Multifunctional peptide nanofiber coatings enhance bone regeneration on xenograft materials

Wait 5 sec.

IntroductionBone regeneration is a cornerstone of implant-based dental rehabilitation, especially in cases of advanced alveolar bone resorption due to premature tooth loss, periodontal disease, odontogenic cysts, trauma, or sinus pneumatization. These clinical scenarios often necessitate bone grafting procedures such as socket preservation, ridge augmentation, and sinus floor elevation to re-establish an anatomically and functionally favorable site for implant placement1,2.While autogenous bone grafts are considered the gold standard due to their osteogenic, osteoconductive, and osteoinductive capabilities, their use is frequently constrained by donor site morbidity, surgical complexity, and inconsistent long-term success rates3,4. Consequently, clinicians increasingly rely on allogeneic, xenogeneic, and synthetic alternatives for reconstructing critical-size defects (CSDs). However, these materials lack intrinsic biological activity, offering passive osteoconduction with limited ability to promote cell recruitment, differentiation, or matrix mineralization5.To overcome these limitations, next-generation biomaterials are being developed to mimic the extracellular matrix (ECM) at the nanoscale and deliver instructive cues for bone regeneration. Self-assembling peptide amphiphile (PA) nanofibers have emerged as a powerful platform in this regard. These molecules consist of a hydrophobic alkyl tail conjugated to a functional peptide sequence, allowing them to form ECM-like nanofibrous networks under physiological conditions6,7,8. Their modular design enables the rational incorporation of multiple biologically active peptide sequences within a single supramolecular framework, allowing for the concurrent presentation of distinct regenerative functions. In the present study, four peptide amphiphiles were co-assembled into a multifunctional nanofiber interface, each selected to address a critical biological process involved in bone healing. The DGEA motif, derived from type I collagen, facilitates integrin-mediated osteogenic differentiation by engaging α2β1 integrins, thereby supporting early commitment of mesenchymal stem cells to the osteoblastic lineage9. The EEE sequence, composed of a negatively charged glutamic acid triplet, mimics calcium-binding domains of native bone matrix proteins and enhances mineralization by promoting hydroxyapatite nucleation on the scaffold surface10. To ensure robust surface adhesion under physiological and wet conditions, DOPA—a catechol-containing residue inspired by mussel adhesive proteins—was incorporated, providing strong binding to the irregular and moist topography of graft materials through hydrogen bonding and metal coordination11,12. Finally, GL13K, a synthetic cationic peptide derived from salivary proteins, was included for its broad-spectrum antimicrobial activity, which has been shown to inhibit bacterial adhesion and biofilm formation, thereby reducing the risk of infection-driven graft failure13,14,15.While surface modification approaches in dentistry have traditionally relied on single-function peptides or have been tested on non-clinical materials, few have been evaluated on clinically approved xenograft, allograft, or synthetic substrates widely used in oral surgery16. Moreover, these commercial grafts—although effective in space maintenance—often exhibit limited regenerative performance, including slow resorption (xenografts), biological variability (allografts), or low osteogenic capacity (synthetics), particularly in compromised clinical scenarios17,18,19,20. Our multifunctional nanofiber interface directly addresses these limitations by providing a tunable, bioactive coating compatible with real-world grafts, aiming to enhance biological performance across multiple regenerative domains. The multifunctional peptide nanofiber platform used herein has previously shown both osteoinductive and antimicrobial effects, as reported in our earlier study21.Materials and methodsMaterialsAll Fmoc-protected amino acids, rink amide resin (0.65 mmol/g), and coupling reagents including HBTU, HOBt, and DIPEA used for peptide synthesis were obtained from Iris Biotech GmbH (Marktredwitz, Germany). Lauric acid (≥ 98%) was purchased from Sigma-Aldrich (St. Louis, MO, USA). Analytical grade solvents such as N, N-dimethylformamide (DMF), dichloromethane (DCM), and diethyl ether were sourced from Merck (Darmstadt, Germany). Trifluoroacetic acid (TFA) and triisopropylsilane (TIS) were also obtained from Merck.Peptide purification was carried out using a preparative reverse-phase high-performance liquid chromatography (HPLC) system (Shimadzu, Japan) equipped with a C18 column (Grace Vydac, USA). The identity and purity of synthesized peptides were confirmed by electrospray ionization mass spectrometry (ESI-MS) using a Bruker amaZon SL ion trap mass spectrometer. Peptide self-assembly behavior and secondary structure were analyzed using a Jasco J-1500 circular dichroism (CD) spectropolarimeter (Tokyo, Japan). Fourier-transform infrared (FTIR) spectra were recorded with a Bruker Tensor II ATR-FTIR spectrometer, and static contact angles were measured using a Dataphysics OCA goniometer (Filderstadt, Germany).Graft materials included bovine-derived xenografts, human-derived allografts, and synthetic hydroxyapatite particles, all obtained from Bioland (Istanbul, Türkiye). Human dental pulp stem cells (DPSCs), pre-characterized for mesenchymal markers, were supplied by Genkök Biotechnology (Ankara, Türkiye). For cell culture, Dulbecco’s Modified Eagle Medium (DMEM, high glucose), fetal bovine serum (FBS), penicillin-streptomycin, and trypsin-EDTA were procured from Gibco, Thermo Fisher Scientific (Waltham, MA, USA).Fluorescent staining of viable cells was performed using Calcein-AM (Thermo Fisher Scientific), and metabolic activity was evaluated using the MTT assay reagent (Sigma-Aldrich). Alkaline phosphatase activity was measured using p-nitrophenyl phosphate (pNPP; Sigma-Aldrich), while extracellular matrix mineralization was assessed with Alizarin Red S and cetylpyridinium chloride, both purchased from Sigma-Aldrich. Total RNA was isolated using TRIzol reagent (Invitrogen, Thermo Fisher), and cDNA synthesis was carried out with the RevertAid First Strand cDNA Synthesis Kit (Thermo Fisher Scientific). Gene expression was analyzed using SYBR Green-based qPCR reagents from Applied Biosystems (Foster City, CA, USA).Micro-computed tomography was conducted with a SkyScan 1176 system (Bruker, Kontich, Belgium), and morphometric analysis was performed using CTAn and CTVol software (Bruker). For histological analysis, decalcification was performed with EDTA (Merck), and sections were stained using 0.1% toluidine blue (Sigma-Aldrich). All other general laboratory reagents and solutions were obtained from Sigma-Aldrich or Merck unless otherwise specified.Peptide amphiphile synthesis and characterizationPeptide amphiphile (PA) molecules were synthesized via standard solid-phase peptide synthesis (SPPS) using Fmoc (9-fluorenylmethyloxycarbonyl) chemistry22. The syntheses were performed on a rink amide resin (0.65 mmol/g loading capacity) using an automated peptide synthesizer (e.g., Liberty Blue, CEM Corporation). Fmoc-protected amino acids were sequentially coupled using HBTU/HOBt activation in the presence of DIPEA. After completion of chain assembly, lauric acid (C12:0) was conjugated to the N-terminus to confer amphiphilic properties, followed by cleavage and side chain deprotection using a TFA: TIS: water (95:2.5:2.5, v/v) cocktail for 2 h at room temperature. Crude products were precipitated with cold diethyl ether and lyophilized.Four distinct peptide amphiphile sequences were synthesized:DGEA-PA (Lauryl–Val–Val–Ala–Gly–Lys–Lys–Gly–Asp–Gly–Glu–Ala–amide) – collagen I mimeticEEE-PA (Lauryl–Val–Val–Ala–Gly–Lys–Lys–Gly–Glu–Glu–Glu–Ala–amide) – hydroxyapatite mineralization promotingDOPA-PA (Lauryl–Val–Val–Ala–Gly–Lys–Lys–Gly–DOPA–Gly–Glu–Ala–amide) – inspired by mussel adhesive proteinGL13K-PA (Lauryl–Val–Val–Ala–Gly–Lys–Lys–Gly–Gly–Leu–Lys–Lys–amide) – antimicrobial sequencePurification was performed via reverse-phase HPLC using a C18 preparative column under a linear water–acetonitrile gradient with 0.1% TFA. Molecular identity and purity (> 95%) were confirmed by electrospray ionization mass spectrometry (ESI-MS). To investigate secondary structures and self-assembly behavior, peptide amphiphiles were dissolved in deionized water or phosphate buffer (10 mM, pH 7.4) at a final concentration of 1 wt%. Solutions were sonicated and incubated at room temperature for 24 h to promote nanofiber formation under physiological ionic conditions (150 mM NaCl, pH 7.4). Circular dichroism (CD) spectra were acquired between 190 and 260 nm to evaluate β-sheet content. The zeta potential of individual peptide amphiphile solutions and the assembled nanofiber system was measured to evaluate surface charge characteristics. Measurements were performed at 25 °C using a Malvern Zetasizer Nano ZS (Malvern Instruments Ltd., UK) equipped with a 633 nm He-Ne laser and a detection angle of 173°. Peptide solutions were prepared at a final concentration of 0.1% (w/v) in 10 mM phosphate-buffered saline (PBS, pH 7.4) and filtered through a 0.22 μm syringe filter prior to analysis. Each sample was measured in triplicate, and average zeta potential values were reported with standard deviation.Graft materials and surface functionalization with peptide nanofibersCommercially available xenografts (Xeno; bovine-derived), allografts (Allo; human-derived), and synthetic hydroxyapatite-based grafts (Synth) were used as base scaffold materials. Prior to functionalization, all grafts were sterilized by UV irradiation for 30 min and stored under sterile conditions. Peptide amphiphile (PA) powders—DGEA-PA, EEE-PA, DOPA-PA, and GL13K-PA—were mixed in equimolar ratios to form a bioactive cocktail. The combined mixture was dissolved in sterile deionized water or phosphate-buffered saline (PBS, pH 7.4) at a total concentration of 1% (w/v) and incubated for 24 h at room temperature under physiological salt conditions (150 mM NaCl) to promote spontaneous nanofiber self-assembly.Table 1 Physical and structural properties of the graft materials Used23.Full size tableEach graft type (100 mg) was immersed in 1 mL of the assembled peptide solution and incubated overnight at 4 °C under gentle agitation to ensure homogeneous coating. Excess solution was removed by centrifugation (500 × g, 5 min), and grafts were washed once with sterile PBS and air-dried under aseptic conditions.Experimental groups were defined as follows:Xeno, Allo, Synth: uncoated graftsXeno-P, Allo-P, Synth-P: grafts coated with the DGEA-PA + EEE-PA + DOPA-PA + GL13K-PA nanofiber mixtureSuccessful peptide immobilization was confirmed via Fourier-transform infrared (FTIR) spectroscopy by detecting characteristic amide I and II absorption bands, indicating the presence of peptide secondary structures on graft surfaces.Morphological and chemical characterization of functionalized graftsSurface morphology of functionalized and unmodified graft materials was examined using scanning electron microscopy (SEM). Samples were mounted on aluminum stubs using carbon adhesive tape, sputter-coated with a 10 nm layer of gold, and imaged using a scanning electron microscope (e.g., Zeiss EVO LS10) operated at 5–15 kV. Multiple fields and magnifications were recorded to evaluate nanofiber coverage and surface architecture. Chemical characterization of the peptide coatings was performed using attenuated total reflectance Fourier-transform infrared (ATR-FTIR) spectroscopy. Spectra were acquired in the range of 4000–600 cm⁻¹ using a spectrometer (e.g., Bruker Tensor II) with 32 scans per sample at 4 cm⁻¹ resolution. Particular attention was given to amide I and II bands, which are indicative of peptide backbone structure. In addition, phosphate- and carbonate-associated absorption bands were monitored to assess interactions with the mineral components of the grafts. Optional analysis of surface wettability was performed using contact angle goniometry (e.g., Dataphysics OCA system). A sessile drop of distilled water (5 µL) was placed on the surface of each sample, and static contact angles were measured at room temperature. Measurements were repeated in triplicate for each group, and values were recorded for comparative assessment of surface hydrophilicity following peptide coating.In vitro cell culture and experimental designHuman dental pulp stem cells (DPSCs) were utilized for in vitro evaluation of graft biocompatibility and osteogenic potential. DPSCs were obtained from a commercial cell supplier (Genkök Biotechnology, Ankara, Türkiye), pre-characterized for mesenchymal stem cell surface markers (CD73⁺, CD90⁺, CD105⁺, CD34⁻, CD45⁻) and confirmed for multipotent differentiation potential24.Cells were cultured in standard growth medium composed of Dulbecco’s Modified Eagle Medium (DMEM, high glucose; Gibco) supplemented with 10% fetal bovine serum (FBS; Biowest), 1% penicillin-streptomycin (Gibco), and maintained in a humidified incubator at 37 °C with 5% CO₂. Media was refreshed every 2–3 days.For experimental studies, DPSCs at passages 3–5 was detached using 0.25% trypsin-EDTA, counted, and seeded onto the graft samples at a density of 1 × 10⁵ cells per well in 24-well tissue culture plates. Each well contained one graft sample (either unmodified or peptide-functionalized).The experimental groups were organized as follows:Unmodified graft groups (negative controls)Xeno (uncoated xenograft), Allo (uncoated allograft), Synth (uncoated synthetic scaffold).Peptide-coated graft groups (test groups)Xeno-P, Allo-P, Synth-P — each coated with a multifunctional peptide nanofiber blend (DGEA-PA + EEE-PA + DOPA-PA + GL13K-PA).In this setup, the unmodified grafts served as negative controls to assess baseline cell behavior, while the peptide-coated groups were used to evaluate the bioactivity and osteoinductive potential of the nanofiber system.All groups were cultured in standard growth medium for initial adhesion and viability studies. For osteogenic differentiation studies, cells were cultured in osteogenic induction medium (DMEM supplemented with 10% FBS, 10 mM β-glycerophosphate, 50 µg/mL ascorbic acid, and 100 nM dexamethasone) for up to 14 days, with medium changes every 3 days.Cell adhesion assayCell adhesion was evaluated 24 h after seeding using a live-cell fluorescence staining method. DPSCs (1 × 10⁵ cells/well) were seeded onto each graft sample in 24-well plates and incubated under standard culture conditions (37 °C, 5% CO₂). After 24 h, non-adherent cells were gently removed by washing the samples twice with phosphate-buffered saline (PBS, pH 7.4). Adherent viable cells were stained using the Calcein-AM live-cell labeling dye (Thermo Fisher Scientific). A working solution of Calcein-AM (2 µM in PBS) was added to each well and incubated for 30 min at 37 °C in the dark. After incubation, excess dye was removed by PBS washing, and the samples were immediately imaged using an inverted fluorescence microscope (e.g., Zeiss Axio Observer) with FITC filter settings. Fluorescence images were captured from three predefined regions per sample. Quantitative analysis of adherent cell density was performed using ImageJ software (National Institutes of Health, USA)25. by thresholding and counting fluorescent cell areas per field. Cell adhesion efficiency was expressed as mean fluorescence area per field (n = 3 per group), providing a relative estimate of the number of viable adherent cells.Cell viability and proliferation assayTo evaluate the metabolic activity and proliferation of DPSCs cultured on graft surfaces, the MTT [3-(4,5-dimethylthiazol-2-yl)−2,5-diphenyltetrazolium bromide] assay was employed at 24-, 48-, and 72-hours post-seeding. Cells were seeded at a density of 1 × 10⁵ cells per well in 24-well plates containing either uncoated or peptide-functionalized grafts and incubated under standard conditions. At each time point, culture medium was aspirated and replaced with 500 µL of serum-free DMEM containing 0.5 mg/mL MTT (Sigma-Aldrich). Samples were incubated for 4 h at 37 °C in the dark to allow for the formation of purple formazan crystals by metabolically active cells. Following incubation, the MTT solution was carefully removed, and the formed formazan crystals were solubilized in 500 µL of dimethyl sulfoxide (DMSO) per well. The resulting solution was mixed thoroughly and transferred to a 96-well plate for absorbance measurement. Optical density was recorded at 570 nm using a microplate reader (e.g., BioTek Epoch 2), with a reference wavelength of 630 nm. All experiments were conducted in triplicate (n = 3 per group per time point). Cell viability was expressed as mean absorbance values, which are directly proportional to the number of viable cells.The antimicrobial activity of peptide nanofiber components (DOPA, GL3K, DGEA, and EEE) was evaluated using the MTT-based viability assay, adapted for bacterial cells. Escherichia coli (ATCC 25922) and Staphylococcus aureus (ATCC 29213) were used as representative Gram-negative and Gram-positive strains, respectively. Bacterial suspensions were adjusted to 1 × 10⁸ CFU/mL using the McFarland standard. Peptide nanofiber solutions were prepared at final concentrations of 50, 100, and 200 µg/mL in sterile PBS. For each condition, 500 µL of the bacterial suspension was mixed with 500 µL of the corresponding peptide solution in sterile microtubes and incubated at 37 °C for 4 h under shaking. After incubation, 100 µL from each mixture was transferred to a 96-well plate, followed by the addition of 10 µL of MTT reagent (5 mg/mL in PBS). The plate was incubated for 2 h at 37 °C in the dark. Formazan crystals were solubilized using 100 µL of DMSO, and absorbance was measured at 570 nm using a microplate reader. Untreated bacteria were used as the viability control. All experiments were performed in triplicate, and data are presented as mean ± standard deviation (Table S2-3).Osteogenic differentiation and gene expression analysisTo assess osteogenic commitment at the molecular level, real-time quantitative polymerase chain reaction (qRT-PCR) was used to analyze the expression of key osteogenesis-associated genes. DPSCs were cultured on unmodified and peptide-functionalized grafts in osteogenic induction medium (DMEM supplemented with 10% FBS, 10 mM β-glycerophosphate, 50 µg/mL ascorbic acid, and 100 nM dexamethasone) for up to 14 days. Medium was refreshed every 3 days. At day 3 and day 14, total RNA was extracted from the cell-graft complexes using a phenol-chloroform-based method (e.g., TRIzol reagent, Invitrogen), according to the manufacturer’s protocol. RNA concentration and purity were determined spectrophotometrically (NanoDrop™ 2000, Thermo Scientific). Reverse transcription was performed using a cDNA synthesis kit (e.g., RevertAid First Strand cDNA Synthesis Kit, Thermo Fisher), employing 1 µg of total RNA per reaction. Gene expression analysis was carried out using SYBR Green-based qRT-PCR (e.g., Applied Biosystems™ StepOnePlus™ system). Specific primers were designed for the following target genes: Runt-related transcription factor 2 (RUNX2), Osteopontin (OPN), Type I collagen alpha 1 (COL1A1). GAPDH was used as the internal reference gene. The relative expression of each gene was calculated using the ΔΔCt method26, and results were expressed as fold change relative to control samples at each time point. All qPCR reactions were run in triplicate, and negative controls (no template) were included to confirm specificity. Primer sequences and annealing temperatures are provided in Supplementary Table 1.Alkaline phosphatase (ALP) activity and mineralization assayTo evaluate functional osteogenic differentiation, alkaline phosphatase (ALP) activity and extracellular matrix mineralization were assessed at day 14 of induction.Alkaline Phosphatase (ALP) Activity: DPSCs cultured on grafts were lysed using ALP assay lysis buffer (e.g., 0.1% Triton X-100 in Tris-HCl, pH 7.6). Cell lysates were collected and incubated with p-nitrophenyl phosphate (pNPP; Sigma-Aldrich) substrate solution at 37 °C for 30 min. The enzymatic reaction was terminated by adding 3 N NaOH, and the absorbance was measured at 405 nm using a microplate reader (BioTek Epoch 2). Protein concentrations in lysates were determined by BCA protein assay (Thermo Scientific), and ALP activity was normalized to total protein content (U/mg protein) (Figure S2). All measurements were performed in triplicate.Calcium Deposition by Alizarin Red S (ARS) Staining: To assess matrix mineralization, samples were fixed with 4% paraformaldehyde for 15 min and rinsed with distilled water. Fixed samples were stained with 2% (w/v) Alizarin Red S (ARS; Sigma-Aldrich) solution (pH 4.2) for 20 min at room temperature. Excess dye was removed by repeated washing with distilled water. Stained calcium deposits were visualized under a stereomicroscope and imaged. For quantification, the bound ARS dye was extracted using 10% (w/v) cetylpyridinium chloride in 10 mM sodium phosphate buffer (pH 7.0) for 30 min, and the absorbance was measured at 550 nm. Quantitative values were reported as mean ± standard deviation (n = 3 per group). (Standard curve can be found in Supplemantary Information). To quantify calcium deposition following osteogenic differentiation, Alizarin Red S-stained samples were imaged under identical exposure settings using a brightfield microscope. For each sample, representative images were acquired from at least three independent fields per group. Red channel intensity was then quantified using ImageJ (NIH) software. Briefly, images were converted to RGB stack, and the red channel was isolated. Mean red intensity values were measured using standardized ROI selections, ensuring consistent area and thresholding across all groups. The resulting values were averaged for each group and presented as mean ± standard deviation (n = 3). This method provides a semi-quantitative assessment of mineralized matrix content, complementing the visual ARS staining results.In vivo calvarial bone defect model and treatment groupsTo evaluate the bone regenerative capacity of peptide-functionalized grafts in vivo, a critical-sized calvarial defect model was utilized in adult male New Zealand White rabbits (Oryctolagus cuniculus). A bilateral critical-sized calvarial defect (8 mm in diameter) was selected, as such defects do not heal spontaneously and are widely accepted for assessing bone regeneration strategies in preclinical studies5,27.All animals used were adult male New Zealand White rabbits, weighing between 2.5 and 3.0 kg and approximately 5–6 months old at the time of surgery. Animals were obtained from Kobay Deney Hayvanlari Laboratuvari (Ankara, Turkey). All animals were clinically healthy with no known immunodeficiencies or genetic modifications, and no previous surgical or experimental procedures had been performed prior to this study.Upon arrival, animals were individually housed in standard cages under controlled ambient temperature conditions with a 12-hour light/dark cycle. They were fed a standard commercial rabbit chow and had ad libitum access to fresh water throughout the study. Animals were acclimatized to the housing environment for seven days prior to surgical procedures to minimize stress-induced variability.All experimental procedures were approved by the Local Animal Ethics Committee of Kobay Deney Hayvanlari Laboratuvari (Protocol No: 709, Approval Date: 02/2024) and conducted in accordance with Turkish national regulations, the European Directive 2010/63/EU on the protection of animals used for scientific purposes and reported in compliance with the ARRIVE guidelines (Animal Research: Reporting of In Vivo Experiments).Inclusion criteria required that all animals be healthy, adult male New Zealand White rabbits (2.5–3.0 kg) without any clinical signs of systemic disease prior to surgery. Furthermore, only male rabbits were used in this study to avoid the potential confounding effects of hormonal fluctuations associated with the estrous cycle in females. Estrogen and other sex hormones are known to influence bone turnover, inflammatory responses, and soft tissue remodeling, which could introduce variability into regenerative outcomes. By limiting the cohort to skeletally mature males, we aimed to ensure a more uniform biological background, enhancing the reproducibility and interpretability of the observed effects. Exclusion criteria included intraoperative complications, postoperative infections, significant surgical site morbidity, or systemic illness that could impact bone healing. No animals or defects met exclusion criteria; therefore, all defects were included in the final analysis. Each experimental group (Blank, Xeno, Xeno-P, PA-NF) consisted of 6 defects (n = 6). The sample size was determined based on standard practice in preclinical rabbit calvarial defect models. Six animals were used, each contributing four defects, resulting in 24 defects total. No formal a priori sample size calculation was performed; the selected sample size is consistent with previous studies that reported sufficient statistical power for detecting significant differences in bone regeneration outcomes28.Under general anesthesia induced by intramuscular injection of xylazine (5 mg/kg) and ketamine (35 mg/kg), the calvarial region was shaved, disinfected, and a midline sagittal incision was made to expose the parietal bone. Bilateral full-thickness flaps were reflected, and circular, critical-sized defects (8 mm in diameter) were created on both sides of the parietal bone using a trephine bur under sterile saline irrigation27,29. The following experimental conditions were randomly assigned to the defects:Blank: untreated critical-sized defect (empty control)Xeno: defect filled with unmodified xenograft particlesXeno-P: defect filled with peptide-functionalized xenograft (coated with DGEA-PA, EEE-PA, DOPA-PA, and GL13K-PA nanofibers)PA-NF: defect filled with self-assembled peptide nanofiber hydrogel alone (no graft material)The experimental unit for all analyses was the individual defect. To minimize potential confounding effects, defects were created bilaterally, and treatment allocations were randomized using a computer-generated random number. Randomization of defects to treatment groups was performed using a computer-generated random number table; the individual performing the allocation was aware of group assignments. The surgical procedures were conducted by personnel who were aware of the treatment groups due to the nature of the interventions. However, all outcome assessments, including micro-CT imaging and histological evaluations, were performed by investigators blinded to the group allocations. Data analysis was conducted using anonymized datasets to maintain blinding during statistical evaluation. Following implantation, surgical sites were closed in anatomical layers using resorbable sutures. Postoperative analgesia (meloxicam, 0.2 mg/kg) and prophylactic antibiotic therapy (enrofloxacin, 10 mg/kg) were administered subcutaneously once daily for three consecutive days. At six weeks post-surgery, animals were euthanized with an overdose of anesthesia, and the calvarial bones were harvested for micro-computed tomography (micro-CT) and histological analyses. The primary outcome measure was bone volume fraction (BV/TV) assessed via micro-CT imaging. Secondary outcome measures included trabecular thickness (Tb.Th), trabecular spacing (Tb.Sp), and histological evaluation of new bone formation and tissue integration.Micro-CT imaging and bone morphometric analysisQuantitative evaluation of newly formed bone within calvarial defects was performed using high-resolution micro-computed tomography (micro-CT)27. Harvested calvarial specimens were fixed in 10% neutral-buffered formalin for 48 h, then scanned using a micro-CT imaging system (e.g., SkyScan 1176, Bruker) at 9 μm isotropic voxel resolution, with 50 kV voltage, 500 µA current, and a 0.5 mm aluminum filter. Regions of interest (ROIs) were defined based on the full defect diameter. All groups—including peptide-functionalized grafts, unmodified grafts, and the blank group (defect without graft material)—were analyzed under identical scanning and processing conditions.Thresholding was applied to distinguish mineralized tissue from soft tissue and background.Quantitative morphometric parameters included:Bone volume fraction (BV/TV, %)Trabecular thickness (Tb.Th, µm)Trabecular number (Tb.N, 1/mm)Trabecular separation (Tb.Sp, µm)Bone surface to volume ratio (BS/TV, 1/mm)For each experimental group, a minimum of three independent biological replicates (n = 3) were analyzed, and group-level comparisons were performed based on the mean values derived from these replicates. Outcome assessors were blinded to group allocation during analysis. Three-dimensional renderings were generated to visualize the extent and spatial pattern of bone regeneration within the defect area.Histological evaluationFollowing micro-CT analysis, calvarial bone specimens were decalcified in 10% ethylenediaminetetraacetic acid (EDTA, pH 7.4) at room temperature for 3 weeks with regular solution changes. After complete decalcification, samples were dehydrated through a graded ethanol series, cleared in xylene, and embedded in paraffin. Serial Sect. (5 μm thickness) were obtained in the coronal plane using a microtome (e.g., Leica RM2235). Sections were stained with toluidine blue (0.1% w/v, pH 2.5) to visualize cellular and extracellular components, including newly formed bone tissue and residual graft material. Slides were examined under a light microscope (e.g., Olympus BX51), and digital images were captured for qualitative assessment29. Regions of interest (ROI) were delineated manually around the original defect margins (Figure S7). ImageJ software (NIH, USA) was used to quantify three key parameters: graft retention (% area occupied by residual graft material), new bone formation (% area occupied by newly formed trabecular bone), and void/soft tissue area (% area lacking mineralized tissue or graft). For each sample, three non-overlapping fields were analyzed, and values were averaged per animal (n = 3 per group). Statistical comparisons were conducted using one-way ANOVA followed by Tukey’s post hoc test. Results were reported as mean ± standard deviation (SD), and significance thresholds were set at p