While numerous hydrogel dressings are available for treating chronically infected wounds, their clinical application is impeded by intricate preparation processes, low mechanical strength, and frequent reliance on exogenous antimicrobial agents. The latter often leads to antibiotic misuse and compromises the bioactivity of cell growth-promoting substances. Therefore, the field of therapeutic treatment is faced with a pressing need to prepare high-mechanical-strength hydrogels through a facile procedure, achieving endogenous antibacterial characteristics and long-term healing abilities for chronically infected wounds. In this study, employing the Michael addition reaction principle, we conducted the conjugate addition of the natural antimicrobial poly amino acid, ε-polylysine (ε-PL), to gelatin methacrylate. This not only augmented the hydrogel’s mechanical strength but also preserved its antimicrobial efficacy. Subsequently, platelet-rich plasma (PRP), capable of releasing diverse growth factors, was introduced. Injectable and degradable hydrogels with high mechanical strength and water absorption were prepared through UV curing while retaining PRP bioactivity. The combination of PRP and ε-PL substantiated the enhanced antimicrobial properties and promotion of human umbilical vein endothelial cell growth, as validated through bacterial inhibition experiments, such as live-dead bacterial staining and cellular assays, including cell proliferation. Therefore, the as-developed PL-PRP hydrogel presents as a promising hydrogel dressing for the treatment of chronically infected wounds.

As the primary protective barrier of the human body, the skin is the most delicate organ, vulnerable to damage from various factors such as mechanical trauma and pathological ulcers.1 Notably, skin wounds, being open, are prone to infection by common pathogenic bacteria, triggering intense inflammatory reactions. This not only significantly heightens infection-related complications but also considerably impedes wound healing. Consequently, chronically infected wounds present a formidable challenge in the realm of skin trauma.2 Wound dressing emerges as an integral component in the clinical management of chronically infectious wounds.3 Traditional dressings, like gauze and cotton, have limitations owing to their absence of hemostatic and antibacterial properties. Moreover, the act of refreshing or changing these dressings can lead to reinjury, including wound tearing, inflammation, or pain induction.4,5 Hydrogel, with its superior biocompatibility, degradability, and moisture retention properties, emerges as a promising material for wound dressings.6–11 Furthermore, hydrogel dressings foster the healing of chronically infectious wounds by providing three-dimensional support structures and attachment points for cell growth. They alleviate pain and improve hypoxic or anaerobic environments.12,13 Hydrogels with diverse functionalities, such as antibacterial effects, hemostatic characteristics, robust adhesion, oxidation resistance, anti-inflammatory properties, controlled drug release, and stimulus-responsive behavior, have been developed to cater to distinct medical requirements.14–20 Consequently, these hydrogel-based dressings are gradually supplanting traditional dressings in clinical treatments.

Despite the increasing utilization of hydrogel dressings in chronically infected wound healing, some hydrogels are intricate and costly to prepare. They also exhibit low strength and poor stability under external environmental stimuli.21 This affects the proximity of newly generated proteins in the microenvironment to the cells,22,23 thereby influencing the fate of the cells.24–26 Additionally, hydrogels are typically loaded with antimicrobial drugs or metal ions (Ag+) to enhance their antimicrobial properties. However, this practice may lead to antibiotic abuse, drug resistance, metal ion deposition,27 and the generation of a significant amount of reactive oxygen species (ROS) that accumulate in damaged wounds. This accumulation deteriorates the activity of fibroblasts, impeding skin repair.28–30 Consequently, these challenges have restrained the widespread adoption and application of hydrogel dressings in the market. Among the hydrogels, Gelatin methacryloyl (Gel-MA), a semi-synthetic hydrogel with photoactive methacrylate groups interspersed in a natural gelatin framework, has gained prominence.31 Gel-MA is widely recognized for its injectability, high biocompatibility, ability to enhance cellular adhesion and growth, biodegradability, photocrosslinking capability, relative ease of synthesis, and cost-effectiveness.32,33 However, Gel-MA also exhibits limitations, including low strength, lack of antimicrobial properties, and a restricted ability to promote the rapid proliferation of cells and tissues.34,35 Previous studies often involve adding antimicrobial substances, such as metal ions, antibiotic drugs, or herbal extracts, to the Gel-MA skeleton to confer antimicrobial properties.36–38 Alternatively, a single growth factor, stem cells, or their derivatives are added to facilitate the proliferation and development of cells and tissues.39,40 Nevertheless, these modifications alone fail to address the issues of antibiotic abuse, metal ion deposition, and simultaneous antimicrobial and cell tissue proliferation and development. Even when these active substances are added simultaneously, exogenous antimicrobial substances inhibit the activity and growth of stem cells and peptide-like substances.41,42 Furthermore, directly adding the aforementioned active substances to Gel-MA does not enhance its mechanical strength,43 limiting its efficacy as a wound dressing. Consequently, there is an urgent clinical need to develop hydrogels with high mechanical strength, ease of processing, endogenous antimicrobial properties, and long-term promotion of wound healing.

ε-Polylysine (ε-PL), a natural antimicrobial agent, serves as a positively charged peptide containing –NH2 and has gained widespread recognition for its application as an endogenous antimicrobial material in the formulation of hydrogel dressings.44,45 Nevertheless, hydrogels solely composed of ε-PL exhibit weak mechanical properties and prove inadequate in withstanding external pressure on wounds.46 Additionally, PRP, derived from the patient’s blood and characterized by a high concentration of platelets, is abundant in essential growth factors and cytokines,47 including exosomes, nerve growth factor (NGF), and platelet-derived growth factor (PDGF), all of which contribute to promoting cell and tissue regeneration.48–50 Incorporating PRP into fibrin glue hydrogel (PRP gel) yielded promising outcomes in both clinical practice and research, particularly in the treatment of chronic wounds.51–53 However, PRP gel has inherent limitations, such as poor stability, rapid degradation, and a lack of antibacterial properties. In some instances, it may even inadvertently promote bacterial growth, leading to suboptimal clinical results.54,55

In this study, a hydrogel was formulated by conjugating ε-PL with gelatin methacrylate and incorporating platelet-rich plasma (PL-PRP hydrogel). Our design addresses the challenges associated with the low mechanical strength of Gel-MA and ε-PL, the absence of antimicrobial properties in Gel-MA hydrogel, and the potential for antibiotic resistance. This is achieved by introducing positively charged ε-PL to Gel-MA through conjugate addition, all while retaining the favorable attributes of Gel-MA, such as high biocompatibility, injectability, degradation, and rapid hydrogel formation through light curing. Simultaneously, the hydrogel can electrostatically adsorb negatively charged proteins in the extracellular matrix, providing an optimal cellular microenvironment for cell growth. The inclusion of PRP in ε-PL-modified Gel-MA is informed by the findings that PRP can facilitate rapid cell growth and elongated cell synapse under ultraviolet light (300–500 nm). This approach prolongs the release of growth factors,56 thereby addressing the dual challenge of antimicrobial efficacy and sustained promotion of cell tissue proliferation. Figure 1 illustrates the synthesis and characterization of methacryloyl gelatin (Gel-MA), ε-PL-modified Gel-MA (Gel-PL), and platelet-rich plasma-conjugated Gel-PL (PL-PRP). Consequently, our study presents a hydrogel with a straightforward preparation process, injectability, degradability, high mechanical strength, antimicrobial properties, and long-term promotion of cell proliferation. This is achieved through the utilization of natural amino acids and self-extracted PRP, making it highly promising for application in chronically infected wounds.

FIG. 1.

Preparation process and properties of PL-PRP hydrogel. (a) Synthesis process of Gel-MA polymer. (b) Synthesis process of Gel-PL polymer. (c) Preparation process and properties of PL-PRP hydrogel.

FIG. 1.

Preparation process and properties of PL-PRP hydrogel. (a) Synthesis process of Gel-MA polymer. (b) Synthesis process of Gel-PL polymer. (c) Preparation process and properties of PL-PRP hydrogel.

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Before delving into the characterization of PL-PRP hydrogels, we conducted an assessment of the synthesized Gel-MA and Gel-PL using Fourier Transform Infrared Spectroscopy (FTIR), as depicted in Fig. 2(a). The Gel-MA polymer exhibited an amide I band at 1654 cm−1 and an amide II band at 1542 cm−1, indicative of the reaction between gelatin and methacrylic anhydride. This suggests the successful incorporation of methacrylamide groups into the gelatin backbone.57 The band at approximately 2900 cm−1, representing CH stretching due to ε-PL, signifies the successful integration of ε-PL onto the Gel-MA backbone. Further analysis of the chemical structures of Gel-MA, ε-PL, and Gel-PL through 1H NMR revealed two strong peaks at δ 5.3 and 5.6 ppm corresponding to acrylic protons, with peaks at δ 2.0 and 2.9 ppm representing methyl groups. The 1H NMR spectrum of the Gel-MA polymer exhibited a significant peak for acrylic protons, confirming successful synthesis. The diminished resonance signal for acrylic protons at δ 5.3 and 5.6 ppm, coupled with heightened peaks at δ 3.79, 3.09, 1.74, 1.45, and 1.27 ppm, indicated the successful synthesis of the Gel-PL polymer [Fig. 2(b)]. Zeta potential results indicated the Gel-MA polymer’s negative charge (−10.7 mV), while the Gel-PL polymer possessed a positive charge of +9.3 mV after reaction, reaffirming the success of the process [Fig. 2(c)].

FIG. 2.

Characterization of Gel-MA and Gel-PL polymers. (a) FTIR spectrum of ε-PL, Gel-MA, and Gel-PL polymer. (b) 1H NMR spectra of ε-PL, Gel-MA, and Gel-PL polymers in D2O. (c) Zeta potential of ε-PL, Gel-MA, and Gel-PL polymer. (d) Infrared spectra and (e) 1H NMR spectra of Gel-PL polymers synthesized by feeding different quantities of ε-PL. (f) Representative stress-strain curves of Gel-PL hydrogels synthesized with different quantities of ε-PL. Scanning electron microscopy (SEM) of freeze-dried hydrogels synthesized by feeding (g) 0.005 mmol, (h) 0.01 mmol, and (i) 0.02 mmol ε-PL. The scale bars represent 10 µm.

FIG. 2.

Characterization of Gel-MA and Gel-PL polymers. (a) FTIR spectrum of ε-PL, Gel-MA, and Gel-PL polymer. (b) 1H NMR spectra of ε-PL, Gel-MA, and Gel-PL polymers in D2O. (c) Zeta potential of ε-PL, Gel-MA, and Gel-PL polymer. (d) Infrared spectra and (e) 1H NMR spectra of Gel-PL polymers synthesized by feeding different quantities of ε-PL. (f) Representative stress-strain curves of Gel-PL hydrogels synthesized with different quantities of ε-PL. Scanning electron microscopy (SEM) of freeze-dried hydrogels synthesized by feeding (g) 0.005 mmol, (h) 0.01 mmol, and (i) 0.02 mmol ε-PL. The scale bars represent 10 µm.

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To optimize the performance of PL-PRP hydrogels, we meticulously selected Gel-PL with superior performance, factoring in both porosity and mechanical strength. The impact of ε-PL grafting on the Gel-MA skeleton on Gel-PL’s porosity and mechanical strength was assessed. Following a previously proposed method,58 we gauged the grafting degree of ε-PL by analyzing peak areas at 2900 cm−1 in the FTIR spectrum and at δ 5.6 ppm in the 1H NMR spectrum [Figs. 2(d) and 2(e)]. Using Gel-MA spectra as the baseline, ε-PL grafting degrees were determined as 28%, 44%, and 68% at ε-PL quantities of 0.005, 0.01, and 0.02 mmol, respectively. Gel-PL hydrogels were then fabricated through light curing, and their mechanical strength was assessed. Results indicated an increase in compression modulus with higher quantities of added ε-PL [Fig. 2(f)]. Additionally, freeze-dried Gel-PL hydrogels exhibited a porous structure, with pore size decreasing as more ε-PLs were incorporated into the Gel-MA backbone [Figs. 2(g), 2(h), and 2(i)]. The determined porosity values were 55% ± 1%, 61% ± 1%, and 65% ± 1% at ε-PL quantities of 0.005, 0.01, and 0.02 mmol, respectively (Fig. S1, supplementary material). The significance of porosity lies in its internal structure, often linked to hydrogel crosslinking density. Prior studies have indicated that higher porosity is advantageous for tissue growth, cell adhesion, proliferation, and oxygen exchange.59 However, it may result in decreased strain stress. Consequently, Gel-PL fed with 0.01 mmol ε-PL was selected for subsequent characterization, striking a balance between high porosity and mechanical properties.

PL-PRP hydrogels were formulated by introducing PRP and a photoinitiator to the 0.01 mmol Gel-PL polymer solution, as selected in Sec. II A, and subsequently cured under UV light. The detailed preparation method is outlined in the experimental section. Figure 3(a) illustrates the PL-PRP hydrogel alongside comparative Gel-MA, Gel-PRP, and Gel-PL hydrogels post-UV curing. A supplementary material video capturing changes in each hydrogel group before and after UV curing is available. Notably, this affirms that the conjugation of ε-PL or the addition of PRP does not influence the gelation process. The morphological features of the hydrogel cross-sections for each group were examined using scanning electron microscopy (SEM), revealing a three-dimensional (3D) porous structure with interconnected internal pores [Fig. 3(b)]. This architecture facilitates tissue growth, cell adhesion, proliferation, and oxygen exchange.60 Subsequently, we measured the gelation time of each hydrogel group using the syringing bottle tilt method. In contrast, PL-PRP hydrogels exhibited the shortest gelation time and superior rapid remodeling performance [Fig. 3(c)]. The presence of attractive electrostatic interactions proves pivotal in expediting the gelation process. Comparative analysis with Gel-MA revealed an increase in the porosity of Gel-PRP, Gel-PL, and PL-PRP hydrogels. The augmentation was more pronounced for PL-PRP and Gel-PL hydrogels, with the pore size of PL-PRP hydrogel surpassing that of Gel-PL hydrogel. This resulted in a more homogeneous porous structure with relatively thinner pore walls and a more compact network [Fig. 3(d)]. Such a property is particularly advantageous for cell proliferation, blood vessel formation, and tissue regeneration.61,62

FIG. 3.

Morphology, gelation time, porosity, mechanical properties of PL-PRP hydrogel. (a) Images of Gel-MA, Gel-PRP, Gel-PL, and PL-PRP hydrogels after UV curing. (b) Scanning electron microscopy (SEM) images of the cross-sections of the freeze-dried Gel-MA, Gel-PRP, Gel-PL, and PL-PRP hydrogel. The scale bars represent 10 µm. (c) Gelation time, (d) porosity, and (e) stress–strain curves of different hydrogels.

FIG. 3.

Morphology, gelation time, porosity, mechanical properties of PL-PRP hydrogel. (a) Images of Gel-MA, Gel-PRP, Gel-PL, and PL-PRP hydrogels after UV curing. (b) Scanning electron microscopy (SEM) images of the cross-sections of the freeze-dried Gel-MA, Gel-PRP, Gel-PL, and PL-PRP hydrogel. The scale bars represent 10 µm. (c) Gelation time, (d) porosity, and (e) stress–strain curves of different hydrogels.

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To assess the mechanical properties of PL-PRP hydrogels, compressive stress–strain tests were conducted on each hydrogel group. Results revealed that Gel-MA and Gel-PRP hydrogels could endure approximately 20% strain, reaching maximum stresses of about 0.20 and 0.12 MPa, respectively. The incorporation of ε-PL significantly enhanced the maximum stress, reaching 0.80 MPa. In contrast, PL-PRP hydrogels demonstrated the capacity to withstand nearly 50% strain, achieving a maximum stress of 1.00 MPa [Fig. 3(e)], a notably higher value than the compression modulus of subcutaneous tissue (4.5–8 kPa).63 Given PRP’s gel-forming properties,64 the synergy of a dual gel network enhances the mechanical strength of PL-PRP hydrogels.

Subsequently, the rheological properties of PL-PRP hydrogels under oscillatory shear were examined to further elucidate their mechanical characteristics. We explored the relationship between storage and loss modulus in response to strain through oscillating amplitude scanning, determining the linear viscoelastic region of the hydrogel. Figure 4(a) illustrates that the storage modulus (G′) and loss modulus (G″) of hydrogels in each group remained constant until 30%. Beyond this point, the intersection of G′ and G″ appeared in the Gel-MA and Gel-PRP hydrogel, indicating a transition from gel to sol state in Gel-MA and Gel-PRP hydrogels due to internal structural damage. In Gel-PL and PL-PRP hydrogels, G′ exceeded G″ by nearly two orders of magnitude, signifying that over 99% of energy was stored in the hydrogel network as elastic potential energy during deformation. PL-PRP exhibited a higher G′, indicating superior elastic properties. Frequency scanning measurements in the linear elastic region [Fig. 4(b)] further revealed that G′ and G″ values remained constant in the 0.1–100 Hz frequency range, aligning with the rheological characteristics of polysaccharide hydrogels.65 This suggested that the incorporation of ε-PL and PRP did not alter the fundamental structure of the Gel-MA backbone. With G′ surpassing G″, indicative of the predominance of the elastic component, PL-PPR hydrogel demonstrated the highest G′, reflecting enhanced resistance to external disturbances and superior mechanical properties. Thereafter, we explored the viscosity of PL-PRP hydrogels as a function of shear rate, as depicted in Fig. 4(c). The viscosity displayed a decreasing trend with an increasing shear rate, showcasing shear-thinning properties.

FIG. 4.

Rheological properties, swelling rate, and in vitro degradation behavior of PL-PRP hydrogels. (a) Amplitude sweep, (b) frequency sweep, and (c) viscosity sweep of different hydrogels in the rheological test. (d) Swelling rate and (e) degradation rate of different hydrogels. (f) Images of different hydrogels during degradation.

FIG. 4.

Rheological properties, swelling rate, and in vitro degradation behavior of PL-PRP hydrogels. (a) Amplitude sweep, (b) frequency sweep, and (c) viscosity sweep of different hydrogels in the rheological test. (d) Swelling rate and (e) degradation rate of different hydrogels. (f) Images of different hydrogels during degradation.

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To validate the remarkable water absorption capabilities of PL-PRP hydrogel, we calculated its in vitro swelling rate. The results, depicted in Fig. 4(d), revealed that PL-PRP exhibited the highest swelling rate at 388%, surpassing the other three hydrogel groups by 2.2, 1.5, and 1.4 times, respectively. This can be attributed to the increased hydrophilicity resulting from the reduction of double bonds in the polymer, thereby enhancing the interaction between the hydrogel and water molecules.60 Additionally, PRP extracted from blood contributes to excellent hydrophilicity. Following the addition of PRP, the adsorption between the hydrogel and water molecules is enhanced, facilitating the penetration and diffusion of water within the hydrogel and augmenting its swelling ability.66 

To assess the degradability of PL-PRP hydrogels, we explored the in vitro degradation behavior by immersing each hydrogel group in PBS containing type II collagenase (0.2 U/ml) and monitoring weight changes over time. The results indicated that the degradation rate followed the sequence PL-PRP < Gel-PL < Gel-PRP < Gel-MA hydrogels [Figs. 4(e) and 4(f)]. This finding aligns with the notion that the degradation rate of Gel-MA is primarily contingent on the degradation rate of collagen to gelatin.67 However, our previous experimental results demonstrated that increases in the porosity and crosslink density of PL-PRP, Gel-PL, and Gel-PRP hydrogels resulted in a decline in substances available for degradation, leading to a decrease in the degradation rate. Despite this, the PL-PRP hydrogel exhibited complete degradation within 12 days. This suggests that the slow-degrading nature of PL-PRP hydrogel is particularly advantageous for the prolonged reconstruction of chronically infected wounds.

The antimicrobial efficacy of PL-PRP was assessed through inhibition rate tests, inhibition circles, bacterial plate counts, bacterial live-dead tests, and SEM for bacterial morphology. Results from the bacterial inhibition rate test revealed that Gel-MA and Gel-PRP hydrogels exhibited no antimicrobial effect against Escherichia coli and Staphylococcus aureus. Intriguingly, Gel-PRP hydrogel even promoted bacterial growth due to the presence of PRP. In contrast, PL-PRP and Gel-PL hydrogels demonstrated inhibition rates of 60% ± 2% and 70% ± 4% against E. coli and 58% ± 4% and 73% ± 1% against S. aureus, respectively [Figs. 5(a) and 5(b)]. The presence of ε-PL in Gel-PL and PL-PRP hydrogels played a pivotal role in exerting their biological effect to inhibit bacterial growth.68 The lower inhibition rate of PL-PRP compared to Gel-PL is attributed to the inclusion of PRP, which can release various growth factors providing nutrition to bacteria. Importantly, this indirectly confirms that the natural endogenous antimicrobial agent, ε-PL, does not compromise the biological activity of proteins and peptides.

FIG. 5.

Antibacterial properties of PL-PRP hydrogel. (a) Antibacterial rate of different hydrogels against E. coli. (b) Antibacterial rate of different hydrogels against S. aureus one-way ANOVA, ***p ≤ 0.001, n = 3. Scanning electron microscopy (SEM) images of E. coli (c) and S. aureus (d) co-cultured with different hydrogels. Scale bar: 1 µm. Plate count of E. coli (e) and S. aureus (f) co-cultured with different hydrogels.

FIG. 5.

Antibacterial properties of PL-PRP hydrogel. (a) Antibacterial rate of different hydrogels against E. coli. (b) Antibacterial rate of different hydrogels against S. aureus one-way ANOVA, ***p ≤ 0.001, n = 3. Scanning electron microscopy (SEM) images of E. coli (c) and S. aureus (d) co-cultured with different hydrogels. Scale bar: 1 µm. Plate count of E. coli (e) and S. aureus (f) co-cultured with different hydrogels.

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SEM results of bacteria post-inhibition [Figs. 5(c), 5(d), and S2, supplementary material] demonstrated that bacteria co-cultured with Gel-MA hydrogel maintained intact cell walls, comparable to the control group. Gel-PRP hydrogel, on the other hand, led to an increased bacterial population due to the proliferative effect of PRP, in addition to maintaining intact cell walls. In contrast, bacteria co-cultured with Gel-PL and PL-PRP hydrogels not only experienced significant growth inhibition but also suffered notable damage to their cell walls. This damage can be attributed to the electrostatic interaction between the positively charged groups of Gel-PL and PL-PRP and the negatively charged bacterial cell membrane, resulting in membrane damage. Consequently, intracellular fluid is released, ultimately leading to bacterial death.66,69 The results of bacterial plate counting [Figs. 5(e) and 5(f)] and the inhibition circle experiments [Fig. S3, supplementary material] corroborated these findings.

Live-dead fluorescence staining experiments (Fig. 6) visualized the destruction of bacterial structures co-cultured with PL-PRP and Gel-PL hydrogels, stained red by the PI staining solution. In conclusion, these antimicrobial experiments demonstrated that PL-PRP hydrogels could disrupt bacterial structures, effectively inhibit bacterial proliferation, and possess antimicrobial properties. Although the antibacterial effect is weaker than that of Gel-PL, PL-PRP hydrogel is more suitable for the clinical application of chronically infected wounds, considering its mechanical strength, water absorption, and other comprehensive properties.

FIG. 6.

Live and dead fluorescence staining images of (a) E. coli and (b) S. aureus co-cultured with different hydrogels. The scale bars represent 100 µm. Dead bacteria ratio of E. coli (c) and S. aureus (d) co-cultured with different hydrogels. SYT09 represents live bacteria, while PI represents dead bacteria. One-way ANOVA, ***p ≤ 0.001, n = 3.

FIG. 6.

Live and dead fluorescence staining images of (a) E. coli and (b) S. aureus co-cultured with different hydrogels. The scale bars represent 100 µm. Dead bacteria ratio of E. coli (c) and S. aureus (d) co-cultured with different hydrogels. SYT09 represents live bacteria, while PI represents dead bacteria. One-way ANOVA, ***p ≤ 0.001, n = 3.

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To validate the cell and tissue growth promotion capability of PL-PRP hydrogel, human umbilical vein endothelial cells (HUVEC) were co-cultured with PL-PRP hydrogel [Fig. S(4), supplementary material], with DMEM medium serving as the control. Observations on the effects of PL-PRP hydrogel on cell survival, vitality, and morphology were conducted using the CCK-8 assay, fluorescence staining, and cytoskeleton staining. Initial verification confirmed the continuous and stable release of vascular endothelial growth factor by PL-PRP hydrogel through an in vitro growth factor release assay [Fig. 7(a)]. Subsequently, the cell viability assay results indicated that Gel-MA and Gel-PL hydrogels, while not cytotoxic, lacked the ability to promote cell proliferation compared to the control group. In contrast, both PL-PRP and Gel-PRP hydrogels exhibited enhanced cell viability [Fig. 7(b)]. Live-dead staining assay results revealed high cell viability across all hydrogels with minimal dead cells compared to the control group. Moreover, PL-PRP and Gel-PRP hydrogels demonstrated a significant increase in viable cells compared to other groups [Figs. 7(c) and S(5), supplementary material]. This enhancement is attributed to their PRP content, capable of releasing various growth factors, including vascular endothelial growth factor, fostering vessel wall permeability, and promoting the growth and proliferation of endothelial and perivascular cells. Consequently, angiogenesis is enhanced, facilitating tissue regeneration in vivo.70–72 Although Gel-PRP is comparable to PL-PRP in promoting cell proliferation, its lack of antimicrobial properties and poor mechanical strength render PL-PRP the preferred choice—a high-mechanical-strength hydrogel with endogenous antimicrobial and long-term wound-healing-promoting functions, aligning with current clinical requirements.

FIG. 7.

Promotion of cell proliferation by PL-PRP hydrogel. (a) Growth factor release over time for PL-PRP hydrogel. (b) Cell viability of HUVEC co-cultured with different hydrogels. One-way ANOVA, ***p ≤ 0.001, n = 5. (c) Live staining images of HUVEC co-cultured with different hydrogels. The fluorescent dye Calcein-M labeled living cells, while PI stained dead cells. The scale bars represent 100 µm.

FIG. 7.

Promotion of cell proliferation by PL-PRP hydrogel. (a) Growth factor release over time for PL-PRP hydrogel. (b) Cell viability of HUVEC co-cultured with different hydrogels. One-way ANOVA, ***p ≤ 0.001, n = 5. (c) Live staining images of HUVEC co-cultured with different hydrogels. The fluorescent dye Calcein-M labeled living cells, while PI stained dead cells. The scale bars represent 100 µm.

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Cytoskeleton staining experiments illustrated that cells in all groups exhibited spindle shapes and good stretching compared to the control group. However, the number of HUVEC co-cultured with PL-PRP was significantly increased, indicating the ability of HUVEC cells to spread and adhere effectively in the PL-PRP hydrogel [Fig. 8]. This cytoskeletal extension allowed cells to form an adaptive structural scaffold, facilitating nutrient and oxygen exchange while reducing the communication distance between cells. These factors play a crucial role in regulating cellular function and adaptation to the surrounding environment.73 In conclusion, these findings suggest that PL-PRP hydrogel rapidly promotes the long-term proliferation and growth of vascular endothelial cells, thereby contributing to effective wound healing.

FIG. 8.

Fluorescent images illustrating cytoskeleton staining of HUVEC cells co-cultured with various hydrogels. Rhodamine B (RhB) marked cellular actin, while 4′,6-diamidino-2-phenylindole (DAPI) labeled the nucleus. The scale bars represent 100 µm.

FIG. 8.

Fluorescent images illustrating cytoskeleton staining of HUVEC cells co-cultured with various hydrogels. Rhodamine B (RhB) marked cellular actin, while 4′,6-diamidino-2-phenylindole (DAPI) labeled the nucleus. The scale bars represent 100 µm.

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The management of chronically infected wounds poses a formidable challenge in cutaneous trauma—difficult to treat, prolonged in duration, and economically burdensome for patients and the healthcare system. In response, we have engineered an injectable, degradable, high-strength hydrogel with high water absorption incorporating the natural antimicrobial agent ε-PL through conjugate addition with Gel-MA. This hybridization not only augments the mechanical robustness of the hydrogel but also preserves the antimicrobial and non-detrimental cellular and bioactive properties of ε-PL. Furthermore, the infusion of autologous PRP ensures a sustained release of diverse growth factors, fostering prolonged cell proliferation. The hydrogel developed herein can also promote long-term cellular tissue proliferation, addressing the pressing clinical need for high-mechanical-strength hydrogel wound dressings with facile preparation processes, endogenous antimicrobial properties, and the ability to promote the healing of infected wounds in the long term, expected to fulfill an unmet need in chronic wound care.

To formulate the PL-PRP hydrogel, 100 µl of PRP was introduced into the Gel-PL solution (15%, w/v) at 37 °C. Following thorough stirring, Irgacure (0.3%, w/v) was dissolved in the solution. Subsequently, the mixture was placed in a mold and exposed to UVA (365 nm, 610 mW/cm−2, Guangzhou, China) light for 30 s from a distance of 10 cm to initiate photocrosslinking of the hydrogel.56 Using an identical preparation protocol, Gel-PRP hydrogel was prepared by adding 100 µl of PRP to the Gel-MA solution.

The gelation time was determined using the tilt method. After ultraviolet irradiation, the penicillin bottle was tilted to observe solution flow. The gelation time was determined based on the time when the solution stopped flowing upon bottle tilting.

The internal morphologies of the Gel-MA, Gel-PL, and PL-PRP hydrogels were observed using samples sprayed with a thin gold layer after freeze-drying through scanning electron microscopy (SEM, Zeiss Gemini Sigma 300, Germany).

The mechanical properties of hydrogels were assessed using a universal testing machine (INSTRON, 6800, USA). Specifically, 400 µl of hydrogels, shaped into cylinders with a height of 10 mm and a diameter of 16 mm, was compressed at a controlled rate of 5 mm/min until fracture. Each sample was tested three times.

Rheological measurements were conducted using an oscillatory rheometer (Anton Paar, H-PTD200, Austria). (1) In the amplitude sweep (γ = 0.1%–1000%), the gel–sol point and linear viscoelastic region were determined at a constant frequency of 0.1 Hz at 25 °C. Before data collection, the hydrogel disk (15.4 mm diameter, 400 µl) was placed between 20 mm parallel plates with a gap of 1200 µm. (2) Employing the same configuration, the viscoelasticity of hydrogels was measured in a frequency sweep with a constant strain of 1% at 25 °C. (3) The shear-thinning effect was assessed using 400 µl of the polymer mixture between 20 mm parallel plates under a constant strain of 1% at 25 °C, with a gap of 900 mm.

A 400 µl volume of hydrogel was placed in a mold, and its initial mass (Ws) was measured. Subsequently, the hydrogel was extracted at 30-min intervals after immersion in a PBS solution at 37 °C. The mass, post-absorbing residual moisture with absorbent paper, was recorded as Wt. The experiment concluded when the weight of the hydrogel stabilized. The swelling ratio (SR) was calculated using the following equation:
SR(%)=WtWs/Ws×100%.
(1)

Ws is the initial hydrogel mass, and Wt is the mass after swelling at each interval. The experiment was conducted thrice.

For in vitro degradation assessment, hydrogels at swelling equilibrium (weighed as Wa) were placed in 15 ml PBS with collagenase Type II (0.2 U/ml) at 37 °C. Extracted at predetermined intervals, the hydrogels’ weight (Wt) was recorded after removing water with filter paper. All data were triply tested, and the degradation rate equation is defined as
DR (%)=WaWt/Wa×100%,
(2)
where Wa is the initial weight, and Wt is the weight remaining at different times.

HUVECs obtained from the Research Center of China-Japan Union Hospital of Jilin University were selected for the experiment. They were resuscitated from a liquid nitrogen tank and cultured for subsequent experiments.

  1. Cell viability assay: Hydrogel (100 µl) was prepared on a 96-well plate by photocrosslinking. Subsequently, 6 × 103 cells per well were seeded on the hydrogel surface. After 24 h of culture, the medium was replaced with 100 µl of DMEM containing 10 µl of CCK-8 reagent. After incubation at 37 °C in the dark for 1 h, the absorbance (OD) at 450 nm was measured using an enzyme-labeled instrument, and the data were recorded and statistically analyzed to generate a histogram of cell viability.

  2. Cell living staining assay: Hydrogel (100 µl) was prepared on a 24-well plate by photocrosslinking. Then, 6 × 103 cells per well were seeded on the surface of hydrogel and cultured for 24 h. Then, live/dead working solution under dark conditions was prepared according to the ratio of 2 ml of buffer +4 µl of Calcein-AM + 6 µl of PI. The medium in each well was discarded and rinsed twice with PBS. Subsequently, 300 µl of the working solution was added to each well and incubated at 37 °C for 20 min under dark conditions, and the excess working solution was rinsed with PBS. A certain amount of culture solution was added again and observed using a confocal microscope (C2, Nikon, Japan). Semi-quantitative analysis was performed using Image J software.

  3. Cell stretching experiment: Hydrogel (100 µl) was prepared on a 96-well plate by photocrosslinking. Then, 3 × 103 cells per well were seeded on the surface of the hydrogel. After 24 h of culture, the cells were stained with Rhb-phalloidin (phalloidin labeled with RhB) and DAPI to observe the cytoskeleton and nucleus. The cell morphology was observed using a confocal microscope.

  1. Inhibition rate test: Ninety microliters of bacterial liquid (1 × 106 CFU/ml) were added to a 96-well plate, with six groups, each containing five wells. The blank control group received 10 µl of PBS solution, the negative control group received 10 µl of ε-PL (0.04 g/ml), and the remaining groups were treated with 10 µl of hydrogel solution (PBS). After 24 h of culture, the absorbance (OD) at 600 nm was measured using an enzyme-labeled instrument. The collected data were statistically analyzed to generate a histogram representing the bacteriostatic rate.

  2. Bacteriostatic ring test: Approximately 20 ml of plate culture medium was poured into a sterilization plate, left to solidify horizontally, and coated with 0.1 ml of bacterial liquid (1 × 106 CFU/ml). Filter paper pieces (diameter of ∼1 cm) soaked in various hydrogel solutions (PBS) for 30 min were evenly placed on the culture medium coated with bacterial liquid. After 24 h of culture, the diameter of each bacteriostatic ring was measured.

  3. Bacterial live staining test: A 12-well plate containing 1 ml of bacterial solution (1 × 106 CFU/ml) was placed in a bacterial incubator at 37 °C for 24 h. The medium was then removed, centrifuged at 10 000 g for 15 min, and washed three times with 0.9% sodium chloride solution. A working solution for live-dead bacteria staining was prepared by diluting 10 µl SYT09 and 6 µl PI into 2 ml of 0.9% sodium chloride solution. Bacteria were resuspended in 500 µl of staining solution, incubated in the dark for 20 min at room temperature, and then washed three times with 0.9% sodium chloride solution. Bacteria were suspended in 0.9% NaCl solution and photographed with a confocal microscope.

  4. Observation of bacteria morphology and number after hydrogel inhibition: A bacterial suspension (100 µl, 1 × 106 CFU/ml) was dropped on the hydrogel surface and incubated at 37 °C for 24 h. Subsequently, 10 µl of the bacterial suspension was transferred to a silicon wafer, fixed with a 2.5 wt. % glutaraldehyde solution for 1 h, and dehydrated in ethanol solutions with different concentrations for 10 min. Finally, the samples were gold-sprayed for SEM inspection.

  5. Bacterial plate counting assay: The bacterial suspension (100 µl, 1 × 106 CFU/ml) was dropped on the hydrogel surface and incubated at 37 °C for 24 h. The antibacterial bacterial suspension (100 µl) was dropped on a Petri dish and cultured in an incubator for 24 h to form colonies.

Summary statistics for analysis variables are presented as the mean ± standard deviation (SD) unless otherwise noted. Statistical significance was determined through the one-way ANOVA test. A significance level of p < 0.05 was considered indicative of statistically significant differences.

See the supplementary material for a comprehensive understanding of the materials, methods, and additional results referred to in this manuscript, and detailed information is available online.

This research project was financially supported by the Special Project of Health Research Talents in Jilin Province (Grant No. 2020slz05). We thank the Research Center of China-Japan Friendship Hospital of Jilin University for providing the experimental platform for us.

The authors declare no conflicts of interest.

The Ethics Committee of China-Japan Union Hospital of Jilin University approved the procedure adopted for the extraction and application of human platelet-rich plasma (20231101).

Peiyu Yan: Conceptualization (equal); Data curation (equal); Formal analysis (equal); Investigation (equal); Methodology (equal); Validation (equal); Visualization (equal); Writing – original draft (equal). Xiangru Chen: Data curation (supporting); Formal analysis (supporting); Investigation (supporting). Xin He: Formal analysis (supporting); Methodology (supporting). Zhaoyang Liu: Formal analysis (supporting); Methodology (supporting). Jing Sun: Conceptualization (equal); Data curation (equal); Funding acquisition (supporting); Investigation (equal); Methodology (equal); Supervision (lead); Validation (equal); Visualization (equal); Writing – review & editing (lead).

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

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