An immunomodulatory bioactive glass orchestrates early bone healing via coordinated macrophage polarization and osteogenesis-osteolysis balance

Abstract

Effective repair of large bone defects remains a major challenge in regenerative medicine. Conventional strategies often emphasize enhancing osteogenesis, while the role of the early immune microenvironment remains insufficiently addressed. Bone regeneration is not merely a structural filling process but a complex physiological program orchestrated by the immune system, requiring a dynamic balance between osteogenesis and osteolysis. Here, we propose an immune-instructive strategy aimed at modulating the early immune response to establish a favorable microenvironment for efficient bone regeneration. Based on this concept, a pH-neutral bioactive glass functionalized with Arginine-Glycine-Aspartic Acid-Serine (RGDS) peptides, designated as NBG@RGDS, was designed and fabricated to achieve synchronized and multi-targeted regulation of the osteoimmune microenvironment. This system concurrently orchestrates three pivotal stages in osteoimmunology including polarizing macrophages toward a pro-repair M2 phenotype, inhibiting excessive osteoclastogenesis and osteoclast activity, and promoting the osteogenic differentiation of bone marrow mesenchymal stem cells (BMSCs). Transcriptomic analysis suggested the potential mechanisms through which NBG@RGDS regulates macrophages. In a rat femoral condyle defect model, NBG@RGDS optimized the early immune landscape, effectively attenuated inflammation, and markedly accelerated early-stage new bone formation. This study demonstrates that modulating the initial immune crosstalk represents a more fundamental and efficient strategy for bone regeneration than merely stimulating osteogenesis. It provides a novel perspective for the development of next-generation immunomodulatory bone repair materials.

Graphical abstract

1. Introduction

The regeneration of large bone defects, particularly in patients suffering from osteoporosis [1] or diabetes [[2], [3], [4]], remains a formidable clinical challenge [5,6]. Autografts, traditionally considered the gold standard, are limited by donor site availability and morbidity; allografts face risks of immune rejection and ethical concerns [7]; while xenografts generally lack osteoinductive activity and exhibit slow degradation, restricting their clinical application. With the advantage of tissue engineering, strategies based on constructing scaffolds combined with seed cells and growth factors have shown promising application prospects by creating a favorable microenvironment to promote cell proliferation and osteogenic differentiation. While the development of bioactive scaffolds has advanced significantly, conventional strategies are often constrained by a predominant focus on providing mechanical support or solely promoting osteoblast differentiation [8]. However, increasing evidence indicates that dysregulated early inflammation and excessive osteoclast activation critically impair bone healing [9,10], suggesting that osteogenesis-centered approaches alone are insufficient for achieving effective regeneration.

In fact, bone regeneration is far from a simple matter of osteogenesis. It is a complex biological program meticulously orchestrated by the immune system, involving a dynamic balance between osteogenesis and osteolysis [9,11]. Recent advances in osteoimmunology have further revealed that successful bone repair depends not only on osteogenic activity but also on the timely regulation of bone metabolic balance by the immune microenvironment [12]. Therefore, moving beyond the traditional pro-osteogenic single target approach and developing novel biomaterials capable of actively guiding early immune responses while synergistically regulating the osteogenesis–osteolysis balance has become key to advancing the field [13].

Achieving this goal faces dual and interrelated regulatory challenges. First, trauma and the implantation procedure itself trigger a strong acute inflammatory response, which can be exacerbated by the implant itself [14]. Macrophages, as the central coordinators of inflammation and repair, determine the trajectory of the microenvironment through their functional polarization [15]. While the initial M1 macrophage-dominated response is essential for clearing necrotic tissue [16], its persistence or overactivation leads to chronic inflammation that hinders the regeneration process [17]. Therefore, proactively regulating macrophage polarization to facilitate a timely transition from an early pro-inflammatory state to a pro-repair state is crucial for managing the inflammatory microenvironment [18,19]. Second, the quality of repair fundamentally depends on the dynamic balance between osteogenic and osteoclastic activities [13]. In pathological microenvironments, the abnormal activation of osteoclasts leads to excessive bone resorption [20,21], while the insufficient osteoblast activity hinders effective bone filling [22]. An imbalance between these processes directly results in repair failure. Thus, direct regulation of the osteogenesis–osteolysis balance is the cornerstone of high-quality structural regeneration [23]. Although appropriate immune responses can indirectly influence this balance [15], direct and precise regulation of osteoblasts and osteoclasts, complementing macrophage immunomodulation, is indispensable for the successful repair of complex bone defects. An ideal bone repair material may possess these multi-target synergistic capabilities: capable of guiding and optimizing the post-traumatic and post-implantation immune microenvironment, while simultaneously intervening in the core processes of bone metabolism [24].

Bioactive glass has emerged as a multifunctional inorganic material with excellent bioactivity and osteoinductive properties [25]. In recent years, it has been widely applied in orthopedic and dental fields. This material is capable of forming a bone-like carbonate hydroxyapatite layer on its surface under physiological conditions, thereby promoting integration with the host bone. Meanwhile, the release of ions such as Si, Ca and P activates the expression of osteogenesis-related genes and induces the osteogenic differentiation of bone marrow mesenchymal stem cells (BMSCs) [26]. Beyond ion release, bioactive glass can also regulate cell behavior and tissue regeneration through its structural features [27], surface functionalization [28] or the loading of bioactive factors [29]. In addition to its osteogenic capacity, bioactive glass has also demonstrated pro-angiogenic and antibacterial properties. In recent years, its potential in immunomodulation has gradually been uncovered, and modulating the immune system to promote tissue repair has become a new research focus. Advances in this field suggest that multifunctional platforms based on bioactive glass scaffolds hold promise for addressing multiple regulatory challenges. For example, the design of multifunctional bioactive glass scaffolds has enabled synergistic effects, such as combining bone tumor therapy with bone regeneration [[30], [31], [32]]. Collectively, bioactive glass has evolved from a traditional bone graft substitute into a versatile platform capable of simultaneously modulating the immune microenvironment and bone metabolism.

Guided by the dual-regulation concept, we designed a multifunctional bone repair material which is capable of synergistic immune-osteogenic regulation. A pH-neutral bioactive glass (NBG), previously shown to possess favorable osteoinductive properties [33,34], was selected as the base material and functionalized with covalently immobilized Arginine-Glycine-Aspartic Acid-Serine (RGDS) peptides to generate NBG@RGDS. This design targets three critical stages of early bone healing: modulating trauma-induced inflammation, suppressing aberrant osteoclastic activity, and robustly initiating osteogenic programs. By coordinating immune regulation with osteogenesis–osteolysis balance, NBG@RGDS aims to establish a favorable microenvironment for efficient bone regeneration, providing a rational design strategy for next-generation immunomodulatory bone repair materials (Scheme 1).

Scheme 1.

Scheme illustration of the preparation of NBG@RGDS and its promotion of bone defect repair via coordinating macrophage polarization and osteogenesis-osteolysis balance.

2. Materials and methods

2.1. NBG@RGDS preparation

2.1.1. NBG preparation

In this study, NBG with a chemical composition of 10.8% P₂O₅–54.2% SiO₂–35% CaO (mol.%) was synthesized via the sol–gel method using phytic acid as the phosphorus precursor. The synthesis procedure was as follows [34]: Tetraethyl orthosilicate (TEOS), calcium nitrate tetrahydrate, and phytic acid were used as silicon, calcium, and phosphorus precursors, respectively. First, the phytic acid solution was thoroughly mixed with water. TEOS was then added and hydrolyzed for 1 h under stirring. Subsequently, calcium nitrate tetrahydrate was introduced into the mixture and stirred magnetically until a clear and transparent solution was obtained. The solution was aged at room temperature to form a gel, followed by drying at 60 °C for 1 week and further aging at 120 °C for 2 weeks. The dried gel was then calcined at 400 °C for 2 h, ground in a ball mill, and sieved to obtain NBG particles with a size below 74 μm.

2.1.2. Surface amination modification of NBG

Surface amination of NBG was performed using a chemical coupling method. The procedure was as follows: 1 g of NBG particles was added to 100 mL of anhydrous toluene and ultrasonicated for 10 min, followed by magnetic stirring for 1 h to ensure uniform dispersion. Subsequently, 3 mL of 3-aminopropyl triethoxysilane (APTES) was added, and the mixture was heated under reflux in an oil bath for 24 h under nitrogen protection. After the reaction, the suspension was collected, and the powder was recovered by high-speed centrifugation. The obtained powder was washed three times each with anhydrous toluene, anhydrous ethanol, and deionized water, then dried at 60 °C to yield amino-functionalized bioactive glass (NBG-NH₂). The product was stored in a desiccator for further use.

2.1.3. Surface grafting of RGDS sequence

The RGDS peptide was grafted onto the surface of NBG-NH₂ through a chemical conjugation procedure. The modification process was as follows: 3 g of NBG-NH₂ particles were dispersed in 100 mL of 2-(N-morpholino)ethanesulfonic acid buffer (pH 5.5) and magnetically stirred at room temperature until uniformly mixed. Subsequently, 4 mM 1-(3-dimethylaminopropyl)-3-ethylcarbodiimide hydrochloride and 2 mM N-hydroxysuccinimide were sequentially added, and the mixture was stirred for 15 min at room temperature. Then, 300 mg of RGDS peptide was introduced, and the reaction was allowed to proceed under magnetic stirring at 4 °C for 24 h. After completion, the precipitate was collected by high-speed centrifugation, the supernatant was discarded, and the product was washed three times with deionized water. The final NBG@RGDS material was obtained by freeze-drying and stored at -20 °C for subsequent use.

2.2. Material characterization

2.2.1. Physical and chemical properties of NBG@RGDS

The morphological characteristics of NBG and NBG@RGDS were observed using scanning electron microscopy (SEM, SU8010, Hitachi). Characteristic functional groups in NBG and NBG@RGDS were detected by Fourier transform infrared spectroscopy (FTIR). The surface elemental composition was analyzed using energy-dispersive X-ray spectroscopy (EDS, D/max-2500, Rigaku).

Thermogravimetric analysis (TGA) was conducted using a Pyris-1 thermogravimetric analyzer. The measurements were carried out under an air flow rate of 20 mL/min with a heating rate of 5 °C/min to 1000 °C. The grafting efficiency was calculated based on the weight loss difference between NBG and NBG@RGDS.

Nitrogen adsorption-desorption isotherms of NBG and NBG@RGDS were measured using an ASAP 2020 apparatus. The specific surface area was determined by the Brunauer-Emmett-Teller (BET) method.

2.2.2. Immersion experiment in vitro

2 mg/mL NBG or NBG@RGDS was immersed in simulated body fluid (SBF) at 37 °C and 150 rpm. The pH of the extract was measured after 1, 2, 3, 6, 9, 18, 24, 48 and 72 h of immersion. After 3 days, the precipitates were collected by centrifugation, washed three times with Milli-Q water under ultrasonication, and dried in an oven at 60 °C to obtain the SBF-treated particles. The morphological characteristics of NBG and NBG@RGDS after SBF immersion were observed by SEM. To evaluate the long-term mineralization capacity and mineralized layer stability, the same immersion procedure was performed for extended durations. After 3, 7 and 14 days of SBF immersion, the crystalline structures of NBG and NBG@RGDS before and after immersion were analyzed using X-ray diffraction (XRD).

2 mg/mL NBG or NBG@RGDS were immersed in α-minimum essential medium (α-MEM) for 3 days at 37 °C and 150 rpm. After centrifugation, the extracts were obtained by passing through a 0.22 μm filter. The concentrations of Si, Ca and P ions in the extracts were measured using inductively coupled plasma mass spectrometry (ICP-MS).

2.3. Biocompatibility test

2.3.1. Cell culture

THP-1 were purchased from Procell (China) and cultured in RPMI 1640 medium containing 10% fetal bovine serum (FBS) and 1% Penicillin-Streptomycin (P/S). Human BMSCs were purchased from Sciencell (USA) and cultured in Mesenchymal stem cell medium (MSCM). Cells at passage 3-5 were used for the experiments. RAW264.7 cells were purchased from Procell (China) and cultured in α-MEM containing 10% FBS and 1% P/S.

2.3.2. Cell counting kit-8 (CCK-8) assay

THP-1 were seeded in 96-well plates at a density of 6 × 10⁴ cells per well and induced into M0 macrophages with 100 ng/mL Phorbol 12-myristate 13-acetate (PMA) for 24 h. The medium was then replaced with suspension medium containing different concentrations (0.01, 0.1, 1, 2 mg/mL) of NBG or NBG@RGDS. Cell viability was assessed by the CCK-8 assay at 1 and 2 days after seeding, and the absorbance was measured at 450 nm.

BMSCs or RAW264.7 were seeded in 96-well plates at a density of 2 × 10³ cells per well. After 24 h of attachment, the medium was replaced with medium prepared using 2 mg/mL NBG or NBG@RGDS extract. Cell viability was assessed by the CCK-8 assay at 1, 3, 5 and 7 days, and the absorbance was measured at 450 nm.

2.3.3. Live-dead staining

THP-1 were seeded in 24-well plates at a density of 1.3 × 10⁵ cells per well and induced into M0 macrophages with 100 ng/mL PMA for 24 h. The medium was then replaced with suspension medium containing 0.1 mg/mL of NBG or NBG@RGDS. Cell viability was observed by live-dead staining after 1 or 2 days.

2.4. The effect of NBG@RGDS on the polarization of macrophages in vitro

2.4.1. M1 macrophage induction

THP-1 cells were seeded at a density of 8 × 10⁵ cells per well in a six-well plate. PMA at a concentration of 100 ng/mL was used to induce the formation of M0 macrophages. After 24 h, the medium was replaced with an M1 macrophage induction medium containing 1 μg/mL lipopolysaccharide (LPS, Escherichia coli) and 20 ng/mL interferon-gamma (IFN-γ), and the cells were cultured for another 24 h to form M1 macrophages.

2.4.2. Gene expression related to macrophage polarization

After inducing THP-1 to form M0 macrophages, the cells were cultured in M1 macrophage induction medium containing 0.1 mg/mL NBG or NBG@RGDS for 24 h. The cells were lysed using TRIzol and total RNA was extracted. The expression levels of M1 phenotype macrophage markers (IL-1β, IL-6, TNF-α) and M2 phenotype macrophage markers (CD206, CCL22, TGF-β) were assessed by quantitative reverse transcription polymerase chain reaction (qRT-PCR). The forward and reverse primer sequences are listed in Table 1. GAPDH levels served as internal controls.

Table 1.

The primer sequence of all genes used in the qRT-PCR.

Gene	Forward(5′-3′)	Reverse(5′-3′)
GAPDH	GAAGGTGAAGGTCGGAGTC	GAGATGGTGATGGGATTTC
IL-1β	GCCAGTGAAATGATGGCTTATT	AGGAGCACTTCATCTGTTTAGG
IL-6	CACTGGTCTTTTGGAGTTTGAG	GGACTTTTGTACTCATCTGCAC
TNF-α	CAATGGCGTGGAGCTGAGAGATAAC	TTGAAGAGGACCTGGGAGTAGATGAG
CD206	CGCTACTAGGCAATGCCAATG	GCAATCTGCGTACCACTTGTTT
CCL22	TGCCGTGATTACGTCCGTTA	AAGGTTAGCAACACCACGCC
TGF-β	CTGTACATTGACTTCCGCAAG	TGTCCAGGCTCCAAATGTAG
ALP	AGCACTCCCACTTCATCTGGAA	GAGACCCAATAGGTAGTCCACATTG
COL-1	CGAAGACATCCCACCAATCAC	TGTCGCAGACGCAGAT
OPN	ACGCCGACCAAGGAAAACTC	GTCCATAAACCACACTATCACCTCG
RUNX-2	TCACCTCAGGCATGTCCCTCGGTAT	TGGCTTCCATCAGCGTCAACACC
Gapdh	GTGAAGGTCGGTGTGAACGGATT	GGAGATGATGACCCTTTTGGCTC
Mmp9	AGTTTGGTGTCGCGGAGCAC	TACATGAGCGCTTCCGGCAC
β3	GGAAGGCTGGCAGGCATTGTC	GAGGCAGGTGGCATTGAAGGAC
Ctsk	CTCGGCGTTTAATTTGGGAGA	TCGAGAGGGAGGTATTCTGAGT

2.4.3. Measurement of pro-inflammatory cytokine levels by ELISA

After inducing THP-1 to form M0 macrophages, the cells were cultured in M1 macrophage induction medium containing 0.1 mg/mL NBG or NBG@RGDS for 24 h. The supernatant was collected and the protein expression levels were detected using ELISA kits (TNF-α, IL-6).

2.4.4. Western blot analysis of macrophage polarization markers and PI3K-Akt signaling pathway

After inducing THP-1 cells to form M0 macrophages, the cells were treated with 0.1 mg/mL NBG or NBG@RGDS in M1 induction medium for 24 h. Cells were lysed in radioimmunoprecipitation assay (RIPA) buffer (Beyotime, China) with protease and phosphatase inhibitors. Equal amounts of protein were subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), transferred to polyvinylidene fluoride (PVDF) membranes, and blocked. Membranes were probed with primary antibodies against CD86, CD206, PI3K, p-PI3K, Akt1, p-Akt1 and GAPDH, followed by horseradish peroxidase (HRP)-conjugated secondary antibodies. Protein bands were visualized by enhanced chemiluminescence (ECL) and quantified using ImageJ.

2.4.5. Macrophage polarization by flow cytometry (FCM) analysis

After inducing THP-1 to form M0 macrophages, the cells were cultured in M1 macrophage induction medium containing 0.1 mg/mL NBG or NBG@RGDS for 24 h. Cells were collected and double-stained for CD86 and CD206 using specific primary antibodies and corresponding secondary antibodies (Alexa Fluor® 488 for CD86 and Alexa Fluor® 647 for CD206). Permeabilization was applied for CD206 staining. The cell polarization state was analyzed using the flow cytometer (Agilent Novocyte, USA) and NovoExpress (version 1.5.0, USA) software.

2.4.6. Transcriptome analysis

The experiment comprised three groups. After induction into M0 macrophages, THP-1 were further cultured for 24 h in M1 macrophage induction medium supplemented with either 0.1 mg/mL NBG (M1+NBG group), 0.1 mg/mL NBG@RGDS (M1+NBG@RGDS group), or no bioactive glass (M1 group). Total RNA was extracted using TRIzol reagent. Samples that passed quality control were subjected to transcriptome sequencing, which was performed by Shanghai OE Biotech Co., Ltd. Differential expression analysis was conducted, followed by Gene Ontology (GO) analysis, trend analysis, Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis, and Gene Set Enrichment Analysis (GSEA). Gene expression heatmaps, along with KEGG and GO enrichment plots, were generated to visualize the results.

2.5. The effect of NBG@RGDS on osteoblastogenesis in vitro

2.5.1. Cell culture and osteogenic differentiation of BMSCs

BMSCs were seeded into 12-well plates at a density of 4 × 10⁵ cells per well. After 24 h of incubation for cell attachment, the medium was replaced with osteogenic induction medium(OM). The experimental groups were cultured in OM prepared using the extract of NBG or NBG@RGDS. The OM consisted of α-MEM, 10% FBS, 1% P/S, 10 mM β-glycerophosphate, 100 μg/mL ascorbic acid, and 10 nM dexamethasone. The medium was changed every 2 days.

2.5.2. ALP staining and quantitative analysis

After 7 days of osteogenic induction, BMSCs were stained using a BCIP/NBT alkaline phosphatase color development kit according to the manufacturer's instructions. After observing the coloration results under the microscope, the stained samples were de-colored using DMSO, and the absorbance at 620 nm was measured as the quantitative indicator.

2.5.3. Alizarin Red S (ARS) staining and quantitative analysis

After 14 or 21 days of osteogenic induction, BMSCs were fixed with 4% paraformaldehyde and stained with 1% Alizarin Red S (pH 4.2). The staining results were observed under a microscope. Subsequently, the stained samples were decolorized with 10% cetylpyridinium chloride, and the absorbance at 562 nm was measured as the quantitative indicator.

2.5.4. Gene expression related to osteoblastogenesis

After 7 days of osteogenic induction in BMSCs, the mRNA expression levels of OPN, ALP, RUNX-2, and COL-1 were detected using qRT-PCR. The method was consistent with section 2.4.2. The forward and reverse primer sequences are listed in Table 1. GAPDH levels served as internal controls.

2.6. The effect of NBG@RGDS on osteoclastogenesis in vitro

2.6.1. Cell culture and differentiation of osteoclasts

RAW264.7 were seeded at a density of 3 × 10⁴ cells per well in a 12-well plate. After 4 h of culture for cell attachment, the medium was replaced with osteoclast induction medium containing 100 ng/mL Receptor Activator of Nuclear Factor Kappa-B Ligand (RANKL). The experimental groups were cultured in osteoclast induction medium prepared using the extract of NBG or NBG@RGDS. The medium was changed every 2 days. On the fourth day, the formation of osteoclasts can be observed.

2.6.2. Tartrate-resistant acid phosphatase (TRAP) staining

After 4 days of osteoclast induction, cells were fixed with 4% paraformaldehyde and stained using a TRAP staining kit at 37 °C for 3 h. The formation of osteoclasts was observed under a microscope.

2.6.3. F-actin ring staining

After 4 days of osteoclast induction, cells were fixed with 4% paraformaldehyde, permeabilized with 0.1% Triton X-100, and stained with Actin-Tracker Green-488 for 30 min. Subsequently, the nuclei were labeled with DAPI, and the formation of F-actin rings was observed under a fluorescence microscope.

2.6.4. Gene expression associated with osteoclastogenesis

After 4 days of osteoclast induction, the mRNA expression levels of osteoclast marker genes (β3, Mmp9, Ctsk) in cells were detected using qRT-PCR. The method was consistent with section 2.4.2. The forward and reverse primer sequences are listed in Table 1. Gapdh levels served as internal controls.

2.7. The therapeutic effect of NBG@RGDS in vivo

2.7.1. Fabrication of animal models

To evaluate the therapeutic efficacy of NBG@RGDS as a bone graft material, a femoral condyle defect model was established in Sprague-Dawley (SD) rats. The experimental protocol was approved by XXXXXXXX. Eight-week-old male SD rats weighing 250–300 g, provided by Spafu Company, were used in this study. A total of 5 bone defect replicates were set for each group (n = 5). Each rat received different material implants in the bilateral femoral condyle defects to exclude individual differences. The experimental groups included blank control, NBG, and NBG@RGDS. After anesthesia via intraperitoneal injection of pentobarbital sodium solution, a longitudinal incision was made on the lateral side of both knee joints. The skin and fascial layer were incised to expose the lateral femoral condyle. Following periosteal stripping, a cylindrical bone defect with a diameter of 3 mm and depth of 3 mm was created using a trephine. The defect site was rinsed with saline, after which NBG or NBG@RGDS was implanted into the defect area. In the blank control group, the defect was filled with blood. Finally, the muscle, fascia, and skin were sutured layer by layer. All animals were allowed free access to food and water postoperatively.

2.7.2. Determination of biosafety in vivo

We evaluated the in vivo biosafety of NBG@RGDS through histopathological analysis of the major organs. Hematoxylin-Eosin (H&E) staining was performed on the hearts, livers, spleens, lungs and kidneys of each experimental group of rats to observe the tissue morphology and structure.

2.7.3. Micro-CT measurement

The harvested rat femurs at 4 weeks and 9 weeks post-surgery were scanned using micro-CT (SkyScan1276, Bruker, Germany) with a 10 μm resolution. Following the acquisition of three-dimensional images, CTvox (version3.3.0, Bruker, Germany), CTAn (version1.20.8.0, Bruker, Germany), CTvol (version2.3.2.0, Bruker, Germany), and DataViewer (version1.7.0.1, Bruker, Germany) software were used for image format conversion, three-dimensional reconstruction analysis of the region of interest (ROI), and subsequent measurement of morphometric parameters based on the reconstructed images.

2.7.4. Histological analysis

The rats were sacrificed at 3 days, 2 weeks and 9 weeks after surgery. The femurs were isolated and fixed, and then the samples were decalcified using ethylenediaminetetraacetic Acid (EDTA). After paraffin embedding, the sections were cut along the long axis of the vertical cylindrical bone defect. The sections were stained with H&E staining, Masson's trichrome staining and TRAP staining. Quantitative analysis was performed using ImageJ (version 2.16.0) software to determine the number of osteoclasts, and evaluate the area of new bone formation.

2.7.5. Immunofluorescence (IF) staining

The expression levels of CD86 and CD206 were assessed by IF staining. Briefly, sections were blocked with goat serum, followed by incubation with primary antibodies overnight at 4 °C. Sections were further incubated with the corresponding secondary antibodies and stained with 4′,6-diamidino-2-phenylindole (DAPI). The bone tissue at the edge of the defect was defined as the ROI for evaluating the positive expression of CD86 and CD206.

2.7.6. Immunohistochemical (IHC) staining

IHC staining was performed to evaluate the expression of IL-1β (3 days) and RUNX-2 and OCN (2 and 9 weeks) following NBG@RGDS implantation. After deparaffinization, rehydration, and antigen retrieval with 0.1% trypsin, sections were treated with H₂O₂ to block endogenous peroxidase activity and then blocked with goat serum. The sections were then incubated with primary antibodies at 4 °C overnight, followed by incubation with a corresponding secondary antibody. Color development was performed using 3,3′-diaminobenzidine (DAB), and the nuclei were counterstained with hematoxylin. Stained sections were examined under a microscope.

2.8. Statistical analysis

Data were analyzed using SPSS software (version 24.0). All experiments were performed with three independent replicates (n = 3), and data are presented as the mean ± standard deviation. Intergroup comparisons were evaluated by one-way analysis of variance (ANOVA), with the Bonferroni correction applied for multiple comparisons. Significance was set at p < 0.05 and denoted as follows: ns, p ≥ 0.05; *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001.

3. Results and discussion

3.1. Design and fabrication of NBG@RGDS

Traditional bioactive glasses, despite their favorable bioactivity and osteoinductive potential, often induce a significant local pH increase during degradation [35], which may adversely affect surrounding cells and tissues [36]. More importantly, they generally lack the ability to actively modulate the osteoimmune microenvironment and are particularly ineffective in directly regulating osteoclast activity.

To address these limitations, we engineered a multi-functional system based on NBG. Our design strategy utilizes NBG as a stable substrate to maintain a physiological pH environment during degradation [33]. To functionalize the surface, we grafted APTES onto the NBG surface through a high-temperature condensation reaction to generate aminated NBG (NBG-NH₂), followed by the covalent immobilization of the multifunctional RGDS peptide. In this integrated NBG@RGDS system, the NBG core serves as an osteoconductive scaffold, while stabilizing the local microenvironment, whereas the surface-grafted RGDS peptide exerts a triple regulatory function: guiding macrophage polarization toward a pro-repair M2 phenotype [37,38], inhibiting osteoclast activity [39,40] and enhancing biomimetic mineralization [41,42]. This integrated design enables NBG@RGDS to simultaneously and actively coordinate immune responses and bone metabolic balance, offering a multi-functional, single-material solution for the healing of bone defects.

3.2. Characterization of NBG@RGDS

Material characterization results confirmed the successful synthesis and surface functionalization of the composite. Morphological observations via SEM showed that both unmodified NBG and NBG@RGDS consisted of irregular particles, with RGDS grafting causing no significant alteration in physical morphology (Fig. 1A). FTIR further verified successful grafting (Fig. 1B), both NBG and NBG@RGDS exhibited characteristic asymmetric stretching vibration peaks of Si–O–Si in the range of 1000-1100 cm⁻¹, along with an O–H vibrational absorption peak at 3386 cm⁻¹. After RGDS grafting, NBG@RGDS displayed a vibration peak near 1130 cm⁻¹ attributed to the C-N bond, as well as an additional amide II band absorption peak at 1649 cm⁻¹ originating from N–H bending vibrations. EDS of C, N, and Si elements further revealed changes in surface composition (Fig. 1C). Following amination and RGDS grafting, the mass fractions of C and N in NBG@RGDS increased significantly due to the carbon and nitrogen rich peptide chains, whereas the Si content remained largely unchanged. This elemental distribution shift indirectly confirmed the successful grafting of RGDS onto the NBG surface. TGA provided quantitative evidence to confirm the grafting process. The weight loss curve of NBG and NBG@RGDS showed that upon heating to 1000 °C, the mass decreased by 6.1% and 19.5% compared with their initial masses, respectively. The weight loss difference was attributed to the grafted polypeptides and the coupling agent (Fig. 1D). The grafting efficiency was reproducible across different batches by strictly controlling the reaction time during material preparation. Despite the changes in surface elements and chemical groups, no significant difference in specific surface area was observed between NBG and NBG@RGDS (Fig. 1E and F, Table S1). Together, the morphological, spectroscopic, and elemental analysis results demonstrate the successful modification of NBG surfaces with the functional RGDS peptide.

Fig. 1.

Characterization of NBG@RGDS. A) SEM images of NBG, NBG@RGDS, and their respective morphologies after immersion in SBF. B) Characterization of functional groups in NBG and NBG@RGDS by FTIR. C) Characterization of surface elemental composition of NBG and NBG@RGDS by EDS. D) TGA of NBG and NBG@RGDS; E) Specific surface area of NBG; F) Specific surface area of NBG@RGDS; G) The pH variation pattern of the extract solutions of NBG and NBG@RGDS over time. H) XRD pattern of the crystal structure of NBG and NBG@RGDS before and after immersion in SBF for 3 days. I) Release profiles of Si, Ca and P ions in extracts of NBG or NBG@RGDS. Data are presented as mean ± SD (n = 3). Statistical significance was determined by one-way ANOVA followed by Bonferroni post hoc test. ns, p ≥ 0.05; *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001.

Performance tests demonstrated that NBG@RGDS retains the critical bioactive advantages of the NBG substrate. When incubated in SBF, the extract maintained a stable pH within the physiological neutral range (∼7.5) (Fig. 1G). Moreover, after 3 days of immersion, a large number of lamellar aggregates formed on its surface (Fig. 1A), exhibiting the typical morphological characteristics of hydroxyapatite. This result highlights the rapid mineralization capacity of the material, as a mineralized layer formed within this short period. To evaluate long-term mineralization capacity and mineralized layer stability, the SBF immersion was extended to 7 and 14 days. XRD patterns further confirmed that both NBG and NBG@RGDS displayed characteristic diffraction peaks of hydroxyapatite at 25.9°, 31.7°, 39.7°, 46.6°, 49.4° and 53.2° after 3, 7 and 14 days of immersion (Fig. 1H, Fig. S1), indicating that the deposited surface crystals were hydroxyapatite and demonstrating favorable in vitro biomineralization activity. Additionally, the release rates of Si, Ca, and P ions from NBG@RGDS in α-MEM were elevated compared to NBG (Fig. 1I). These results suggest that RGDS grafting not only preserves the pH-stabilizing and mineralization properties of NBG, but also enhances ion exchange kinetics, which is beneficial for promoting local cellular activity and inducing bone tissue regeneration [43,44].

3.3. NBG@RGDS promotes the polarization of M1 macrophages toward the M2 phenotype

The regulation of macrophages is central to the immunomodulatory function of NBG@RGDS [18,19]. We first evaluated the biocompatibility of the material with macrophages. CCK-8 results showed that neither NBG nor NBG@RGDS at concentrations of 0.1 mg/mL or lower exhibited cytotoxicity (Fig. 2A). Live-dead staining further confirmed that 0.1 mg/mL of NBG or NBG@RGDS did not affect the viability of M0 macrophages (Fig. 2B and Fig. S2). Therefore, 0.1 mg/mL was selected as the working concentration for subsequent macrophage experiments. Notably, at higher concentrations, NBG particles tended to form a dense physical barrier on the culture dish surface, which restricted cell access to the substrate and hindered adhesion. This issue was effectively addressed by covalent grafting of RGDS, which provides specific RGD motifs that act as ligands for integrin receptors, thereby transforming the particles into adhesion anchors (Fig. 2A). This observation aligns with the well-established function of the RGD sequence as an integrin ligand that promotes cell adhesion and spreading [45,46]. Furthermore, as previously described, this modification also significantly altered the ion release profile, thereby optimizing the local microenvironment for cell survival (Fig. 1I). These synergistic factors, combining biochemical recognition with an improved ionic microenvironment, collectively enhanced interfacial biocompatibility and cellular metabolic activity.

Fig. 2.

In vitro modulation of macrophage polarization from M1 to M2 phenotype by NBG@RGDS. A) Cell viability of M0 macrophages cultured with different concentrations of NBG or NBG@RGDS measured by the CCK-8 kit. B) Live-dead staining of M0 macrophages cultured with suspension medium for 1 and 2 days. C-H) Relative mRNA expression levels of M1 macrophage differentiation-related markers (IL-6, IL-1β, TNF-α) (C-E), and M2 macrophage differentiation-related markers (CD206, CCL-22, TGF-β) (F-H) at 24h. These data are relative to the average level of GAPDH and normalized to the expression level in the control group. I-J) ELISA for detecting the protein expression levels of TNF-α and IL-6 in the supernatant of macrophages. K-M) Representative images of western blot and quantitative analyses of CD86 and CD206. N-P) FCM for detecting the expression of CD86 and CD206 on macrophage surfaces. Data are presented as mean ± SD (n = 3). Statistical significance was determined by one-way ANOVA followed by Bonferroni post hoc test. ns, p ≥ 0.05; *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001.

Following confirmation of biosafety, we further investigated the regulatory capacity of NBG@RGDS on macrophage polarization. In LPS and IFN-γ-induced M1 polarization model, NBG@RGDS exhibited stronger regulatory effects compared to NBG. qRT-PCR results showed that both NBG and NBG@RGDS significantly suppressed the gene expression of pro-inflammatory M1 markers (IL-6, IL-1β, and TNF-α), with NBG@RGDS demonstrating more pronounced inhibition (Fig. 2C–E). Concurrently, both materials significantly upregulated anti-inflammatory M2 markers (CD206, CCL-22, and TGF-β), with NBG@RGDS showing a more prominent effect on CD206 and CCL-22 (Fig. 2F–H). Consistent with the gene expression, ELISA results further confirmed that NBG@RGDS substantially reduced TNF-α and IL-6 protein secretion (Fig. 2I and J). Western blot analysis provided additional protein-level evidence, revealing that NBG@RGDS more effectively downregulated CD86 and upregulated CD206 compared to NBG (Fig. 2K–M). Flow cytometry analysis further corroborated these observations, showing that both NBG and NBG@RGDS significantly decreased the percentage of CD86+CD206-cells while increasing CD86⁻CD206+ cells, with the NBG@RGDS group exhibiting a more potent effect (Fig. 2N–P).

During the early phase of bone defect healing, persistent M1 macrophages infiltration can lead to chronic inflammation, thereby prolonging the initial inflammatory response and inhibiting osteoblast activity [15]. Our findings indicate that RGDS grafting significantly enhances the ability of NBG to regulate macrophage phenotypes, not only more effectively suppressing M1 polarization but also further promoting their transition toward the M2 phenotype. M2 macrophages, in turn, secrete various bioactive factors (e.g., TGF-β, osteoprotegerin, BMP-2) that induce extracellular matrix deposition and promote osteogenesis [47,48]. In summary, by driving macrophage polarization from M1 to M2, NBG@RGDS reshapes the local osteoimmune microenvironment, thereby creating favorable conditions for bone regeneration.

3.4. Transcriptomic analysis reveals the molecular mechanism of NBG@RGDS-mediated immunomodulation

To further elucidate the regulatory mechanism underlying NBG@RGDS-mediated macrophage regulation, we performed RNA-seq analysis on M1-polarized macrophages treated with NBG or NBG@RGDS, utilizing untreated M1 macrophage as a control. Compared with the M1 group, differentiation expression analysis identified 546 differentially expressed genes in the M1+NBG group and 459 in the M1+NBG@RGDS group (Fig. 3A). The results indicate that, compared with the M1 group, both the M1+NBG and M1+NBG@RGDS groups showed downregulation of selected pro-inflammatory genes (Fig. 3B). Trend analysis was conducted to examine the dynamic changes in gene expression in the order of the M1, M1+NBG and M1+NBG@RGDS groups. The results showed that, compared with the M1+NBG group, the M1+NBG@RGDS group more significantly upregulated biological processes such as cell-cell adhesion and regulation of substrate adhesion-dependent cell spreading (Fig. 3C and D). These changes align closely with the known biological functions of the RGD sequence [45,46], indicating that RGDS grafting significantly enhances cell adhesion and chemotactic behavior. Cell adhesion serves as a critical initial step for cell proliferation, migration, and differentiation [49]. The strong adhesive properties conferred by RGDS lay an important foundation for subsequent cellular regulation and tissue regeneration.

Fig. 3.

Transcriptomic analysis of the effects of NBG and NBG@RGDS on M1 macrophages. A) Venn diagram of pairwise comparisons among the three sample groups. B) Heatmap analysis of differentially expressed genes among the three sample groups. C-D) Trend analysis among the three sample groups. E-F) GO enrichment analysis comparing the M1+NBG@RGDS group with the M1 group, showing upregulated (E) and downregulated (F) terms. G) GSEA analysis reveals positive regulation of the PI3K-Akt signaling pathway. H-J) Representative images of western blot and quantitative analyses of p-PI3K, PI3K, p-Akt1 and Akt1. K) GSEA analysis reveals positive regulation of the BMP signaling pathway in the M1+NBG@RGDS group compared to the M1 group. Data are presented as mean ± SD (n = 3). Statistical significance was determined by one-way ANOVA followed by Bonferroni post hoc test. ns, p ≥ 0.05; *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001.

GO and KEGG enrichment analyses further suggested the possible pathway mechanisms underlying NBG@RGDS-mediated regulation of macrophage polarization. GO analysis showed that NBG@RGDS treatment significantly upregulated genes involved in tissue-repair processes such as cell adhesion (Fig. 3E), while downregulating inflammatory response genes (Fig. 3F). KEGG pathway enrichment analysis revealed that differentially expressed genes in the NBG@RGDS group were enriched in multiple signaling pathways associated with macrophage polarization (Fig. S3). Consistently, GSEA confirmed that the PI3K-Akt signaling pathway was significantly upregulated (Fig. 3G). The PI3K-Akt pathway support M2 polarization thereby enhancing cell survival, proliferation, and tissue-repair gene expression [50,51]. To validate these transcriptomic findings at the protein level, western blot analysis was performed to assess the expression and phosphorylation levels of key proteins in the PI3K-Akt pathway. The results showed that the levels of p-PI3K and p-Akt1 were significantly increased in the NBG@RGDS group, confirming activation of this pathway (Fig. 3H–J). These findings suggest that this pathway may mediate the regulatory effect of NBG@RGDS on macrophage polarization. Direct evidence for the binding between the RGDS peptide and integrin receptors on macrophages is not provided in the current study, which constitutes a limitation. We recognize that further validation using integrin blocking assays, specific pathway inhibitors or gene knockdown experiments would provide stronger evidence, and this will be an important direction for our future research.

Notably, GO analysis indicated that NBG@RGDS significantly upregulates genes involved in the positive regulation of osteoblast proliferation (Fig. 3E). Furthermore, GSEA results revealed a marked upregulation of the BMP signaling pathway in the M1+NBG@RGDS group (Fig. 3K and S4). The BMP pathway plays a central role in skeletal development, formation, and repair [52]. Activated BMP signaling amplifies the secretion of multiple growth factors from macrophages, thereby activating osteoblast differentiation pathways and accelerating bone defect healing [48]. In summary, these findings suggest that NBG@RGDS not only induces macrophage polarization from M1 to M2 phenotype through coordinated regulation of signaling networks, but may also influence osteoblast proliferation and differentiation by modulating macrophages, thereby promoting bone regeneration.

3.5. NBG@RGDS simultaneously orchestrates osteogenesis and suppresses osteoclastogenesis

To evaluate the regulatory effect of NBG@RGDS on the balance between osteogenesis and osteoclastogenesis, we designed separate experiments to evaluate its impact on RAW264.7-induced osteoclast formation and on the osteogenic differentiation of BMSCs.

First, CCK-8 assay demonstrated that RAW264.7 cell viability was not compromised by treatment with either 0.1 mg/mL NBG or NBG@RGDS (Fig. S5). RANKL was employed to induce the differentiation of RAW264.7 into osteoclasts, intervening with NBG or NBG@RGDS extracts. TRAP staining results showed that both the control and NBG groups exhibited numerous of large, multinucleated cells characteristic of mature osteoclasts, whereas NBG@RGDS treatment significantly reduced both the number and size of TRAP-positive multinucleated osteoclasts (Fig. 4A and B). This indicates that NBG alone has a limited anti-resorptive capability, highlighting the critical role of RGDS modification. Further observation of the F-actin ring—a key structure for osteoclast function [53]—revealed that control and NBG groups formed complete and distinct F-actin rings after 4 days of RANKL induction (Fig. 4C). In contrast, NBG@RGDS treatment disrupted these structures, significantly reduced both the size and number of F-actin rings (Fig. 4C and D). At the molecular level, qRT-PCR confirmed that NBG@RGDS significantly downregulated the mRNA levels of osteoclast maturation markers (Mmp9, β3, and Ctsk) compared with the control and NBG groups (Fig. 4E–G). These results collectively demonstrate that the introduction of RGDS endows NBG with a novel, active function of inhibiting osteoclast formation. The underlying mechanism lies in the RGD motif, which competitively blocks the α_vβ₃ integrin receptor essential for osteoclast function [53,54]. This aligns with studies showing RGD-containing peptides [55,56] inhibit osteoclast differentiation by binding to α_vβ₃, validating our surface engineering.

Fig. 4.

NBG@RGDS bidirectionally regulates osteogenic and osteoclastic metabolism. A-B) TRAP staining of osteoclasts and quantitative analysis. C-D) F-actin ring staining of osteoclasts and quantitative analysis. E-G) qRT-PCR analysis showing the gene expression levels of Mmp9, β₃ and Ctsk. H-I) ALP staining and quantitative analysis of BMSCs under different treatments. J-K) ARS staining and quantitative analysis of BMSCs under different treatments. L-O) qRT-PCR analysis showing the gene expression levels of ALP, OPN, COL-1 and RUNX-2 in BMSCs. Data are presented as mean ± SD (n = 3). Statistical significance was determined by one-way ANOVA followed by Bonferroni post hoc test. ns, p ≥ 0.05; *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001.

On the other side of the balance, to evaluate the regulatory effect of NBG@RGDS on osteogenesis, its extract was used to culture BMSCs. CCK-8 assays demonstrated favorable biocompatibility of both NBG and NBG@RGDS toward BMSCs (Fig. S6). Further osteogenic induction experiments were conducted to analyze their influence on cell differentiation. Staining for ALP (an early osteogenic differentiation marker) revealed stronger expression in the NBG@RGDS group compared to the NBG group at day 7 (Fig. 4H), a trend confirmed by quantitative analysis (Fig. 4I). Late-stage mineralization capacity was assessed by ARS staining at 14 and 21 days. Under osteogenic induction medium, both NBG and NBG@RGDS promoted the formation of mineralized nodules, with the most significant mineralization observed in the NBG@RGDS group (Fig. 4J and K). Notably, even in the absence of exogenous mineralization supplements, NBG@RGDS group still induced mineral deposition, whereas NBG did not (Fig. 4J and K). Together, the ALP and ARS staining results indicate that NBG@RGDS possesses superior mineralization-inducing ability. Further qRT-PCR analysis of osteogenesis-related gene expression elucidated its regulatory role. The results showed that both NBG and NBG@RGDS significantly upregulated the expression levels of key osteogenic markers including ALP, OPN, COL-1, and RUNX2, with NBG@RGDS exerting a stronger promoting effect on ALP and OPN than NBG (Fig. 4L–O). The inherent release of Ca and Si ions from NBG provides basal osteogenic signals [35], while the enhanced effect of NBG@RGDS arises from two aspects. First, grafting of RGDS on the NBG surface alters the ion release kinetics, as demonstrated in the preceding sections, leading to an enhanced capacity to induce osteogenic differentiation. Second, the serine residue at the C-terminus of RGDS may serve as a potential phosphorylation site [41], enabling Ca ion binding and guiding hydroxyapatite deposition, thereby promoting the formation of well-ordered hydroxyapatite crystals [57]. This mimics the key role of non-collagenous proteins in biomineralization and directly strengthens the mineralization-inducing capacity.

In summary, the incorporation of the RGDS peptide endows the material with the dual capability to synergistically regulate osteogenesis and osteoclastogenesis. This study employs an elegant design in which the same RGDS sequence delivers distinct biological signals to BMSCs and osteoclast precursors. The cell-type-specific responses allows NBG@RGDS to directly and simultaneously intervene on both sides of bone metabolism. Consequently, this study successfully transforms NBG, which originally possessed only unidirectional osteogenic activity, into an intelligent material capable of actively and bidirectionally regulating bone metabolism. This achievement enables the coordinated management of the osteogenesis–osteolysis balance and provides a key material-based solution for repairing complex bone-defect microenvironments.

3.6. NBG@RGDS accelerates early-stage bone healing through synergistic immunomodulation and bone metabolism in vivo

To evaluate the in vivo bone defect repair efficacy of NBG@RGDS, we established a cylindrical bone defect model (3 mm in diameter and 3 mm in depth) in the lateral femoral condyle of rat. Defects were implanted with either NBG or NBG@RGDS, with untreated defects serving as a blank control group. Post-operative evaluation was conducted at 3 days, 2 weeks, 4 weeks and 9 weeks. This time series enabled us to track callus formation, monitor remodeling processes and delineate the early healing pattern of NBG@RGDS. In addition, inflammatory infiltration, osteoclast aggregation, as well as overall bone repair progression were assessed.

At 3 days post-implantation, the immune microenvironment was assessed. H&E staining revealed well-defined bone defect margins with inflammatory cell infiltration and fibrous tissue ingrowth across all groups (Fig. S7). No signs of suppuration, necrosis, or infection was detected. In the material-implanted groups, voids resulting from the detachment of bioactive glass during sectioning were visible (Fig. S7). IF staining analysis demonstrated that NBG@RGDS significantly reduced the proportion of CD86⁺ M1 macrophages while increasing the proportion of CD206⁺ M2 macrophages compared with the blank control and NBG groups (Fig. 5A–G and H). This indicates effective polarization of macrophages from a pro-inflammatory toward a pro-repair phenotype. IHC staining and quantitative analysis showed that IL-1β were mainly expressed in immune cells surrounding the defect margins and the adjacent extracellular matrix (Fig. 5B), with the NBG@RGDS group exhibiting the lowest expression levels (Fig. 5I), further confirming its ability to suppress excessive early-stage inflammatory responses. These results indicate that NBG@RGDS effectively polarizes macrophages from the pro-inflammatory M1 phenotype toward the pro-regenerative M2 phenotype, thereby modulating the secretion of inflammatory cytokines and remodeling the local immune microenvironment. TRAP staining showed abundant osteoclast aggregation at the defect margins in the control group, in contrast to a marked decrease in osteoclast numbers in the NBG@RGDS group (Fig. 5C and J). These results indicate that NBG@RGDS not only suppresses the local inflammatory response but also inhibits osteoclast activation and aggregation during the early implantation phase, thereby creating favorable conditions for the rapid initiation of bone repair in the early stage.

Fig. 5.

Evaluation of in vivo immunomodulation, osteoclastic regulation and osteogenic performance of NBG@RGDS implantation in femoral condyle defects. A,G,H) IF images (A) and corresponding quantitative analysis (G,H) of CD86 and CD206 expression at the edge of bone defects under different treatment conditions at 3 days post-operation, revealing the polarization state of macrophages in vivo. B,I) IHC staining of IL-1β and corresponding quantitative analysis. C, J) TRAP staining of osteoclasts and corresponding quantitative analysis at the defect margin. D,K) Masson's trichrome staining showing newly formed bone (D) and corresponding quantitative analysis (K) for each group at 2 weeks post-operation. E) H&E staining of each group at 2 weeks post-operation. F,L) IHC staining images of OCN (F) and corresponding quantitative analysis (L) for each group. NB: new bone, F: fibrous tissue. The dotted line indicates the original boundary of the bone defect. Data are presented as mean ± SD (n = 5). Statistical significance was determined by one-way ANOVA followed by Bonferroni post hoc test. ns, p ≥ 0.05; *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001.

At 2 weeks post-implantation, the advantage of proactively orchestrating the early immune microenvironment became evident in bone formation. H&E and Masson's trichrome staining revealed limited new bone formation at the defect margins in the blank control group, whereas the NBG@RGDS group exhibited significant bone ingrowth extending from the periphery toward the center (Fig. 5D, E and K), demonstrating substantially accelerated osteogenesis compared to other groups. IHC analysis of osteogenic markers OCN further confirmed the strongest expression of bone-related proteins in the NBG@RGDS group (Fig. 5F and L), indicating its efficacy in initiating and accelerating the early osteogenic differentiation program. These findings demonstrate that NBG@RGDS, by simultaneously regulating macrophage polarization and inhibiting osteoclast aggregation, rapidly suppresses inflammation and bone resorption while activating osteogenic pathways. This coordinated multi-cellular regulation ensures a rapid transition from the inflammatory phase to the regenerative phase. Such multi-cellular regulatory capacity highlights the promise of NBG@RGDS in facilitating the early functional recovery of bone defects.

Micro-CT was employed to evaluate the long-term repair outcomes. At 4 weeks post-implantation, slightly hyperdense images of undegraded material were observed within the defect area (Fig. 6A). Quantitative analysis revealed significantly higher BV/TV and BMD in both NBG and NBG@RGDS groups compared to the control group (Fig. 6F and G). Notably, the NBG@RGDS group exhibited a more pronounced increase than the NBG group, along with significantly enhanced Tb. Th and Tb. N, indicating its superior capacity to accelerate bone regeneration (Fig. 6H and I). At 9 weeks post-implantation (Fig. 6B, Fig. S8), both material-implanted groups continued to show elevated BV/TV and BMD relative to the blank control (Fig. S9), confirming the osteogenic potential of the bioactive glass. However, no significant differences in Tb.Th and Tb.N were observed between the material-implanted groups and the control group (Fig. S9), suggesting that while the materials increased bone volume, they did not alter the microstructure of the newly formed trabecular bone, reflecting good osteoconductivity. Interestingly, at this terminal time point, no statistically significant differences were observed between NBG and NBG@RGDS groups. The results from H&E and Masson's trichrome staining further confirmed this finding (Fig. 6C, Fig. S10). IHC analysis further demonstrated prominent OCN expression at the osteogenic fronts in both NBG and NBG@RGDS groups (Fig. 6D and J), with notably enhanced RUNX-2 expression specifically in the NBG@RGDS group (Fig. 6E and K).

Fig. 6.

Promotion of in vivo bone defects regeneration with NBG@RGDS. A,F-I) Micro-CT images and 3D reconstruction images of the original bone defect area of the femoral condyles in each group at 4 weeks post-operation (A). The arrows indicate undegraded material. Quantitative analysis of BV/TV (F), BMD (G), Tb.Th (H) and Tb.N (I), reflecting bone regeneration effects under different treatments. B) 3D reconstruction images of the original bone defect area at 9 weeks post-operation. C) Masson's trichrome staining images of femoral condyles. D,E,J,K) IHC staining images of OCN (D) and RUNX-2 (E) with corresponding quantitative analysis (J,K) for each group at 9 weeks post-operation. Data are presented as mean ± SD (n = 5). Statistical significance was determined by one-way ANOVA followed by Bonferroni post hoc test. ns, p ≥ 0.05; *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001.

Through this longitudinal observation across multiple time points, the results precisely highlight the strategic advantages of NBG@RGDS as an accelerator of the healing process. It rapidly initiates bone repair at an early stage, exhibiting significantly enhanced new bone formation at 2 and 4 weeks. While the inherent bioactivity of NBG allows it to eventually achieve comparable bone volume, RGDS grafting is critical for shortening the vulnerable early healing window. In clinical scenarios, achieving rapid bony bridging is often more critical than marginal gains in final bone volume, as it significantly reduces the risks of implant loosening, infection, and non-union [58,59]. By actively remodeling the immune microenvironment and balancing bone metabolism at an early stage, NBG@RGDS achieves an efficient initiation-enhanced repair strategy. Besides histological analysis of major organs in rats revealed no pathological abnormalities, indicating favorable biocompatibility of both NBG and NBG@RGDS (Fig. S11).

4. Conclusions

This study demonstrates that surface-grafting RGDS peptide onto pH-neutral bioactive glass establishes a material-based platform enabling integrated, multi-target regulation of the osteoimmune microenvironment. The developed NBG@RGDS not only preserves the inherent osteoinductive capacity of the NBG substrate but also leverages the RGDS motif to orchestrate three key biological processes: proactively modulating macrophage polarization from the pro-inflammatory M1 phenotype toward the pro-repair M2 phenotype; effectively inhibiting osteoclastogenesis and excessive bone resorption; and synergistically enhancing the osteogenic differentiation and mineralization of stem cells. In vivo, this synergistic mechanism successfully remodeled the early repair microenvironment, effectively suppressing excessive inflammation and osteoclast activation while significantly accelerating the onset of new bone formation, thereby laying the foundation for high-quality bone regeneration. This work suggests a shift in design strategy for bone repair material design: shifting from the traditional passive support or unidirectional bone formation to an active management of early immune responses and intercellular crosstalk, thereby systematically initiating and coordinating the entire regeneration process. This strategy provides a theoretical foundation and design direction for developing next-generation bone repair materials with immunomodulatory functions.

Ethics approval statement

This study was approved by the Laboratory Animal Welfare and Ethics Subcommittee of the Peking University Biomedical Ethics Committee (PKUIRB-LA2022051).

CRediT authorship contribution statement

Xueying Li: Formal analysis, Investigation, Methodology, Writing – original draft. Jilin Wu: Investigation, Methodology. Jingyi Li: Investigation, Validation. Guibin Huang: Formal analysis, Investigation. Peipei Jia: Investigation. Yanmei Dong: Conceptualization, Funding acquisition, Supervision, Writing – review & editing.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Appendix A. Supplementary data

The following is the Supplementary data to this article:

Data availability

Data will be made available on request.