A 3D Prevascularized Calcium Phosphate Cement Scaffold for Accelerated Alveolar Bone Regeneration and Angiogenesis in Rats

A 3D Prevascularized Calcium Phosphate Cement Scaffold for Accelerated Alveolar Bone Regeneration and Angiogenesis in Rats | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Research Article A 3D Prevascularized Calcium Phosphate Cement Scaffold for Accelerated Alveolar Bone Regeneration and Angiogenesis in Rats Yaxi Sun, Zeqing Zhao, Qingchen Qiao, Wenting Yu, Yuxing Bai This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-8260921/v1 This work is licensed under a CC BY 4.0 License Status: Under Review Version 1 posted 15 You are reading this latest preprint version Abstract Objectives. The objective of this study is to develop a load-bearing prevascularized construct by combining calcium phosphate cement (CPC) with cells within a three-dimensional (3D) hydrogel culture system, to accelerate the regeneration of alveolar bone defects. Methods. A 3D co-culture system was established by encapsulating human periodontal ligament stem cells (hPDLSCs) and human umbilical vein endothelial cells (hUVECs) within a gelatin methacryloyl (GelMA) hydrogel on 3D-printed porous CPC scaffolds. The mechanical properties, pore structure and angiogenic potency were determined in vitro. In vivo performance was evaluated using a nude rat subcutaneous implantation model and a rat alveolar bone defect model. Four groups were tested: (1) Blank group (surgery-only group); (2) CPC+GelMA group (non-prevascularized group)；(3) CPC+GelMA-cell group (prevascularized group)；(4) Natural Periodontiumgroup. Results. The novel construct had good mechanical properties and biocompatibility. The 3D co-culture in GelMA successfully induced microvascular formation in vitro. Subcutaneous implantation in nude rats showed that the CPC+GelMA-cell group exhibited markedly greater angiogenic capacity than the CPC+GelMA group after 6 weeks, with a neovascular density 1.93-fold higher than that of the non-prevascularized group. Among all groups, the CPC+GelMA-cell group exhibited the strongest capacity for repairing rat alveolar bone defects. Compared to CPC+GelMA group, CPC+GelMA-cell group significant enhanced bone regeneration in rats by 1.23-1.37 folds, and increased vascularization by 2.65 folds (p＜0.05). Conclusions: The novel three-dimensional prevascularized CPC construct combined appropriate mechanical properties with great efficacy for alveolar bone regeneration and vascularization in vivo in an animal model. Human periodontal ligament stem cells Human umbilical vein endothelial cells Three-dimensional printed Calcium phosphate cement Gelatin methacryloyl hydrogel Osteogenesis Angiogenesis rat model Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Figure 7 Figure 8 Figure 9 1. Background Orthodontic treatment carries the risk of periodontal tissue damage. Studies indicate that over one-third of adult patients exhibit anterior alveolar bone recession exceeding 2 mm post-treatment [1]. Improper orthodontic design may also lead to alveolar bone defects such as fenestration and dehiscence [1]. Tissue engineering scaffolds are a critical tool for reconstructing alveolar bone defects, yet their regenerative potential is critically constrained by inadequate and delayed vascularization [2]. Insufficient blood supply can cause hypoxia, impair osteogenesis, and lead to graft necrosis and failure [2, 3]. Therefore, overcoming insufficient vascularization is essential for bone-defect repair. Prevascularized scaffolds enables their vascular networks to anastomose with the host’s vasculature after implantation, markedly reducing the risk of ischemic necrosis, and substantially increasing the success rate of bone regeneration [4]. Calcium phosphate cement (CPC) is an ideal osteogenic material due to its biocompatibility, osteoconductivity, and bone-like composition [5]. However, conventional CPC is predominantly microporous and lacks interconnected macroporous structures, hindering vascular ingrowth and limiting its utility in large bone defects [6]. Although prevascularized CPC scaffolds have been studied for cranial and long bone repair [7-10], a three-dimensional (3D) prevascularized CPC scaffold tailored for alveolar bone regeneration has not yet been reported. Calcium phosphate cement (CPC) is a self-setting osteogenic scaffold material, resembling the inorganic composition of natural bone [5]. CPC regards as a highly promising scaffold material for craniofacial and dental repairs，due to its good biocompatibility, osteoconductivity, and mechanical properties [5]. Nevertheless, unmodified CPC is predominantly microporous and lacks interconnected macroporous structures, hindering vascular ingrowth and limiting its utility in large bone defects [6]. Prevascularized CPC scaffolds may significantly enhance bone repair capabilities. Currently, research on prevascularized calcium-phosphate osteogenic scaffolds primarily focuses on cranial and long bone repair [7-10]. To date, there has been no report of constructing three-dimensional (3D) prevascularized CPC scaffolds for alveolar bone tissue engineering. Prevascularization of CPC scaffolds is currently achieved through vascular implantation [7], in vivo culturing [6], or cell-based in vitro prevascularization [11]. However, vascular implantation and in vivo culturing involve invasive procedures, limiting their clinical applicability. Therefore, cell-based in vitro prevascularization is now in the spotlight [12]. Human umbilical vein endothelial cells (hUVECs) readily self-assemble into microcapillaries, yet endothelial monocultures fail to generate stable, mature vascular structures [13]. Endothelial cell migration and neovessel formation require specific pro-angiogenic factors, which are insufficiently produced by endothelial cells in monoculture [4, 14]. Mesenchymal stem cells (MSCs) enhance angiogenesis by secreting pro-angiogenic factors and stabilizing nascent vessels as pericytes [13, 15, 16]. Numerous studies demonstrate that co-culturing MSCs with endothelial cells yields stable vascular structures [16]. Human periodontal ligament stem cells (hPDLSCs) are a seed cell source that can be harvested from the extracted wisdom teeth or the teeth extracted for orthodontic purpose without additional invasive surgery for the patient. Previous studies have shown positive results in using hPDLSCs for tissue regeneration, especially in bone and periodontal tissue repairment [17, 18]. In addition, hPDLSCs can differentiate into bone, nerve, connective tissue, and cementum under specific conditions [19, 20]. Therefore, hPDLSCs are a potent cell source in stem cell delivery via scaffolds for bone regeneration, especially for alveolar bone repair. Studies showed that co-culturing hPDLSCs and hUVECs on CPC surfaces can form microvascular-like structures, suggesting their potential for CPC scaffold prevascularization [11]. To date, a literature search revealed no report on the prevascularized CPC scaffolds seeded with hPDLSCs and hUVECs for alveolar bone defect repair. Beyond seed cell selection, scaffold structural properties also influence prevascularization efficacy. Three-dimensional (3D) printing technology enables precise control over pore size and interconnectivity [21, 22]. 3D-printed grid-like CPC scaffolds exhibit uniform, interconnected micropores that readily conducive to microvascular growth support microvessel ingrowth and anastomosis [23]. Additionally, the culture method for seed cells is crucial for prevascularization. Traditional two-dimensional (2D) culture involves seeding cells directly onto scaffold surfaces, where restricted area and contact inhibition curtail expansion and capillary morphogenesis [24, 25]. Research indicated that 3D microenvironments enhance cell responsiveness to biochemical signals during angiogenesis [24]. Compared to 2D culture, 3D culture better mimics natural cell growth conditions, promoting cell-cell and cell-matrix interactions [26]. Encapsulating cells in hydrogel-based materials can simulate extracellular matrix structures, providing 3D support [24]. Gelatin methacryloyl (GelMA) hydrogel offers excellent biocompatibility and enables 3D cell encapsulation, simulating a natural extracellular matrix conducive to microvascular formation [27]. However, hydrogels lack sufficient mechanical strength for load-bearing applications like alveolar bone repair [28]. Recent studies combine hydrogels with high-strength scaffolds to create composite materials with both biological and mechanical advantages [28]. To address the challenges of slow vascularization and inadequate mechanical strength in current bone grafts, we aim to design a CPC-based, cell-laden 3D hydrogel composite. The primary goals are to engineer a scaffold with improved load-bearing capacity and to establish a preformed vascular network within it, ultimately boosting the regenerative outcomes in alveolar bone defects. 2. Methods 2.1. Harvesting hPDLSCs from extracted teeth Periodontal ligament (PDL) tissues were harvested from healthy premolars extracted from patients aged 18–26 years for orthodontic purpose. The hPDLSCs were isolated as described previously [29]. The procedures were approved by the Medical Ethics Committee of Beijing Stomatological Hospital, Capital Medical University (NO. CMUSH-IRB-KJ-PJ-2024-28). The written informed consent was obtained from each participant before the study. The study was carried out in accordance with the Declaration of Helsinki. The PDL tissues were enzymatically digested with 3 mg/mL collagenase type I (Gibco BRL, Grand Island, NY, USA) and 4 mg/mL dispase (Gibco BRL) at 37°C for 45 minutes. After digestion, the cell suspension was collected and transferred to culture dishes (Costar, Cambridge, MA, USA) with dulbecco’s modified Eagle’s medium (DMEM, Gibco BRL) supplemented with 1% penicillin/streptomycin (P.S, Gibco BRL) and 20% fetal bovine serum (FBS, Gibco BRL). Upon reaching 70–80% confluence, cells were passaged using 0.25% trypsin-EDTA (Gibco BRL). Cells at passages 3–5 were used in subsequent experiments. 2.2. Identification of hPDLSCs The expression of surface antigen profiles (CD34, CD45, CD90, CD105, and STRO-1) of passage 3-5 cells were analyzed by flow cytometry as described previously [29]. PE-conjugated antibodies against CD34, CD105, and STRO-1 (Thermo Fisher Scientific, Rockford, IL, USA), along with FITC-labeled CD45 and CD90 antibodies (Thermo Fisher Scientific), were employed for immunophenotyping. Cells were digested, centrifuged and resuspended in cold PBS to achieve a concentration of 1×10 5 cells/cube. For staining, the cell suspension was mixed with antibody and incubated on ice in dark. After being washed and resuspended, cell surface antigen expression was tested using a BD Vantage flow cytometer (BD Biosciences). 2.3. Culturing of hUVECs Primary hUVECs were acquired from Sciencell (Carlsbad, CA, USA). The cells were cultured in 10 cm culture dishes with endothelial cell medium (ECM, Sciencell) under standard culture conditions [30]. Subsequent passages were performed when confluency reached 70% to 80%. Cells between passage 2–3 were used in this study. 2.4. Fabrication of 3D prevascularized CPC scaffold The CPC paste was manufactured by InnoTERE GmbH (Radebeul, Germany) and fabricated with the BioScaffolder 2.1 (GeSiM mbH, Radeberg, Germany) operated in a laminar flow workbench [31]. CPC paste was plotted in 60° configuration (the layer orientation changed after every second layer by 60°) and a filament diameter of 300 μm with designed pore size of 300 μm. Scaffolds used for a rat alveolar defect model were plotted a rectangular external structure with a size of 4 mm×2 mm×1 mm. A cylindrical outer geometry with a height of 3 mm and a diameter of 12 mm were used for the following experiments. The hUVECs and hPDLSCs were detached and mixed at a ratio of 3:1 (hUVECs:hPDLSCs) as described previously [30]. The co-cultured cells were suspended into 5% GelMA solution at a total density of 1 ×10 6 cells/ml at 37℃ in dark. Then, the suspension was placed in a 1 ml syringe (with 100 µm inner diameter) and injected into the pores inside the 3D printed CPC scaffold. Expose the construct to a 405 nm wavelength light source to facilitate its curing process. The 3D co-culture scaffold was cultured with the endothelial cell medium for 21 days. 2.5. Scanning electron microscopy (SEM) of 3D printed CPC scaffolds After 21 days of culturing, the composite constructs were fixed overnight. The next day, each cylindrical scaffold was bisected through its mid-plane and the resulting cross-sections were dehydrated and examined under a scanning electron microscope (Quanta 200, FEI, Hillsboro, OR, USA). Filament widths and pore sizes were measured from representative micrographs. Six specimens were analyzed as described previously [19]. 2.6. Mechanical properties Three-point flexural tests were performed on the composite scaffolds using a universal testing machine for mechanical testing [29]. The span is 20 mm and the displacement speed of test head is 1 mm/min. Flexural strength (σ) and elastic modulus (E) were derived from the load–displacement curves. σ=3FmaxL/2bh 2 , E=(F/d) (L 3 /4bh 3 ), where F max is the peak load, L is the span, b and h are the specimen width and thickness, respectively, and F/d represents the slope of the linear-elastic region. Six specimens were tested. 2.7. Viability of encapsulated hDPLSCs After 1, 4, 7 and 14 days of culturing, cellular viability within the hydrogel constructs was evaluated using a live/dead viability assay kit (Sigma-Aldrich). Epifluorescence microscope (Sigma-Aldrich) was used for observation. The percentage of live cells was calculated by Image J software (NIH) as described previously [29]. A cell counting kit (CCK-8 assay, Dojindo, Tokyo, Japan) was used to evaluate cell viability in the 3D culture system at 1, 4, 7, 14 and 21 days. The working solution was prepared with endothelial cell medium containing 10% CCK-8 solution, followed by a 1-hour incubation at 37°C. The cell proliferative rate was determined by measuring the absorbance at an optical density of 450 nm using microplate reader (SpectraMax M5, Molecular Devices, Sunnyvale, CA) as described previously [29]. 2.8. Observing hUVECs via CD31 immunofluorescence staining After 1, 4, 7 and 14 days of culturing, the resulting microvascular-like structures were then visualized and assessed by CD31 (PECAM-1) immunofluorescence staining as described previously [11]. The samples were fixed and incubated with CD31 mouse mAb (1:500, Cell signaling technology, Pudong District, Shanghai, China) overnight. After washing with PBS, Alexa Fluor 488-conjugated goat anti-mouse IgG (1:1000, goat anti-mouse Alexa Fluor 488, green fluorescence, Cell signaling technology) was applied, followed by DAPI counterstaining (1:1000, Beyotime) at room temperature. The samples were observed with confocal laser scanning microscopy (OLS5100, Olympus, Tokyo, Japan). For each time-point, three random regions from each of five specimens were recorded. Image J (National Health Institute, Bethesda, MA, USA) was used to obtain the vessel length per area and junction number per area (n=5). Quantification was performed in duplicate by a double blinding protocol (n = 5). 2.9. Rat complete periodontal defect model The animal protocol was approved by the Committee for Animal Experiments of Beijing Stomatological Hospital, Capital Medical University (NO. 2024-82301117). Male athymic nude rats (8 weeks old, 200–250 g, SPF Biotechnology, Haidian District, Beijing, China) were anesthetized with an intraperitoneal injection of Zoletil 50 (Virbac, Carros, France) at 50 mg/kg body weight (n=6). All surgical procedures were performed under strict aseptic conditions. After sterilization of rats, a critical-size alveolar defect (4 mm × 2 mm × 1 mm) was prepared buccal to the mesial root of the mandibular second molar to establish a rat alveolar bone defect model as described previously [32]. The mesial root was exposed, and all residual periodontal ligament and cementum were meticulously curetted. Scaffolds were press-fit into the defects. Additionally, the rats received flunixin meglumine (2 mg/kg, s.c.; Shanghai Yuanye Bio-Technology, Shanghai, China) and penicillin G benzathine (24,000 IU/kg, i.m.; Pengdi, Henan, China) once daily for 3 consecutive days. Animals were monitored daily for signs of pain, wound integrity, and normal ambulation. Four groups were set: (1) Blank group: underwent the surgical procedure without any scaffold implantation； (2) CPC+GelMA group: received an acellular scaffold, which was preconditioned in ECM for 21 days prior to implantation, serving as the non-prevascularized control； (3) CPC+GelMA-cell group: received the experimental intervention—a prevascularized scaffold seeded with a co-culture of hPDLSCs and hUVECs and matured in ECM for 21 days； (4) Natural Periodontium group: consisted of the contralateral non-operated sites, provided the baseline native tissue. At 4 weeks post-surgery, six rats per group were euthanized. After deep anesthesia with an overdose of intraperitoneal pentobarbital (150 mg/kg), animals were sacrificed by CO₂ asphyxiation followed by exsanguination via bilateral thoracotomy. The implants were then retrieved and immediately fixed in 4% paraformaldehyde at 4 °C for 24 h. 2.10. Micro computed tomography （ Micro-CT ） Micro-CT (SkyScan 1276, Bruker BioSpin, Germany) was employed to scan the bone-defect regions and perform three-dimensional reconstructions [29]. The ROI was defined as the buccal alveolar bone encircling the mesial root of the mandibular second molar and encompassed a volume of 4 mm×2 mm×1 mm. Bone thickness and bone volume fraction was calculated to quantify the percentage of newly formed bone within the defect. Additionally, a cross-sectional slice at the mid-root level of the mesial root was selected for qualitative evaluation. An initial grayscale threshold of 150–1000 HU was applied to distinguish newly formed bone. 2.11. Histomorphometric analyses Specimens were decalcified in 10% ethylene diamine tetraacetic acid (EDTA) (Solarbio Science & Technology, Beijing, China) for 2 months, processed routinely, and embedded in paraffin. The central part of the implant and defect was cut into 5 μm-thick sections for hematoxylin and eosin (H&E) staining, Masson’s staining and immunohistochemistry (IHC) staining. 2.11.1. H&E staining The samples were decalcified and embedded in paraffin. Serial 5 µm-thick sections were prepared and stained with H&E. New bone area, total defect area and the number of new vessels were quantified in each section by image J as described previously [19]. New bone area fraction was calculated by dividing the area of new bone with the area of the total defect. New vessels density was expressed as the number of new vessels divided by total defect area (n=6). 2.11.2. Masson’s staining Deparaffinized sections were stained with Weigert’s iron hematoxylin, sequentially incubated in Biebrich scarlet-acid fuchsin and phosphomolybdic/phosphotungstic acid, and finally differentiated in aniline blue to visualize collagen. Blue-stained bone area and total defect area were measured in each section [30]. Osteoid area fraction was calculated by dividing the area of blue-stained bone with the area of the total defect (n=6). 2.11.3. IHC staining Immunodetection of human CD31 and RUNX2 was performed on 5 µm paraffin sections. Sections were incubated with rabbit anti-human CD31 (1:500, Abcam) and anti-human RUNX2 (5 µg/mL, Abcam), followed by HRP-conjugated secondary antibody (1:500, Abcam). Signals were developed with DAB and counterstained with hematoxylin. For histomorphometry, one mid-sagittal section per animal was analyzed (n=6). The density of TRAP-positive cells was calculated. CD31 and RUNX2 expression was assessed as integrated optical density (IOD) from 6 random fields per section as described previously [32]. 2.12. S ubcutaneous transplantation model in nude rats For subcutaneous transplantation, CPC+GelMA group and CPC+GelMA-cell group were selected. After induction of general anesthesia and sterile preparation, scaffolds were implanted into subcutaneous pockets on the back nude rats (8 weeks, 200–250 g) as described previously [33]. After 6 weeks of implantation, all scaffolds were obtained and fixed for stereomicroscope observations (Olympus, Tokyo, Japan) and H&E staining. Vessels were enumerated in three randomly selected fields per section, and vascular density was calculated as vessel number per unit area. 2.13 ． Statistical analysis All statistical analyses were conducted using SPSS 22.0 (IBM Corp., Armonk, NY, USA). Data are expressed as mean ± standard deviation (SD). Group comparisons were performed by one-way analysis of variance (ANOVA) followed by Tukey’s post-hoc test for multiple comparisons. p-value < 0.05 was considered statistically significant. 3. Results 3.1. Identification of hPDLSCs Figure 1. A plots flow cytometry result of isolated hPDLSCs. CD105, CD90, STRO-1 were highly expressed to 99.6%, 76.4% and 88.6%. While, CD34 and CD45 were weakly expressed to 0.3% and 0.6%, respectively. 3.2. Fabrication of three-dimensionally prevascularized CPC scaffold Figure 1. B, C illustrates the fabrication of a three-dimensional grid-like CPC scaffold. Figure 1. D plots the injection of an hPDLSCs–hUVECs-laden GelMA hydrogel into the 3D-printed CPC scaffold to generate a three-dimensionally prevascularized CPC construct. 3.3. Physical properties of scaffolds Figure 2. A shows CPC filaments intersecting at defined angles. Cross-sectional images (Figure 2. B, C) reveal spherical crystalline structures and abundant pores within the CPC matrix. Quantitative measurements yielded filament diameters of 285.21 ± 1.4 μm, pore sizes of 302.31 ± 3.4 μm, and filament intersection angles of 60.12 ± 0.4° for the 3D-printed CPC scaffold. The flexural strength (Figure 2. D) and elastic modulus (Figure 2. E) of CPC construct had significant difference with cancellous bone. The average value of flexural strength (6.78±0.68 MPa) and elastic modulus (0.41±0.02 GPa) was significantly higher than that of cancellous bone. Values indicated by dissimilar letters are significantly different from each other (p<0.05). 3.4. Viability and cell proliferation of cells co-cultured within GelMA-CPC constructs Live/dead staining images for the encapsulated cells (Figure 3. A) represented numerous live cells (green staining) and a few dead ones (red staining) at each time points tested. At day 1, cells were embedded within the hydrogel, predominantly spherical and clustered, with relatively low cell numbers visible. From day 4 to 7, cells began to aggregate, the number of released cells increased continuously, which extended well, showing spindle or polygonal shape. At day 14, cells were evenly distributed throughout the hydrogel. At day 21, all cells had fully extended into long spindle shapes, forming pseudopodia and interconnecting into branched, network-like structures. The percentages of live cells (Figure 3. B) exceeded 80% on day 1 and significantly increased to approximately 90% from day 4 to 21. As shown in CCK-8 assessment (Figure 3. C), co-cultured hPDLSCs and hUVECs showed 9.2 folds increase of proliferation from day 1 to 21. Values indicated by dissimilar letters are significantly different from each other (p <0.05). 3.5. CD31 immunofluorescence staining Imaging of hPDLSCs–hUVECs co-cultured in CPC-GelMA scaffolds was performed by CD31 immunostaining (Figure 4. A–C). HUVEC membranes were stained green for the endothelial marker CD31, while nuclei were counterstained blue with DAPI. Branch-like structures gradually increased from day 7 to day 21. The co-cultured cells within the GelMA hydrogel progressively formed vessel-like networks with prolonged culture. Figure 4. D demonstrates that the cumulative vessel-like branch length increased significantly over time (p < 0.05). Compared to day 7, the cumulative vessel length had risen 18-fold at day 21, reaching 17.12 ± 2.34 mm/mm². Figure 4. E reveals that the number of vessel junctions increased from day 7 to day 21 (p < 0.05). By day 21, the junction density had risen to 37.25 ± 3.14 vessels/mm², representing a 3.4-fold increase over the value recorded on day 7. 3.6. Complete periodontal tissue regeneration in vivo At 4 weeks post-surgery, the cross-sections view of rat alveolar bone defects were observed by Micro-CT (Figure 5. A-D). The buccal bone thickness adjacent to the mesial root of the second molar in all groups were observed. As shown in Figure 5. E, the CPC+GelMA-cell group (0.549±0.01 mm) showed the greatest new formed bone thickness, which were 1.3 folds that of CPC-GelMA group (0.421±0.02 mm) and 2.57 folds that of the blank group (0.214±0.02 mm). The new bone area fraction (Figure 5. F) was also highest in the CPC+GelMA-cell group (44.33±3.01 %), reaching 1.23 folds that of the CPC-GelMA group (35.92±2.52 %) and 1.43 folds that of the blank group (30.99±1.78 %). For H&E staining, all groups newly formed bone (NB) and blood vessels (V) in defects, with no evident inflammation or immune reaction (Figure 6). New bone with a typical organized bone morphology was formed. New blood vessels were observed around the new bone. Osteoblasts with blue cytoplasm and round-to-oval nuclei lined the surfaces of the new bone. Compared with the Natural Periodontium group (Figure 6. G, H), the blank group (Figure 6. A, B) showed only sparse alveolar bone regeneration on the buccal side of the mesial root of the second molar. The CPC+GelMA group (Figure 6. C, D) exhibited moderate, irregularly distributed new bone and vessels. The CPC+GelMA-cell group (Figure 6. E, F) displayed the most abundant and irregularly distributed new bone and vessels, with active osteoblasts. As shown in Figure 7. I, CPC+GelMA-cell group (57.99±4.78 %) formed the highest new bone area that was 1.37 folds that of the CPC+GelMA group (42.26±5.48 %) and 2.29 folds that of the blank group (25.33±5.25 %). The new vessel density in CPC+GelMA-cell group (41.32±2.95 vessels/mm 2 ) were about 2.65 folds that of CPC-GelMA group (41.32±2.95 vessels/mm 2 ) and 6.2 folds that of the Blank group (41.32±2.95 vessels/mm 2 ) (Figure 6. J). Masson staining effectively reveals the maturity of collagen within bone tissue: mature collagen stains red, whereas newly formed bone or immature bone stains blue. Relative to the Natural Periodontium group (Figure 7. G, H), all three intervention groups exhibited immature bone within the defect. The Blank group (Figure 7. A, B) showed sparse blue trabeculae embedded in abundant fibrous tissue. In contrast, the CPC+GelMA-cell group (Figure 7. E, F) and CPC+GelMA group (Figure 7. C, D) produced extensive, irregularly arranged immature bone that appeared predominantly blue, with scattered foci of mature red-stained matrix. The area of blue-stained immature bone (Figure 7. I) in CPC+GelMA-cell group was highest (56.38±3.47 %), which was 1.29 folds that of CPC+GelMA group (43.59±2.77 %) and 2.38 folds that of Blank group (23.7±1.33 %). For immunohistochemical images of rat alveolar bone defects, both angiogenic marker CD31 (Figure 8. A) and osteogenic marker Runx2 (Figure 8. B) were expressed in the periodontal defects in all groups at 4 weeks after transplantation, compared with the negative-control group. The proportion of CD31-positive cells in the CPC+GelMA-cell group (2.387±0.027 per slice) was also much higher than those in the CPC+GelMA (2.061±0.035 per slice) and Blank (1.639±0.028 per slice) groups, reaching 1.16-fold and 1.46-fold higher values (Figure 8. C). Moreover, the newly formed periodontal tissues in the CPC+GelMA-cell group contained the highest proportion of Runx2-positive cells (2.471±0.022 per slice), being 1.17-fold and 1.91-fold greater than in the control (2.112 ± 0.038 per slice) and blank (1.675 ± 0.027 per slice) groups, respectively (Figure 8. D). 3.7. S ubcutaneous transplantation in vivo Figure 9 plots the images of stereomicroscopic view and H&E staining after subcutaneous implantation in nude rats at 6 weeks. Neovessels were visible on the surface of both groups. Histological evaluation revealed newly formed vessels within the macropores of both scaffolds. The vessels density of CPC+GelMA-cell group (Figure 9. B) was markedly higher than CPC+GelMA group (Figure 9. A). Morphometric analysis (Figure 9. C) demonstrated that the vessels density in the CPC+GelMA-cell group reached 81.33 ± 3.54 vessels/mm², a 1.93-fold increase over the 42.21 ± 5.84 vessels/mm² measured in the CPC+GelMA group (p<0.05). 4. Discussion The present study is the first to combine a 3D printed CPC framework with a GelMA-hPDLSCs-hUVECs system to fabricate a prevascularized CPC scaffold for alveolar bone regeneration. The hypotheses were proven that the novel construct had good mechanical properties, pore structure, biocompatibility and angiogenic capability in vitro. Compared with non-prevascularized CPC controls, the prevascularized scaffold significantly enhanced new bone and vessels formation in vivo. Our results showed that the isolated cells highly expressed STRO-1, CD90, and CD105, with low expression of CD34 and CD45, consistent with the characteristics of MSCs and thus considered as hPDLSCs [34]. Yeasmin et al [35] found that hPDLSCs can secrete angiogenesis-related factors such as VEGF and FGF, and provide stable support for endothelial cell networks. These findings further demonstrated that hPDLSCs, as seed cells, have significant vascularization potential in co-culture systems. Co-culture systems, through heterotypic cell–cell interactions, demonstrate a synergistic augmentation of pro-angiogenic growth factor output compared to monocultures [35]. MSCs not only provide trophic support to ECs but are also reciprocally activated by EC-derived signals, leading to further upregulation of their own growth factor secretion [35]. Monocultured MSCs are unable to self-assemble into microvascular structures, whereas co-culture with ECs rapidly initiates capillary-like network formation [10]. In the present study, hUVECs and hPDLSCs were co-seeded at a fixed ratio of 3:1. It has been demonstrated that a 1:1 ratio of hUVECs to MSCs generates stable vascular networks while simultaneously facilitating osteoid matrix deposition [15]. Conversely, hPDLSCs have been reported to proliferate significantly faster than hUVECs [11]. To offset this imbalance, initial EC proportions have been raised to 3:1 or even 5:1 (hUVEC:hPDLSC) in prior work [36]. Longitudinal profiling confirms a progressive decline in the relative abundance of hUVECs during extended co-culture [37]. Therefore, the 3:1 seeding ratio chosen here is expected to counterbalance the differential expansion rates and stabilize the two populations as the assay proceeds. Nevertheless, systematic optimization studies are still required to definitively establish the ideal hUVECs:hPDLSCs numerical ratio for maximal vasculogenic output in co-culture systems. GelMA hydrogels provide a cytocompatible extracellular environment, yet their bioactivity is dictated by GelMA concentration, photoinitiator content, and UV dose [38, 39]. Increasing GelMA density elevates cross-linking efficiency and network tightness. When the GelMA concentration exceeds 10% (w/v), pore size and swelling ratio decline, restricting cell proliferation and hindering the diffusion of oxygen, ultimately impairing cellular metabolism and phenotypic function [40]. Conversely, GelMA below 5% (w/v) remains high biocompatibility but lacks adequate mechanical integrity [40]. Comparative studies reveal that 5% (w/v) GelMA yields larger pores than its 10% (w/v) counterpart, facilitating Ca²⁺ flux and diffusion without compromising viability, and confers superior in vitro osteogenic potential [41]. Therefore, 5% (w/v) GelMA hydrogel was selected as the cell-laden matrix in the present study. CCK-8 assays revealed a robust and sustained proliferative profile of the co-cultured cells within the 5% GelMA matrix. Additionally, the live cell percentage was 80% on day 1 and exceeded 90% at every subsequent time point. The brief drop in live-cell number on day 1 is likely attributable to residual unreacted functional groups or small-molecular by-products released during GelMA cross-linking [42]. Future work should therefore optimize cross-linking parameters such as lowering photoinitiator concentration or shortening UV exposure to curb radical formation and improve early survival. Moreover, GelMA’s intrinsic drug-loading capacity and interconnected porosity enable sustained, tunable release of bioactive cargos [43]. With this benefit, future studies should encapsulate pro-angiogenic factors within GelMA-based co-culture systems, thereby integrating controlled growth-factor delivery with cellular paracrine signaling to enhance vascularization. Nevertheless, the inherently low mechanical strength of GelMA hydrogel limits its standalone use in load-bearing alveolar defects [29]. Consequently, recent studies have focused on integrating hydrogels with high-strength scaffolds to generate composites that unite excellent bioactivity with sufficient mechanical competence [29]. The calcium-phosphate cement (CPC) paste used in the present study has been extensively validated as a biocompatible and osteoconductive bone substitute [44-48]. Therefore, we combined GelMA hydrogel co-encapsulating hPDLSCs and hUVECs with a 3D-printed CPC scaffold to achieve sufficient mechanical competence to withstand physiological masticatory loads. Flexural strength and elastic modulus are critical mechanical indices for evaluating scaffold performance. Flexural strength and elastic modulus are critical design parameters [49]. Flexural strength (σ) is defined as the maximum stress a material can sustain before fracture, whereas the elastic modulus (E) quantifies the stress required per unit strain in the elastic region [49]. In human cancellous bone, flexural strength is approximately 3.5 MPa and the elastic modulus is 0.30 GPa, respectively [50] . Three-point bending test indicates that the scaffold showed better flexural strength and elastic modulus than those of cancellous bone. Thus, the scaffold offers adequate mechanical strength and dimensional stability for load-bearing alveolar repairs. In 3D printed scaffolds designed for vascular regeneration, filament diameter, pore size and filament intersection angle are determinants of pro-angiogenic performance [51]. Pores between 200 and 400 µm allow efficient cell migration and nutrient exchange [52]. Pores below 100 µm hinder cell infiltration and capillary ingrowth [53]. Intrinsic micropores of CPC are typically less than 50 µm and poorly interconnected, which limiting the migration and interaction of cells [54, 55]. Given that human microvessels range from 5 to 200 µm in diameter [51] , the 302.31 ± 3.4 µm pore size generated in the printed CPC scaffold readily accommodate nascent microvessels and their interconnection, facilitating 3D vascular network assembly. Notably, 300 µm pores have been specifically identified as optimal for alveolar bone regeneration [19]. Moreover, scaffolds of 300 µm-pore size honeycomb architecture concurrently drive bone and vessel formation in vivo [56]. Moreover, similar filament diameter and pore size have been shown to homogenize stress distribution and markedly improve scaffold strength [57]. In the present work, the printed CPC displays comparable filament and pore sizes, providing a regular architecture conducive to cell attachment and growth. In addition, the intersection angle between filaments influences both mechanical integrity and cell alignment [23] . An appropriate angle can enhance scaffold strength and stability while promoting uniform cell distribution. For bone engineering, CPC plots between 15° and 90° configurations are commonly investigated [23]. Previous study demonstrated that a 60° intersection angle yielded higher compressive modulus and strength than a 45° pattern [58]. Scaffolds printed with a 60° filament intersection angle likewise exhibited superior mechanical performance in rat femoral defects [44]. This advantage is attributed to the angle’s balanced stability, which simultaneously supports multidirectional cell signaling and nutrient exchange [44]. The CPC scaffolds described here were printed at 60.12 ± 0.4°, closely matching the reported optimum. Nevertheless, the definitive intersection angle for 3D-printed CPC remains to be established and demands systematic exploration. CD31 immunostaining was used to quantify in vitro angiogenesis of hUVECs co-cultured with hPDLSCs inside GelMA-CPC scaffolds. Cumulative vessel length and junction number are standard metrics of neovascularization: length reflects network expansion, whereas junction density mirrors architectural complexity and functional anastomosis [59]. Confocal imaging revealed microcapillary-like networks that elongated and interconnected over time. By day 21, the 3D GelMA-CPC co-culture system exhibited significantly greater cumulative vessel length and junction number than previously reported 2D CPC surface co-culture [11], underscoring the superior pro-angiogenic potential of the 3D co-culture system. The present in vivo studies demonstrated that prevascularized scaffolds have significant advantages in promoting angiogenesis and accelerating alveolar bone repair. Previous study demonstrated that prevascularized scaffolds can set up a functional blood supply by speeding up microvessel formation and stably connecting with host blood vessels, which greatly improving implant performance [60]. Currently, scaffold prevascularization mainly uses two strategies: in vivo and in vitro. In vivo prevascularization involves implanting the scaffold into a well-vascularized site (e.g. muscle) for several weeks to form a microvessel network, but it requires an invasive procedure [61]. In contrast, in vitro prevascularization avoids a second operation and donor-site morbidity, offering a minimally invasive, surgeon-friendly route that is more suitable for clinical use [61]. Researchers first co-cultured endothelial cells with osteoprogenitor cells and generated a 3D vascular network in vitro [62]. Inspired by this, we built a prevascularized scaffold in a 3D co-culture system and tested its angiogenic potential by subcutaneous implantation in nude rat. Because nude rats are immunocompromised, xenograft rejection is sharply reduced, which improving graft survival and the likelihood of vessel formation [63]. At week 6, vessel density inside the scaffolds reached 81.33 ± 3.54 vessels/mm², matching earlier reports that prevascularized constructs accelerate perfusion [60]. In the previous report, the fully prevascularized implants typically reach 80% vascular coverage within 6 weeks, whereas non-prevascularized controls achieve only 50% [60]. This advantage stems from the pre-built vessels acting as “highways” that pull in more capillary sprouts, culminating in a denser and more durable microvascular network throughout the scaffold [60]. Alveolar bone regeneration was evaluated in all groups in vivo. By week 4, new bone was evident in every defect. However, the CPC+GelMA-cell group produced significantly more bone than either CPC+GelMA or Blank groups. Micro-CT gave a mineralized-bone fraction of 44.33 ± 3.01 % in the CPC+GelMA-cell group—23 % higher than CPC+GelMA group—whereas semiquantitative histomorphometry of H&E sections yielded 57.99 ± 4.78 % total new bone area, 1.37-fold that of CPC+GelMA group. The discrepancy reflects the fact that Micro-CT records only mineralized tissue, whereas H&E staining also captures unmineralized osteoid [64]. Newly formed trabeculae stained with H&E appear as irregular nets or cords. The 3D scaffold apparently influenced this trabecular arrangement. Nevertheless, H&E cannot distinguish immature bone from mature lamellar bone, so Masson staining was additionally performed to verify the maturity of the regenerated tissue [65]. Masson staining reliably visualizes collagen remodeling in bone tissues [65]. Newly formed bone is rich in type I collagen and stains blue. As mineralization proceeds, the stained color shifts toward red. This color transition is routinely exploited to gauge maturation, with the nascent-to-mature conversion typically occurring between 4 and 8 weeks post-operatively [65]. We therefore harvested specimens at week 4 and used the area fraction of blue-stained bone as a surrogate for new bone formation. Masson images revealed that the CPC+GelMA-cell group contained the largest amount of immature bone (56.38 ± 3.47 %), representing 1.29- and 2.38-fold increases over the CPC+GelMA and Blank groups, respectively, indicating that the defect area was undergoing active osteogenesis and that the experimental intervention accelerated this process. Due to Masson and H&E staining provide limited information on cellular and vascular components, IHC staining was employed for a more comprehensive evaluation. CD31, an endothelial-specific marker, was used to quantify angiogenesis [66], whereas Runx2, the master transcription factor driving osteoblast differentiation, was used to assess osteoblastic activity and maturity [67]. Semi-quantitative IHC demonstrated that Runx2 expression in the CPC+GelMA-cell group was 1.17-fold that of CPC+GelMA group, corroborating the superior osteogenic potential observed histologically. Similarly, a previous study demonstrated that scaffold pre-seeded with cells generated significantly more new bone than its cell-free counterpart, at 4 weeks in a rat cranial bone defect model [68]。 Moreover, a significant positive correlation between osteogenesis and angiogenesis was evident [68] . H&E morphometry revealed a neovessel density of 41.32 ± 2.95 vessels/mm² in the CPC+GelMA-cell group, 2.65-fold that of CPC+GelMA group, accompanied by a 16% increase in CD31 immunostained area. These data corroborate earlier reports that bone formation rate rises in parallel with vascular ingrowth [69, 70]. This coupling is likely initiated by the hypoxic milieu that develops immediately after scaffold implantation. Under low oxygen tension, co-cultured cells activate hypoxia-inducible factor-1α (HIF-1α) pathway, which transcriptionally up-regulates vascular endothelial growth factor (VEGF) and fibroblast growth factor-2 (FGF-2). While promoting angiogenesis, these cytokines simultaneously deliver the nutrients and osteoinductive signals required for effective bone repair [71, 72]. Additionally, PI3K/Akt [73] and MAPK [74] signaling pathways participate in hypoxia-induced angiogenesis, which can modulate cell proliferation, survival, and differentiation to accelerate bone repair and regeneration. In addition to the contribution of co-cultured cells, the intrinsic properties of the scaffold itself critically influence osteogenesis and angiogenesis [75]. In the present study, the CPC+GelMA group exhibited substantially more de novo bone than the empty defect（Blank group）. This effect is attributable to the osteoinductive capacity of CPC, which sequesters endogenous growth factors and directs multipotent stem cells toward an osteoblastic phenotype [76]. In earlier work, traditional 2D CPC placed in cranial defects for 8 weeks regenerated only 13.89 ± 2.95 % new bone [77]，while traditional 2D CPC packed into alveolar socket yielded less than 5% repair by week 12 [78]. In contrast, micro-CT of our 3D CPC scaffold revealed 35.92 ± 2.52 % new alveolar bone within only 4 weeks. This pronounced disparity is presumably attributable to differences in defect site and internal scaffold structure [79]. In addition, porogen-leavened macroporous CPC [79], collagen-sponge CPC [78] and CPC fortified with SP [78] or rhBMP-2 [80] have respectively doubled to quintupled new bone output within 4 weeks. Thus, compositional and architectural refinement can readily enhance CPC-mediated alveolar regeneration. In the field of vascularized scaffolds, bioactive glass [81] and silk fibroin [82] have attracted intense attention because of their superior pro-angiogenic capacity. Concurrently, microfluidic technologies are opening new routes for the fabrication of individualized constructs [83]. Smart stimulus-responsive drug-delivery systems that release therapeutics under endogenous or exogenous factors further enhance therapeutic efficacy [84]. Collectively, these innovations offer great clinical potential for more effective tissue repair and regeneration. The periodontium’s intricate architecture reveals that alveolar bone, periodontal ligament and cementum are structurally and functionally interdependent to ensure tooth stability [85]. Therefore, future research should focus on engineering hierarchically layered multi-phase scaffolds that faithfully replicate the native periodontium. 5. Conclusion This study developed a novel 3D printed CPC scaffold to establish a prevascularized 3D construct for alveolar bone engineering. The resulting construct had proper mechanical properties and pore structure of the CPC skeleton. The delivered hPDLSCs-hUVECs exhibited excellent cell proliferation and angiogenic potential in GelMA hydrogel in vitro. When implanted in vivo, the prevascularized CPC+GelMA-cell group elicited significantly greater neovascularization and accelerated alveolar-bone regeneration compared with the cell-free CPC+GelMA control. The intrinsic properties of CPC also enhance osteogenesis and angiogenesis. Therefore, the novel CPC-GelMA-hPDLSCs-hUVECs construct is highly promising for concurrent bone and vascular regeneration in dental applications. Declarations Ethics approval and consent to participate Capital Medical University Ethics Committee approved this study (CMUSH-IRB-KJ-PJ-2024-28). All participants agreed to participate in this study and signed an informed consent form. Consent for publication Not applicable. Availability of data and materials All data generated or analysed during this study are included in this published article [and its supplementary information files]. Competing interests The authors declare no competing interests. Funding This study was supported by National Natural Science Foundation of China (No. 82301117), the Innovation Research Team Project of Beijing Stomatological Hospital, Capital Medical University (Grant No. CXTD202203) and Beijing Stomatological Hospital, Capital Medical University Young Scientist Program (No. YSP202510). Authors' contributions Y. Sun conducted the investigation, performed visualization and validation. Z. Zhao contributed to methodology and data curation, and drafted the original manuscript. Q. Qiao was responsible for visualization and methodology. conceptualized the study. W. Yu contributed to validation. Y. Bai participated in writing - review and editing, project administration, funding acquisition. All authors read and approved the final manuscript. Acknowledgement We are grateful to the patients who are willing to participate in the study. References Wishney M. 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PI3K/Akt and HIF‑1 signaling pathway in hypoxia‑ischemia (Review) [J]. Mol Med Rep. 2018; 18: 3547-54. https://doi.org/10.3892/mmr.2018.9375. Chen J, Chen J, Cheng Y, et al. Mesenchymal stem cell-derived exosomes protect beta cells against hypoxia-induced apoptosis via miR-21 by alleviating ER stress and inhibiting p38 MAPK phosphorylation [J]. Stem Cell Res Ther. 2020; 11: 97. https://doi.org/10.1186/s13287-020-01610-0. Thein-Han W, Xu H H. Prevascularization of a gas-foaming macroporous calcium phosphate cement scaffold via coculture of human umbilical vein endothelial cells and osteoblasts [J]. Tissue Eng Part A. 2013; 19: 1675-85. https://doi.org/10.1089/ten.TEA.2012.0631. Tsai Y H, Tseng C C, Lin Y C, et al. Novel artificial tricalcium phosphate and magnesium composite graft facilitates angiogenesis in bone healing [J]. Biomed J. 2024: 100750. https://doi.org/10.1016/j.bj.2024.100750. Zhu C, Chang Q, Zou D, et al. LvBMP-2 gene-modified BMSCs combined with calcium phosphate cement scaffolds for the repair of calvarial defects in rats [J]. J Mater Sci Mater Med. 2011; 22: 1965-73. https://doi.org/10.1007/s10856-011-4376-6. Wang T, Wu D, Li Y, et al. Substance P incorporation in calcium phosphate cement for dental alveolar bone defect restoration [J]. Mater Sci Eng C Mater Biol Appl. 2016; 69: 546-53. https://doi.org/10.1016/j.msec.2016.07.014. Lee K, Weir M D, Lippens E, et al. Bone regeneration via novel macroporous CPC scaffolds in critical-sized cranial defects in rats [J]. Dent Mater. 2014; 30: e199-207. https://doi.org/10.1016/j.dental.2014.03.008. Francis C S, Mobin S S N, Lypka M A, et al. rhBMP-2 with a demineralized bone matrix scaffold versus autologous iliac crest bone graft for alveolar cleft reconstruction [J]. Plast Reconstr Surg. 2013; 131: 1107-15. https://doi.org/10.1097/PRS.0b013e3182865dfb. Xu H, Zhu Y, Hsiao A W, et al. Bioactive glass-elicited stem cell-derived extracellular vesicles regulate M2 macrophage polarization and angiogenesis to improve tendon regeneration and functional recovery [J]. Biomaterials. 2023; 294: 121998. https://doi.org/10.1016/j.biomaterials.2023.121998. Durán-Rey D, Brito-Pereira R, Ribeiro C, et al. Development of Silk Fibroin Scaffolds for Vascular Repair [J]. Biomacromolecules. 2023; 24: 1121-30. https://doi.org/10.1021/acs.biomac.2c01124. Quintard C, Tubbs E, Jonsson G, et al. A microfluidic platform integrating functional vascularized organoids-on-chip [J]. Nat Commun, 2024; 15: 1452. https://doi.org/10.1038/s41467-024-45710-4. Guo Z, Bo D, He P, et al. Sequential controlled-released dual-drug loaded scaffold for guided bone regeneration in a rat fenestration defect model [J]. J Mater Chem B. 2017; 5: 7701-10. https://doi.org/10.1039/c7tb00909g. Qin W, Li L, Mu Z, et al. A hierarchical Bilayered scaffold for periodontal complex structure regeneration [J]. J Biomed Mater Res A. 2025; 113: e37793. https://doi.org/10.1002/jbm.a.37793. Additional Declarations No competing interests reported. Cite Share Download PDF Status: Under Review Version 1 posted Editorial decision: Revision requested 26 Feb, 2026 Reviews received at journal 18 Feb, 2026 Reviews received at journal 13 Feb, 2026 Reviewers agreed at journal 11 Feb, 2026 Reviewers agreed at journal 05 Feb, 2026 Reviews received at journal 04 Feb, 2026 Reviewers agreed at journal 04 Feb, 2026 Reviewers agreed at journal 27 Dec, 2025 Reviews received at journal 26 Dec, 2025 Reviewers agreed at journal 25 Dec, 2025 Reviewers invited by journal 25 Dec, 2025 Editor assigned by journal 24 Dec, 2025 Editor invited by journal 15 Dec, 2025 Submission checks completed at journal 12 Dec, 2025 First submitted to journal 12 Dec, 2025 You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. 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Also discoverable on Platform About Our Team In Review Editorial Policies Advisory Board Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {"props":{"pageProps":{"initialData":{"identity":"rs-8260921","acceptedTermsAndConditions":true,"allowDirectSubmit":false,"archivedVersions":[],"articleType":"Research Article","associatedPublications":[],"authors":[{"id":565629408,"identity":"16973c5f-e70f-419f-8581-d71780b07431","order_by":0,"name":"Yaxi Sun","email":"","orcid":"","institution":"Capital Medical University","correspondingAuthor":false,"prefix":"","firstName":"Yaxi","middleName":"","lastName":"Sun","suffix":""},{"id":565629414,"identity":"062a6ea7-fee8-45e4-9d8e-b730489d88a1","order_by":1,"name":"Zeqing Zhao","email":"","orcid":"","institution":"Capital Medical University","correspondingAuthor":false,"prefix":"","firstName":"Zeqing","middleName":"","lastName":"Zhao","suffix":""},{"id":565629419,"identity":"049c0abb-9863-43c4-8a27-2ec9e00ce53c","order_by":2,"name":"Qingchen Qiao","email":"","orcid":"","institution":"Capital Medical University","correspondingAuthor":false,"prefix":"","firstName":"Qingchen","middleName":"","lastName":"Qiao","suffix":""},{"id":565629428,"identity":"69cbb336-f0d3-4be4-ab9f-93a7a732f15b","order_by":3,"name":"Wenting Yu","email":"","orcid":"","institution":"Capital Medical University","correspondingAuthor":false,"prefix":"","firstName":"Wenting","middleName":"","lastName":"Yu","suffix":""},{"id":565629429,"identity":"a482b8c4-a2c7-410f-a567-bfb211a9d8bf","order_by":4,"name":"Yuxing Bai","email":"data:image/png;base64,iVBORw0KGgoAAAANSUhEUgAAAZAAAAAyAQMAAABI0h/eAAAABlBMVEX///8AAABVwtN+AAAACXBIWXMAAA7EAAAOxAGVKw4bAAAA0ElEQVRIiWNgGAWjYBACAxCRwMAgB+UzE6/FmEQtQJDYQLQWc4kcM4mHO2rT+9tPp0kwVFgnNrCfPYBXi+WMHGODxDPHc2ecyd0mwXAmPbGBJy8Bv8Nu5Bg+SGw7lrtBgnebBGPb4cQGCR4DQloMDgC1pBuAtfwjTgvIlpoEiJYGIrRY9jwrNkhsO2AI9Mtmi4Rj6cZtPDn4tZizJ2+T/NlWJ8/ffnbjjQ811rL97Gfwa2EQSACRhyEcEJsNv3og4D8AIusIqhsFo2AUjIIRDADlhUTigb2mkQAAAABJRU5ErkJggg==","orcid":"","institution":"Capital Medical University","correspondingAuthor":true,"prefix":"","firstName":"Yuxing","middleName":"","lastName":"Bai","suffix":""}],"badges":[],"createdAt":"2025-12-02 13:21:14","currentVersionCode":1,"declarations":"","doi":"10.21203/rs.3.rs-8260921/v1","doiUrl":"https://doi.org/10.21203/rs.3.rs-8260921/v1","draftVersion":[],"editorialEvents":[],"editorialNote":"","failedWorkflow":false,"files":[{"id":99259711,"identity":"7972664f-054d-4b26-b4ec-50e5664b3e94","added_by":"auto","created_at":"2025-12-31 01:13:15","extension":"png","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":10694043,"visible":true,"origin":"","legend":"\u003cp\u003e(A) Flow cytometry results of the isolated hPDLSCs: 99.6% CD105-positive cells, 76.4% CD90-positive cells, 88.6% STRO-1-positive cells, 0.3% CD34-positive cells, 0.6% CD45-positive cells. (B)-(C) Three-dimensionally printed CPC scaffold. (D) Three-dimensionally prevascularized CPC scaffold.\u003c/p\u003e","description":"","filename":"Figure1.png","url":"https://assets-eu.researchsquare.com/files/rs-8260921/v1/be6f092e73c55155c89eeb7f.png"},{"id":99259715,"identity":"b4431436-2c7b-485e-a582-ba01a13ff4be","added_by":"auto","created_at":"2025-12-31 01:13:15","extension":"png","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":6009390,"visible":true,"origin":"","legend":"\u003cp\u003ePhysical properties of 3D-printed CPC constructs. (A)-(C) SEM images of 3D-printed CPC constructs (n=6). (D) Flexural strength. (E) Elastic modulus. Values with dissimilar letters are significantly different from each other (p \u0026lt;0.05). Each value is mean±SD (n=6).\u003c/p\u003e","description":"","filename":"Figure2.png","url":"https://assets-eu.researchsquare.com/files/rs-8260921/v1/cacb8274678cf04db703f437.png"},{"id":99259712,"identity":"bbf5758e-e338-4927-81cd-15a3939077d2","added_by":"auto","created_at":"2025-12-31 01:13:15","extension":"png","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":11607760,"visible":true,"origin":"","legend":"\u003cp\u003eViability and cell proliferation of cells co-cultured within GelMA-CPC constructs versus time. (A) Live/dead staining and fluorescence microscopy of cell release from GelMA. At day 1, cells were embedded within the hydrogel, predominantly spherical and clustered. At day 4, cell number slightly increased. At day 7, cells began to aggregate, with a noticeable increase in number and a predominance of short spindle shapes. At day 14, cells were evenly distributed throughout the hydrogel and displayed long spindle or polygonal morphologies. At day 21, all cells had fully extended into long spindle shapes, forming network-like structures. At each time points tested, there were numerous live cells (green staining) and only a few dead ones (red staining). (B) Representative the percentages of live cells in 21 days. (C) CCK-8 assessment in 21 days. Values indicated by dissimilar letters are significantly different from each other (p \u0026lt;0.05). Each value is mean ±SD (n=6).\u003c/p\u003e","description":"","filename":"Figure3.png","url":"https://assets-eu.researchsquare.com/files/rs-8260921/v1/fdf069ebcbefec62bfcc2fd1.png"},{"id":99318374,"identity":"7b52b540-ec74-40bd-a278-11e1f0aeb96a","added_by":"auto","created_at":"2025-12-31 16:32:57","extension":"png","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":11748739,"visible":true,"origin":"","legend":"\u003cp\u003eCD31 immunostaining of hPDLSCs–hUVECs co-cultured in CPC-GelMA scaffolds. (A)–(C) The formation of branch-like structures formed by hUVECs and hPDLSCs co-cultured in GelMA hydrogel. HUVECs were identified by immunostaining with endothelial marker CD31 in green on the cell membrane, and the nuclei were stained with DAPI in blue. HPDLSCs were depicted by nuclei counterstaining with DAPI in blue but without green stains on the cell membrane. (D) Quantification the vessel length of co-culture cells. (E) Quantification the vessel junctions of co-culture cells. Microcapillary-like structures increased with culture time. The vessel length and vessel junction number of co-culture cells increased with time. Values indicated by dissimilar letters are significantly different from each other (p \u0026lt;0.05).\u003c/p\u003e","description":"","filename":"Figure4.png","url":"https://assets-eu.researchsquare.com/files/rs-8260921/v1/3311e7032f696d1bafff0d76.png"},{"id":99259714,"identity":"85ce7f78-4988-4918-96d8-159769aa7c87","added_by":"auto","created_at":"2025-12-31 01:13:15","extension":"png","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":5923537,"visible":true,"origin":"","legend":"\u003cp\u003eMicro-CT evaluation of rat alveolar-bone defect repair. (A)-(D) Representative cross-sectional views of the defect region in each group. (E) Newly formed bone thickness within the defect. (F) The new bone area fraction within the defect. Values indicated by dissimilar letters are significantly different from each other (p\u0026lt;0.05).\u003c/p\u003e","description":"","filename":"Figure5.png","url":"https://assets-eu.researchsquare.com/files/rs-8260921/v1/9768e08995d0f253e539b77f.png"},{"id":99259719,"identity":"ceab2524-f83b-4385-ab80-467ea6881edc","added_by":"auto","created_at":"2025-12-31 01:13:16","extension":"png","order_by":6,"title":"Figure 6","display":"","copyAsset":false,"role":"figure","size":30977461,"visible":true,"origin":"","legend":"\u003cp\u003eRepresentative H\u0026amp;E images of alveolar bone defects in rats. Compared to the Natural Preiodontium group (G), limited new bone and vessels were found In Blank group (A). The CPC+GelMA group (C) exhibited moderate, irregularly distributed new bone and vessels in the defects. The CPC+GelMA-cell group (E) had the largest amount of new bone and vessels. (B), (D), (F), (H) is a higher magnification image of the defect section in (A), (C), (E), (G), respectively. (I) Fraction of new bone areas. (J) New vessel density. Values with dissimilar letters are significantly different from each other (p \u0026lt;0.05). Each value is mean ±SD (n=5).\u003c/p\u003e","description":"","filename":"Figure6.png","url":"https://assets-eu.researchsquare.com/files/rs-8260921/v1/d556a0556f3704196b447785.png"},{"id":99259718,"identity":"6f180ee3-c1a3-4f8d-ba45-0f2549eababf","added_by":"auto","created_at":"2025-12-31 01:13:16","extension":"png","order_by":7,"title":"Figure 7","display":"","copyAsset":false,"role":"figure","size":35517993,"visible":true,"origin":"","legend":"\u003cp\u003eRepresentative Masson staining images of alveolar bone defects in rats. Relative to the Natural Periodontium group (G), Blank group (A) showed sparse blue trabeculae embedded in abundant fibrous tissue. In contrast, the CPC+GelMA-cell group (E) and CPC+GelMA group (C) produced extensive, irregularly arranged immature bone that appeared predominantly blue, with scattered foci of mature red-stained matrix. (B), (D), (F), (H) is a higher magnification image of the defect section in (A), (C), (E), (G), respectively. NB indicates newly formed bone, R indicates root, blue staining indicates immature bone, red staining indicates mature bone. (I) Fraction of immature bone area. Values indicated by dissimilar letters are significantly different from each other (p \u0026lt;0.05).\u003c/p\u003e","description":"","filename":"Figure7.png","url":"https://assets-eu.researchsquare.com/files/rs-8260921/v1/885678c2ea31017dd5cf7da1.png"},{"id":99259716,"identity":"659b19e1-ddd2-449d-8705-8a26f3a0dd00","added_by":"auto","created_at":"2025-12-31 01:13:16","extension":"png","order_by":8,"title":"Figure 8","display":"","copyAsset":false,"role":"figure","size":18572193,"visible":true,"origin":"","legend":"\u003cp\u003e(A) Immunohistochemical staining of CD31 after 4 weeks implantation. (B) Immunohistochemical staining of Runx2 after 4 weeks implantation. (C) Semi-quantitative analysis of CD31-positive cell numbers per slice of (A). (D) Semi-quantitative analysis of Runx2-positive cell numbers per slice of (B). Values indicated by dissimilar letters are significantly different from each other (p \u0026lt;0.05).\u003c/p\u003e","description":"","filename":"Figure8.png","url":"https://assets-eu.researchsquare.com/files/rs-8260921/v1/e1ae5dc1f8ef4a7f1690bf01.png"},{"id":99259717,"identity":"9e977072-0f7c-48ad-b7b9-b48d12fee548","added_by":"auto","created_at":"2025-12-31 01:13:16","extension":"png","order_by":9,"title":"Figure 9","display":"","copyAsset":false,"role":"figure","size":28740549,"visible":true,"origin":"","legend":"\u003cp\u003eIn vivo subcutaneous transplantation in rats at 6 weeks (A) Stereomicroscopic view and H\u0026amp;E staining images of CPC+GelMA group. (B) Stereomicroscopic view and H\u0026amp;E staining images of the CPC+GelMA-cell group. (C) Semi-quantitative analysis of the vessel density of CPC+GelMA and CPC+GelMA-cell groups. Values indicated by dissimilar letters are significantly different from each other (p \u0026lt;0.05).\u003c/p\u003e","description":"","filename":"Figure9.png","url":"https://assets-eu.researchsquare.com/files/rs-8260921/v1/99f570dd7fc1b7caabd252a1.png"},{"id":99324311,"identity":"cfb8f625-78fc-42e9-b6f4-126062a74a86","added_by":"auto","created_at":"2025-12-31 16:47:15","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":144217959,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-8260921/v1/8c930c89-1f70-4f32-aa16-29c3fbe9fde8.pdf"}],"financialInterests":"No competing interests reported.","formattedTitle":"A 3D Prevascularized Calcium Phosphate Cement Scaffold for Accelerated Alveolar Bone Regeneration and Angiogenesis in Rats","fulltext":[{"header":"1. Background","content":"\u003cp\u003eOrthodontic treatment carries the risk of periodontal tissue damage. Studies indicate that over one-third of adult patients exhibit anterior alveolar bone recession exceeding 2 mm post-treatment\u0026nbsp;[1]. Improper orthodontic design may also lead to alveolar bone defects such as fenestration and dehiscence\u0026nbsp;[1]. Tissue engineering scaffolds are a critical tool for reconstructing alveolar bone defects, yet their regenerative potential is critically constrained by inadequate and delayed vascularization [2]. Insufficient blood supply can cause hypoxia, impair osteogenesis, and lead to graft necrosis and failure [2, 3]. Therefore, overcoming insufficient vascularization is essential for bone-defect repair.\u003c/p\u003e\n\u003cp\u003ePrevascularized scaffolds enables their vascular networks to anastomose with the host’s vasculature after implantation, markedly reducing the risk of ischemic necrosis, and substantially increasing the success rate of bone regeneration\u0026nbsp;[4].\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eCalcium phosphate cement (CPC) is an ideal osteogenic material due to its biocompatibility, osteoconductivity, and bone-like composition [5]. However, conventional CPC is predominantly microporous and lacks interconnected macroporous structures, hindering vascular ingrowth and limiting its utility in large bone defects [6]. Although prevascularized CPC scaffolds have been studied for cranial and long bone repair [7-10], a three-dimensional (3D) prevascularized CPC scaffold tailored for alveolar bone regeneration has not yet been reported.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eCalcium phosphate cement (CPC) is a self-setting osteogenic scaffold material, resembling the inorganic composition of natural bone\u0026nbsp;[5]. CPC regards as a highly promising scaffold material for craniofacial and dental repairs，due to its good biocompatibility, osteoconductivity, and mechanical properties\u0026nbsp;[5]. Nevertheless, unmodified CPC is predominantly microporous and lacks interconnected macroporous structures, hindering vascular ingrowth and limiting its utility in large bone defects [6]. Prevascularized CPC scaffolds may significantly enhance bone repair capabilities. Currently, research on prevascularized calcium-phosphate osteogenic scaffolds primarily focuses on cranial and long bone repair [7-10]. To date, there has been no report of constructing three-dimensional (3D) prevascularized CPC scaffolds for alveolar bone tissue engineering.\u003c/p\u003e\n\u003cp\u003ePrevascularization of CPC scaffolds is currently achieved through vascular\u0026nbsp;implantation [7], in vivo culturing [6], or cell-based in vitro prevascularization [11]. However, vascular implantation and in vivo culturing involve invasive procedures, limiting their clinical applicability. Therefore, cell-based in vitro prevascularization is now in the spotlight [12].\u0026nbsp;Human umbilical vein endothelial cells (hUVECs) readily self-assemble into microcapillaries, yet endothelial monocultures fail to generate stable, mature vascular structures [13].\u0026nbsp;Endothelial cell migration and\u0026nbsp;neovessel formation\u0026nbsp;require specific pro-angiogenic factors, which are insufficiently produced by endothelial cells in monoculture [4, 14]. Mesenchymal stem cells (MSCs) enhance angiogenesis by secreting pro-angiogenic factors and stabilizing nascent vessels as pericytes [13, 15, 16]. Numerous studies demonstrate that co-culturing MSCs with endothelial cells yields stable vascular structures [16].\u003c/p\u003e\n\u003cp\u003eHuman periodontal ligament stem cells (hPDLSCs) are a seed cell source that can be harvested from the extracted wisdom teeth or the teeth extracted for orthodontic purpose without additional invasive surgery for the patient. Previous studies have shown positive results in using hPDLSCs for tissue regeneration, especially in bone and periodontal tissue repairment [17, 18]. In addition,\u0026nbsp;hPDLSCs can differentiate into bone, nerve, connective tissue, and cementum under specific conditions\u0026nbsp;[19, 20]. Therefore, hPDLSCs are a potent cell source in stem cell delivery via scaffolds for bone regeneration, especially for alveolar bone repair.\u0026nbsp;Studies showed that co-culturing hPDLSCs and hUVECs on CPC surfaces can form microvascular-like structures, suggesting their potential for CPC scaffold prevascularization\u0026nbsp;[11].\u0026nbsp;To date, a literature search revealed no report on the\u0026nbsp;prevascularized CPC scaffolds seeded with hPDLSCs and hUVECs for alveolar bone defect repair.\u003c/p\u003e\n\u003cp\u003eBeyond seed cell selection, scaffold structural properties also influence prevascularization efficacy. Three-dimensional (3D) printing technology enables precise control over pore size and interconnectivity [21, 22]. 3D-printed grid-like CPC scaffolds\u0026nbsp;exhibit\u0026nbsp;uniform, interconnected micropores that readily\u0026nbsp;conducive to microvascular growth\u0026nbsp;support microvessel ingrowth and anastomosis [23]. Additionally, the culture method for seed cells is crucial for prevascularization. Traditional two-dimensional (2D) culture involves seeding cells directly onto scaffold surfaces, where restricted area and contact inhibition curtail expansion and capillary morphogenesis\u0026nbsp;[24, 25].\u0026nbsp;Research indicated that 3D microenvironments enhance cell responsiveness to biochemical signals during angiogenesis\u0026nbsp;[24].\u0026nbsp;Compared to 2D culture, 3D culture better mimics natural cell growth conditions, promoting cell-cell and cell-matrix interactions\u0026nbsp;[26].\u0026nbsp;Encapsulating cells in hydrogel-based materials can simulate extracellular matrix structures, providing 3D support\u0026nbsp;[24].\u0026nbsp;Gelatin methacryloyl (GelMA) hydrogel offers excellent biocompatibility and enables 3D cell encapsulation, simulating a natural extracellular matrix conducive to microvascular formation\u0026nbsp;[27].\u0026nbsp;However, hydrogels lack sufficient mechanical strength for load-bearing applications like alveolar bone repair\u0026nbsp;[28].\u0026nbsp;Recent studies combine hydrogels with high-strength scaffolds to create composite materials with both biological and mechanical advantages\u0026nbsp;[28].\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eTo address the challenges of slow vascularization and inadequate mechanical strength in current bone grafts, we aim to design a CPC-based, cell-laden 3D hydrogel composite. The primary goals are to engineer a scaffold with improved load-bearing capacity and to establish a preformed vascular network within it, ultimately boosting the regenerative outcomes in alveolar bone defects.\u003c/p\u003e"},{"header":"2. Methods","content":"\u003cp\u003e\u003cstrong\u003e\u003cem\u003e2.1. Harvesting hPDLSCs from extracted teeth\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003ePeriodontal ligament (PDL) tissues were\u0026nbsp;harvested from healthy premolars extracted from patients aged 18–26 years for orthodontic purpose. The hPDLSCs were isolated as described previously [29]. The procedures were approved by the Medical Ethics Committee of Beijing Stomatological Hospital, Capital Medical University (NO. CMUSH-IRB-KJ-PJ-2024-28). The written informed consent was obtained from each participant before the study. The study was carried out in accordance with the Declaration of Helsinki.\u003c/p\u003e\n\u003cp\u003eThe PDL tissues were enzymatically digested with 3 mg/mL collagenase type I (Gibco BRL, Grand Island, NY, USA) and 4 mg/mL dispase (Gibco BRL) at 37°C for 45 minutes.\u0026nbsp; After digestion, the cell suspension was collected and transferred to culture dishes (Costar, Cambridge, MA, USA) with dulbecco’s modified Eagle’s medium (DMEM, Gibco BRL) supplemented with 1% penicillin/streptomycin (P.S, Gibco BRL) and 20% fetal bovine serum (FBS, Gibco BRL). Upon reaching 70–80% confluence, cells were passaged using 0.25% trypsin-EDTA (Gibco BRL). Cells at passages 3–5 were used in subsequent experiments.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e2.2. Identification of hPDLSCs\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe expression of surface antigen profiles (CD34, CD45, CD90, CD105, and STRO-1) of passage 3-5 cells were analyzed by flow cytometry as described previously [29]. PE-conjugated antibodies against CD34, CD105, and STRO-1 (Thermo Fisher Scientific, Rockford, IL, USA), along with FITC-labeled CD45 and CD90 antibodies (Thermo Fisher Scientific), were employed for immunophenotyping. Cells were digested, centrifuged and resuspended in cold PBS to achieve a concentration of 1×10\u003csup\u003e5\u003c/sup\u003e cells/cube. For staining, the cell suspension was mixed with antibody and incubated on ice in dark. After\u0026nbsp;being washed and resuspended, cell surface antigen expression was tested using a BD Vantage flow cytometer (BD Biosciences).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e2.3. Culturing of hUVECs\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003ePrimary hUVECs were acquired from Sciencell (Carlsbad, CA, USA). The cells were cultured in 10 cm culture dishes with endothelial cell medium (ECM, Sciencell) under standard culture conditions [30].\u0026nbsp;Subsequent passages were performed\u0026nbsp;when confluency reached 70% to 80%.\u0026nbsp;Cells between passage 2–3 were used in this study.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e2.4.\u0026nbsp;\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003eFabrication of 3D prevascularized CPC scaffold\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe CPC paste was manufactured by InnoTERE\u0026nbsp;GmbH (Radebeul, Germany) and fabricated with the BioScaffolder 2.1 (GeSiM mbH, Radeberg, Germany) operated in a laminar flow workbench [31]. CPC paste was plotted in 60° configuration (the layer orientation changed after every second layer by 60°) and\u0026nbsp;a filament diameter of 300 μm with designed pore size of 300 μm.\u0026nbsp;Scaffolds used for a rat alveolar defect model were plotted a rectangular external structure with a size of\u0026nbsp;4 mm×2 mm×1 mm. A cylindrical outer geometry with a height of 3 mm and a diameter of 12 mm\u0026nbsp;were used for\u0026nbsp;the following experiments.\u003c/p\u003e\n\u003cp\u003eThe hUVECs and hPDLSCs were detached and mixed at a ratio of 3:1 (hUVECs:hPDLSCs) as described previously [30]. The co-cultured cells were suspended into 5% GelMA solution at a total density of 1 ×10\u003csup\u003e6\u003c/sup\u003e cells/ml at 37℃ in dark.\u0026nbsp;Then, the suspension was placed in a 1 ml syringe (with 100 µm inner diameter) and injected into the pores inside the 3D printed CPC scaffold.\u0026nbsp;Expose the construct to a 405 nm wavelength light source to facilitate its curing process.\u0026nbsp;The 3D co-culture scaffold\u0026nbsp;was cultured with the endothelial cell medium for 21 days.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e2.5. Scanning electron microscopy (SEM) of 3D printed CPC scaffolds \u0026nbsp;\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAfter 21 days of culturing, the composite constructs were fixed overnight. The next day, each cylindrical scaffold was bisected through its mid-plane and the resulting cross-sections were dehydrated and examined under a scanning electron microscope (Quanta 200, FEI, Hillsboro, OR, USA).\u0026nbsp;Filament\u0026nbsp;widths and pore sizes were measured from representative micrographs. Six specimens were analyzed as described previously [19].\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e2.6.\u0026nbsp;\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003eMechanical properties\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThree-point flexural tests were performed on the composite scaffolds using a universal testing machine for mechanical testing [29]. The span is 20 mm and the displacement speed of test head is 1 mm/min. Flexural strength (σ) and elastic modulus (E) were derived from the load–displacement curves. σ=3FmaxL/2bh\u003csup\u003e2\u003c/sup\u003e, E=(F/d) (L\u003csup\u003e3\u003c/sup\u003e/4bh\u003csup\u003e3\u003c/sup\u003e), where F\u003csub\u003emax\u0026nbsp;\u003c/sub\u003eis the peak load, L is the span, b and h are the specimen width and thickness, respectively, and F/d represents the slope of the linear-elastic region. Six specimens were tested.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e2.7. Viability of encapsulated hDPLSCs\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAfter 1, 4, 7 and 14 days of culturing,\u0026nbsp;cellular viability within the hydrogel constructs was evaluated using a live/dead viability assay kit (Sigma-Aldrich). Epifluorescence microscope (Sigma-Aldrich) was used for observation. The percentage of live cells was calculated by Image J software (NIH) as described previously [29].\u003c/p\u003e\n\u003cp\u003eA cell counting kit (CCK-8 assay, Dojindo, Tokyo, Japan) was used to evaluate cell viability in the 3D culture system at 1, 4, 7, 14 and 21 days. The working solution was prepared with endothelial cell medium containing 10% CCK-8 solution, followed by a 1-hour incubation at 37°C. The cell proliferative rate was determined by measuring the absorbance at an optical density of 450 nm using microplate reader (SpectraMax M5, Molecular Devices, Sunnyvale, CA) as described previously [29].\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e2.8. Observing hUVECs via CD31 immunofluorescence staining\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAfter 1, 4, 7 and 14 days of culturing,\u0026nbsp;the resulting microvascular-like structures were then visualized and assessed by CD31 (PECAM-1) immunofluorescence staining as described previously [11]. The samples were fixed and incubated with CD31 mouse mAb (1:500, Cell signaling technology, Pudong District, Shanghai, China) overnight. After washing with PBS, Alexa Fluor 488-conjugated goat anti-mouse IgG (1:1000, goat anti-mouse Alexa Fluor 488, green fluorescence, Cell signaling technology) was applied, followed by DAPI counterstaining (1:1000, Beyotime) at room temperature. The samples were observed with confocal laser scanning microscopy (OLS5100, Olympus, Tokyo, Japan). For each time-point, three random regions from each of five specimens were recorded. Image J (National Health Institute, Bethesda, MA, USA) was used to obtain the vessel length per area and junction number per area (n=5). Quantification was performed in duplicate by a double blinding protocol (n = 5).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e2.9. Rat complete periodontal defect model\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe animal protocol was approved by the Committee for Animal Experiments of Beijing Stomatological Hospital, Capital Medical University (NO. 2024-82301117). Male athymic nude rats (8 weeks old, 200–250 g, SPF Biotechnology, Haidian District, Beijing, China) were anesthetized with an intraperitoneal injection of Zoletil 50 (Virbac, Carros, France) at 50 mg/kg body weight (n=6). All surgical procedures were performed under strict aseptic conditions. After sterilization of rats, a critical-size alveolar defect (4 mm × 2 mm × 1 mm) was prepared buccal to the mesial root of the mandibular second molar to establish a rat alveolar bone defect model as described previously [32]. The mesial root was exposed, and all residual periodontal ligament and cementum were meticulously curetted. Scaffolds were press-fit into the defects. Additionally, the rats received flunixin meglumine (2 mg/kg, s.c.; Shanghai Yuanye Bio-Technology, Shanghai, China) and penicillin G benzathine (24,000 IU/kg, i.m.; Pengdi, Henan, China) once daily for 3 consecutive days. Animals were monitored daily for signs of pain, wound integrity, and normal ambulation. Four groups were set:\u003c/p\u003e\n\u003cp\u003e(1) Blank group: underwent the surgical procedure without any scaffold implantation；\u003c/p\u003e\n\u003cp\u003e(2) CPC+GelMA group: received an acellular scaffold, which was preconditioned in ECM for 21 days prior to implantation, serving as the non-prevascularized control；\u003c/p\u003e\n\u003cp\u003e(3) CPC+GelMA-cell group: received the experimental intervention—a prevascularized scaffold seeded with a co-culture of hPDLSCs and hUVECs and matured in ECM for 21 days；\u003c/p\u003e\n\u003cp\u003e(4) Natural Periodontium group: consisted of the contralateral non-operated sites, provided the baseline native tissue.\u003c/p\u003e\n\u003cp\u003eAt 4 weeks post-surgery, six rats per group were euthanized. After deep anesthesia with an overdose of intraperitoneal pentobarbital (150 mg/kg), animals were sacrificed by CO₂ asphyxiation followed by exsanguination via bilateral thoracotomy. The implants were then retrieved and immediately fixed in 4% paraformaldehyde at 4 °C for 24 h.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e2.10. Micro computed tomography\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003e（\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003eMicro-CT\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003e）\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eMicro-CT (SkyScan 1276, Bruker BioSpin, Germany) was employed to scan the bone-defect regions and perform three-dimensional reconstructions [29]. The ROI was defined as the buccal alveolar bone encircling the mesial root of the mandibular second molar and encompassed a volume of 4 mm×2 mm×1 mm. Bone thickness and bone volume fraction was calculated to quantify the percentage of newly formed bone within the defect. Additionally, a cross-sectional slice at the mid-root level of the mesial root was selected for qualitative evaluation. An initial grayscale threshold of 150–1000 HU was applied to distinguish newly formed bone.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e2.11. Histomorphometric analyses \u0026nbsp;\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eSpecimens were\u0026nbsp;decalcified in 10% ethylene diamine tetraacetic acid (EDTA) (Solarbio Science \u0026amp; Technology, Beijing, China) for 2 months, processed routinely, and embedded in paraffin. The central part of the implant and defect was cut into 5 μm-thick sections for hematoxylin and eosin (H\u0026amp;E) staining, Masson’s staining and immunohistochemistry (IHC) staining.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e2.11.1. H\u0026amp;E staining\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe samples were decalcified and embedded in paraffin. Serial 5 µm-thick sections were prepared and stained with H\u0026amp;E. New bone area, total defect area and the number of new vessels were quantified in each section by image J as described previously [19]. New bone area fraction was calculated by dividing the area of new bone with the area of the total defect. New vessels density was expressed as the number of new vessels divided by total defect area (n=6).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e2.11.2. Masson’s staining\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eDeparaffinized sections were stained with Weigert’s iron hematoxylin, sequentially incubated in Biebrich scarlet-acid fuchsin and phosphomolybdic/phosphotungstic acid, and finally differentiated in aniline blue to visualize collagen. Blue-stained bone area and total defect area were measured in each section [30]. Osteoid area fraction was calculated by dividing the area of blue-stained bone with the area of the total defect (n=6).\u0026nbsp;\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e2.11.3. IHC staining\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eImmunodetection of human CD31 and RUNX2 was performed on 5 µm paraffin sections. Sections were incubated with rabbit anti-human CD31 (1:500, Abcam) and anti-human RUNX2 (5 µg/mL, Abcam), followed by HRP-conjugated secondary antibody (1:500, Abcam). Signals were developed with DAB and counterstained with hematoxylin. For histomorphometry, one mid-sagittal section per animal was analyzed (n=6). The density of TRAP-positive cells was calculated. CD31 and RUNX2 expression was assessed as integrated optical density (IOD) from 6 random fields per section as described previously [32].\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e2.12. S\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003eubcutaneous transplantation model\u0026nbsp;\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003ein nude rats\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eFor subcutaneous transplantation, CPC+GelMA group and CPC+GelMA-cell group were selected. After induction of general anesthesia and sterile preparation, scaffolds were\u0026nbsp;implanted into subcutaneous pockets on the back nude rats (8 weeks, 200–250 g) as described previously [33]. After 6 weeks of implantation, all scaffolds were obtained and fixed for\u0026nbsp;stereomicroscope\u0026nbsp;observations\u0026nbsp;(Olympus, Tokyo, Japan) and H\u0026amp;E staining.\u0026nbsp;Vessels were enumerated in three randomly selected fields per section, and vascular density was calculated as vessel number per unit area.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e2.13\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003e．\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003eStatistical analysis \u0026nbsp;\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAll statistical analyses were conducted using SPSS 22.0 (IBM Corp., Armonk, NY, USA). Data are expressed as mean ± standard deviation (SD). Group comparisons were performed by one-way analysis of variance (ANOVA) followed by Tukey’s post-hoc test for multiple comparisons. p-value \u0026lt; 0.05 was considered statistically significant.\u003c/p\u003e"},{"header":"3. Results","content":"\u003cp\u003e\u003cstrong\u003e\u003cem\u003e3.1. Identification of hPDLSCs\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eFigure 1. A plots flow cytometry result of isolated hPDLSCs. CD105, CD90, STRO-1 were highly expressed to 99.6%, 76.4% and 88.6%. While, CD34 and CD45 were weakly expressed to 0.3% and 0.6%, respectively.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e3.2.\u0026nbsp;\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003eFabrication of\u0026nbsp;\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003ethree-dimensionally prevascularized CPC scaffold\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eFigure 1. B, C illustrates the fabrication of a three-dimensional grid-like CPC scaffold. Figure 1. D plots the injection of an hPDLSCs–hUVECs-laden GelMA hydrogel into the 3D-printed CPC scaffold to generate a three-dimensionally prevascularized CPC construct.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e3.3. Physical properties of scaffolds\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eFigure 2. A shows CPC filaments intersecting at defined angles. Cross-sectional images (Figure 2. B, C) reveal spherical crystalline structures and abundant pores within the CPC matrix. Quantitative measurements yielded filament diameters of 285.21 ± 1.4 μm, pore sizes of 302.31 ± 3.4 μm, and filament intersection angles of 60.12 ± 0.4° for the 3D-printed CPC scaffold.\u003c/p\u003e\n\u003cp\u003eThe flexural strength (Figure 2. D) and elastic modulus (Figure 2. E) of CPC construct had significant difference with cancellous bone. The average value of flexural strength (6.78±0.68 MPa) and elastic modulus (0.41±0.02 GPa) was significantly higher than that of cancellous bone. Values indicated by dissimilar letters are significantly different from each other (p\u0026lt;0.05).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e3.4. Viability and cell proliferation of cells\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003e\u0026nbsp;co-cultured within GelMA-CPC constructs\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eLive/dead staining images for the encapsulated cells (Figure 3. A) represented numerous live cells (green staining) and a few dead ones (red staining) at each time points tested. At day 1, cells were embedded within the hydrogel, predominantly spherical and clustered, with relatively low cell numbers visible. From day 4 to 7, cells began to aggregate, the number of released cells increased continuously, which extended well, showing spindle or polygonal shape. At day 14, cells were evenly distributed throughout the hydrogel. At day 21, all cells had fully extended into long spindle shapes, forming pseudopodia and interconnecting into branched, network-like structures. The percentages of live cells (Figure 3. B) exceeded 80% on day 1 and significantly increased to approximately 90% from day 4 to 21. As shown in CCK-8 assessment (Figure 3. C),\u0026nbsp;co-cultured hPDLSCs and hUVECs showed 9.2 folds increase of proliferation from day 1 to 21. Values indicated by dissimilar letters are significantly different from each other (p \u0026lt;0.05).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e3.5. CD31 immunofluorescence staining\u0026nbsp;\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eImaging of hPDLSCs–hUVECs co-cultured in CPC-GelMA scaffolds was performed by CD31 immunostaining\u0026nbsp;(Figure 4. A–C).\u0026nbsp;HUVEC membranes were stained green for the endothelial marker CD31, while nuclei were counterstained blue with DAPI. Branch-like structures gradually increased from day 7 to day 21. The co-cultured cells within the GelMA hydrogel progressively formed vessel-like networks with prolonged culture. Figure 4. D demonstrates that the cumulative vessel-like branch length increased significantly over time (p \u0026lt; 0.05). Compared to day 7, the cumulative vessel length had risen 18-fold at day 21, reaching 17.12 ± 2.34 mm/mm². Figure 4. E reveals that the number of vessel junctions increased from day 7 to day 21 (p \u0026lt; 0.05). By day 21, the junction density had risen to 37.25 ± 3.14 vessels/mm², representing a 3.4-fold increase over the value recorded on day 7.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e3.6.\u0026nbsp;\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003eComplete periodontal tissue regeneration in vivo\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAt 4 weeks post-surgery, the cross-sections view of rat alveolar bone defects were observed by Micro-CT (Figure 5. A-D). The buccal bone thickness adjacent to the mesial root of the second molar in all groups were observed. As shown in Figure 5. E, the CPC+GelMA-cell group (0.549±0.01 mm) showed the greatest new formed bone thickness, which were 1.3 folds that of CPC-GelMA group (0.421±0.02 mm) and 2.57 folds that of the blank group (0.214±0.02 mm). The new bone area fraction (Figure 5. F) was also highest in the CPC+GelMA-cell group (44.33±3.01 %), reaching 1.23 folds that of the CPC-GelMA group (35.92±2.52 %) and 1.43 folds that of the blank group (30.99±1.78 %).\u003c/p\u003e\n\u003cp\u003eFor H\u0026amp;E staining, all groups\u0026nbsp;newly formed bone (NB) and blood vessels (V) in defects, with no evident inflammation or immune reaction (Figure 6). New bone with a typical organized bone morphology was formed. New blood vessels were observed around the new bone. Osteoblasts with blue cytoplasm and round-to-oval nuclei lined the surfaces of the new bone. Compared with the Natural Periodontium group (Figure 6. G, H), the blank group (Figure 6. A, B) showed only sparse alveolar bone regeneration on the buccal side of the mesial root of the second molar. The CPC+GelMA group (Figure 6. C, D) exhibited moderate, irregularly distributed new bone and vessels. The CPC+GelMA-cell group (Figure 6. E, F) displayed the most abundant and irregularly distributed new bone and vessels, with active osteoblasts. As shown in Figure 7. I, CPC+GelMA-cell group (57.99±4.78 %) formed the highest new bone area that was 1.37 folds that of the CPC+GelMA group (42.26±5.48 %) and 2.29 folds that of the blank group (25.33±5.25 %). The new vessel density in CPC+GelMA-cell group (41.32±2.95 vessels/mm\u003csup\u003e2\u003c/sup\u003e) were about 2.65 folds that of CPC-GelMA group (41.32±2.95 vessels/mm\u003csup\u003e2\u003c/sup\u003e) and 6.2 folds that of the Blank group (41.32±2.95 vessels/mm\u003csup\u003e2\u003c/sup\u003e) (Figure 6. J).\u003c/p\u003e\n\u003cp\u003eMasson staining effectively reveals the maturity of collagen within bone tissue: mature collagen stains red, whereas newly formed bone or immature bone stains blue. Relative to the Natural Periodontium group (Figure 7. G, H), all three intervention groups exhibited immature bone within the defect. The Blank group (Figure 7. A, B) showed sparse blue trabeculae embedded in abundant fibrous tissue. In contrast, the CPC+GelMA-cell group (Figure 7. E, F) and CPC+GelMA group (Figure 7. C, D) produced extensive, irregularly arranged immature bone that appeared predominantly blue, with scattered foci of mature red-stained matrix. The area of blue-stained immature bone (Figure 7. I) in CPC+GelMA-cell group was highest (56.38±3.47 %), which was 1.29 folds that of CPC+GelMA group (43.59±2.77 %) and 2.38 folds that of Blank group (23.7±1.33\u0026nbsp;%).\u003c/p\u003e\n\u003cp\u003eFor immunohistochemical images of rat alveolar bone defects, both angiogenic marker CD31\u0026nbsp;(Figure 8. A) and osteogenic marker Runx2 (Figure 8. B) were expressed in the periodontal defects in all groups at 4 weeks after transplantation, compared with the negative-control group. The proportion of CD31-positive cells in the CPC+GelMA-cell group (2.387±0.027 per slice) was also much higher than those in the CPC+GelMA (2.061±0.035 per slice) and Blank (1.639±0.028 per slice) groups, reaching 1.16-fold and 1.46-fold higher values (Figure 8. C). Moreover, the newly formed periodontal tissues in the CPC+GelMA-cell group contained the highest proportion of Runx2-positive cells (2.471±0.022 per slice), being 1.17-fold and 1.91-fold greater than in the control (2.112 ± 0.038 per slice) and blank (1.675 ± 0.027 per slice) groups, respectively (Figure 8. D).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003e3.7. S\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003eubcutaneous transplantation\u0026nbsp;\u003c/em\u003e\u003c/strong\u003e\u003cstrong\u003e\u003cem\u003ein vivo\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eFigure 9 plots the images of stereomicroscopic view and H\u0026amp;E staining after subcutaneous implantation in nude rats at 6 weeks. Neovessels were visible on the surface of both groups. Histological evaluation revealed newly formed vessels within the macropores of both scaffolds. The vessels density of CPC+GelMA-cell group (Figure 9. B) was markedly higher than CPC+GelMA group (Figure 9. A). Morphometric analysis (Figure 9. C) demonstrated that the vessels density in the CPC+GelMA-cell group reached 81.33 ± 3.54 vessels/mm², a 1.93-fold increase over the 42.21 ± 5.84 vessels/mm² measured in the CPC+GelMA group (p\u0026lt;0.05).\u0026nbsp;\u003c/p\u003e"},{"header":"4. Discussion","content":"\u003cp\u003eThe present study is the first to combine a 3D printed CPC framework with a GelMA-hPDLSCs-hUVECs system to fabricate a prevascularized CPC scaffold for alveolar bone regeneration. The hypotheses were proven that the novel construct had good mechanical properties, pore structure, biocompatibility and angiogenic capability in vitro. Compared with non-prevascularized CPC controls, the prevascularized scaffold significantly enhanced new bone and vessels formation in vivo.\u003c/p\u003e\n\u003cp\u003eOur results showed that the isolated cells highly expressed STRO-1, CD90, and CD105, with low expression of CD34 and CD45, consistent with the characteristics of MSCs and thus considered as hPDLSCs [34]. Yeasmin et al [35] found that hPDLSCs can secrete angiogenesis-related factors such as VEGF and FGF, and provide stable support for endothelial cell networks. These findings further demonstrated that hPDLSCs, as seed cells, have significant vascularization potential in co-culture systems. Co-culture systems, through heterotypic cell–cell interactions, demonstrate a synergistic augmentation of pro-angiogenic growth factor output compared to monocultures [35].\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eMSCs not only provide trophic support to ECs but are also reciprocally activated by EC-derived signals, leading to further upregulation of their own growth factor secretion\u003c/strong\u003e[35]. Monocultured MSCs are unable to self-assemble into microvascular structures, whereas co-culture with ECs rapidly initiates capillary-like network formation [10]. In the present study, hUVECs and hPDLSCs were co-seeded at a fixed ratio of 3:1. It has been demonstrated that a 1:1 ratio of hUVECs to MSCs generates stable vascular networks while simultaneously facilitating osteoid matrix deposition [15]. Conversely, hPDLSCs have been reported to proliferate significantly faster than hUVECs [11]. To offset this imbalance, initial EC proportions have been raised to 3:1 or even 5:1 (hUVEC:hPDLSC) in prior work [36]. Longitudinal profiling confirms a progressive decline in the relative abundance of hUVECs during extended co-culture [37]. Therefore, the 3:1 seeding ratio chosen here is expected to counterbalance the differential expansion rates and stabilize the two populations as the assay proceeds. Nevertheless, systematic optimization studies are still required to definitively establish the ideal hUVECs:hPDLSCs numerical ratio for maximal vasculogenic output in co-culture systems.\u003c/p\u003e\n\u003cp\u003eGelMA hydrogels provide a cytocompatible extracellular environment, yet their bioactivity is dictated by GelMA concentration, photoinitiator content, and UV dose [38, 39]. Increasing GelMA density elevates cross-linking efficiency and network tightness. When the GelMA concentration exceeds 10% (w/v), pore size and swelling ratio decline, restricting cell proliferation and hindering the diffusion of oxygen, ultimately impairing cellular metabolism and phenotypic function [40]. Conversely, GelMA below 5% (w/v) remains high biocompatibility but lacks adequate mechanical integrity [40]. Comparative studies reveal that 5% (w/v) GelMA yields larger pores than its 10% (w/v) counterpart, facilitating Ca²⁺ flux and diffusion without compromising viability, and confers superior in vitro osteogenic potential [41]. Therefore, 5% (w/v) GelMA hydrogel was selected as the cell-laden matrix in the present study. CCK-8 assays revealed a robust and sustained proliferative profile of the co-cultured cells within the 5% GelMA matrix. Additionally, the live cell percentage was 80% on day 1 and exceeded 90% at every subsequent time point. The brief drop in live-cell number on day 1 is likely attributable to residual unreacted functional groups or small-molecular by-products released during GelMA cross-linking [42]. Future work should therefore optimize cross-linking parameters such as lowering photoinitiator concentration or shortening UV exposure to curb radical formation and improve early survival. Moreover, GelMA’s intrinsic drug-loading capacity and interconnected porosity enable sustained, tunable release of bioactive cargos [43]. With this benefit, future studies should encapsulate pro-angiogenic factors within GelMA-based co-culture systems, thereby integrating controlled growth-factor delivery with cellular paracrine signaling to enhance vascularization.\u003c/p\u003e\n\u003cp\u003eNevertheless, the inherently low mechanical strength of GelMA hydrogel limits its standalone use in load-bearing alveolar defects [29]. Consequently, recent studies have focused on integrating hydrogels with high-strength scaffolds to generate composites that unite excellent bioactivity with sufficient mechanical competence [29]. The calcium-phosphate cement (CPC) paste used in the present study has been extensively validated as a biocompatible and osteoconductive bone substitute [44-48]. Therefore, we combined GelMA hydrogel co-encapsulating hPDLSCs and hUVECs with a 3D-printed CPC scaffold to achieve sufficient mechanical competence to withstand physiological masticatory loads. Flexural strength and elastic modulus are critical mechanical indices for evaluating scaffold performance. Flexural strength and elastic modulus are critical design parameters [49]. Flexural strength (σ) is defined as the maximum stress a material can sustain before fracture, whereas the elastic modulus (E) quantifies\u0026nbsp;the stress required per unit strain in the elastic region [49]. In human cancellous bone, flexural strength is approximately 3.5 MPa and the elastic modulus is 0.30 GPa, respectively [50] . Three-point bending test indicates that the scaffold showed better flexural strength and elastic modulus than those of cancellous bone. Thus, the scaffold offers adequate mechanical strength and dimensional stability for load-bearing alveolar repairs.\u003c/p\u003e\n\u003cp\u003eIn 3D printed scaffolds designed for vascular regeneration, filament diameter, pore size and filament intersection angle are determinants of pro-angiogenic performance [51]. Pores between 200 and 400 µm allow efficient cell migration and nutrient exchange [52]. Pores below 100 µm hinder cell infiltration and capillary ingrowth [53]. Intrinsic micropores of CPC are typically less than 50 µm and poorly interconnected, which limiting the migration and interaction of cells [54, 55]. Given that human microvessels range from 5 to 200 µm in diameter [51] , the 302.31 ± 3.4 µm pore size generated in the printed CPC scaffold readily accommodate nascent microvessels and their interconnection, facilitating 3D vascular network assembly. Notably, 300 µm pores have been specifically identified as optimal for alveolar bone regeneration [19]. Moreover, scaffolds of 300 µm-pore size honeycomb architecture concurrently drive bone and vessel formation in vivo [56]. Moreover, similar filament diameter and pore size have been shown to homogenize stress distribution and markedly improve scaffold strength [57]. In the present work, the printed CPC displays comparable filament and pore sizes, providing a regular architecture conducive to cell attachment and growth. In addition, the intersection angle between filaments influences both mechanical integrity and cell alignment [23] . An appropriate angle can enhance scaffold strength and stability while promoting uniform cell distribution. For bone engineering, CPC plots between 15° and 90° configurations are commonly investigated [23]. Previous study demonstrated that a 60° intersection angle yielded higher compressive modulus and strength than a 45° pattern [58]. Scaffolds printed with a 60° filament intersection angle likewise exhibited superior mechanical performance in rat femoral defects [44]. This advantage is attributed to the angle’s balanced stability, which simultaneously supports multidirectional cell signaling and nutrient exchange [44]. The CPC scaffolds described here were printed at 60.12 ± 0.4°, closely matching the reported optimum. Nevertheless, the definitive intersection angle for 3D-printed CPC remains to be established and demands systematic exploration.\u003c/p\u003e\n\u003cp\u003eCD31 immunostaining was used to quantify in vitro angiogenesis of hUVECs co-cultured with hPDLSCs inside GelMA-CPC scaffolds. Cumulative vessel length and junction number are standard metrics of neovascularization: length reflects network expansion, whereas junction density mirrors architectural complexity and functional anastomosis [59]. Confocal imaging revealed microcapillary-like networks that elongated and interconnected over time. By day 21, the 3D GelMA-CPC co-culture system exhibited significantly greater cumulative vessel length and junction number than previously reported 2D CPC surface co-culture [11], underscoring the superior pro-angiogenic potential of the 3D co-culture system.\u003c/p\u003e\n\u003cp\u003eThe present in vivo studies demonstrated that prevascularized scaffolds have significant advantages in promoting angiogenesis and accelerating alveolar bone repair. Previous study demonstrated that prevascularized scaffolds can set up a functional blood supply by speeding up microvessel formation and stably connecting with host blood vessels, which greatly improving implant performance [60]. Currently, scaffold prevascularization mainly uses two strategies: in vivo and in vitro. In vivo prevascularization involves implanting the scaffold into a well-vascularized site (e.g. muscle) for several weeks to form a microvessel network, but it requires an invasive procedure [61]. In contrast, in vitro prevascularization avoids a second operation and donor-site morbidity, offering a minimally invasive, surgeon-friendly route that is more suitable for clinical use [61].\u003c/p\u003e\n\u003cp\u003eResearchers first co-cultured endothelial cells with osteoprogenitor cells and generated a 3D vascular network in vitro [62]. Inspired by this, we built a prevascularized scaffold in a 3D co-culture system and tested its angiogenic potential by subcutaneous implantation in nude rat. Because nude rats are immunocompromised, xenograft rejection is sharply reduced, which improving graft survival and the likelihood of vessel formation [63]. At week 6, vessel density inside the scaffolds reached 81.33 ± 3.54 vessels/mm², matching earlier reports that prevascularized constructs accelerate perfusion [60]. In the previous report, the fully prevascularized implants typically reach 80% vascular coverage within 6 weeks, whereas non-prevascularized controls achieve only 50% [60]. This advantage stems from the pre-built vessels acting as “highways” that pull in more capillary sprouts, culminating in a denser and more durable microvascular network throughout the scaffold\u0026nbsp;[60].\u003c/p\u003e\n\u003cp\u003eAlveolar bone regeneration was evaluated in all groups in vivo. By week 4, new bone was evident in every defect. However, the CPC+GelMA-cell group produced significantly more bone than either CPC+GelMA or Blank groups. Micro-CT gave a mineralized-bone fraction of 44.33 ± 3.01 % in the CPC+GelMA-cell group—23 % higher than CPC+GelMA group—whereas semiquantitative histomorphometry of H\u0026amp;E sections yielded 57.99 ± 4.78 % total new bone area, 1.37-fold that of CPC+GelMA group. The discrepancy reflects the fact that Micro-CT records only mineralized tissue, whereas H\u0026amp;E staining also captures unmineralized osteoid [64].\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eNewly formed trabeculae stained with H\u0026amp;E appear as irregular nets or cords. The 3D scaffold apparently influenced this trabecular arrangement. Nevertheless, H\u0026amp;E cannot distinguish immature bone from mature lamellar bone, so Masson staining was additionally performed to verify the maturity of the regenerated tissue [65].\u003c/p\u003e\n\u003cp\u003eMasson staining reliably visualizes collagen remodeling in bone tissues [65]. Newly formed bone is rich in type I collagen and stains blue. As mineralization proceeds, the stained color shifts toward red. This color transition is routinely exploited to gauge maturation, with the nascent-to-mature conversion typically occurring between 4 and 8 weeks post-operatively [65]. We therefore harvested specimens at week 4 and used the area fraction of blue-stained bone as a surrogate for new bone formation. Masson images revealed that the CPC+GelMA-cell group contained the largest amount of immature bone (56.38 ± 3.47 %), representing 1.29- and 2.38-fold increases over the CPC+GelMA and Blank groups, respectively, indicating that the defect area was undergoing active osteogenesis and that the experimental intervention accelerated this process. Due to Masson and H\u0026amp;E staining provide limited information on cellular and vascular components, IHC staining was employed for a more comprehensive evaluation. CD31, an endothelial-specific marker, was used to quantify angiogenesis [66], whereas Runx2, the master transcription factor driving osteoblast differentiation, was used to assess osteoblastic activity and maturity [67]. Semi-quantitative IHC demonstrated that Runx2 expression in the CPC+GelMA-cell group was 1.17-fold that of CPC+GelMA group, corroborating the superior osteogenic potential observed histologically. Similarly, a previous study demonstrated that scaffold pre-seeded with cells generated significantly more new bone than its cell-free counterpart, at 4 weeks in a rat cranial bone defect model [68]。\u003c/p\u003e\n\u003cp\u003eMoreover, a significant positive correlation between osteogenesis and angiogenesis was evident [68] . H\u0026amp;E morphometry revealed a neovessel density of 41.32 ± 2.95 vessels/mm² in the CPC+GelMA-cell group, 2.65-fold that of CPC+GelMA group, accompanied by a 16% increase in CD31 immunostained area. These data corroborate earlier reports that bone formation rate rises in parallel with vascular ingrowth [69, 70].\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eThis coupling is likely initiated by the hypoxic milieu that develops immediately after scaffold implantation. Under low oxygen tension, co-cultured cells activate hypoxia-inducible factor-1α (HIF-1α) pathway, which transcriptionally up-regulates vascular endothelial growth factor (VEGF) and fibroblast growth factor-2 (FGF-2). While promoting angiogenesis, these cytokines simultaneously deliver the nutrients and osteoinductive signals required for effective bone repair [71, 72].\u0026nbsp;Additionally, PI3K/Akt [73] and MAPK [74] signaling pathways participate in hypoxia-induced angiogenesis, which can modulate cell proliferation, survival, and differentiation to accelerate bone repair and regeneration.\u003c/p\u003e\n\u003cp\u003eIn addition to the contribution of co-cultured cells, the intrinsic properties of the scaffold itself critically influence osteogenesis and angiogenesis [75]. In the present study, the CPC+GelMA group exhibited substantially more de novo bone than the empty defect（Blank group）. This effect is attributable to the osteoinductive capacity of CPC, which sequesters endogenous growth factors and directs multipotent stem cells toward an osteoblastic phenotype [76]. In earlier work, traditional 2D CPC placed in cranial defects for 8 weeks regenerated only 13.89 ± 2.95 % new bone [77]，while traditional 2D CPC packed into alveolar socket yielded less than 5% repair by week 12 [78]. In contrast, micro-CT of our 3D CPC scaffold revealed 35.92 ± 2.52 % new alveolar bone within only 4 weeks. This pronounced disparity is presumably attributable to differences in defect site and internal scaffold structure [79].\u003c/p\u003e\n\u003cp\u003eIn addition, porogen-leavened macroporous CPC [79], collagen-sponge CPC [78] and CPC fortified with SP [78] or rhBMP-2 [80] have respectively doubled to quintupled new bone output within 4 weeks. Thus, compositional and architectural refinement can readily enhance CPC-mediated alveolar regeneration.\u003c/p\u003e\n\u003cp\u003eIn the field of vascularized scaffolds, bioactive glass [81] and silk fibroin [82] have attracted intense attention because of their superior pro-angiogenic capacity. Concurrently, microfluidic technologies are opening new routes for the fabrication of individualized constructs [83]. Smart stimulus-responsive drug-delivery systems that release therapeutics under endogenous or exogenous factors further enhance therapeutic efficacy [84]. Collectively, these innovations offer great clinical potential for more effective tissue repair and regeneration.\u003c/p\u003e\n\u003cp\u003eThe periodontium’s intricate architecture reveals that alveolar bone, periodontal ligament and cementum are structurally and functionally interdependent to ensure tooth stability [85].\u0026nbsp;Therefore, future research should focus on engineering hierarchically layered multi-phase scaffolds that faithfully replicate the native periodontium.\u003c/p\u003e"},{"header":"5. Conclusion","content":"\u003cp\u003eThis study developed a novel 3D printed CPC scaffold to establish a prevascularized 3D construct for alveolar bone engineering. The resulting construct had proper mechanical properties and pore structure of the CPC skeleton. The delivered hPDLSCs-hUVECs exhibited excellent cell proliferation and angiogenic potential in GelMA hydrogel in vitro. When implanted in vivo, the prevascularized CPC+GelMA-cell group elicited significantly greater neovascularization and accelerated alveolar-bone regeneration compared with the cell-free CPC+GelMA control. The intrinsic properties of CPC also enhance osteogenesis and angiogenesis. Therefore, the novel CPC-GelMA-hPDLSCs-hUVECs construct is highly promising for concurrent bone and vascular regeneration in dental applications.\u003c/p\u003e"},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eEthics approval and consent to participate\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eCapital Medical University Ethics Committee approved this study (CMUSH-IRB-KJ-PJ-2024-28). All participants agreed to participate in this study and signed an informed consent form.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eConsent for publication\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eNot applicable.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAvailability of data and materials\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAll data generated or analysed during this study are included in this published article [and its supplementary information files].\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eCompeting interests\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors declare no competing interests.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eFunding\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThis study was supported by National Natural Science Foundation of China (No. 82301117), the Innovation Research Team Project of Beijing Stomatological Hospital, Capital Medical University (Grant No. CXTD202203) and Beijing Stomatological Hospital, Capital Medical University Young Scientist Program (No. YSP202510).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAuthors' contributions\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eY. Sun conducted the investigation, performed visualization and validation. Z. Zhao contributed to methodology and data curation, and drafted the original manuscript. Q. Qiao was responsible for visualization and methodology. conceptualized the study. W. Yu contributed to validation. Y. Bai participated in writing - review and editing, project administration, funding acquisition. All authors read and approved the final manuscript.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAcknowledgement\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eWe are grateful to the patients who are willing to participate in the study.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\n \u003cli\u003eWishney M. Potential risks of orthodontic therapy: a critical review and conceptual framework [J]. Aust Dent J. 2017; 62 Suppl 1: 86-96. https://doi.org/10.1111/adj.12486.\u003c/li\u003e\n \u003cli\u003eHan X, Sun M, Chen B, et al. 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J Biomed Mater Res A. 2025; 113: e37793. https://doi.org/10.1002/jbm.a.37793.\u003c/li\u003e\n\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":false,"highlight":"","institution":"","isAcceptedByJournal":true,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"bmc-oral-health","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"ohea","sideBox":"Learn more about [BMC Oral Health](http://bmcoralhealth.biomedcentral.com/)","snPcode":"","submissionUrl":"https://www.editorialmanager.com/ohea/default.aspx","title":"BMC Oral Health","twitterHandle":"BMC_series","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"em","reportingPortfolio":"BMC Series","inReviewEnabled":true,"inReviewRevisionsEnabled":true},"keywords":"Human periodontal ligament stem cells, Human umbilical vein endothelial cells, Three-dimensional printed, Calcium phosphate cement, Gelatin methacryloyl hydrogel, Osteogenesis, Angiogenesis, rat model","lastPublishedDoi":"10.21203/rs.3.rs-8260921/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-8260921/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003cp\u003e\u003cem\u003e\u003cstrong\u003eObjectives.\u003c/strong\u003e\u003c/em\u003e The objective of this study is to develop a load-bearing prevascularized construct by combining calcium phosphate cement (CPC) with cells within a three-dimensional (3D) hydrogel culture system, to accelerate the regeneration of alveolar bone defects.\u003cbr\u003e\n \u003cem\u003e\u003cstrong\u003eMethods.\u003c/strong\u003e\u003c/em\u003e A 3D co-culture system was established by encapsulating human periodontal ligament stem cells (hPDLSCs) and human umbilical vein endothelial cells (hUVECs) within a gelatin methacryloyl (GelMA) hydrogel on 3D-printed porous CPC scaffolds. The mechanical properties, pore structure and angiogenic potency were determined in vitro. In vivo performance was evaluated using a nude rat subcutaneous implantation model and a rat alveolar bone defect model. Four groups were tested: (1) Blank group (surgery-only group); (2) CPC+GelMA group (non-prevascularized group)；(3) CPC+GelMA-cell group (prevascularized group)；(4) Natural Periodontiumgroup.\u003c/p\u003e\n\u003cp\u003e\u003cem\u003e\u003cstrong\u003eResults. \u003c/strong\u003e\u003c/em\u003eThe novel construct had good mechanical properties and biocompatibility. The 3D co-culture in GelMA successfully induced microvascular formation in vitro. Subcutaneous implantation in nude rats showed that the CPC+GelMA-cell group exhibited markedly greater angiogenic capacity than the CPC+GelMA group after 6 weeks, with a neovascular density 1.93-fold higher than that of the non-prevascularized group. Among all groups, the CPC+GelMA-cell group exhibited the strongest capacity for repairing rat alveolar bone defects. Compared to CPC+GelMA group, CPC+GelMA-cell group significant enhanced bone regeneration in rats by 1.23-1.37 folds, and increased vascularization by 2.65 folds (p＜0.05).\u003c/p\u003e\n\u003cp\u003e\u003cem\u003e\u003cstrong\u003eConclusions:\u003c/strong\u003e\u003c/em\u003e The novel three-dimensional prevascularized CPC construct combined appropriate mechanical properties with great efficacy for alveolar bone regeneration and vascularization in vivo in an animal model.\u003c/p\u003e","manuscriptTitle":"A 3D Prevascularized Calcium Phosphate Cement Scaffold for Accelerated Alveolar Bone Regeneration and Angiogenesis in Rats","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2025-12-31 01:13:11","doi":"10.21203/rs.3.rs-8260921/v1","editorialEvents":[{"type":"communityComments","content":0},{"type":"decision","content":"Revision requested","date":"2026-02-26T09:02:19+00:00","index":"","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2026-02-18T12:52:45+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2026-02-13T15:12:05+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"112975522434200825287522111986646655307","date":"2026-02-12T01:16:55+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"85927552302037013141592714302293803515","date":"2026-02-05T09:17:44+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2026-02-04T12:54:00+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"53292343350410252208484091239141595285","date":"2026-02-04T12:00:10+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"120748432009260263717519653230044545899","date":"2025-12-27T08:56:00+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-12-26T15:05:27+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"147727583098472165414439246381317017867","date":"2025-12-26T02:58:56+00:00","index":"hide","fulltext":""},{"type":"reviewersInvited","content":"","date":"2025-12-25T08:32:46+00:00","index":"","fulltext":""},{"type":"editorAssigned","content":"","date":"2025-12-25T04:06:39+00:00","index":"","fulltext":""},{"type":"editorInvited","content":"","date":"2025-12-15T16:49:38+00:00","index":"","fulltext":""},{"type":"checksComplete","content":"","date":"2025-12-12T15:31:52+00:00","index":"","fulltext":""},{"type":"submitted","content":"BMC Oral Health","date":"2025-12-12T15:21:14+00:00","index":"","fulltext":""}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"bmc-oral-health","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"ohea","sideBox":"Learn more about [BMC Oral Health](http://bmcoralhealth.biomedcentral.com/)","snPcode":"","submissionUrl":"https://www.editorialmanager.com/ohea/default.aspx","title":"BMC Oral Health","twitterHandle":"BMC_series","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"em","reportingPortfolio":"BMC Series","inReviewEnabled":true,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"e7c7415a-3894-4f86-90da-f667436c6346","owner":[],"postedDate":"December 31st, 2025","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"under-review","subjectAreas":[],"tags":[],"updatedAt":"2026-05-08T12:40:02+00:00","versionOfRecord":[],"versionCreatedAt":"2025-12-31 01:13:11","video":"","vorDoi":"","vorDoiUrl":"","workflowStages":[]},"version":"v1","identity":"rs-8260921","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-8260921","identity":"rs-8260921","version":["v1"]},"buildId":"8U1c8b4HqxoKbykW_rLl7","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}