration routes of curcumin on wound healing. The results of this investigation indicated that the rates of cell migration and angiogenesis were significantly enhanced in cells treated with PADM@MgC, which is advantageous for the healing of wounds, although the exact mechanism requires further exploration.

Fig. 6
figure 6

In vitro cell migration and angiogenesis behavior of various PADM-based hydrogels. (a) Different migration abilities of HUVECs after coculturing with the extract of various PADM-based hydrogels. Scale bar: 100 μm. (b) Quantification of gap closure ratio of different groups. Scale bar: 100 μm. (c) Fluorescence images of tube formation experiment via coculturing HUVECs with the extract of various PADM-based hydrogels. (d) Quantifications of junctions and tube length in each group. Results are presented as mean ± SD (N = 3). ****P < 0.0001

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In vitro biocompatibility

The biocompatibility of hydrogels is a vital parameter for their adoption in clinical settings as wound dressings. To this end, cell viability assays were conducted after culturing cells on various hydrogel compositions for a 24-hour period followed by live/dead staining procedures. Live cells exhibited green fluorescence, while the nuclei of nonviable cells emitted red fluorescence. Observations indicated a predominance of viable cells post-incubation on hydrogels-control, PADM, PADM@Mg, and PADM@MgC (Fig. 7a-b). Subsequent quantification via the CCK-8 assay reaffirmed the absence of significant cytotoxicity attributed to the hydrogels (Fig. 7c). SEM imaging further corroborated these findings by displaying cells in a state of healthy morphology upon hydrogel substrates (Fig. 7d).

Fig. 7
figure 7

The cytocompatibility of different PADM-based hydrogels in vitro. (a) Confocal microscopy images of a live/dead assay of cells co-cultured with several PADM-based hydrogels for 24 h. Scale bar: 200 μm. (b) Quantitative analysis of rusults from (a). (c) Cell proliferation of PADM-based hydrogels. (d) SEM results of RAW 264.7 co-cultured with PADM-based hydrogels for 24 h

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In vivo investigation on PADM@MgC for Infected Wound Healing

Prolonged non-healing of wounds may increase the risk of bacterial infection. Furthermore, this type of infection may compromise the immune function, thereby adversely affecting wound healing and ultimately leading to systemic infection. Therefore, expediting wound healing is imperative for preventing and treating infections. Next, we will utilize a murine excisional skin infection model to determine the impact of PADM@MgC hydrogel on wound healing (Fig. 8a).

With the progression of time, the infected wounds in each treatment group gradually underwent healing (Fig. 8b-c). However, significant variations were observed in the infection severity and rate of healing. Four days ago, the wounds of the control group, PADM group, and PADM@Mg group all exhibited evident infection, tissue necrosis, and discharge of pus. In contrast, the wounds of the PADM@MgC group remained consistently clean and dry throughout the entire observation period, showing no signs of obvious infection or suppuration. Furthermore, the mice in the PADM@MgC group demonstrated quicker wound healing (Fig. 8d), with only 17.1 ± 2.5% of the wounds remaining unhealed on the 10th day, followed by the PADM@Mg group (33.2% ± 1.6%), PADM group (42.5% ± 2.6%), and the control group (44.1% ± 2.1%). Additionally, on the 4th day, spreading plate method (SPM) was employed to evaluate the bacterial survival rate on the wounds after different treatments, showing a significant decrease in the bacterial survival rate following treatment with PADM@MgC hydrogel (Fig. 8e).

Fig. 8
figure 8

PADM@MgC demonstrates increased effectiveness in facilitating the healing of infected wounds in vivo. (a) Illustration of the model for mouse excisional wound infection and the experimental methods. (b) Representative photographic images of wounds in different groups after 0, 1, 4, 7, or 10 days of treatment. (c) Day 0 (blue), day 1 (orange), day 4 (purple), day 7 (yellow), and day 10 (green) explanatory photos of the wounds, as well as superimposed photos of treatment with various PADM-based hydrogels. (d) Area of unhealed wounds in each group at different time periods. (e) Bacterial counts in the wounds of the different treatment groups. Results are presented as mean ± SD (N = 3). **P < 0.01; ***P < 0.001; ****P < 0.0001

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Histological evaluations were undertaken through HE staining (Fig. 9a), with findings aligning with the statistical patterns observed in wound surface diminution. Moreover, the extent of granulation tissue formation among the mouse cohorts was quantified (Fig. 9b). Notably, the PADM@MgC-treated group exhibited accelerated wound closure, a trend that was most evident during the initial healing phases. Quantitative assessment of the granulation tissue thickness indicated a superior contraction rate in the PADM@MgC group relative to its counterparts. Effective wound repair is contingent upon vascularization to supply essential nutrients and oxygen. Consequently, angiogenic sufficiency is a critical component of the healing process. To gauge angiogenic activity, we assessed the expression levels of endothelial cell marker, cell adhesion molecule 1 (CD31), within the wound beds (Fig. S2). Comparative analysis indicated a heightened presence of CD31 in the wound tissues treated with PADM@MgC hydrogel. Collectively, these data point towards the notable wound-healing capabilities of PADM@MgC hydrogel. Further immunofluorescence analysis revealed that the PADM@MgC group had the lowest proportion of M1 macrophages (iNOS) (Fig. 9d and e) and the highest proportion of M2 macrophages (CD206) (Fig. 9d and f), confirming the in vitro results. These findings suggest that PADM@MgC triggers macrophage polarization towards an M2 phenotype, inducing an anti-inflammatory microenvironment, thereby promoting wound healing.

Fig. 9
figure 9

PADM@MgC hydrogel-induced acceleration of the wound repair. (a) Images of HE and Masson’s trichromatic staining of the infectious wounds of different groups on day 10. (b) Corresponding quantitative analysis of the granulation tissue width on Day 10. (c) Images of Giemsa staining of the infectious wounds of different groups on day 10. (d) Immunofluorescence staining for iNOS (M1-labelled) and CD206 (M2-labelled) was performed on tissue sections at the wound site on days 4 post-infection e, (f) Statistical analyses regarding the proportion of (e) iNOS- and (f) CD206-positive macrophages. Results are presented as mean ± SD (N = 3). ****P < 0.0001

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Conclusion

Here, we developed a PADM@MgC hydrogel to enhance post-infectious wound healing by sequentially releasing Cur and Mg2+. The therapeutic effectiveness of PADM@MgC was optimized through (i) improved stability and bioavailability of Cur; (ii) anti-inflammatory, antimicrobial, and cell proliferation-promoting effects of Cur; (iii) Mg2+ improving cell migration and angiogenesis; (iv) Mg2+ and curcumin synergistically promote repolarization of M2 macrophages; (v) the hybridized hydrogel forming a physical barrier to prevent secondary damage; and (vi) the hybridized hydrogel achieving exudate absorption and antimicrobial PADM@MgC. Various functions of PADM@MgC have been demonstrated in vivo and in vitro. However, our study did not comprehensively examine the mobilization and function of other immune cells (e.g. DC cells, T cells, NK cells, etc.) in addition to macrophages during the initial stages of infected wound healing. Additionally, the molecular mechanisms of these processes have not been explored. In the next phase of our research, we will focus on these aspects in order to better understand the factors that promote infected wound healing through the use of biomaterials. Nevertheless, this novel diachronic bidirectional immunomodulatory hydrogel has good antimicrobial and wound healing effects and can be used to design and synthesize biomaterials that modulate immune responses according to the immune profile of the target disease.

Materials and methods

Synthesis of PADM@MgC hydrogel

The preparation protocol for Mg-MBG and Mg-MBG@Cur is comprehensively outlined in the Supporting Information. The porcine skin tissue, after a 3-hour rinse in sterile water, underwent multiple freeze/thaw cycles using liquid nitrogen. This was followed by agitation at 120 rpm and 25 °C to facilitate the removal of subcutaneous layers. The tissue samples were then treated with 0.1% Triton X-100 for 12 h and subsequently with 0.1% sodium dodecyl sulfate for 6 h. The final steps involved lyophilization, grinding into a fine powder, and digestion with pepsin in an acidic milieu (pH 2–3) for 10 min. Mg@C nanoparticles were introduced and rapidly mixed into the solution. The resulting gel was refrigerated at 4 °C and enzymatically digested over a 2-hour period until it achieved a translucent, viscous consistency. Subsequent steps included the addition of PBS to regulate osmotic pressure, followed by pH normalization to 7–8 using chilled 10 M NaOH. PADM@MgC hydrogels were then synthesized by incubating these gels at 37 °C for 20 min, followed by freeze-drying and compression. The final PADM@MgC hydrogels, with concentrations of 1%, 5%, and 10%, were determined based on the mass differential between the added nanoparticles and the freeze-dried hydrogel powder. In parallel, PADM@Mg hydrogels were synthesized using an identical process, except Mg-MBG were used in place of Mg@C nanoparticles for PADM@Mg preparation.

Characterization of PADM@MgC hydrogels

Initially, Hematoxylin and Eosin (HE) and DAPI staining were conducted to evaluate the removal of cellular and nuclear debris. The DNA content of each set of skin tissue samples was analyzed using a DNA extraction kit, following the instructions in the manual. The DNA content of each set was measured using a UV spectrophotometer. Following this, the samples underwent dehydration through a critical point drying process. The structural characteristics of the hydrogels were then examined via scanning electron microscopy. Subsequently, the chemical composition of these materials was investigated using X-ray Photoelectron Spectroscopy (XPS), X-ray Diffraction (XRD), and Fourier-Transform Infrared (FTIR) spectroscopy.

Water retention

The hydrogels PADM, PADM@Mg, and PADM@MgC were incubated at 37 °C, and their masses were periodically recorded until a stable mass was noted. The water retention capacity of these hydrogels was determined using the formula:

$${\rm{Water}}\,{\rm{retention}}\,{\rm{rate }} = {\rm{ W}}2/{\rm{W}}1{\rm{ }} \times {\rm{ }}100\%$$
(1)

Here, W2 denotes the hydrogel’s weight at each measured time point, while W1 represents its initial weight.

Biodegradability

The biodegradability of PADM, PADM@Mg, and PADM@MgC hydrogels was determined under different conditions. The hydrogels were solidified in PBS and then incubated at 37 °C with or without trypsin. The solution was removed from the centrifuge tube, and the hydrogels were washed with PBS daily. Finally, the samples were dried and weighed.

Hygroscopic properties

To investigate the hygroscopic properties of the hydrogel, 1 mL of the synthesized hydrogel was divided into thirds and subjected to lyophilization to obtain the initial dry weight (W0). Subsequently, the hydrogel was immersed in a 10 mL PBS solution and incubated at 37 °C for different time intervals (10 min, 30 min, 30 min, 30 min, and 30 min). At each time point, the hydrogel was removed, and its weight (Wt) was measured. The swelling ratio was then calculated using the formula:

$${\rm{Swelling}}\,{\rm{ratio }} = {\rm{ }}\left( {{\rm{Wt }} - {\rm{ W}}0} \right){\rm{ }}/{\rm{ W}}0{\rm{ }} \times {\rm{ }}100\%$$
(2)

Ion release assessment

The investigation of ion release from PADM@MgC hydrogels involved placing an equal mass of hydrogel in 10 mL of simulated body fluid (SBF, PH1820, China). The medium was entirely replaced daily with fresh SBF to maintain consistent conditions. Copper and zinc ion concentrations were quantified using an inductively coupled plasma atomic emission spectrometer (X Series 2, Thermo Fisher Scientific, USA). For comparative analysis, control hydrogels PADM and PADM@Mg were subjected to identical testing procedures.

Cells and bacteria

This study utilized mouse-derived macrophages (RAW264.7) and human umbilical vein vascular endothelial cells (HUVEC), both acquired from the Shanghai Institute of Cell Biology. These cells were cultured in DMEM medium, enriched with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin. Additionally, Methicillin-resistant Staphylococcus aureus (MRSA, ATCC 43,300) was procured from the American Type Culture Collection and propagated in tryptic soy broth (TSB; Hopebio).

Preparation of sample extracts

The PADM, PADM@Mg, and PADM@MgC hydrogels were sectioned into 10 × 10 × 5 mm dimensions and submerged in DMEM supplemented with 10% FBS and 1% penicillin/streptomycin. The cultures were incubated at 37 °C for 24 h, after which the extracts were obtained and preserved at 4 °C for subsequent analysis.

In vitro antibacterial tests

Frozen MRSA cultures were revived and incubated overnight in TSB at 37 °C on a shaker. This was followed by a dilution step at a 1:10,000 ratio, leading to further incubation until the cultures reached the logarithmic growth phase. Subsequently, bacterial suspensions with a concentration of 1 × 106 CFU/mL, in a volume of 500 µL, were introduced to a variety of hydrogels for incubation. The influence of these hydrogels on bacterial growth was evaluated through growth curve assays. These experiments were conducted in triplicate and replicated thrice to ensure the robustness of the data. Following this, the bacteria were co-cultured with the different hydrogels for 24 h. Post-incubation, SPM and colony-forming unit (CFU) quantification were carried out on agar plates to further assess the interaction between the bacteria and the hydrogels. Additionally, we analyzed the antibacterial activity by live/dead staining. Briefly, we added 500 µL of combined dye (Syto9 and PI) in different samples and cultured them for 15 min. Bacteria were collected on glass slides and visualized under a fluorescence microscope.

Membrane damage and leakage assessment

To detect membrane damage and leakage, bacteria and hydrogels were co-cultured for 24 h. Post-co-culture, SEM was employed to examine the morphological characteristics of the bacteria. In addition, the samples were stained with Propidium Iodide (PI) at a concentration of 2.5 µg/mL for 30 min. The fluorescence intensity of these stained samples was then measured at an excitation/emission wavelength of 535 nm/615 nm using a microplate reader. Subsequently, cell content leakage under various treatment conditions was evaluated. This involved centrifuging the bacterial supernatant post-co-culture at 8000 × g for 10 min, followed by measuring the absorbance at 260 nm for nucleic acids (A260) and 280 nm for proteins (A280) using a spectrophotometer. All experiments were conducted in triplicate to ensure consistency and accuracy.

In vitro immunomodulatory evaluation

Immunofluorescence

Briefly, RAW264.7 cells were incubated with PADM, PADM@Mg, or PADM@MgC hydrogels for one day. Subsequently, the cells were collected, fixed, and blocked before staining with antibodies against iNOS (ab210823) and CD206 (ab64693). Next, cells were incubated with corresponding secondary antibodies and counterstained with DAPI for 1 h. The cells were observed and photographed using CLSM.

Flow cytometry assay

Flow cytometry, employing antibodies against CD86 and CD206, was utilized to assess the ratios of M1 and M2 macrophages. Macrophages were initially cultured with various hydrogels for a 24-hour period. This was followed by a 40-minute blocking phase and staining with allophycocyanin (APC)-conjugated CD206 and phycoerythrin (PE)/Cy7-conjugated CD86. The final steps involved washing the cells, resuspending them in 400 µL of PBS, and their subsequent evaluation using flow cytometry.

Gene expression

To investigate gene expression profiles in macrophages, we undertook transcriptome sequencing analyses. RAW 264.7 cells, initially inoculated at 5 × 105 cells per well, were subjected to a 24-hour incubation with diverse hydrogels in an LPS-enriched environment. Following this, total cellular RNA was meticulously extracted using TRIzol reagent, in strict accordance with the manufacturer’s instructions. The subsequent RNA-sequencing and bioinformatics analyses were proficiently conducted by Novel Biotech, Shanghai. This process facilitated the examination of differentially expressed genes (DEGs) for their involvement in KEGG pathways and gene ontology enrichment.

RT-PCR and ELISA

RAW 264.7 cells were cultivated following the above outlined method and harvested after a 24-hour period. Total RNA was then meticulously extracted utilizing TRIzol. For accuracy in gene expression analysis, normalization was conducted against the housekee** gene ACTB. Details of the primer sequences used are comprehensively listed in the Supplementary Information (Table S1). Furthermore, culture supernatants were harvested from macrophages post-incubation with various hydrogels. The concentrations of cytokines IL-1β, TNF-α, IL-6, and IL-10 in these supernatants were quantitatively assessed employing Enzyme-Linked Immunosorbent Assay (ELISA).

Cell viability analysis

Cellular cytotoxicity was evaluated using a live/dead staining kit, applied as per the manufacturer’s guidelines. Additionally, to ascertain the biocompatibility of the hydrogels, a Cell Counting Kit-8 (CCK-8) assay was conducted. In this process, cells were co-cultured with different hydrogels at a density of 2 × 104 cells per well. After 24 h, for live cell staining, cells were rinsed and incubated with calcein and PI for dead cells, for 30 min. Fluorescence microscopy was utilized to capture images of the stained cells. Concurrently, at designated intervals (24 and 48 h), the culture supernatant was removed, and 500 µL of prepared CCK-8 solution was added. Following an additional 2-hour incubation, absorbance readings at 450 nm were taken using a BioTek microplate reader. This procedure was replicated thrice for each experimental group to ensure consistency of results. Moreover, post-incubation with hydrogels, cells were harvested, centrifuged at 1500 rpm for 5 min, and subsequently fixed in 2.5% glutaraldehyde at 4°C overnight. The samples then underwent a meticulous dehydration process through a graded ethanol series (30%, 50%, 70%, 80%, 90%, and 100%), followed by freeze-drying. After being sputter-coated with platinum to enhance conductivity, the morphological characteristics of the cells were meticulously examined using scanning electron microscopy.

In vitro migration study

Initially, HUVEC cells were seeded into 6-well plates and allowed to incubate for 24 h. Following this period, a precise line was drawn through each well’s center, after which the wells were gently washed twice with PBS. The cells were then exposed to extracts of PADM@MgC, PADM@Mg, and PADM, as well as complete medium for comparative purposes. To monitor cellular migration, images were captured at the outset and again after 12 h of treatment.

Tube formation assay

Initially, each well of a 96-well plate was prepped with 50 µL of growth factor-reduced Matrigel, followed by a 30-minute incubation to ensure proper solidification of the Matrigel. Subsequently, 2 × 104 HUVECs were carefully inoculated into each well containing the solidified Matrigel substrate. These wells were then divided and co-cultured with high-glucose DMEM infused with extracts from PADM@MgC, PADM@Mg, and PADM, alongside a complete medium serving as the control. Twelve hours post-incubation, the cells were stained using a calcein staining kit. Representative images capturing the tube formation were taken 6 h after treatment to document the cellular responses.

Mouse wound model and treatment procedures

The Animal Ethics and Welfare Committee of Chang Zheng Hospital granted approval for all the animal experiments involved in this study, as documented by approval number 2023 − 564. All surgical interventions were meticulously executed in alignment with the established animal care guidelines. Male ICR mice, aged between 4 and 5 weeks and weighing approximately 18–20 g, were selected to establish a model for wound infection research. The procedure initiated with the careful removal of fur from the mice’s backs. This was followed by the creation of an 8 mm diameter wound on each mouse, which was subsequently inoculated with 10 µL of a bacterial solution at a concentration of 2 × 108 CFU/mL. On the second day post-wounding, the mice were systematically categorized into four experimental groups: control (Ctrl), PADM, PADM@Mg, and PADM@MgC, with each group comprising eight mice. In brief, MRSA-infected wounds were administered a single treatment of either control (PBS), PADM, PADM@Mg, or PADM@MgC during the entire experimental period. The mice were provided unrestricted access to food and water. They were housed in microinsulators under regulated conditions of humidity, temperature, and a 12-hour light-dark cycle. Daily digital photographs of the wounds were captured, and the wound area was quantitatively assessed using Image J software. The percentage of wound closure was calculated using the following formula:

$$\eqalign{& {\rm{Scratch}}\,{\rm{open}}\,{\rm{area }}\left( \% \right){\rm{ }} \cr & = {\rm{ }}\left( {{\rm{remaining}}\,{\rm{wound}}\,{\rm{area/original}}\,{\rm{wound}}\,{\rm{area}}\,{\rm{vehicle}}} \right){\rm{ }} \times {\rm{ }}100 \cr}$$
(3)

Microbiological and histological examination

Upon euthanization of the mice on day 4 post-surgery, the infected tissues were meticulously harvested and homogenized in 5 mL of PBS for bacterial load assessment, with bacterial quantification executed as previously described. Subsequently, both wound and adjacent tissues were procured for comprehensive histological analysis. This included the application of Hematoxylin and Eosin (HE) staining and Giemsa staining to evaluate tissue morphology and infection status. Additionally, immunofluorescence techniques were employed to ascertain the expression levels of endogenous markers iNOS and CD206, providing insights into the inflammatory and vascular responses within the tissue.

Statistical analysis

Quantitative data were presented as the mean ± standard deviation (SD), except where noted differently. For the comparison between two distinct groups, a two-tailed Student’s t-test was employed. In cases involving multiple groups, analyses were performed using one-way analysis of variance (ANOVA), complemented by Tukey’s post hoc test for in-depth comparisons. All data calculations and statistical analyses were meticulously conducted using Excel 2016 and GraphPad Prism 9. The threshold for statistical significance was established at P < 0.05.