Immunoregulative coating for scarless healing in anterior cruciate ligament reconstruction

Abstract

Polyethylene terephthalate (PET) artificial ligaments are widely used in anterior cruciate ligament (ACL) reconstruction due to their high tensile strength. However, bone tunnel enlargement around PET ligaments poses a risk for surgical failure. PET's inert surface, lower bioactivity, and mechanical abrasion trigger an M1 macrophage-mediated inflammatory response, leading to excessive, disorganized scar tissue. This scar tissue creates a space-occupying effect at the interface, obstructing graft-bone integration and contributing to bone tunnel enlargement. To address this issue, we developed a multi-layered immune-regulating hydrogel coating for scar-free PET-bone integration. Comprising gelatin methacrylate (GelMA), polyethyleneglycol diacrylate (PEGDA), and sulfated polysaccharide (SCS), the hydrogel forms a hydrogen-bonded lubricating layer to reduce friction. The sustained release of SCS also down-regulates M1 macrophage polarization, inhibiting early scar formation. By eliminating the space-occupying effect of scar tissue, SCS subsequently promotes M2 macrophage polarization. This shift releases endogenous factors that enhance blood vessel formation and new bone growth, ultimately achieving high-quality graft-bone integration. The application of this multi-layered, inflammation-modulating hydrogel coating not only removes scar tissue barriers but also improves graft-bone integration through enhanced angiogenesis and osteogenesis. Moreover, it avoids the overuse of exogenous growth factors and potential complications, offering a more convenient and feasible therapeutic strategy.

Graphical abstract

Highlights

• •Hydrogel coating reduces friction and inflammation in ACL reconstruction with immunomodulatory effects.
• •Degrading hydrogel releases SCS to promote M2 polarization, aiding angiogenesis and osteogenesis.
• •Reduces PET-bone friction, modulates inflammation, limits scar tissue, and enhances graft-bone integration.

1. Introduction

The incidence of anterior cruciate ligament (ACL) injuries has been increasing in recent years [1]. ACL injuries result in knee joint instability, pain, and functional impairment, escalating the risk of cartilage and meniscus damage and subsequently raising the prevalence of osteoarthritis. The ACL forms a robust mechanical connection between the ligament and bone through a structural gradient in the extracellular matrix (ECM) [2,3]. However, due to the complex nature of the native ligament-bone interface, it lacks an effective self-repair mechanism post-injury. For athletes, ACL reconstruction is the gold standard for treating ACL injuries. Common graft options in clinical practice include autografts, allografts, and polyethylene terephthalate (PET) artificial ligaments. PET artificial ligaments are favored in ACL reconstruction due to their ample supply, superior mechanical strength, and potential for rapid postoperative recovery. Nonetheless, post-implantation, there is a documented phenomenon of bone tunnel enlargement surrounding the graft, which is currently recognized as a potential risk factor for fixation loosening and surgical failure [4]. Furthermore, bone tunnel enlargement complicates revision surgery and increases the risk of associated complications. Therefore, in considering the safety and efficacy of PET artificial ligaments for ACL reconstruction, particularly with respect to long-term outcomes, enhancing the osteointegration of PET artificial ligaments and preventing bone tunnel enlargement remains a critical clinical challenge [5].

Investigations into clinical failure cases have indicated that bone tunnel enlargement primarily results from the formation of disorganized fibrous scar tissue at the graft-bone interface, as observed through various levels of monitoring methods [6,7]. This scar tissue exhibits poor mechanical properties and creates a space-occupying effect, hindering the growth of blood vessels and bone tissue, leading to poor graft integration with the bone [8]. PET, as a non-degradable and inert hydrophobic material, lacks cellular biocompatibility. During the surgical process of pulling the graft into the bone tunnel, continuous friction with the hard bone tissue generates wear debris, which is believed to induce sustained activation of pro-inflammatory M1 macrophages (Mφs), resulting in a chronic inflammatory microenvironment [9,10]. Research has shown that the early phenotypic transition of local Mφs from pro-inflammatory M1 to anti-inflammatory M2 is crucial for tissue repair [11,12]. Therefore, inhibiting scar formation and regulating the local inflammatory microenvironment postoperatively are critical for promoting the integration of PET grafts with bone. The surface properties of biomaterials play a crucial role in shaping their interactions with adjacent cells and tissues, profoundly affecting tissue repair outcomes [13]. Optimizing the surface characteristics of PET to enhance its bioactivity and promote interactions with tissues has been shown to have positive clinical significance. Researchers have proposed various surface modification methods, including β-TCP, silk fibroin [14], graphene [15] and hydroxyapatite [16]. Unfortunately, these strategies focus solely on optimizing osteogenic capacity, neglecting the impact of scar tissue. Clearly, designing interface coatings with immunomodulatory capabilities and low friction coefficients to modify PET grafts, thereby preventing scar formation and inducing vascular and new bone growth to promote graft integration with bone, is a superior strategy for addressing clinical bone tunnel enlargement.

Glycosaminoglycan (GAG) is abundant in mammalian tissues and can influence macrophage activity by binding to specific carbohydrate receptors on their surface, resulting in decreased secretion of pro-inflammatory cytokines [17,18]. GAG also affects the bioactivity of endogenous cytokines by interacting with the positively charged amino acid residues of chemokines via its negatively charged sulfate groups [19]. The heterogeneity in GAGs' chemical structures limits their practical application. In our previous studies, we synthesized sulfated polysaccharide (SCS), a GAG analog, which significantly modulates Mφ polarization towards the M2 phenotype and enhances endogenous VEGF secretion, inducing angiogenesis in a mouse hind limb ischemia model [20]. Therefore, SCS-based engineered materials may regulate chemokine levels, thus potentially reducing inflammation and preventing tissue scar formation. Hydrogel materials, by forming topological micro/nanostructures or covalent bonds on the substrate surface, reduce interfacial friction between tissues and materials and serve as a slow-release platform. As a result, they have garnered significant attention as a promising surface coating [21,22].

In this study, we developed a multi-tiered immune-regulating hydrogel coating to control early inflammation at the graft-bone interface during healing (Scheme 1). This functional coating comprises two key modules: (i) an anti-inflammatory sugar module where SCS modulates macrophage polarization and binds growth factors, thereby reducing the release of inflammatory cytokines and preventing scar tissue formation. (ii) The lubrication module, composed of GelMA hydrogel, mimics the extracellular matrix (ECM) [23] and supports cell biocompatibility [24] while enabling the controlled release of SCS. Additionally, the native ACL surface features a hydrated lubrication layer that mitigates mechanical irritation under typical physiological conditions. Inspired by this, we incorporated 8-arm PEGDA with enhanced cross-linking density and more cross-linking points into GelMA hydrogel's molecular network. Hydrogen bonding between the PEGDA hydrogel and water molecules stabilizes the lubrication area, lowers mechanical friction, and diminishes initial inflammatory irritation. Additionally, it boosts the mechanical strength and stability of the GelMA hydrogel. This anti-inflammatory approach is expected to significantly impact ACL reconstruction and holds potential for broader applications in biomedicine.

Scheme 1.

Schematics of multi-hierarchy immunity regulating hydrogel coating for scarless healing in anterior cruciate ligament reconstruction. Following O₂ plasma treatment, the SCS@PGH hydrogel system adheres strongly to PET film surfaces. The hydrogel system forms a lubricating layer enriched with SCS, modulating macrophage polarization to the M2 phenotype and regulating factor secretion. Consequently, it suppresses scar formation and enhances osteointegration at the graft-bone interface.

2. Materials and methods

2.1. Materials

Eight arm-PEG (molecular weight, M_W = 20000), porcine skin gelatin, and lithium phenyl-2,4,6-Trimethylbenzoyl phosphinate (LAP) photoinitiator were procured from Sigma-Aldrich. Chitosan (Mw: 30 × 10⁴ Da approx, degree of de-acetylation: 92 %) was obtained from Zhongfayuan Biological Technology Co. Ltd. (Shenzhen, China). Polystyrene sulfonate (PSS) (M_W = 1.7 × 10⁴) was purchased from Sigma-Aldrich (St. Louis, MO, USA). Trizol reagent, PrimeScript RT reagent kit, and TB Green Premix Ex Taq™ were acquired from Takara Biotechnology Co., Ltd. (Dalian, China). Gene primers were synthesized by Shanghai Bioengineering Technology Co. Ltd. All cell culture-related reagents were sourced from Gibco (Grand Island, NY, USA). The PET LARS artificial ligament was obtained from Huashan Hospital (Fudan University, Shanghai).

2.2. Fabrication of hydrogels and coating preparation

We first synthesized GelMA to form the primary network, then mixed it with PEGDA to form PGH, and finally added SCS into the PGH solution, resulting in the formation of SCS@PGH hydrogel under the action of the LAP crosslinker. The synthesis of GelMA proceeded as outlined below. Initially, methyl methacrylate (MMA) was gradually added dropwise to a solution of porcine gelatin in PBS at 50 °C, and the resulting mixture was stirred for 2 h. Subsequently, the solution was diluted with preheated PBS (50 °C) to a final volume of 100.0 mL, followed by dialysis at 40 °C for one week to eliminate any residual MMA and anhydride. The mixture was prepared with the following component ratios:10 % (w/v) GelMA, 10 % (v/v) eight-arm PEGDA, 0.5 % (w/v) SCS, and 0.05 % (w/v) LAP in phosphate-buffered saline was prepared to generate PEG-GelMA (PGH) and SCS@PEG-GelMA (SCS@PGH) hydrogels.

The preparation method of SCS was based on a previously reported method in the literature [25]. The sulfating reagent was prepared by slowly adding HClSO₃ to 50 mL of N, N-dimethylformamide (DMF) cooled to 0–4 °C, followed by stirring for 15 min to stabilize the mixture. The stabilized mixture was then transferred to a three-neck flask containing 2.5 g of chitosan, 50 mL of formamide, and 2 mL of formic acid. The reaction was conducted at 45–55 °C for 2 h under mechanical agitation and an argon atmosphere, producing a homogeneous pale yellow solution.

The PET membrane was ultrasonically cleaned in deionized water for 15 min and dried under a nitrogen stream. The cleaned PET was subjected to plasma treatment (Femto A, Diener, Germany) in a pure oxygen atmosphere (100 W, 5 min) to activate and introduce oxygen-containing functional groups. The PET underwent immersion in the precursor polymer solution for the hydrogel treatment. Afterward, the PET was subjected to UV light (365 nm, 30 W) for 15 s to ensure complete curing of the hydrogel layer. For cell experiments, the PET membrane was cut into 1 × 1 cm square pieces. In contrast, the PET was rolled into 0.5 mm-wide threads for animal experiments before coating. Sterilization procedures were carried out before utilization.

2.3. Hydrogel characterization and coating characterization

The microstructure was observed using scanning electron microscopy (SEM; S-3400; Hitachi, Japan). Surface zeta potential changes during hydrogel assembly were measured with a nanoparticle tracking analyzer (Zeta viewer, Particle Metrix, Germany). All rheological experiments were conducted at room temperature using a rheometer (Kinexus, Malvern). Hydrogels with a diameter of 1 cm and a thickness of 5 mm were loaded onto the parallel plate. A strain amplitude of 0.5 % was applied to ensure that the hydrogel networks remained within the linear viscoelastic range during measurements. The swelling behaviour of the hydrogel was assessed by determining its equilibrium swelling ratio. The dry hydrogel was initially weighed (Wd) and then immersed in phosphate-buffered saline (PBS) at 37 °C for 24 h. Afterward, the hydrogel was removed from the solution, excess liquid was absorbed using filter paper, and the resulting weight was recorded as Ws. The swelling ratio (SR) was calculated using the formula SR = (Ws - Wd)/Wd × 100 %. The mechanical properties of the hydrogel were evaluated at room temperature utilizing an Instron 2344 micro-test machine. Initially, the hydrogel were prepared into cylindrical samples (5 mm × 5 mm) were subjected to compression testing. The compression strength was directly determined from the stress-strain curve, while the compression modulus was derived from the slope of the linear region (10%–20 % strain) of the stress-strain curve. Quantitative determination of SCS content in hydrogel coatings using Toluidine Blue (TB) solution. Hydrogels were immersed in 1 mL of PBS solution, to which 1 mL of TB solution was added. The reaction was conducted at 37 °C for 3 h. Following this, an equal volume of n-hexane was added, and the mixture was thoroughly shaken. After allowing for phase separation, the aqueous layer was collected, and the absorbance of the solution at 630 nm was measured using a spectrophotometer. A standard curve was established based on known concentrations of SCS. The release of SCS from the hydrogels was assessed on days 1, 3, 5, 7, 9, 11, and 14. Cumulative release was quantified as a percentage of the total SCS adsorbed onto the samples.

The morphology of the hydrogel coating and PET substrate was observed using scanning electron microscopy. The surface roughness of the wet coating was observed using atomic force microscopy (AFM; MFP-3D; Oxford Instruments, UK). Surface hydrophilicity was determined via a contact angle measurement system using the static drop method with deionized water. The surface elemental composition of the hydrogel was analyzed using energy-dispersive X-ray spectroscopy (EDS; QUANTAX 400-30, Bruker, Germany). The chemical composition of the hydrogel-coated PET membrane was assessed via X-ray photoelectron spectroscopy (ESCALAB 250Xi, Thermo Fisher, USA). The functional groups of freeze-dried hydrogel coatings and PET membranes were examined using infrared spectroscopy (Nicolet 6700, Thermo Fisher, USA). Mechanical property tests were performed using an Instron 5543A automated testing system (UTM5105, China). The tensile strength, and elongation at the break of the membrane material were determined at a loading rate of 10 mm/min. The lubricating properties of the hydrogel coating were characterized using a Universal Micro-Tribotester (UMT2, Bruker, Germany). The prepared PET coating was placed on a sample stage with temperature controlled at 37 °C, a friction frequency of 1 Hz, an amplitude of 4 mm, and a load of 1 N, running for 1500 s, and the experimental data were recorded.

2.4. Cell isolation and culture

Murine bone marrow-derived Mφs (BMDMs) were isolated from male C57BL/6 mice aged eight weeks. Tibias and femurs were dissected to collect bone marrow, which underwent red blood cell lysis. The obtained mMφs were cultured for 5 days in DMEM medium containing 10 % fetal bovine serum (FBS), 1 % penicillin/streptomycin, and 30 ng/mL M-CSF (Novoprotein, China). HUVECs were procured from the American Type Culture Collection (ATCC) and were cultured in DMEM medium containing 10 % FBS and 1 % penicillin/streptomycin. All culturing procedures were conducted in a humidified environment at 37 °C with 5 % CO₂.

2.5. Mφs polarization and CM collection from Mφs

To investigate the impact of the hydrogel coating on the polarization of Mφs, immunofluorescence qualitative analysis and flow cytometry quantitative analysis were carried out. Initially, Mφs induced by M-CSF were seeded on different coated materials at a cell density of 1.0 × 10⁶ and stimulated continuously for 3 days. Following PBS washing, cells underwent fixation with 4 % paraformaldehyde for 15 min, and their membranes were subsequently permeabilized with 0.1 % Triton X-100 for 20 min. A 5 % goat serum was then used for blocking for 1 h, followed by overnight incubation with CD197 and CD206 antibodies at 4 °C. After removing the primary antibodies, corresponding secondary antibodies were applied and incubated for 1 h at room temperature. After PBS washing, cells were stained with DAPI for 10 min. After another round of PBS washing, image acquisition was performed using a confocal microscope. To investigate the impact of Mφs on angiogenesis, Mφs were stimulated for three days following the procedure. Subsequently, they were further cultured for an additional day in fresh complete culture medium, and the supernatant was collected and stored at −80 °C for use as the Mφs CM for subsequent experiments. Mφs CM containing 10 % FBS was utilized for rat aortic ring angiogenesis assay, while Mφs CM with 2 % FBS was employed for cell migration assays.

2.6. Flow cytometric analysis

The extracted bone marrow-derived macrophages were co-cultured with the coating material and PET membrane for three days. After digestion with trypsin and centrifugation, cells were resuspended in staining buffer and stained at 4 °C for 45 min with antibodies. The cell suspension was assessed using a CytoFLEX flow cytometer system (Beckman) and analyzed using FlowJo software (Tree Star). The strategy to analyze inflammatory cells is diagrammed in Fig. S14.

2.7. Scratch wound assay and rat aortic ring angiogenesis assay

The scratch wound healing assay was conducted to examine the influence of Mφs CM on the migratory activity of HUVECs. Initially, HUVECs were seeded in 24-well plates at a density of 5 × 10⁴ cells per well and allowed to reach 90 % confluence. Scratch wounds were then created in each well using sterile 200 μL pipette tips, followed by triple washing with PBS and replacing the medium with Mφs CM. Images of the wells were captured at specified time intervals, and the wound area was quantitatively analyzed using ImageJ software.

A rat aortic ring angiogenesis assay was performed to investigate the influence of Mφs on arterial endothelial cells. Aortas were harvested from 8-week-old Sprague Dawley rats and sectioned into 1–1.5 mm rings in this procedure. These rings were then embedded within neutral rat tail collagen I hydrogel and cultured with Mφs CM every two days. After seven days of incubation, several outgrowths sprouted. Subsequently, the rings were fixed with a 4 % neutralized paraformaldehyde solution and labelled with FITC-phalloidin and DAPI for visualization and subsequent quantification.

2.8. RNA analysis by qRT-PCR

To investigate the influence of the materials on macrophage-mediated gene expression regulation, quantitative real-time polymerase chain reaction (qRT-PCR) assays were performed following the manufacturer's protocols. Total RNA was initially extracted from Trizol reagent, which was subsequently reverse-transcribed into cDNA utilizing the PrimeScript RT kit. qPCR experiments were conducted using Takara's TB Green Premix Ex Taq on the Bio-Rad CFX96 real-time PCR system, involving 44 cycles. Relative gene expression was determined using the 2-ΔΔCt method, and Gapdh was utilized as the reference gene. The primers employed in this study were synthesized by Sangon Biotech (Shanghai), and their sequences have been provided in Supplementary Table S1. To explore the influence of macrophage polarization on gene expression associated with vascular neogenesis, HUVECs were seeded in a 12-well plate and treated with Mφs CM. Following the same procedures outlined previously, mRNA extraction and subsequent qPCR analysis were performed. The sequences have been provided in Supplementary Table S2.

2.9. Mouse ACL reconstruction

All animal experiments adhered to the regulations of the East China University of Science and Technology (Ethics Approval Number: ECUST-2022-053) and followed the National Institute of Health Guide for the Care and Use of Laboratory Animals. The knee ACL was transected, and tunnels were created in the tibia and femur for reconstruction. The knee joint was kept flexed, and a PET artificial ligament (0.5 mm diameter) was threaded through the tunnels, and ensure that both ends of the ligament are securely fixed. The wound was disinfected and sutured under strict aseptic conditions. One side of the tibia from each mouse was used for biomechanical experiments (n = 3), and the other side for CT imaging and histological examination (n = 3). Histological sections were prepared after CT imaging.

2.10. Micro-CT scanning and mechanical performance

At the eight-week post-surgery mark, intact femur-graft-tibia complexes were obtained from euthanized mice. Soft tissues were removed, and the complexes were fixed in 4 % paraformaldehyde for 24 h. Subsequently, the tibiae underwent scanning using a Skyscan 1174 micro-CT system (Bruker, Germany) at a resolution of 16 μm. The axial images acquired were then imported into NRecon and CTvox software for visualization and analysis. The freshly harvested tissues for biomechanical testing were preserved at −80 °C and allowed to thaw at room temperature for 24 h before testing. Tensile testing was conducted using a universal materials testing machine (AGS-X; Shimadzu Corporation, Kyoto, Japan). A pre-load force of 1 N was applied for 5 min, followed by uniform stretching of the ligament at a rate of 2 mm/min until failure or rupture, with the results recorded.

2.11. In vitro and in vivo enzyme-linked immunosorbent assay

The cell culture supernatant used for in vitro ELISA detection was derived from conditioned media of macrophages. Tissue samples were retrieved from the surgical site on days 14 and 28 after ACLR. These samples were homogenized in 1 mL of cold PBS, followed by centrifugation to isolate the supernatant from the implantation site, which was then stored at −80 °C until ELISA analysis. The levels of cytokines in the supernatant from the implantation site were quantified using a series of ELISA kits (Neobioscience), including TNF-α, IL-1β, IL-6, IL-4, IL-10, and TGF-β1. All ELISA assays were performed according to the manufacturer's instructions. Following centrifugation, the harvested tissues were digested, red blood cells were lysed in the cell suspension, and total RNA was extracted using a TRIzol reagent. Subsequently, the RNA was reverse transcribed and subjected to quantitative real-time polymerase chain reaction (qRT-PCR) analysis, following the manufacturer's instructions.

2.12. Mφs depletion

To evaluate the influence of Mφs on tissue repair, intraperitoneal injections of clodronate liposomes (100 μL per mouse) were initiated 3 days before surgery and administered every two days to ensure complete depletion of circulating Mφs. PBS liposomes served as the control. The injections were administered post-surgery, after which the mice were monitored until specific time points when tissue samples were collected to assess the recovery of the injured site. This experimental strategy was devised to examine the role of Mφs in the dynamics of tissue repair.

2.13. Histological staining and immunofluorescent staining

For histological analysis, tissues were fixed in 4 % neutral buffered formalin, decalcified, dehydrated, embedded, and sectioned into 5 μm-thick slices. These sections were subjected to hematoxylin, eosin (H&E), and Masson's trichome staining for microscopic examination of neoangiogenesis and tissue structure. Sirius Red staining was carried out, and polarized light microscopy was used to capture images. Immunohistochemical staining was conducted to observe the expression of TGF-β and pSmad3.

For immunofluorescence staining, fresh implants were extracted from the mice and promptly fixed in 4 % cold paraformaldehyde for 4 h. Subsequently, the implants underwent PBS washing, followed by dehydration through immersion in 20 % sucrose and 2 % polyvinylpyrrolidone for 24 h. They were then embedded and frozen in an OCT compound (Leica). Frozen sections were cut using a Leica CM1950 cryostat. Immunofluorescence staining involved permeabilizing sections in 0.3 % Triton X-100 for 20 min, blocking with 5 % goat serum at room temperature for 1 h, and overnight incubation at 4 °C with primary antibodies. After primary antibody incubation, the sections were washed three times with PBST and then incubated with appropriate secondary antibodies at room temperature for 1 h. Finally, sections were mounted with DAPI (Cell Signaling Technology) for imaging using a Leica confocal microscope. All antibody information is in Table S3.

2.14. Statistical analysis

The results were analyzed using GraphPad PRISM software (version 9.0; GraphPad, La Jolla, CA, USA). All data was presented as mean ± SD. The 'N' values represent the biological replicates of experiments, which were conducted at least three times unless otherwise specified. Statistical analyses involving multiple experimental groups, where appropriate, were conducted using either one-way ANOVA or two-way ANOVA. A significance threshold of P < 0.05 was considered statistically significant, indicated as follows: ∗P < 0.05; ∗∗P < 0.01; ∗∗∗P < 0.005.

3. Results

3.1. Preparation and characterization of the hydrogel

We developed a dual-network hydrogel, specifically a GelMA-PEGDA hydrogel (PGH), designed to serve as a carrier for the controlled release of SCS(SCS@PGH) (Fig. 1A). First, GelMA is prepared to form the primary network. The results of the ¹H NMR spectroscopy show that the appearance of new signals at 5.4 and 5.6 ppm indicates the modification by methacrylic anhydride (MA) (Fig. S1), confirming the successful conjugation of methacrylate vinyl group to gelatin. The degree of methacrylation was calculated based on the reduction of the peak at 2.9 ppm compared to gelatin [26], and it was determined to be approximately 39 %. Then, it is combined with PEGDA to further reinforce the structure of the hydrogel, forming PGH. Finally, SCS is added, and with the addition of the photoinitiator (LAP), the secondary network SCS@PGH is constructed under visible blue light. Scanning electron microscope (SEM) images reveal the porous structures of both PGH and SCS@PGH hydrogels, featuring pore sizes of several hundred micrometers. Adding SCS to the biocomposite hydrogels does not significantly alter the pore size (Fig. 1B). The zeta potential of the original PGH hydrogel, initially at −2 mV. After the introduction of SCS into the hydrogel system, the zeta potential slightly decreases, which is attributed to the negative charge of SCS. This modification enhances the hydrogel's capacity for electrostatic interactions with growth factor-binding polysaccharides (Fig. 1C). Rheological properties of the composite hydrogel scaffolds were assessed using frequency scanning across a broad frequency range (0.1–10 Hz) to characterize their viscoelastic behavior. Both hydrogel groups demonstrated similar rheological behaviors. For both hydrogel groups, the storage modulus (G′) was consistently higher than the loss modulus (G″), reaching up to 25 kPa for the SCS@PGH group across the frequency range. This indicates the stability of the network structure within the gel systems (Fig. 1D). Furthermore, a slight increase in the elastic modulus of SCS@PGH compared to PGH hydrogel has been observed, potentially forming more stable structures through hydrogen bonding and electrostatic interactions between SCS and PGH polymers (Fig. 1E). Swelling of the hydrogel-based scaffolds, another critical property, directly affects nutrient and gas exchange. Adding SCS to the hydrogel matrix causes a slight decrease in swelling ratio, from 150 % in PGH to 120 % (Fig. 1F). Nonetheless, this change does not significantly impact the porosity of the hydrogel construct, ensuring adequate nutrient and gas exchange. Compression tests are performed to assess the mechanical properties of the hydrogel (Fig. 1G). The average compressive modulus ranges from 1.0 to 1.5 kPa, respectively (Fig. 1H). We quantitatively analyzed the release of SCS from the SCS@PGH hydrogel using toluidine blue reagent. The results indicated that SCS exhibited a rapid release trend during the initial phase, followed by a gradual decrease in release rate, eventually reaching a plateau (Fig. 1I).

Fig. 1.

Preparation and characterization of the hydrogels. A) Chemical structures of synthesized hydrogel. B) SEM image of the hydrogels. C) Zeta potentials of PGH and SCS@PGH. D) Storage modulus (G′) and loss modulus (G″) of the hydrogels. E) The elastic modulus of the two types of hydrogels under compression at fully swollen state. F) Swelling ratio of the two types of hydrogels. G) Compression curves and H) Compression modulus of the PGH and SCS@PGH hydrogel. I) The Cumulative release profiles of SCS from SCS@PGH.

3.2. Preparation and characterization of the coating

The hydrogel coating preparation starts with O₂ plasma treatment of the PET film. This treatment creates weak cross-links between the hydrogel and PET, ensuring secure attachment to the substrate. Next, different hydrogel solutions are applied to the PET surface, forming coatings with lubricating and anti-inflammatory properties. These coatings are named PGH/PET and SCS@PGH/PET. The surface morphology of the PGH/PET and SCS@PGH/PET coatings was examined via SEM and atomic force microscopy (AFM). SEM analysis of the PET coating surface showed that the SCS@PGH/PET and PGH/PET hydrogel layers were uniform and smooth compared with the PET (Fig. 2A and Fig. S2). Upon lyophilization, the hydrogel coatings revealed a porous structure. Energy dispersive spectroscopy (EDS) mapping verified the even distribution of elements like C, O, N and S elements to be homogeneously distributed on the surface of SCS@PGH/PET (Fig. 2B and Fig. S3), indicating the successful incorporation of SCS into the hydrogel. Atomic force microscopy (AFM) results showed that the surface roughness (Ra) of PET and SCS@PGH/PET was 105 nm and 1.37 nm, respectively (Fig. 2C), illustrating that the hydrogel coating markedly decreased the surface roughness of PET. The water contact angles (WCAs) for the PET, PGH/PET, and SCS@PGH/PET groups were 127.49°, 78.5°, and 55°, respectively (Fig. 2D). These results indicate that hydrogel-coated groups exhibited lower WCAs than untreated PET. The uniformly dispersed hydrogel coating significantly enhanced the hydrophilicity of PET, thereby facilitating cell adhesion and fluid permeation.

Fig. 2.

Preparation and characterization of the coating. A) SEM micrographs of the coating. B) EDS mapping showing the element distribution of SCS@PGH/PET. C) AFM images of PGH/PET and SCS@PGH/PET. D) Water contact angle of different coatings. E) FTIR spectra. F) XPS spectra. G) tensile strength and H) strain at failure of different coatings (n = 3, ns, not significant). I) Coefficient of friction-time curve. J) Cell viability of HUVECs on the different coating (n = 3, ∗∗∗P < 0.005, two-way ANOVA with Tukey's post hoc test). K) Live/dead cellular staining and confocal fluorescence images of HUVECs.

The FTIR spectra of SCS@PGH/PET displayed a small peak at 1240 cm⁻¹, attributed to the stretching vibrations of O=S=O groups, and a new band around 815 cm⁻¹ from the stretching vibrations of C-O-S groups (Fig. 2E). To further analyze the composition and chemical states of the composite, the hydrogel coating was characterized using X-ray photoelectron spectroscopy (XPS). The S₂P peak was observed in the SCS@PGH/PE (Fig. 2F and Fig. S4). Tensile assessments, such as stress-strain curves and tensile strength analysis, showed enhanced mechanical properties of the lubricating hydrogel coating over pure PET (Fig. 2G and H). The friction coefficient of the hydrogel, measured with a micro-dynamic wear tester, demonstrates its lubricity. In the 1500-s test, the pure PET group exhibited a friction coefficient of 0.6, which was reduced upon adding hydrogels. Specifically, the GelMA/PET and SCS@GelMA/PET groups showed a friction coefficient of approximately 0.4 (Fig. S5). Meanwhile, the PGH/PET and SCS@PGH/PET groups stabilized around 0.2. These results indicate that the addition of PGEDA reduced the friction coefficient by 66.67 % compared to PET, contributing to a more effective lubrication coating (Fig. 2I). Besides optimal physicochemical properties, scaffolds for tissue repair also require excellent biocompatibility. MTT assays (Fig. 2J) demonstrated that hydrogel-coated groups had an accelerated cell proliferation rate compared to bare PET, which increased further after SCS@PGH treatment in HUVECs. Considering the crucial role of rapid cellular localization at the wound site in early tissue repair, further assessment of cell adhesion to the coating was performed. Using live/dead and actin filament staining techniques, HUVEC cells cultured on the hydrogel-coated membrane for one day exhibited an enhanced, elongated morphology (Fig. 2K), affirming the hydrogel coating's superior biocompatibility.

3.3. In vitro immune regulation of the hydrogel coating grafts

Given the frequent association of Mφs with ACL implants, these myeloid cells are considered crucial participants. Additionally, macrophage polarization significantly affects the reparative efficacy of ACL reconstruction [27,28]. To investigate the effect of hydrogel coating on macrophage polarization, we examined the polarization of bone marrow-derived macrophages (BMDMs) stimulated by PGH/PET, SCS@PGH/PET, and PET samples in an in vitro cell culture model (Fig. 3A). Initial flow cytometry of Mφs markers showed that SCS significantly influences M1-to-M2 polarization (Fig. 3B and C). Immunofluorescent staining was used to visualize CD206 and CD197 expressions, markers of M2 and M1 Mφs, respectively (Fig. 3D). Mφs treated with SCG@PGH/PET hydrogel displayed more CD206-positive spots and predominantly elongated shapes. Conversely, PET-treated groups exhibited greater CD197 expression with a more rounded, mononuclear appearance of Mφs. The PGH/PET group showed a significant decrease in CD197-positive cells and a minimal level of CD206-positive expression compared to the PET group. These results were further validated by the expression of M1 and M2-related genes (Fig. 3E). The results indicated that SCS significantly enhanced the expression of anti-inflammatory genes associated with reparative anti-inflammatory genes (CD206, TGF-β, VEGF) and downregulated the expression of specific pro-inflammatory genes (Il-β, Il-6, and TNF-α). Therefore, these findings suggest that SCS promotes the transition from M1 to M2 Mφs polarization. Additionally, we conducted a detailed analysis of the inflammatory responses in Mφs exposed to the hydrogel coatings. Cytokine secretion in cells co-cultured for three days was quantified using ELISA kits (Fig. 3F), and a marked decrease in the secretion of factors associated with M1 Mφs (TNF-α, IL-1β, and IL6) was observed upon the inclusion of SCS, accompanied by a significant increase in the secretion of factors associated with M2 Mφs. Importantly, TGF-β showed the most substantial growth. These outcomes suggest that the lubricating properties of hydrogels can ameliorate the pro-inflammatory state of Mφs. Furthermore, the polarization of pro-inflammatory M1 Mφs towards anti-inflammatory M2 Mφs was significantly facilitated by the addition of SCS. At the same time, the secretion of associated inflammatory factors was modulated.

Fig. 3.

In vitro immune responses of Mφs on various coating. A) Schematic representation of Mφs cocultured on different samples. B) Polarization of Mφs and C) Quantitative analysis was evaluated by the flow cytometry of CD197 (M1) and CD206 (M2) after simulating with samples for 3 days (n = 3). D) Fluorescence microscopy images of Mφs polarization on different samples for 3 days. E) Relative mRNA expression levels of M1 and M2 macrophage–related genes (n = 3). F) ELISA analyses of inflammatory cytokines (n = 4). Data represent mean ± SD. ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.005; ns, not significant (one-way ANOVA with Tukey's post hoc test).

3.4. In vitro influence of coating grafts-treated Mφs CM on angiogenic behavior

Implant-induced inflammatory responses frequently trigger angiogenesis, which is vital for providing oxygen and nutrients necessary for tissue repair [29]. This study aimed to assess the impact of macrophages on the neovascularization behavior of endothelial cells. Therefore, Human umbilical vein endothelial cells (HUVECs) were cultured with conditioned medium (CM) from Mφs for subsequent experiments. Given the critical role of endothelial cell migration in neovascularization, Fig. 4A and B illustrate the migratory response of HUVECs to Mφs CM. The wound healing rate in the PGH/PET group was faster than in the PET group, with cell migration significantly enhanced in the SCS@PGH/PET group. After 48 h, the SCS@PGH/PET group exhibited a wound healing rate of approximately 78 %, about 1.6 times higher than that of the PET group.

Fig. 4.

SCS induces Mφ-mediated arteriogenesis in vitro. A) Cell migration assay: optical images (scale bars: 200 μm) and B) quantification of wound closure assay for 24 and 48 h of HUVECs in Mφs CM after stimulating with coating (n = 3). C) Relative angiogenic-related mRNA expressions using HUVECs in Mφs CM. D) Immunofluorescence staining labelled with VEGFR2 (scale bars: 200 μm). E) Representative rat aortic ring outgrowth images and F) quantification of angiogenic sprouts co-cultured with Mφs for 7 days (n = 3). Scale bar, 200 μm. All data represent mean ± SD. ∗P < 0.05, ∗∗∗P < 0.005 (one-way ANOVA with Tukey's post hoc test).

The angiogenic gene profiling showed a significant increase in the VEGF gene expression of the SCS@PGH/PET group (Fig. 4C). Moreover, the expression levels of angiopoietin-1 (ANG-1), a prominent angiogenic factor, and Esm-1, the gene product associated with tip cell marker and VEGF-regulated gene, were significantly upregulated in the SCS@PGH/PET group. Vascular endothelial growth factor receptor 2 (VEGFR2) is crucial for angiogenesis and endothelial cell function. The impact of Mφs on endothelial VEGF expression was explored using immunofluorescent staining of HUVECs with anti-VEGFR2 antibodies (Fig. 4D). After treatment of endothelial cells with Mφs CM, the PGH/PET group showed positive VEGFR2 staining, with even stronger expression observed in the SCS@PGH/PET group. This indicates that the lubricating hydrogel coating and SCS addition to Mφs CM appear to upregulate or activate the receptor, which could enhance endothelial cell proliferation, migration, and related functions. To further investigate Mφs effects on arterial formation, an ex vivo aortic ring angiogenesis model was used. The SCS@PGH/PET treatment group showed significantly enhanced microvessel formation, characterized by obvious sprouting from the arterial rings and prominent tip cells (Fig. 4E and F). These findings suggest that SCS influences endothelial cell neovascularization via Mφs. Previous studies have shown that macrophage polarization not only influences angiogenesis but may also play a significant role in the differentiation of osteoblasts. Therefore, we further evaluated the impact of macrophages on the osteogenic differentiation of BMSCs (Fig. S6). By assessing the expression of specific markers, we found that alkaline phosphatase (ALP), an early osteogenic marker, exhibited the most prominent staining on day 7 in the SCS@PGH/PET group under the stimulation of Mφs CM. Additionally, to assess extracellular matrix mineralization, which is a critical indicator of late-stage osteogenic efficiency, we conducted Alizarin Red S (ARS) staining. The results showed that the SCS@PGH/PET group had the most significant formation of mineralized nodules. This indicates that SCS, through regulating macrophage polarization, not only promotes angiogenesis but also enhances the osteogenic differentiation capacity of BMSCs.

3.5. Reduction of inflammation and Inhibition of scar formation in vivo

Given the excellent immunomodulatory and pro-neovascularization activity observed in vitro, a mouse ACLR model was created to evaluate the effect of hydrogel coating on the tissue repair function at the tendon-bone interface. Histological examination using hematoxylin and eosin (H&E) staining was conducted to evaluate the repair of the graft-bone interface on post-operative days 14 and 28 (Fig. 5AB). On day 14, a significant presence of inflammatory cells was observed around the PET implants. The PGH/PET group showed fewer inflammatory cells than the PET group. The SCS@PGH/PET group exhibited a small number of microvascular structures. By day 28, the PET group still had many inflammatory cells and increased abnormal vascularization and scar tissue formation. In contrast, the SCS@PGH/PET group demonstrated notable integration of bone tissue at the graft-bone interface.

Fig. 5.

Inflammation was attenuated and scar formation was inhibited in vivo. Hematoxylin and eosin (HE) staining from different groups at A) 14 days and B) 28 days postoperatively (P: PET; S: Scar tissue interface; the black arrow indicates new vessels). C) Immunofluorescence staining of CD197⁺CD206^- Mφs (M1) and CD197⁻CD206⁺ Mφs (M2), D) Col 1, E) Ki67 and OSX of the interface of each group on days 14 and 28 postoperatively (n = 3). F) TNF-α, G) IL-1β, H) IL-6, I) TGF-β, J) IL-10, and K) IL-4 levels on 14 and 28 days after postoperation (n = 4). Data represent mean ± SD. ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.005; ns, not significant (Two-way ANOVA with Tukey's post hoc test).

Given the significant role of macrophages in chronic inflammation, immunofluorescence tissue staining was used to evaluate the effects of different coatings on macrophage polarization (Fig. 5C). The results showed no significant differences in the positive areas of F4/80⁺ Mφs among the groups. However, the PET group exhibited a higher presence of CD197⁺ M1-polarized at 28-day interval, indicating prolonged inflammation within this group. In contrast, both the PGH/PET and SCS@PGH/PET groups showed an inclination towards an increased number of CD206⁺ M2-polarized Mφs, with a significant rise observed in the SCS@PGH/PET group. These findings demonstrate the anti-inflammatory properties of the lubricating hydrogel. Furthermore, the inclusion of SCS proved more effective in promoting Mφ polarization towards the M2 phenotype.

Scar formation and regression are closely associated with the continuous remodeling of the extracellular matrix (ECM) [30]. Immunofluorescence analysis shows varied expression patterns of type I collagen (Col 1) and smooth muscle actin (SMA) across different groups. After 14 days, the PET group showed elevated levels of Col 1 and SMA, in contrast to the marked reductions in these markers observed in the SCS@PGH/PET and PGH/PET groups. By day 28, both the SCS@PGH/PET groups displayed increased anisotropically aligned Col 1 (Fig. 5D) and a reduction in SMA-positive cells (Fig. S7). Additionally, tissue osteogenic and proliferative responses were evaluated. The SCS@PGH/PET group demonstrated notably higher expression of OSX and Ki67 on days 14 and 28 than other groups. Similarly, the PGH/PET group exhibited greater expression than the PET group. These results underscore the efficacy of lubricating hydrogel coatings in promoting bone growth and reducing scar formation, with SCS inclusion hastening the transition to proliferative and osteogenic tissue repair stages (Fig. 5E). This emphasizes the significant role of these hydrogel coatings in improving tissue integration and regeneration.

ELISA was employed to measure inflammatory factors in the surgical region to elucidate the effects of various PET coatings on Mφs polarization post-ACLR. Results showed a significant decrease in M1-related inflammatory factors (IL-1β, TNF-α) in both PGH/PET and SCS@PGH/PET groups, with no significant change in IL-6 levels (Fig. 5 F, G, H). Additionally, there was an increase in M2-related healing factors (IL-4, IL-10, TGF-β) (Fig. 5 I, J, K). Initially, TGF-β levels were lower in the PET group than in others, increasing by week 4. In contrast, the SCS@PGH/PET group showed elevated TGF-β secretion at week 2, which declined by week. TGF-β is a critical regulator in the mouse ACLR model, especially during the transition from inflammation to proliferation. These findings indicate that the SCS@PGH/PET group may enhance healing by modulating inflammatory cells and their associated factors.

Furthermore, Masson staining (Fig. 6A) and polarized light microscopy (Fig. 6B) were employed to verify the formation of collagenous tissue at the interface. Minimal regenerated bone and collagen were observed at the interface region of PET fiber and carbon fiber grafts. In contrast, the SCS@PGH/PET group exhibited the development of a mature collagen arrangement, characterized by collagen bundles gradually forming around and within the coating during the initial stages of osseointegration. An anisotropic collagen layer resembling natural bone tissue was observed at the interface region between the material and the host bone. Given the significant expression of TGF-β following ACLR and its essential role in various cellular processes such as proliferation, differentiation, and tissue regeneration, the overall expression of TGF-β and its downstream effector molecule, pSmad3 was further investigated. The results demonstrated a declining trend in TGF-β expression in the SCS@PGH/PET group at week 4 (Fig. 6C). Additionally, pSmad3 accumulation in the nuclei of cells around PET implants indicated active TGF-β signaling (Fig. 6D). Conversely, pSmad3 was almost absent in the nuclei of fibroblasts in collagen tissue around hydrogel-coated implants, indicating minimal TGF-β activation. The quantitative results are consistent with the aforementioned findings (Fig. 6E, F, G).

Fig. 6.

Macrophage influence on collagen formation and scar formation in vivo. A) Representative Masson's trichrome staining images and B) Polarized light at 14 and 28 days post-operative ACL reconstruction. C) Immunohistochemistry staining for TGF-βand D) pSmad3. Quantitative analysis of E) Collagen, F) TGF-β and G) pSmad3-positive nuclei(n = 3). H) Schematic with Mφs depletion (clodronate liposomes) and treatment strategies. I) Representative fluorescent images of Col 1 following recovery in a Mφs depletion model (n = 3). J) Immunofluorescence staining of F4/80, CD31 and Emcn of the interface of each group at day 14 after Mφs depletion (n = 3). Data represent mean ± SD. ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.005; ns, not significant (Two-way ANOVA with Tukey's post hoc test).

To examine the role of Mφs in scar formation, we depleted Mφs using clodronate liposomes after anterior cruciate ligament reconstruction. Treatment outcomes were evaluated on day 14 following the cessation of injections (Fig. 6H). Furthermore, Col 1 immunostaining and quantitative results showed considerably higher collagen expression in the SCS@PGH/PET group than in other groups (Fig. 6I, Fig. S8A). In contrast, the expression of SMA was the opposite (Fig. S9). Immunofluorescent staining revealed an increase in CD31⁺Emcn⁺ vessels in the SCS@PGH/PET group after slightly recovering F4/80⁺ Mφs (Fig. 6J, Figure S8BC). These findings collectively emphasize the crucial involvement of Mφs in the initial stages of scar resolution. Furthermore, the SCS@PGH/PET group efficiently inhibited Mφs-mediated inflammation and scar tissue at the tendon-bone interface.

3.6. Therapeutic effect of SCS@PGH/PET in the mouse model of ACL reconstruction

To assess the therapeutic effect of SCS@PGH/PET in ACL reconstruction, high-resolution micro-computed tomography (μCT) was utilized to precisely quantify changes in tunnel morphology and to visualize new bone formation. Significant reductions in the dimensions of tibia and femoral tunnels were noted in the SCS@PGH/PET group, along with distinct signals of new bone formation in the tibial tunnels (Fig. 7A and Fig. S10). The 3D μCT reconstructions showed new bone tissue infiltrating the tunnel voids and partially integrating into the PET fiber matrix, with the SCS@PGH/PET group exhibited substantially more new tissue formation than the other groups (Fig. 7B). Quantitative analysis showed that, at 8 weeks, the SCS@PGH/PET group had the smallest tibial bone marrow tunnel diameter, measuring approximately 0.446 ± 0.032 mm (Fig. 7C). Furthermore, this group displayed a bone volume fraction (BV/TV) of 34.0 % ± 1.92 %, roughly 4.5 times higher than the control group's 7.4 % ± 0.71 % (Fig. 7D). These results suggest that SCS@PGH/PET offers superior bone regeneration capabilities, forming significant amounts of mature bone tissue, indicative of its potential clinical efficacy.

Fig. 7.

Therapeutic effect of all groups in mouse ACL reconstruction at 8 weeks. A) 2D micro-CT of the cross sections of tibial tunnels at the intermediate and tibial tunnels images. B) Representative 3D reconstruction micro-CT images showing regenerated bone around defects. Quantitative analysis of C) the tunnel cross-sectional diameter at the middle entrances of tibial tunnels in each group and D) BV/TV. E) HE and Masson staining, Scale bars, 200 μm. F) Digital camera image of the biomechanical test for different groups at 8 weeks after surgery and the ultimate failure load. G) Representative fluorescent images stained with OPN, Ki67, DAPI, Scale bars, 200 μm. H) Representative immunostainings of CD31^hiEmcn^hi (Type H) endothelium cells at 8 weeks after surgery. Data represent mean ± SD. ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.005; ns, not significant (Two-way ANOVA with Tukey's post hoc test).

Histological assessments were performed to evaluate graft integration with bone, a key determinant of biomechanical stability at the graft-bone interface. In the PET group, substantial scar tissue and minimal new bone formation were observed at the implant-tissue interface, whereas the SCS@PGH/PET group demonstrated distinct bone tunnels around the implants (Fig. 7E). Sirius red and polarized light evaluations of collagen formation showed neatly aligned collagen structures around the bone tunnels in the SCS@PGH/PET group (Fig. S11), indicative of effective osteointegration. This enhanced integration may improve the biomechanical properties crucial for successful implantation. To quantify the interfacial strength between various hydrogel coatings groups and adjacent natural bone, we measured the pull-out force required to extract implants from bone tunnels, which indicates osseointegration levels (Fig. 7F) [31]. At 8-week after ACL reconstruction, the SCS@PGH/PET group showed the highest pull-out strength at approximately 12N, 3 times that of the control group. Meanwhile, the pull-out strength of the PGH/PET group was slightly higher than that of the pure PET group. Further exploration into the mechanisms of bone formation was conducted using immunofluorescent staining. After eight weeks, a significant increase in OPN-positive osteoblasts and Ki67 expression in the SCS@PGH/PET group (Fig. 7G, Fig. S12AB) indicated notable osteoblast proliferation during bone formation. Additionally, CD31^hi and Emcn^hi vascular structures in the SCS@PGH/PET scaffolds (Fig. 7H, Figure S12C and Figure S13) demonstrated their ability to support a coherent vascular network, further validating the osteoinductive potential of the samples. These findings indicate that adding lubricating hydrogels and SCS enhances the regenerative microenvironment at the implant-bone interface, thus accelerating healing.

4. Discussion

In this investigation, the SCS@PGH/PET hydrogel was successfully engineered, and its viability for anterior cruciate ligament (ACL) reconstruction applications was assessed. The hydrogel's refined physicochemical attributes, such as its optimal porous architecture, augmented lubricative properties, and proficiency in promoting cellular adhesion and proliferation, establish a solid foundation for its potential clinical efficacy. Importantly, the hydrogel's immunomodulatory capacity, particularly in facilitating the phenotypic transition of Mφs from a pro-inflammatory M1 state to an anti-inflammatory M2 state [32], proposes a novel paradigm for mitigating scar formation, attenuating inflammatory responses, and accelerating tissue regeneration. Further, angiogenesis stimulation and substantial osteointegration tend to be induced by this hydrogel, thereby bolstering its applicability in ACL reconstruction.

Research has established that tissue exposure to biomaterials triggers a foreign body response (FBR) [33,34]. This process encompasses inflammation, wound healing, and, if unresolved, tissue scarring and in severe instances, the formation of fibrosis. Effective tissue regeneration relies not only on the extent of regenerative capacity at the injury site but also on the capability to transition immediate fibrosis-based repair into a transient response. Scar tissue primarily arises from excessive extracellular matrix (ECM) collagen deposition [35]. Studies have shown that modulating the extracellular matrix can effectively reduce fibrosis, thereby promoting wound healing and tissue regeneration. The design of the material not only improves the stability of the graft physically but may also optimize the healing process by regulating local cellular activity.Striking a balance between scar formation and maintaining a pro-regenerative environment is imperative for achieving successful regenerative outcomes [36]. Macrophages play a pivotal role as the primary inflammatory cells at the interface between biomaterials and tissues [37,38]. They show either pro-inflammatory or anti-inflammatory behavior in response to inflammation and scar tissue formation triggered by implanted materials. Modifying the structure or surface activity of the biomaterial can impact the equilibrium between pro-inflammatory and anti-inflammatory as well as regulatory macrophages at the graft-bone interface, potentially facilitating functional tissue remodeling [39]. In this study, macrophages were depleted by injecting mice with clodronate after ACL surgery. The findings of this work align with existing literature indicating that clodronate treatment inhibits the recruitment and maturation of Mφs. Current research indicates that excessive deposition of extracellular matrix (ECM) is typically characterized by increased expression of α-SMA and Col 1 [40]. In the normal experimental group without macrophage depletion, the PET group exhibited elevated levels of Col 1 and SMA after 14 days, indicating a close association between scar formation and macrophage activity. In contrast, in the macrophage depletion group, the expression of these markers was significantly reduced in both the SCS@PGH/PET and PGH/PET groups, further demonstrating that macrophage depletion effectively reduced scar formation. These findings collectively highlight the critical role of macrophages in scar formation and tissue repair, suggesting that early inflammatory regulation may be a key pathway to reducing scarring and promoting tissue regeneration. However, the specific mechanisms by which SCS influences scar formation remain to be fully elucidated.

Various microenvironmental features, such as the shape and geometric properties of biomaterials, biochemical surface or composition [41], mechanical stiffness, topology, porosity, and protein or drug release can influence the behavior of Mφs. Hence, lubricating PGH hydrogels hold promise in regulating Mφs reactions and facilitating post-operative recovery [42,43]. Nonetheless, the research outcomes suggest that relying solely on lubricating hydrogels to enhance interfacial biomechanical properties falls short. This observation highlights the insufficiency of improving physical lubrication conditions to modulate the inflammatory response in chronic wounds effectively. Consequently, more targeted strategies are imperative to regulate the inflammatory microenvironment adequately. Another limitation of the current hydrogel coating material is its lack of self-healing capability. Self-healing ability is particularly crucial when dealing with complex bone tunnel environments and revision surgeries, especially in cases of bone tunnel scar proliferation or the presence of internal fixation devices after the initial surgery. Self-healing capability will undoubtedly be a key feature in the development of future artificial ligament coatings and may become an important direction for future research. Flow cytometry and immunofluorescence data were employed in this investigation to illustrate how the SCS@PGH/PET hydrogel fosters Mφs conversion to the pro-healing CD206⁺ M2 phenotype.

Simultaneously, inflammatory chemokines promote the recruitment and activation of immune cells, creating a pro-inflammatory cycle that leads to persistent inflammation and exacerbates scar formation [44]. Hence, targeting Mφs and regulating inflammatory factors might offer a more practical approach to managing chronic inflammation and paving the way for minimizing undesirable scar formation. Prior research has indicated that extracellular matrix glycosaminoglycans (GAGs) [45], like heparin sulfate or heparin, facilitate the binding of growth factors and chemokines via electrostatic interactions [46], heparin-binding domains [47], and spatial conformation. This mechanism involves electrostatic interactions between positively charged amino acid residues of chemokines and negatively charged sulfate groups of GAGs. Our findings confirm that the SCS@PGH hydrogel coating possesses a negative charge. ELISA assessments at days 14 and 28 post-application show that the SCS@PGH/PET hydrogel markedly reduces pro-inflammatory factor secretion and enhances pro-healing factor secretion. Importantly, it regulates the TGF-β signaling pathway. In the SCS@PGH/PET group implants' collagenous tissue, nuclear pSmad3 is sparse, suggesting reduced TGF-β activation [48]. TGF-β is crucial for activating and proliferating myofibroblasts [49], inhibiting collagen degradation, and enhancing extracellular matrix (ECM) synthesis [50]. In summary, our study demonstrates that SCS@PGH/PET promotes M2 Mφs polarization in the early stages of post-surgical inflammation, effectively isolating inflammatory chemokines. This impedes further M1 Mφs polarization, improves the post-operative inflammatory microenvironment, and reduces scarring displacement. Additionally, it enhances osteointegration at the graft-bone interface. According to recent studies, novel biomaterials like SCS-based materials have been shown to promote better bone integration by facilitating the recruitment and polarization of M2 macrophages, which are crucial for tissue repair and remodeling. Moreover, such materials contribute to the formation of a more favorable inflammatory environment, thereby reducing the risk of chronic inflammation and enhancing the long-term stability of the graft [51]. This comprehensive response demonstrates the potential of SCS@PGH/PET in enhancing tissue repair and integration following surgical interventions.

Within bone repair, neovascularization diminishes the development of disorganized connective tissue [52,53]. It achieves this by maintaining efficient cellular communication, influencing the inflammatory response, and regulating cytokine release, all of which contribute to establishing a more structured cellular layout and matrix organization [54]. This orchestrated process deters scarring and fosters optimal healing. Without sufficient angiogenesis, the cartilage template fails to effectively transform into mature bone tissue, potentially leading to scar tissue formation. Our study demonstrated that SCS@PGH/PET promotes the formation of CD31⁺Emcn⁺ vessels and arterioles at the junction between the graft and bone by triggering an immune response that boosts macrophages' secretion of VEGF in the early stages post-ACLR. To summarize, Mφs shape scar tissue formation and maturation through their phenotypic transitions and control of cytokine secretion. By comprehending and guiding these pathways, tissue engineering and regenerative medicine approaches might be improved to reduce undesired scarring and fibrosis while facilitating effective tissue repair [55]. While this study offers valuable insights, it's important to acknowledge certain limitations. For instance, the chosen experimental model and sample size may limit the broader applicability of our findings. Future investigations should aim to validate the results under more diverse conditions and explore additional factors that may impact the performance of hydrogels. Nonetheless, the current study highlights the synergistic relationship between hydrogel materials science and immunoregulatory strategies within intricate biological systems, presenting a novel approach for ACL reconstruction. These findings offer fresh perspectives for future ligament repair procedures and pave the way for further exploration in bone and soft tissue engineering research.

5. Conclusion

In this study, SCS@PGH/PET hydrogel coating was developed to modulate the immune microenvironment after ACLR. This hydrogel displays exceptional features, including an optimized porous structure for cellular infiltration, and increased lubricity crucial for cell adhesion and proliferation. Incorporating sulfated polysaccharide and bioactive molecules has proven vital in modulating Mφs polarization from a pro-inflammatory M1 to a regenerative M2 state, with cytokines like TGF-β being regulated. This transition is essential for reducing scar formation, promoting a healing environment, and accelerating angiogenesis. The significant impact of the hydrogel on tissue repair and regeneration was confirmed through both in vitro and in vivo studies. Overall, the SCS@PGH/PET hydrogel coating findings provide a solid scientific foundation for the design and improvement of synthetic ligaments in regenerative medicine.

CRediT authorship contribution statement

Shuang Wang: Writing – original draft, Visualization, Formal analysis, Data curation, Conceptualization. Chao Xu: Writing – review & editing, Visualization, Methodology. Yuanman Yu: Writing – original draft, Methodology. Jie Li: Software, Methodology. Tianwu Chen: Writing – review & editing, Software, Resources, Methodology, Conceptualization. Jing Wang: Writing – review & editing, Supervision, Project administration. Changsheng Liu: Project administration, Funding acquisition.

Ethics approval and consent to participate

All procedures were approved by the Animal Research Committee of East China University of Science and Technology. The ethical approval number was ECUST-2022-053.

Declaration of competing interest

The authors declare no competing interests.

Appendix A. Supplementary data

The following is the Supplementary data to this article.