Peptide-based Self-adaptive Implant Coating Sequentially Regulates Bone Regeneration to Enhance Interfacial Osseointegration

doi:10.21203/rs.3.rs-5132912/v1

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Peptide-based Self-adaptive Implant Coating Sequentially Regulates Bone Regeneration to Enhance Interfacial Osseointegration

https://doi.org/10.21203/rs.3.rs-5132912/v1

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Aseptic loosening is the primary cause of orthopaedic implant failure. Previous designs for implant coatings failed to follow the physiological process of bone regeneration, leading to inadequate osseointegration at the bone–implant interface. We engineered a novel self-adaptive titanium-based coating (DOPA-P1@P2) by combining a mussel-inspired biomimetic strategy with bioorthogonal click chemistry techniques. Both in vitro and in vivo results confirmed that DOPA-P1@P2 exhibited excellent biocompatibility and effectively enhanced osseointegration. Specifically, the outermost K23 layer of DOPA-P1@P2 promotes M2 macrophage polarization in the first stage of bone regeneration, creating a favourable immune microenvironment. Following the release of K23, the sequential exposure of the K15 and Y5 layers promoted angiogenesis and osteogenesis during the second stage of bone regeneration. In the third stage of bone regeneration, the DOPA-P1@P2 group exhibited a 161% increase in maximal push-out force, a 207% increase in BV/TV, and a remarkable 1409% increase in BIC, compared to the TiO₂ control group. In summary, DOPA-P1@P2 effectively promoted bone regeneration through sequential regulation, thus enhancing osseointegration at the bone–implant interface.

Health sciences/Medical research/Translational research

Biological sciences/Biochemistry/Peptides

Each year, over one million joint arthroplasty procedures are performed in orthopaedic practice worldwide with the ultimate goals to alleviate pain and improve functions¹. Despite the great clinical success of employing bone prostheses, complications remain an issue and account for implant failures and revision surgeries. Aseptic loosening is a major complication that causes over half of these failures due to inadequate osseointegration at the bone–implant interface². General descriptions of osseointegration include direct deposition of regenerated bone on the implant surface and its consequent formation of a solid bone-to-implant anchorage without fibrous interference, highlighting that satisfactory peri-implant bone regeneration is crucial for osseointegration³. In recent decades, titanium and its alloys have become the leading materials for orthopedic implants due to their excellent biocompatibility, mechanical properties, and corrosion resistance^{4, 5}. However, titanium-based implants are considered to be biologically inert that requires surface modifications to provide sufficient bioactivity for osseointegration^{6, 7}. Traditional modifications often involve tuning hierarchical structures and ECM functions towards osteogenesis and bone remodelling via physiochemical and/or biomimetic approaches. However, clinical efficacies of those implants have been impeded due to the discrepancies between in vitro and in vivo observations⁸. Recently, extensive research on osteoimmunology has demonstrated that titanium-based implant surfaces embellished with nanotopography, hydrophilicity or bioactive peptides could facilitate M2 macrophage polarization and promoted peri-implant bone regeneration in vivo^9–12. Although current results suggest a clinical potential, the interfacial osseointegration remains to be accomplished through an enhanced approach¹³. A comprehensive understanding of the mechanisms underlying bone regeneration will offer viable insights for the development of innovative bioactive surfaces, thereby ensuring improved implant osseointegration.

Implant-mediated bone regeneration is a multifaceted physiological process that involves various cells and factors and comprises three stages, namely inflammation, proliferation and remodelling, over temporally consecutive frames^{14, 15}. At the inflammation stage, a haematoma rapidly forms surrounding the implant surface and triggers the release of inflammatory mediators. At the implant site, recruited macrophages produce matrix metalloproteinases (MMPs) and degrade the extracellular matrix (ECM) to facilitate bone reconstruction¹⁶. In response to microenvironmental signals, they undergo distinct M1–M2 polarization^{17, 18}. M1 macrophages secrete high levels of inducible nitric oxide synthase (iNOS) and pro-inflammatory cytokines including TNF-α and IL-1β¹⁹. Prolonged of M1 polarization induces transformation of acute inflammation into a chronic state with the subsequent formation of peri-implant fibrous capsule leading to implant failures²⁰. Conversely, M2 macrophages express high level of anti-inflammatory markers such as CD206, Arg-1 and IL-10²¹. Increase of M2 polarization promotes the inflammation stage to the proliferation stage²². During the proliferation stage, neovascularization serves as a crucial conduit between the bone and surrounding tissues to facilitate the transport of oxygen, nutrients, and bone marrow stromal cells (BMSCs) to the implant site²³. Meanwhile, local release of vascular endothelial growth factor (VEGF) and osteogenic growth peptide (OGP) promotes angiogenesis and BMSCs osteogenic differentiation, respectively^{24, 25}. Once the inflammation and proliferation stages are properly progressed, the remodelling stage will ensue smoothly to complete bone regeneration. The dissection of the physiological process of bone regeneration provides new insights that, in order to achieve ameliorative osseointegration, the implant surface should be functionally designed to align with the temporal sequence of the stages.

Building upon this concept, we develop a self-adaptive multifunctional coating by using four peptides that precisely correspond to the characteristic sequential targets within the aforementioned stages: (1) K23 (KAFAKLAARLYRKALARQLGVAA), an anti-inflammatory peptide for immune regulation during the inflammation stage²⁶; (2) PVGLIG, an MMP-2/9-cleavable peptide for the inflammation stage²⁷; (3) K15 (KLTWQELYQLKYKGI), a VEGF mimetic peptide for promoting angiogenesis at the proliferation stage²⁸; and (4) Y5 (YGFGG), an OGP-derived peptide for promoting osteogenesis at the proliferation stage²⁹. We then assemble them and construct a novel biomimetic peptide-coated titanium-based coating (DOPA-P1@P2) by combining a mussel-inspired biomimetic strategy with bioorthogonal click chemistry techniques. Upon implantation, the outermost K23 layer modulates acute inflammation at the first stage of bone regeneration, facilitating M1 to M2 polarization of macrophages. Meanwhile, MMP-2/9 released by activated macrophages cleaves the PVGLIG sequence, resulting in detachment of K23 layer and exposure of the underlying K15 and Y5 layers to promote angiogenesis and osteogenesis at the second stage followed by bone remodelling. In summary, DOPA-P1@P2 can effectively promote bone regeneration by closely aligning with the physiological process, thereby enhancing osseointegration at the bone–implant interface through sequential regulation.

2.1 Titanium Surface Modification and Material Characterization

To engineer a novel self-adaptive biomimetic titanium surface that can orchestrate bone regeneration, we initially synthesized a mussel-inspired peptide bearing a clickable DBCO group ((DOPA)₄-PEG₅-DBCO) via solid-phase peptide synthesis (Figs. 1A and S1D). Building on previous studies^{30, 31}, we optimized the positioning of catechol to enhance the bonding strength of (DOPA)₄-PEG₅-DBCO on titanium substrates. In this work, we employed readily available Fmoc-DOPA (acetone)-OH to incorporate DOPA into the peptide chain. To ensure that the mussel-inspired peptide adheres to titanium surfaces and allows subsequent bioorthogonal click reactions, we separated the tetrameric DOPA structures with glycine (Gly), lysine (Lys), and PEG-linked DBCO, resulting in the clickable biomimetic peptide Ac-(DOPA)-Gly-(DOPA)-Lys[(PEG₅)-(DBCO-COOH)]-(DOPA)-Gly-(DOPA) (Figure S1A). The chemical structure of (DOPA)₄-PEG₅-DBCO was determined via ¹H NMR spectroscopy, which confirmed the inclusion of a DOPA unit by identifying a distinctive peak at 8.64 ppm associated with catecholic hydrogens (Fig. 1J). In previous studies, surface modifications of bone implants have been achieved predominantly by incorporating biomacromolecules (such as proteins and polysaccharides) and active metal ions^{32, 33}. However, biomacromolecules are challenging to chemically modify and are prone to degradation and deactivation, whereas metal ions are limited by toxic reactions and difficulties in controlling ion release^{34, 35}. In recent years, rapid progress in peptide synthesis technology has increased the use of peptides as preferred candidates for surface modification of implants because of their significant biological activity, increased stability, and simplified chemical modification process³⁶. To meet the functional demands of various stages of bone regeneration, four types of peptides are introduced for surface modification: (1) the anti-inflammatory sequence K23, KAFAKLAARLYRKALARQLGVAA; (2) the angiogenic sequence K15, KLTWQELYQLKYKGI; (3) the osteogenic sequence Y5, YGFGG; and (4) the inflammatory response sequence, PVGLIG. The inflammatory response sequence PVGLIG was placed between K23 and K15, as well as between K23 and Y5, to generate two composite peptide sequences that were subsequently modified with two-azidoacetic acid (-N3). This process yielded two azide-modified composite functional peptides capable of inducing an inflammatory response: P1 (N₃-K15-PVGLIG-K23) and P2 (N₃-Y5-PVGLIG-K23) (Figs. 1B–C, S1B–C and S2A–B).

The synthetic mussel-inspired peptides, P1 and P2 were analysed via high-performance liquid chromatography (HPLC), confirming that both peptides exhibited high purities exceeding 96.6% (Fig. 1D–F). Electrospray ionization mass spectrometry (ESI–MS) evaluated the molecular masses of these peptides. The findings suggested that the monoisotopic masses [M + 2H]²⁺ of (DOPA)₄-PEG₅-DBCO, [M + 6H]⁶⁺ of P1, and [M + 5H]⁵⁺ of P2 were 799.82, 834.32, and 718.65 Da, respectively; these findings are in alignment with the theoretical molecular masses of the peptides (1597.67, 5000.03, and 3588.28 Da, respectively) (Fig. 1G–I). These findings verified the successful production of mussel-inspired peptides and two composite functional peptides. A novel self-adaptive biomimetic titanium surface was then developed in two key steps (Scheme 1A-E). Initially, titanium plates or rods were submerged in a 0.01 mg/ml (DOPA)₄-PEG₅-DBCO solution, enabling coating with mussel-inspired peptides. In the second step, these peptide-coated plates or rods were separately immersed in 0.1 mg/ml solutions of P1, P2, or a 1:1 mixture of both, enabling the creation of a variety of modified surfaces through bioorthogonal click chemistry reactions between -DBCO and -N₃. A 1:1 ratio of P1 to P2 was chosen on the basis of preliminary experimental results (Figure S3A–G), as this ratio enabled the modified surface to exhibit both excellent angiogenic and osteogenic capabilities. Plates or rods immersed in PBS were designated the TiO₂ group and served as the control. Plates or rods immersed only in the mussel-inspired peptide solution were classified as the DOPA group, whereas those further grafted with P1, P2, or their mixture were classified as the DOPA-P1, DOPA-P2, and DOPA-P1@P2 groups, respectively. Up to now, numerous physical and chemical techniques for material modification, such as layer-by-layer assembly, acid etching, silanization, and anodic oxidation, are employed to apply various bioactive substances onto implant surfaces³⁷. Nevertheless, the physical and chemical modification techniques currently in use continue to encounter several challenges, including molecular leakage, fast release, complicated processes, and insufficient long-term activity^{38, 39}. Since the first introduction of a novel surface modification method through the polymerization of dopamine in 2007, biomimetic strategies inspired by the molecular adhesion mechanism of marine mussel foot proteins (Mfps) have gained significant attention as promising methods for surface modification⁴⁰. In these methods, the repeated 3,4-dihydroxy-L-phenylalanine (DOPA) is capable of mediating robust adhesion on virtually all surfaces (which relies on catechol groups to conjugate with biomacromolecules or spontaneously coordinate with metal ions) while also endowing implant surfaces with excellent biocompatibility⁴¹. Nevertheless, DOPA-based biomolecular conjugation primarily relies on covalent interactions that could potentially compromise the bioactivity of the involved biomolecules⁴². Consequently, there is an urgent need to optimize the existing mussel-inspired surface modification strategies. Fortunately, the rise of bioorthogonal click chemistry has opened up a new avenue, as its mild and direct reaction conditions, rapid reaction rates, and excellent specificity make this technique a current research hotspot in the field of implant surface modification^{43, 44}. The surface modification approach employed in this work combines mussel-inspired biomimetic strategies with bioorthogonal click chemistry, overcoming numerous drawbacks of traditional surface modification techniques. This method is straightforward, practical, and economical and holds promise for widespread application in settings requiring multifunctional surface modifications.

Atomic force microscopy (AFM) was employed to assess the topography across various surfaces, revealing a significant increase in the roughness of surfaces modified with peptides compared with that of the TiO₂ surfaces (Fig. 1K–L). Interestingly, additional grafting of P1 or P2 onto DOPA-modified surfaces further enhances their roughness, likely due to variations in the peptide chain length or conformation. Subsequent analysis using static water contact angle measurements to evaluate the hydrophilicity across various surfaces, showed that the peptide-modified surfaces exhibited significantly lower water contact angles compared to the TiO₂ group (Fig. 1M–N). The DOPA-P1@P2 group presented the smallest water contact angle (55.6°), indicating superior surface hydrophilicity. Previous research has demonstrated that increased surface roughness enhances osteoblast adhesion and spreading, which facilitates more robust osseointegration than observed with smooth surfaces⁴⁵. Furthermore, enhancing the hydrophilicity of material surfaces promotes the maintenance of protein structure and function, enhances cellular adhesion, and improves biocompatibility⁴⁶. Hence, the biomimetic surfaces (DOPA-P1@P2) developed in this study, which are characterized by increased roughness and hydrophilicity, are well suited to enhance bone regeneration. The compositions of various modified surface elements were characterized via X-ray photoelectron spectroscopy (XPS). As illustrated in Fig. 1O–P, surfaces modified with mussel-inspired peptides presented a notable increase in the N 1s signal relative to that of the TiO₂ group, with additional grafting of P1, P2, or their mixture further amplifying this signal, consistent with the amino content on these surfaces. To confirm that the peptides grafted onto the titanium surfaces remained stable and retained long-term activity, the DOPA-P1@P2 surfaces were incubated in Dulbecco's modified Eagle’s medium (DMEM) at 37°C for three weeks prior to XPS analysis. The results revealed that the N 1s signal decreased by less than 15% after three weeks (Fig. 1Q), indicating that the modified surface retained long-term biological activity. In the initial stage of bone regeneration, acute inflammation peaks within 24 hours and persists for approximately 1 week, followed by a stage characterized by pronounced neovascularization and new bone formation, lasting 3 to 4 weeks^{15, 47}. Consequently, the sequential regulation of inflammation, angiogenesis, and osteogenesis during specific time windows is key to adapting modified surfaces to the natural bone regeneration process. In this work, we utilized MMP-2 to simulate the local microenvironment during the inflammatory phase of bone regeneration and investigated the responsive release trends of an anti-inflammatory peptide (K23) from surfaces P1 and P2. The findings indicated that treatment with MMP-2 markedly enhances the release of K23 due to the specific enzymatic cleavage of the PVGLIG peptide sequence. Specifically, within the initial 24 h after MMP-2 treatment, the release rate of K23 exceeded 60%; the release rate then significantly slowed after day 3, and by the end of week 1, the release rate was approximately 85% (Fig. 1R).

These findings demonstrate that modified titanium surfaces release substantial amounts of anti-inflammatory peptides early in inflammation to modulate immune responses, with K23 nearly depleted within 1 week. During the process of K23 release, its underlying angiogenic peptide sequence (K15) and osteogenic peptide sequence (Y5) become progressively exposed, thereby contributing to the second stage of bone regeneration. In conclusion, we effectively developed a novel self-adaptive biomimetic titanium surface (DOPA-P1@P2), which holds excellent potential for the sequential regulation of bone regeneration.

2.2 Biocompatibility Evaluation of Modified Titanium Surfaces

The primary prerequisite for the clinical application of implanted materials is non-toxicity to body⁴⁸. In this study, R264.7 cells, human umbilical vein endothelial cells (HUVECs), and bone marrow-derived mesenchymal stem cells (BMSCs) were utilized as research objects to assess the biocompatibility across various groups. Initially, live/dead staining experiments were performed, revealing that all three cell types exhibited robust survival on the various modified surfaces. Furthermore, there were no notable differences in the ratios of live to dead cells across the five groups: TiO₂, DOPA, DOPA-P1, DOPA-P2, and DOPA-P1@P2 (Fig. 2A–C). We subsequently quantified the secretion of lactic dehydrogenase (LDH) from the aforementioned three cell types seeded on various modified surfaces to assess their cytotoxicity. Following a 24-hour incubation period, the detected levels of LDH did not significantly differ among the groups, indicating that the peptide-modified surfaces exhibited no toxicity towards the cells (Fig. 2E–G). Additionally, the viability of three different cell types on various modified surfaces was assessed at distinct time intervals (24 and 72 hours) via CCK-8 assays (Fig. 2H–M). The findings indicated that, by day 3, cell viability on the four modified surfaces (DOPA, DOPA-P1, DOPA-P2, and DOPA-P1@P2) was considerably higher compared to the TiO₂ control surface, suggesting that peptide modifications of titanium surfaces significantly promote cell proliferation. Finally, after a 12-hour cultivation period, the morphological features of BMSCs on various substrates were analyzed via cytoskeletal staining, employing fluorescein isothiocyanate (FITC)-phalloidin and 4',6-diamidino-2-phenylindole (DAPI). The findings demonstrated that on TiO₂ surfaces, BMSCs mostly displayed a spherical shape with scant filopodia, while on peptide-modified surfaces, the cells adopted polygonal forms and showed increased expression of filamentous actin (F-actin), particularly prominent in the DOPA-P1@P2 group (Fig. 2D). The enhanced adhesion and spreading of BMSCs on the DOPA-P1@P2 surfaces are significantly correlated with increased surface roughness and hydrophilicity, which aligns with the results presented in the materials characterization section. Besides in vitro assays, we performed in vivo validation experiments in Sprague–Dawley (SD) rats. We implanted modified titanium rods into rat femurs, and after two months, organs were analysed via haematoxylin and eosin (H&E) staining. The results confirmed that all the modified surfaces were free from visceral toxicity (Figure S4). In brief, the findings derived from in vitro and in vivo studies suggest that the developed peptide-coated surfaces enhance cellular adhesion and proliferation and are nontoxic to organisms, establishing the necessary foundation for regulating bone regeneration.

2.3 Immunomodulatory Effects of Modified Titanium Surfaces

The initial stage of bone regeneration mediated by implant materials in vivo is characterized predominantly by an inflammatory response⁴⁹. During this phase, a substantial haematoma forms on the implant surface, dominated by M1 macrophages, which release a plethora of inflammatory mediators, serving as a protective response against injury^{50, 51}. However, the resolution of acute inflammation during the early postimplantation period, especially the timely transition of proinflammatory M1 macrophages to the anti-inflammatory M2 phenotype, is a critical determinant of the fate of implant materials in the body⁵². The prolonged dominance of M1 macrophages at the implant site leads to the sustained release of inflammatory mediators, which results in a shift from acute inflammation post-implantation to chronic inflammation. This triggers the foreign body response, with the recruitment of fibroblasts to encapsulate the implant material with a fibrous capsule, which impairs osseointegration and potentially leads to implant failure⁵³. Conversely, the timely transition from the M1 to M2 macrophage phenotype can steer bone regeneration in the right direction. By secreting essential chemokines and growth factors, M2 macrophages enhance angiogenesis and the migration, homing, and osteogenic differentiation of BMSCs, thus optimizing conditions for subsequent stages of bone regeneration⁵⁴. To determine whether the novel self-adaptive biomimetic modified surfaces can effectively modulate immune responses, we cultured RAW264.7 cells on various groups of titanium surfaces, then conducted immunofluorescence staining. The results revealed that surfaces devoid of anti-inflammatory peptide modifications (TiO₂ and DOPA) presented a marked increase in the expression of M1 macrophage markers (iNOS⁺, depicted in red), whereas surfaces modified with anti-inflammatory peptides (DOPA-P1, DOPA-P2, and DOPA-P1@P2) showed a marked upregulation of M2 macrophage markers (Arg-1⁺, depicted in red) (Fig. 3A–D). These findings suggest that the DOPA-P1, DOPA-P2, and DOPA-P1@P2 surfaces are capable of effectively regulating immune responses and facilitating macrophage polarization to the M2 phenotype. The proportion of F4/80⁺CD86⁺ cells (M1) was notably higher in the TiO₂ and DOPA groups compared to the other groups, as revealed by flow cytometry analysis (Fig. 3E–F). Conversely, the proportion of F4/80⁺CD206⁺ cells (M2) was significantly greater in the DOPA-P1, DOPA-P2, and DOPA-P1@P2 groups than in the TiO₂ and DOPA groups (Fig. 3H–I). The western blot findings were consistent with the aforementioned findings, revealing high expression of an M1-associated protein (iNOS) in the TiO₂ and DOPA groups, whereas an M2-associated protein (Arg-1) was highly expressed in the DOPA-P1, DOPA-P2, and DOPA-P1@P2 groups (Fig. 3K–M). Moreover, the enzyme-linked immunosorbent assay (ELISA) findings demonstrated a substantial elevation in the concentrations of TNF-α within the TiO₂ and DOPA groups compared with the DOPA-P1, DOPA-P2, and DOPA-P1@P2 groups (Fig. 3G). Conversely, the concentrations of IL-10 were elevated in the DOPA-P1, DOPA-P2, and DOPA-P1@P2 groups (Fig. 3J). Furthermore, in addition to protein-level analyses, analyses were also conducted at the gene level. The M1-associated gene Ccr-7 was highly expressed in the TiO₂ and DOPA groups, whereas the M2-associated gene Cd206 was elevated in the DOPA-P1, DOPA-P2, and DOPA-P1@P2 groups, as confirmed by reverse transcription quantitative polymerase chain reaction (RT-qPCR) analysis (Fig. 3N–O). The combined results from in vitro experiments indicated that the modified surfaces engineered in this study effectively promote M2 polarization of macrophages.

In order to assess the in vivo immunomodulatory effects, various treated titanium rods were inserted into the femurs of rats. Five days post-implantation, the femurs were extracted for histological analysis. The findings of H&E staining demonstrated a substantial decrease in the fibrous layer thickness adjacent to the titanium rods in the DOPA-P1, DOPA-P2, and DOPA-P1@P2 groups compared to the TiO₂ and DOPA groups (Fig. 4A, F). Additionally, immunohistochemical staining demonstrated a substantial enlargement of the area showing positive IL-10 staining in the DOPA-P1, DOPA-P2, and DOPA-P1@P2 groups, whereas a significantly increase in the area positive for TNF-α staining was identified in the TiO₂ and DOPA groups (Fig. 4B–C, G–H). Moreover, immunofluorescence staining was performed to evaluate macrophage polarization phenotypes surrounding the titanium rods. The abundance of M1 macrophages (CD68⁺CD86⁺) was notably elevated in the TiO₂ and DOPA groups compared to the rest, whereas M2 macrophages (CD68⁺CD206⁺) were more abundant in the DOPA-P1, DOPA-P2, and DOPA-P1@P2 groups (Fig. 4D–E, I–J). These results suggest that the developed surfaces modified with anti-inflammatory peptides can effectively modulate immunity during the acute phase of inflammation in vivo, promoting the timely shift from M1 to M2. In summary, data from in vitro and in vivo experiments indicated that the novel self-adaptive biomimetic titanium surfaces developed here (DOPA-P1@P2) can effectively modulate bone immunity, thereby creating a favourable immune microenvironment that supports subsequent stages of bone regeneration.

2.4 Modified Titanium Surfaces Indirectly Promote Angiogenesis and Osteogenesis

Angiogenesis and osteogenesis jointly constitute the proliferation stage of bone regeneration⁵⁵. Neovascularization provides a vital connection between bone and the surrounding tissues, enabling the transport of oxygen, nutrients, and BMSCs to the injured area⁵⁶. While the activation of the VEGF signalling pathway effectively promotes angiogenesis⁵⁷. BMSCs around the injury site differentiate into osteoblasts in the local microenvironment, with osteogenic growth peptide (OGP) significantly increasing their proliferation and differentiation⁵⁸. Importantly, angiogenesis and osteogenesis, which are both critical processes in bone regeneration, do not occur in isolation. Extensive molecular interactions between endothelial cells and osteoblasts, commonly referred to as “osteogenic–angiogenic coupling,” play a significant role in bone regeneration⁵⁹. To verify that the developed modified surfaces can indirectly promote angiogenesis and osteogenesis through immunomodulatory effects, we employed conditioned medium (CM) from macrophages in subsequent experiments (Fig. 5A). Initially, to evaluate the migratory potential of HUVECs, we conducted a wound healing assay (Fig. 5B). Compared with those in the TiO2^CM and DOPA^CM groups, the wound healing rates in the DOPA-P1^CM, DOPA-P2^CM, and DOPA-P1@P2^CM groups, whose implant surfaces possess immunomodulatory properties, were significantly greater (Fig. 5I). Additionally, Transwell assays further confirmed that following 24 hours of incubation, increased cell migration was observed in the DOPA-P1^CM, DOPA-P2^CM, and DOPA-P1@P2^CM groups compared to the TiO₂^CM and DOPA^CM groups (Fig. 5C, J). These results indicate that the CM extracted from surfaces modified with anti-inflammatory peptides effectively promoted the migration of HUVECs. Tube formation assays were conducted to assess vascularization in the different groups at 4 and 24 hours (Fig. 5D). Quantitative analysis revealed that, at both 4 and 24 h, the formation of branches and junctions was significantly greater in the DOPA-P1^CM, DOPA-P2^CM, and DOPA-P1@P2^CM groups than in the TiO₂^CM and DOPA^CM groups (Fig. 5K–L), suggesting that surfaces modified with anti-inflammatory peptides can regulate the immune microenvironment to promote angiogenesis. The western blot results were consistent with these findings, showing significantly higher levels of angiogenesis-related proteins (VEGF and FGF-2) in the groups with immunomodulatory implant surfaces (DOPA-P1^CM, DOPA-P2^CM, and DOPA-P1@P2^CM) than in those with implants without such properties (TiO₂^CM and DOPA^CM) (Fig. 5E, M–N).

In addition to demonstrating that the developed modified surfaces can regulate immunity to enhance angiogenesis, confirming their capacity to indirectly promote osteogenesis is essential. Macrophage CM from different modified surfaces was combined with osteogenic induction medium to culture BMSCs, with alkaline phosphatase (ALP) staining conducted on day 7 and alizarin red S (ARS) staining on day 21. ALP staining demonstrated that the area of positive staining for ALP was significantly greater in the DOPA-P1^CM, DOPA-P2^CM, and DOPA-P1@P2^CM groups than in the TiO₂^CM and DOPA^CM groups (Fig. 5G, Q). Similarly, ARS staining revealed that calcium nodule deposition was significantly greater in the groups with implants with immunomodulatory effects than in the other groups (Fig. 5H, R). Western blot results verified that the levels of COL-I and OPN were markedly higher in the DOPA-P1^CM, DOPA-P2^CM, and DOPA-P1@P2^CM groups compared to the TiO2^CM and DOPA^CM groups. (Fig. 5F, O–P). Additionally, the results of the immunofluorescence staining aligned with the aforementioned findings, indicating increased osteogenic activity in the groups with implants with immunomodulatory effects, as indicated by elevated OPN (depicted in red) expression (Figure S4 A–B). Finally, to validate the indirect angiogenic and osteogenic effects of the modified surfaces, assessments were also conducted at the gene level. RT-qPCR revealed that both angiogenesis-related genes (VEGF and FGF-2) and osteogenesis-related genes (Colla1 and Opn) were expressed at higher levels in the DOPA-P1^CM, DOPA-P2^CM, and DOPA-P1@P2^CM groups than in the TiO₂^CM and DOPA^CM groups (Figure S4C–F). In summary, the experimental results collectively indicated that surfaces modified with anti-inflammatory peptides effectively promote the polarization of macrophages towards the M2 phenotype, creating a favourable immunological microenvironment that indirectly enhances both angiogenesis and osteogenesis.

2.5 Modified Titanium Surfaces Directly Enhance Angiogenesis and Osteogenesis

Although regulating the bone immune microenvironment can promote angiogenesis and osteogenesis to some extent, this indirect enhancement is inherently limited and difficult to sustain over long durations. To achieve robust osseointegration at the implant interface, the surface must possess a direct and potent capacity to induce both angiogenesis and osteogenesis. In this work, we pretreated various modified surfaces with MMP-2, which specifically cleaves the PVGLIG peptide, to facilitate the release of anti-inflammatory peptides from the surface, thereby exposing the underlying sequences of angiogenic and osteogenic peptides (Fig. 6A). The angiogenic peptide K15 (KLTWQELYQLKYKGI) is a mimetic peptide of VEGF that effectively activates the VEGF signalling pathway, enhancing endothelial cell proliferation, migration, and angiogenic capabilities⁶⁰. Moreover, the osteogenic peptide Y5 (YGFGG) is the C-terminal fragment of osteogenic growth peptide (OGP), which enhances osteogenic cell proliferation and differentiation both in vivo and in vitro³⁰. Initially, we conducted wound healing assays with HUVECs on modified titanium surfaces after pretreatment with MMP-2. The results demonstrated that surfaces modified with angiogenic peptides (DOPA-P1 and DOPA-P1@P2) significantly increased wound closure rates compared with those of the other groups (TiO₂, DOPA, and DOPA-P2) (Fig. 6B, I). We conducted HUVEC migration assays using different peptide solutions. Similarly, the results indicated that the groups with surfaces containing angiogenic peptides (DOPA-P1 and DOPA-P1@P2) had notably higher numbers of migrating cells compared to the other groups. (Fig. 6C, J). Tube formation assays were then conducted, and the findings demonstrated that at both the 4-hour and 24-hour time points, the DOPA-P1 and DOPA-P1@P2 groups exhibited a substantially higher number of branches and junctions compared to the other groups (Fig. 6D, K–L). Furthermore, HUVECs were cultured directly on titanium surfaces after pretreatment with MMP-2, followed by extraction of proteins and RNA for western blot and RT-qPCR analyses, respectively. Angiogenesis-related proteins (VEGF and FGF-2) and genes (VEGF and FGF-2) were significantly upregulated in the DOPA-P1 and DOPA-P1@P2 groups compared with the TiO₂, DOPA, and DOPA-P2 groups (Figs. 6E, M–N and S5E–F). These results demonstrate that titanium surfaces modified with angiogenic peptides (DOPA-P1 and DOPA-P1@P2) directly promote angiogenesis.

We subsequently investigated whether modified titanium surfaces could directly promote osteogenesis. BMSCs were directly cultured on modified titanium surfaces after pretreatment with MMP-2 for osteogenic induction, with ALP staining conducted on day 7 and ARS staining on day 21. The findings indicate that surfaces modified with osteogenic peptides (DOPA-P2 and DOPA-P1@P2) presented significantly greater ALP activity and calcium nodule deposition than the other surfaces did (TiO₂, DOPA, and DOPA-P1) (Fig. 6G–H, Q–R). Subsequent western blotting and immunofluorescence assays were performed to examine protein expression levels, revealing a significant elevation of COL-I and OPN in the DOPA-P2 and DOPA-P1@P2 groups compared to the other groups (Figs. 6F, O–P and S5A–B). Finally, the results of gene expression analysis were consistent with the protein-level findings, with increased expression of Colla1 and Opn in the DOPA-P2 and DOPA-P1@P2 groups relative to the TiO₂, DOPA, and DOPA-P1 groups (Figure S5 C–D). These results suggest that surfaces modified with osteogenic peptides (DOPA-P2 and DOPA-P1@P2) possess a strong capacity to directly enhance osteogenesis. In summary, the novel biomimetic surface (DOPA-P1@P2) developed in this study not only modulates the immune microenvironment to indirectly enhance angiogenesis and osteogenesis but also directly promotes these processes.

2.6 Exploration of the Mechanisms Underlying the Increased Angiogenesis and Osteogenesis Observed on Modified Titanium Surfaces

To explore how the novel biomimetic modified surfaces directly enhance angiogenesis and osteogenesis, HUVECs and BMSCs were cultured on TiO₂ and DOPA-P1@P2 surfaces, and samples from both groups were subsequently collected for transcriptomic sequencing. The results of principal component analysis (PCA), box plots of gene expression levels, regional distribution plots, and heatmaps of correlation coefficients among samples, along with clustering analysis, indicated that the sequencing data for HUVECs and BMSCs met the required standards, confirming the reliability of the results (Figs. 7A, G and S6A-H). Volcano plot analysis revealed that, in HUVECs, compared with the TiO₂ group, the DOPA-P1@P2 group presented 411 upregulated genes and 557 downregulated genes (Fig. 7B). In BMSCs, compared with the TiO₂ group, the DOPA-P1@P2 group presented 851 upregulated and 976 downregulated genes (Fig. 7H). As depicted in the heatmaps in Figs. 7C and 7I, notable differences were observed in the expression patterns of differentially expressed genes (DEGs) between the TiO₂ and DOPA-P1@P2 groups. The radar chart of the differentially expressed genes further confirmed the above results (Figure S7A, D). These results indicate that the novel biomimetic modified surface developed here (DOPA-P1@P2) substantially affects the proliferation and differentiation processes of HUVECs and BMSCs.

After sequencing of the HUVEC samples, Gene Ontology (GO) enrichment analysis suggested that a considerable portion of the DEGs were involved in the regulation of angiogenesis (Figs. 7E and S7B). Moreover, findings from Kyoto Encyclopedia of Genes and Genomes (KEGG) enrichment analysis showed that the MAPK signalling pathway may play a pivotal role in modulating angiogenesis (Figs. 7F and S7C). Furthermore, the results from gene set enrichment analysis (GSEA) showed that the MAPK signalling pathway was significantly upregulated in the DOPA-P1@P2 group in comparison to the TiO₂ group (Fig. 7D). However, in the GO enrichment analysis of the BMSC samples, we found that numerous DEGs were enriched in processes such as the extracellular matrix and bone mineralization, which are crucial for osteogenesis (Figs. 7K and S7E). Furthermore, the results from the KEGG enrichment analysis revealed that the differentially expressed gene sets were enriched in the PI3K–Akt signalling pathway, with GSEA further indicating significant activation of this pathway in the DOPA-P1@P2 group in comparison to the TiO₂ group (Figs. 7J, L and S7F). Consequently, it is possible that DOPA-P1@P2 surfaces promote angiogenesis and osteogenesis through activation of the MAPK and PI3K–Akt signalling pathways, respectively.

2.7 Novel Biomimetic-modified Surfaces for Temporal Regulation of Bone Regeneration and Enhanced Osseointegration at the Implant Interface

Our in vitro investigations revealed that DOPA-P1@P2 surfaces are capable of modulating immune responses while facilitating angiogenesis and osteogenesis. However, whether the DOPA-P1@P2 surface can similarly exert these effects in vivo, thereby achieving stable osseointegration at the interface, is a key focus of this study. This question highlights the primary innovation of our research, which is the development of inflammation-responsive modified titanium surfaces that regulate the process of bone regeneration in an orderly manner to achieve optimal osseointegration. Here, we implanted titanium rods with various modifications into the femurs of rats and collected the femurs for micro-CT and histological examinations after 8 weeks (Fig. 8A). Three-dimensional images reconstructed from micro-CT scans clearly demonstrated that the bone mass around the titanium rods was greatest in the DOPA-P1@P2 group, followed by the DOPA-P2 and DOPA-P1 groups (Fig. 8B). The quantitative bone mineral density (BMD) results clearly showed that the DOPA-P1@P2 group presented the highest BMD (1.31 ± 0.09 g/cm³), outperforming the DOPA-P2 (0.96 ± 0.06 g/cm³) and DOPA-P1 (0.86 ± 0.05 g/cm³) groups, whereas the lowest BMD was recorded in the DOPA (0.55 ± 0.04 g/cm³) and TiO₂ (0.55 ± 0.03 g/cm³) groups (Fig. 8E). Additionally, the quantitative results for the trabecular bone parameters (BV/TV, Tb.N, and Tb.Th) further confirmed the superior osseointegration in the DOPA-P1@P2 group (Fig. 8F–H). For example, the bone volume fraction (BV/TV) in the DOPA-P1@P2 group reached 61.44 ± 2.83%, which was 1.47-fold greater than that of the DOPA-P2 group, 1.83-fold greater than that of the DOPA-P1 group, and over 3-fold greater than that of the TiO₂ group.

Following micro-CT analysis, we conducted undecalcified bone slicing and van Gieson (VG) staining of femurs containing titanium rods to assess the bone-implant contact ratio (BIC) across various groups. As expected, the DOPA-P1@P2 group presented a bone-implant contact ratio (BIC) as high as 61.10 ± 5.12%, which was markedly higher than the measurements observed for all other groups, with the BIC values in the DOPA-P1 and DOPA-P2 groups also showing marked enhancements over those in the TiO₂ and DOPA groups (Fig. 8I). Additionally, dual labelling with calcein confirmed that the mineral apposition rate (MAR) of the DOPA-P1@P2 group was markedly greater compared with the other groups, demonstrating its great potential to promote osteogenesis (Fig. 8J). The mechanical push-out test (Fig. 8K) is a critical method for evaluating the integration of an implant with surrounding bone tissue⁶¹. We observed that DOPA-P1@P2 achieved the highest mechanical push-out force (82.12 ± 3.12 N), which was substantially higher than the values observed in the other groups (Fig. 8L). This force was 1.29 times greater than that of the DOPA-P2 (63.52 ± 3.58 N) group, 1.65 times greater than that of the DOPA-P1 (49.76 ± 3.67 N) group, and more than 2.5 times greater than that observed in the DOPA (32.31 ± 2.41 N) and TiO₂ (31.35 ± 2.05 N) groups, suggesting that the osseointegration achieved in the DOPA-P1@P2 group was the most robust. Notably, the mechanical push-out strength of the DOPA-P1@P2 group was notably higher than that of the DOPA-P1 and DOPA-P2 groups, likely due to the synergistic effects of angiogenesis and osteogenesis achieved on the DOPA-P1@P2 surface. Finally, a subset of femurs containing titanium rods was decalcified, after which the titanium rods were carefully removed for subsequent Masson and immunohistochemical staining. In alignment with the aforementioned findings, the DOPA-P1@P2 group demonstrated the highest collagen volume fraction and the greatest levels of COL-I and OPN in comparison to the rest (Figure S8A–F). In summary, the DOPA-P1@P2 surface can regulate the process of bone regeneration in an orderly manner, enhancing the synergistic effects of angiogenesis and osteogenesis and thereby achieving optimal osseointegration at the interface.

In summary, we present a novel design concept for titanium-based coatings, focusing on the sequential regulation of bone regeneration stages to optimize interface osseointegration. Briefly, we developed a novel self-adaptive biomimetic titanium surface (DOPA-P1@P2) by combining a mussel-inspired biomimetic strategy with bioorthogonal click chemistry techniques. Upon implantation, the outermost K23 layer modulates acute inflammation during the first stage of bone regeneration, facilitating macrophage polarization from M1 to M2 phenotype. Activated macrophages continuously secrete MMP-2/9, which specifically cleaves the PVGLIG sequences, resulting in detachment of the outermost K23 layer and subsequent exposure of the underlying K15 and Y5 layers, which promote angiogenesis and osteogenesis during the second stage, followed by bone remodelling. This novel self-adaptive biomimetic surface not only supports immunomodulation, angiogenesis, and osteogenesis during bone regeneration but also coordinates these functions across different stages, achieving superior osseointegration at the bone–implant interface. This novel design concept for multifunctional surface modification holds promise for translational and clinical applications.

4.1 Peptide Synthesis and Surface Modification

The mussel-inspired peptide (DOPA)₄-PEG₅-DBCO, which features multiple DOPA units and DBCO groups, and two azide-modified composite functional peptides, P1 (N₃-K15-PVGLIG-K23) and P2 (N₃-Y5-PVGLIG-K23), were produced using the Fmoc solid-phase synthesis approach by ChinaPeptides Co., Ltd. (QYAOBIO). The corresponding molecular designs are as follows: Ac-(DOPA)-G-(DOPA)-K[(PEG₅)-(DBCO-COOH)]-(DOPA)-G-(DOPA), (2-Azido)-KLTWQELYQLKYKGIPVGLIGKAFAKLAARLYRKALARQLGVAA, and (2-Azido)-YGFGGPVGLIGKAFAKLAARLYRKALARQLGVAA. Here, PEG was used as a spacer to reduce steric hindrance and facilitate reactions, whereas DBCO and Azido functioned as the groups that facilitated bioorthogonal reactions. Titanium plates, 1 mm in thickness and 15 mm in diameter, along with titanium rods, 10 mm long and 1.5 mm in diameter, were procured from Tianjin Zhengtian Medical Devices Co., Ltd., China. After being treated with oxygen plasma, the titanium rods and plates were cleaned three times with acetone, 75% alcohol, and deionized water, followed by sterilization under high temperature and pressure. To minimize amino acid oxidation, all peptide solutions were purged with nitrogen for 15 minutes prior to use. Initially, sterile titanium plates and rods were submerged in a 0.01 mg/ml solution of (DOPA)₄-PEG₅-DBCO for 48 hours to coat the titanium surface with mussel-inspired peptides through the spontaneous coordination of catechol groups and metal ions. The DBCO group in the mussel-inspired peptides provides a click site for the subsequent grafting of peptides. Subsequently, surfaces modified with mussel-inspired peptides (defined as the DOPA group) were separately immersed in 0.1 mg/ml solutions of P1, P2, or a 1:1 mixture of both for 4 hours, forming different modified surfaces (defined as the DOPA-P1, DOPA-P2, and DOPA-P1@P2 groups, respectively) through bioorthogonal click chemistry reactions between -DBCO and -N₃. Plates or rods immersed in PBS were designated the TiO₂ group and served as the control. The surfaces with different modifications were meticulously cleaned with MiniQ water and subsequently dried under nitrogen.

4.2 Material Characterization

The purity of (DOPA)₄-PEG₅-DBCO, P1 (N₃-K15-PVGLIG-K23) and P2 (N₃-Y5-PVGLIG-K23) were evaluated via HPLC, and their corresponding molecular weights were measured via ESI–MS. The presence of specific catechol groups in the mussel-inspired peptides was subsequently detected via proton nuclear magnetic resonance (¹NMR). AFM was utilized to evaluate the surface morphology and quantify the roughness of the different groups. The hydrophilicity of different groups was evaluated via measuring the static water contact angle. XPS was used to determine the elemental compositions of the different modified surfaces. The P1 and P2 surfaces were immersed in solutions without MMP-2 and with 1.5 µg/ml MMP-2, respectively, and incubated at 37°C. Supernatants were collected at various time points, and the release trends of PVGLIG peptide sequences were tracked using HPLC.

4.3 Cell Culture

In this work, three distinct cell types were utilized for experimental investigation. The RAW264.7 cell line and the HUVEC line were supplied by the Institute of Biochemistry and Cell Biology of the Chinese Academy of Sciences (Shanghai, China), while BMSCs were extracted from the femoral bone marrow of 4-week-old Sprague–Dawley (SD) rats. In brief, the extraction of BMSCs required a relatively sterile environment. The bilateral femurs of the rats were collected, and the adjacent soft tissues were carefully removed. Subsequently, scissors were used to carefully cut off both ends of the femur, and a 1 ml syringe was used to flush the marrow cavity with culture medium repeatedly until the flushed medium became clear. The mixture was subsequently gathered and passed through a mesh filter. After filtration, the cell suspension was exposed to a red blood cell lysis solution and then centrifuged. The resulting cells were resuspended and cultured in a dish. When the cell density in the culture dish reached approximately 80%, the cells were digested and passaged with 0.25% trypsin/EDTA solution. RAW264.7 cells and HUVECs were cultured in high-glucose DMEM supplemented with 10% fetal bovine serum (FBS), with cell passages occurring typically every 1‒2 days. However, BMSCs were cultured in low-glucose DMEM supplemented with 10% FBS, with cell passage typically performed every 2‒3 days.

4.4 Evaluation of Biocompatibility

Three types of cells were seeded onto differently modified titanium surfaces and cultured for 24 hours, followed by staining using the Calcein–AM/PI Double Stain Kit (Yeasen, China). Cells were cultured on different modified surfaces for 24 hours, and cytotoxicity was assessed using the LDH Cytotoxicity Assay Kit (Beyotime, China). Furthermore, cells were cultured on diverse titanium surfaces, with cell viability evaluated at 24 and 72 hours using the Cell Counting Kit-8 (Yeasen, China). Finally, we performed cytoskeletal staining with phalloidin (Yeasen, China) to assess the adhesion and spreading characteristics of BMSCs on different modified surfaces. Briefly, BMSCs cultured on various modified surfaces for 12 hours were fixed with 4% paraformaldehyde for a duration of 20 minutes, incubated with FITC-labelled phalloidin for 30 minutes, stained with DAPI for 10 minutes, and subsequently imaged using a confocal microscope (Zeiss, LSM800, Germany). Semi-quantitative analysis was performed using ImageJ software (version 1.54d), with all data analyzed in triplicate.

4.5. Evaluation of Macrophage Polarization

RAW264.7 cells were distributed across various surfaces at 20,000 cells for each and exposed to 8 hours of treatment using 100 ng/mL lipopolysaccharide (LPS) sourced from Sigma-Aldrich. Fresh DMEM was then added after replacing the old medium, and the cells were left to incubate for a further 48-hour period. Following a 48-hour period, supernatants from each group were gathered, and ELISA kits were employed to measure the levels of TNF-α and IL-10. Simultaneously, the collected cells were subjected to immunofluorescence staining to assess their polarization characteristics on different modified surfaces. Concisely, samples from various groups were thoroughly rinsed thrice using PBS before being fixed in 4% paraformaldehyde on ice for 20 minutes. The cells were then treated with 0.2% Triton X-100 (Sigma) to induce permeabilization for 10 minutes. Subsequently, the samples underwent a 10-minute blocking phase using QuickBlock™ Blocking Buffer for Immunol Staining (Beyotime, P0260). Afterward, the samples were incubated overnight at 4°C with primary antibodies against iNOS (Abcam, ab178945) and Arg-1 (Abcam, ab91279). The following day, the cytoskeleton was stained with Phalloidin–iFluor 488 (Abcam, ab176753), while target proteins were labeled using goat anti-rabbit IgG H&L (Abcam, ab150079), followed by a 1-hour incubation at 37°C. Ultimately, the samples were prepared with Antifade Mounting Medium containing DAPI (Beyotime, P0131). The polarization states of macrophages on various modified surfaces were analysed via flow cytometry, western blotting (WB), and RT-qPCR. Detailed descriptions of the methods used for WB and RT-qPCR will be provided in a subsequent section. Here, we briefly introduce the flow cytometry procedure. Cells from every group were collected and incubated for 30 minutes with antibodies specific for F4/80 (Thermo, 11-4801-82), CD206 (Thermo, 17-2061-80), and CD86 (Thermo, 12-0862-81). After rinsing the samples twice using PBS supplemented with 5% BSA, the cells were then resuspended. Finally, the subsets of M1 and M2 macrophages within the different samples were identified using flow cytometry (Thermo Scientific, USA). Flow cytometry data were processed using FlowJo V10 software, and analyses were conducted in triplicate.

4.6 Evaluation of Angiogenesis

This section of the experiment is divided into two parts investigating 1) the indirect angiogenic effects of modified surfaces and 2) the direct angiogenic effects of modified surfaces. To investigate whether various modified surfaces can modulate the immune microenvironment to indirectly promote angiogenesis, supernatants were gathered from RAW264.7 cells that had been cultured on these surfaces. Next, the supernatants collected from the RAW264.7 cells and fresh high-glucose DMEM were mixed in equal proportions to prepare CM. HUVECs were plated in 6-well plates at a concentration of 2 × 10⁵ cells per well and cultured in high-glucose DMEM with 10% FBS over a period of 24 hours. The next day, once the cells reached confluence in the wells, a uniform scratch was made in each well and CM was added. The scratch width was observed under a microscope (Zeiss, Germany) at 0 and 24 hours. A migration assay was performed using a Transwell system (Corning, USA) to evaluate cell movement. HUVECs were plated at a concentration of 1 × 10⁴ cells per well in the upper chamber, while CM from different groups were added to the lower chamber for a 24-hour incubation. To evaluate cell migration, the cells were stained using a 0.1% crystal violet solution (Solarbio, China). Ultimately, we performed an angiogenesis assay. Briefly, the ABW® Matrigel matrix was kept on ice and left to thaw overnight at 4°C. After thawing, the matrix was mixed at a 1:1 ratio with fresh, serum-free DMEM. Fifty microliters of the solution were added to a 96-well plate and placed in a cell culture incubator for 40 minutes to enable solidification. Following this, HUVECs were introduced into the previously mentioned 96-well plate with 20,000 cells per well and were incubated with CM from various modified surfaces. Tube formation in HUVECs was visualized at 4 and 24-hour time points. Additionally, HUVECs were incubated in 6-well plates with CM from different groups. After 24 hours, RNA and proteins were extracted for RT-qPCR and WB analysis.

Before confirming their direct angiogenic effects, the modified titanium surfaces were pretreated with a 1.5 µg/ml MMP-2 solution for 72 hours to eliminate the influence of anti-inflammatory peptide sequences on the experiment. Subsequently, HUVECs were directly seeded onto the pretreated modified titanium surfaces and incubated for 24 hours in DMEM supplemented with 10% FBS. The next day, a uniform scratch was created on the modified titanium surface, followed by calcein–AM staining at 0 and 24 hours, and cell migration was examined using an inverted fluorescence microscope. Since the titanium plates are impenetrable, we incorporated peptides from various modified titanium surfaces into DMEM and subsequently conducted migration and tube formation experiments using this peptide-enriched medium. A density of 1 × 10⁴ cells per well was used to seed HUVECs in the upper chamber, with peptide-enriched medium from different groups in the lower chamber for a 24-hour incubation, and cell migration was assessed using 0.1% crystal violet solution (Solarbio, China). The procedure for the tube formation assay was the same as above, except that the CM was replaced with peptide-enriched medium. Prior to examination, the samples were treated with calcein–AM and analyzed using fluorescence microscopy. Finally, HUVECs were directly seeded onto titanium surfaces pretreated with MMP-2 and incubated for 24 hours. RNA and proteins were then extracted for RT-qPCR and WB analysis.

4.7 Evaluation of Osteogenesis

Similarly, experiments related to osteogenesis were conducted via two separate approaches: indirect and direct. First, to verify the indirect osteogenic effect of the surfaces, the supernatant from RAW264.7 cells incubated on various titanium surfaces was collected and mixed with fresh low-glucose DMEM at a 1:1 ratio to prepare macrophage CM. BMSCs were seeded in plates at a density of 2x10⁴ cells per well and incubated for 36 hours in low-glucose DMEM containing 10% FBS. The initial culture medium was replaced with CM containing osteogenic supplements, specifically 10 mM β-glycerophosphate, 0.1 µM dexamethasone, and 0.25 mM ascorbate. The media was refreshed every 2–3 days. On day 7, ALP staining was performed, and on day 21, ARS staining was carried out. The procedures followed the standard protocols provided with the ALP Staining Kit (Beyotime, C3206) and the ARS Staining Kit (Beyotime, C0148S). After the BMSCs were incubated for 48 hours in CM containing osteogenic factors, immunofluorescence staining was conducted to quantify the expression levels of OPN (Immunoway, YT3467) in different groups. Simultaneously, RNA and proteins were extracted from different groups for RT-qPCR and WB analysis.

After the titanium discs were pretreated with MMP-2 solution for 72 hours, BMSCs were directly seeded onto the pretreated titanium surfaces. The BMSCs were incubated in fresh low-glucose DMEM containing 10% FBS and osteogenic factors, and the medium was changed every 2–3 days. After 48 hours, immunofluorescence staining was performed to assess the levels of COL-I (Abcam, ab270993) in different groups. Simultaneously, RNA and proteins were extracted from different groups for RT-qPCR and WB analysis. Similarly, on days 7 and 21, ALP and ARS staining were performed directly on the incubated cells on the modified titanium surfaces.

4.8 RT-qPCR and Western Blotting

Total RNA was extracted from cells subjected to various treatments using TRIzol™ reagent (Thermo Fisher, 15596026CN). Complementary DNA (cDNA) was synthesized from the extracted RNA through reverse transcription. The resulting cDNA was then amplified using a CFX96™ Real-Time PCR System (Bio-Rad, USA). The primers used for the target genes are detailed in Table S1 (Supporting Information).

Total protein was extracted after various treatments and then measured for quantification. The proteins were subjected to SDS‒PAGE for 90 minutes at 110 V, followed by transfer onto PVDF membranes at 350 mA for 30 minutes. After blocking, the membranes were incubated overnight at 4°C with primary antibodies targeting iNOS (Abcam, ab178945), Arg-1 (Abcam, ab91279), VEGF (Immunoway, YN5444), FGF-2 (Abcam, ab208687), COL-I (Abcam, ab270993), OPN (Immunoway, YT3467), and β-actin (Immunoway, YM3028). After thorough rinsing with TBST, the membranes were treated with appropriate secondary antibodies for one hour.

4.9 Animal Models

All procedures involving animals strictly adhered to animal welfare guidelines and were approved by the Ethics Committee of the First Affiliated Hospital of University of Science and Technology of China, with the corresponding ethics approval number: 2024-N (A)-0205. Ninety 8-week-old Sprague–Dawley (SD) rats were evenly divided into five groups: the TiO₂, DOPA, DOPA-P1, DOPA-P2, and DOPA-P1@P2 groups. The rats were anaesthetized via isoflurane inhalation, followed by the implantation of differently modified titanium rods into both femurs of the rats. The detailed surgical procedure is shown in Figure S10. Five days post-surgery, six rats per group were euthanized and their femurs collected for inflammation assessment. The remaining ten rats in each group were administered an intraperitoneal injection of 10 mg/kg calcein (Sigma) 10 days and 3 days prior to euthanasia. All rats were euthanized two months following surgery, and their bilateral femurs, heart, liver, spleen, lungs, and kidneys were obtained for additional research.

4.10 Micro-CT and Biomechanical Push-Out Experiment

Two months post-surgery, all harvested femurs underwent micro-CT scans using a SkyScan 1176 system (Belgium). The femur of each rat was imaged with a spatial resolution of 18 µm per layer, with the voltage and current set at 50 kV and 500 µA, respectively, and the rotational step set at 0.7°. The micro-CT scan data were analysed using CTAn software, which was used to measure the bone mineral density (BMD, g/cm³), bone volume per tissue volume (BV/TV, %), trabecular number (Tb.N, 1/mm), and trabecular thickness (Tb.Th, µm) across various groups. Three-dimensional reconstruction of the CT data was performed to visually compare bone integration across the different groups.

Five right femurs from each experimental group underwent biomechanical push-out testing. The maximum push-out force was evaluated utilizing the HY1080 material testing system (China). Before the mechanical tests, the femurs were fixed vertically to the base using dental cement. A force parallel to the femoral longitudinal axis was then applied to push the femurs downwards with a speed of 1 mm/min, with the maximum push-out force recorded during the process.

4.11 Histological Evaluation

Following biomechanical testing, the remaining five right femurs from each group, each containing a titanium rod, were dehydrated in 70% ethanol and subsequently used for hard tissue sectioning. The hard tissue sections containing titanium rods were subjected to van Gieson's (VG) staining to evaluate the variations in bone-implant contact (BIC) among the different groups. Concurrently, hard tissue sections labelled with calcein were examined to assess the differences in bone mineralization rates (MARs) between the groups. Ten left samples per group were harvested and subsequently immersed in a 10% ethylenediaminetetraacetic acid (EDTA) solution to facilitate gradual decalcification. The entire process was carried out in a constant-temperature shaking incubator at 37°C for five weeks. Upon completion of the decalcification process, the titanium rods were meticulously extracted from the femurs. The tissues were processed into paraffin sections for further study. Histological sections were meticulously prepared and immunohistochemical staining was performed on these paraffin sections to detect the levels of COL-I and OPN. Collagen deposition around the titanium rods was also assessed by performing Masson staining. Additionally, the femurs of rats from each group, which were extracted on the fifth day post-surgery, were subjected to slow decalcification and processed into paraffin sections. These sections were initially stained with H&E to assess fibroproliferation around the titanium rods. Immunohistochemical staining was subsequently carried out to evaluate the differences in the expression of TNF-α (Servicebio, GB11188) and IL-10 (Servicebio, GB11534) between the groups. Finally, immunofluorescence staining was performed using CD68 (Abcam, ab201340) to label macrophages, CD86 (Immunoway, YT7823) as a marker of M1 polarization, and CD206 (Abcam, ab64693) as a marker of M2 polarization. All staining processes were performed following the standard protocols outlined by the reagent suppliers.

4.12 Transcriptome Sequencing

We conducted two transcriptome sequencing analyses to explore the mechanisms of direct angiogenesis and direct osteogenesis on DOPA-P1@P2 surfaces. HUVECs and BMSCs were individually seeded onto TiO₂ and DOPA-P1@P2 surfaces that had been pretreated with MMP-2 and then incubated for two days. Following the extraction of total RNA, RNA purity and integrity were evaluated using an Agilent 2100 Bioanalyzer (Agilent Technologies, USA). Libraries were subsequently constructed using a VAHTS Universal V6 RNA-seq Library Prep Kit in accordance with the manufacturer's instructions. Transcriptome sequencing and analysis were carried out by OE Biotech Co., Ltd. (China).

4.13 Statistical Analysis

The data are expressed as means ± standard deviations, derived from a minimum of three replicates for each experimental sample. Differences between two groups were evaluated using the Student's t test, while the one-way ANOVA was applied for comparisons among multiple groups. Statistical significance was defined as p < 0.05 (*) and strong significance as p < 0.01 (**). All statistical evaluations were conducted with GraphPad Prism version 10.1.0.

Author contributions

WZ and WH contributed to the concept and design. Data acquisition was carried out by WZ, YL, and XN. Data analysis and interpretation were conducted by WZ, YL, and XN. WZ, CZ, Lm X, JZ, and WH were involved in drafting the manuscript. CZ, Lm X, JZ, and WH critically revised the manuscript and approved the final article.

Competing interests

The authors state that they have no conflicts of interest.

Acknowledgments

This work was supported by the National Natural Science Foundation of China (Grant No. 82472437), the Provincial Key R&D Programmes of Anhui (Grant No. S2022e07020018) and the Hefei Natural Science Foundation (Grant No. 2022045).

Data availability

All data can be obtained from the authors upon request. Source data for select figures and supplementary figures are included in a source data file.

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Scheme 1 is available in the Supplementary Files section

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image1.png
Scheme 1: Schematic diagram of a novel smart biomimetic implant coating that optimizes osseointegration at the implant interface through temporal regulation of bone regeneration. (A) Mussel-inspired peptide modified with a bioclickable DBCO group: (DOPA)₄-PEG₅-DBCO. (B) Azide-modified multifunctional peptide P1 with angiogenic and anti-inflammatory properties: N₃-K15-PVGLIG-K23. (C) Azide-modified multifunctional peptide P2 with osteogenic and anti-inflammatory properties: N₃-Y5-PVGLIG-K23. (D) Mussel-inspired peptides bind to titanium surfaces through spontaneous coordination between catechol groups and titanium ions, whereas P1 and P2 are grafted onto biomimetic titanium surfaces via a bioorthogonal click chemistry reaction between N₃ and DBCO. (E) Synthesis of a novel smart biomimetic titanium coating: DOPA-P1@P2. (F) DOPA-P1@P2 promoted M2 macrophage polarization during the first stage of bone regeneration. (G) DOPA-P1@P2 promoted angiogenesis and osteogenesis during the second stage of bone regeneration. (H) In the third stage of bone regeneration, the implant interface achieves robust osseointegration.
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Peptide-based Self-adaptive Implant Coating Sequentially Regulates Bone Regeneration to Enhance Interfacial Osseointegration

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Abstract

Figures

1. Introduction

2. Results and Discussion

2.1 Titanium Surface Modification and Material Characterization

2.2 Biocompatibility Evaluation of Modified Titanium Surfaces

2.3 Immunomodulatory Effects of Modified Titanium Surfaces

2.4 Modified Titanium Surfaces Indirectly Promote Angiogenesis and Osteogenesis

2.5 Modified Titanium Surfaces Directly Enhance Angiogenesis and Osteogenesis

2.6 Exploration of the Mechanisms Underlying the Increased Angiogenesis and Osteogenesis Observed on Modified Titanium Surfaces

3. Conclusion

4. Methods and Materials

4.1 Peptide Synthesis and Surface Modification

4.2 Material Characterization

4.3 Cell Culture

4.4 Evaluation of Biocompatibility

4.5. Evaluation of Macrophage Polarization

4.6 Evaluation of Angiogenesis

4.7 Evaluation of Osteogenesis

4.8 RT-qPCR and Western Blotting

4.9 Animal Models

4.10 Micro-CT and Biomechanical Push-Out Experiment

4.11 Histological Evaluation

4.12 Transcriptome Sequencing

4.13 Statistical Analysis

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References

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Additional Declarations

Supplementary Files

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