The Role and Mechanism of Dental Pulp Stem Cells in Highly Porous Microcarriers Forming New Micro-tissues to Accelerate Mandibular Bone Defect Repair

The Role and Mechanism of Dental Pulp Stem Cells in Highly Porous Microcarriers Forming New Micro-tissues to Accelerate Mandibular Bone Defect Repair | 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 Help Center Sign In Submit a Preprint Cite Share Download PDF Research Article The Role and Mechanism of Dental Pulp Stem Cells in Highly Porous Microcarriers Forming New Micro-tissues to Accelerate Mandibular Bone Defect Repair RenZe Shen, YiCheng Wang, JiaCheng Huang, YongMei Tan, SongXia Ke, and 4 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-3921403/v1 This work is licensed under a CC BY 4.0 License Status: Posted Version 1 posted You are reading this latest preprint version Abstract Objective Dental pulp stem cells (DPSCs) exhibit strong differentiation and proliferation capabilities but are often discarded. This experiment aimed to prepare PLGA (poly lactic-co-glycolic acid) highly open porous microspheres (HOPMS) and form microscale cell carriers with DPSCs, exploring their efficacy and mechanism in promoting alveolar bone defect repair. Methods Rat dental pulp stem cells were cultured and identified. PLGA HOPMS were prepared and their porosity, pore size, degradation rate, and PH change were evaluated. Cytological experiments examined the effects of PLGA HOPMS on cell proliferation, mineralized nodule formation, mineralization gene, and inflammatory factor expression. Cells were transduced with lentivirus for fluorescence marking and co-cultured with PLGA HOPMS to form microscale cell carriers. Subcutaneous and intraosseous implantations were performed to investigate their biocompatibility, angiogenesis, and inflammatory response. Results The size and pore diameter of PLGA HOPM were adjustable. PLGA HOPMS promoted the formation of mineralized nodules, mineralization gene expression, and exhibited no cytotoxicity. Subcutaneous experiments demonstrated that microscale cell carriers had excellent biocompatibility with no significant immune rejection. Intraosseous experiments confirmed that this material promoted osteogenesis and angiogenesis. Conclusion: PLGA HOPMS loaded with DPSCs effectively promoted bone and vascular formation in alveolar bone defect repair, proving to be an excellent stem cell transplantation carrier. PLGA Dental pulp stem cells Osteogenesis GBR Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Figure 7 Introduction Inadequate alveolar bone is a significant factor affecting the prognosis of periodontal therapy, implant surgery outcomes, and the stability of orthodontically treated teeth. However, once lost, the alveolar bone cannot naturally regenerate its original height and shape. Under natural conditions, bone marrow mesenchymal stem cells (BMSCs) are recruited to the defect area and differentiate into osteoblasts to form bone tissue. Nevertheless, in such case, due to the lack of structural support in the defect area, the restored height of the alveolar bone is challenging to achieve. Scaffold-supported stem cell transplantation techniques are key to resolving this issue. Mesenchymal stem cell transplantation has several advantages over other cell transplantation methods. First, these cells can migrate to the injury site and modulate the immune response, enhancing immunity when it is low and downregulating it when it is high. Second, mesenchymal stem cell transplantation is expected to increase osteoblast differentiation, inhibit osteoclast activation, and rebalance bone formation and absorption [1] . Third, mesenchymal stem cells can differentiate in multiple directions. For instance, in burn treatment, transplanting these cells onto burned skin significantly reduces local inflammation and promotes angiogenesis [2] . In treating liver cirrhosis, mesenchymal stem cell transplantation can foster liver tissue regeneration [3] . BMSCs are also crucial for new bone tissue formation and are often the preferred choice for repairing bone tissue defects. Dental pulp stem cells (DPSCs), as a type of mesenchymal stem cell, share embryonic homology with bone marrow mesenchymal stem cells. In current stem cell transplantation practices, DPSCs are often overlooked. In reality, throughout a person's life, numerous teeth are extracted, ranging from deciduous teeth to wisdom teeth and teeth removed due to orthodontic treatment or periodontitis in adults. Unfortunately, these extracted teeth are typically discarded. A single tooth can contain enough DPSCs for 2–3 stem cell transplantations, and these cells exhibit potent proliferation and differentiation abilities [4] . Their strong paracrine effects also play a significant role in injury repair [5] . Autologous DPSCs, used as tool cells, avoid immune rejection and eliminate the need for additional surgeries required for harvesting other stem cells. Research indicates that DPSC transplantation can promote the recovery of hematopoietic function in aplastic anemia, enhancing treatment for hematopoietic failure and providing blood protection [6] . Additionally, when co-cultured with PLCL scaffolds to form micro-tissues, DPSCs can be used in nerve injury repair, leading to the formation of new neurons [7] . These applications are based on the multi-directional differentiation capabilities and immunomodulatory functions of DPSCs. Extensive experiments have also verified the effectiveness and safety of DPSCs as tool cells. Microscale cell carriers have demonstrated significant advantages in tissue injury repair. Comprising scaffold structures and stem cells, these carriers provide a 3D environment conducive to localized cell proliferation and differentiation. The differentiation and paracrine actions of stem cells create a microenvironment favorable for tissue repair. Our preliminary work involved preparing PLGA highly open porous microspheres (HOPMS) and co-culturing them with cells to form micro-tissues. We observed that their implantation promotes tissue vascularization and myofibrosis [8, 9] . The structure of PLGA HOPMS is loosely porous with large interstitial diameters, avoiding the drawbacks of insufficient internal nutrient supply and poor cell migration or internal formation in the scaffold. The spherical 3D structure also provides ample growth space for stem cells. Based on this theories, we propose using PLGA HOPMS co-cultured with dental pulp stem cells (DPSCs) to create microscale cell carriers for repairing alveolar bone defects (Fig. 1 ). Figure 1 A involves preparing PLGA HOPMS; culturing Sprague Dawley (SD) rat DPSCs and performing their identification; and forming microscale cell carriers by co-culturing DPSCs with PLGA HOPMS. Figure 1 B illustrates the implantation of these carriers into SD rat alveolar bone defects, utilizing the scaffold function of the carriers for tissue growth and the tissue repair properties of dental pulp stem cells to promote local calcification and vascular formation for bone tissue repair. Figure 1 C is a schematic of the experimental process, involving the physicochemical characterization of prepared PLGA HOPMS, such as degradation performance and characterization. This is followed by cell experiments to verify the material's cytotoxicity, mineralization ability, and cell immunity. Finally, subcutaneous and bone tissue implantation in animals are used to validate the material's osteogenic and angiogenic properties and immune response, exploring its function and mechanism. Materials and Methods Materials Gelatin (from porcine skin, Type A), DMEM (HyClone), FBS (Thermo Fisher Scientific)。PEGDA (Mw: 6–7 kDa), polyvinyl alcohol (PVA), SYBR Premix DimerEraser kit (Takara)。sodium alginate (SA, low viscosity),DMEM (HyClone). PrimeScript® RT reagent kit (Takara), PLGA (66–107 kDa,lactide:glycolide 75:25) were purchased from SigmaAldrich (St. Louis, USA). RGD was obtained from the Chinese Peptide Company (Hangzhou, China). Methods (1) Preparation of Materials PLGA HOPMs were prepared by initially dissolving gelatin in 925µL of 1% PVA solution, followed by heating and stirring for 6 hours. 0.06g of PLGA was dissolved in 2220µL of dichloromethane. Once fully dissolved, it was mixed with the gelatin solution and processed with an ultrasonic disruptor for 90 seconds to form a uniform emulsion. This emulsion and 1% PVA solution were loaded into a microfluidic device and injected into a beaker containing 1% PVA under ice bath conditions. After microsphere formation, the mixture was left undisturbed for 1 hour, then the upper solution was gently stirred for another hour, and finally left in a refrigerator at 4°C overnight. The PVA was removed multiple times slowly, followed by 4–5 washes with ultrapure water and heating in a water bath for 1 hour. After several more washes, the microspheres were transferred into a tube and frozen at -20℃ overnight, then lyophilized in a vacuum freeze dryer for 24 hours.A factorial experiment was designed using Minitab to investigate the effects of ultrasonic power, gelatin content, and flow rate on the morphology of the prepared microspheres (Table 1 ). Table 1 Minitab factorial experimental design ultrasonic power/W gelatin/g flow velocity/(mL/min) A 200 0.05 0.1 B 200 0.05 0.5 C 200 0.10 0.5 D 200 0.10 0.1 E 400 0.05 0.1 F 400 0.05 0.5 G 300 0.075 0.3 H 400 0.10 0.1 I 400 0.10 0.5 (2) Cell experiments: Cell culture and identification: Dental pulps were extracted from 5-week-old Sprague Dawley rats in a sterile environment, minced and washed in PBS, then digested with Type I collagenase for 3 minutes. Digestion was stopped with 10% fetal bovine serum + culture medium, followed by washing and culturing in dishes. Cells were observed for migration. At the third passage, cells were stained for stro-1 and CD90, and CD34-positive cells were detected using flow cytometry. Cell proliferation was assessed on days 1, 3, 5, and 7 using the MTT assay. Cytotoxicity, proliferation, mineralization response, and migration experiments: Third-generation dental pulp stem cells were co-cultured with 0mg/mL, 0.25mg/mL, 0.50mg/mL, and 1.0mg/mL PLGA HOPMs. Cytotoxic responses were assessed at 0.5 and 1 day using the MTT assay, and effects on cell proliferation were evaluated on days 1, 3, 5, and 7. Cell migration was assessed using a Transwell chamber with DPSCs in the upper chamber and 0-0.15mg/mL PLGA HOPMs in the lower chamber, followed by crystal violet staining after 24 hours. Cells were co-cultured with PLGA HOPMs for 14 days, fixed with paraformaldehyde, and stained with Alizarin Red to assess mineralized nodule formation. RT-PCR was performed at set times to assess mRNA transcription in the material-cell co-cultures. (3) Subcutaneous implantation in animals: Cells were transfected with a GFP-labeled lentivirus provided by Shanghai Jiama Pharmaceutical Technology Co., Ltd. The lentiviral expression vector was pGLV3/Hl/GFP + Puro with the sequence 5 '-ACTACCGTTGTTATAGGTG-3'. The packaging system included pGag/Pol, pRev, and pVSV-G, with 293T cells as the packaging cells. After transduction for one day, GFP + DPSCs were imaged using a fluorescence microscope. GFP + DPSCs were co-cultured with PLGA HOPMs and examined with HE staining after 7 days and scanning electron microscopy on days 3 and 7 for cell growth. SD rats weighing 200-250g were purchased and acclimatized for two weeks. Under general anesthesia, microscale cell carriers formed from co-cultured PLGA HOPMs and GFP + dental pulp stem cells, as well as pure PLGA HOPMs, were implanted subcutaneously. The PLGA + cell group included a cell concentration of 3×10 6 /mL, and the PLGA + 2cell group had a concentration of 6×10 6 /mL. Samples were removed after 7 days; sections were stained with DAPI to observe the growth of GFP + cells inside the PLGA HOPMs and the DAPI staining pattern. Tissues were removed on days 3, 7, 10, and 14 for HE staining to observe the growth of the implanted tissue. At one week post-implantation, samples were taken for Masson staining, silver staining, and immunohistochemical staining to assess collagen formation, nerve tissue development, and the expression of TNF-α and IL-6. (4) Animal bone tissue implantation experiment: Cells were co-cultured with PLGA HOPMs for 7 days, with cell concentrations as previously described. Sprague Dawley rats weighing 200-250g were purchased and acclimatized for two weeks. Under general anesthesia, a dental bur was used to create a bone defect in the mandible, approximately 2 millimeters deep and 2.5 millimeters in diameter. The NC group did not receive any implantation, while the other groups were implanted with either PLGA HOPMs or microscale cell carriers co-cultured with cells. One month post-implantation, specimens were removed for gross observation, micro-CT analysis to calculate bone mineral density (BMD), and bone volume fraction (BV/TV); HE staining to observe tissue healing; TRAP staining to examine osteoclast activity; and to check for hemolysis in the animals. Immunofluorescence was used to observe the expression of CD34 and VEGF for angiogenic reaction and vascular formation location; immunohistochemistry to explore osteogenesis mechanisms through OC and OPG expression; and to assess bone tissue inflammatory response by measuring TNF-α expression. Animal weights were monitored to evaluate the implant's impact on their growth. Results (1) Material preparation and physicochemical properties Figure 2 shows SEM images of microspheres prepared under different experimental conditions, demonstrating that the microspheres had a uniform spherical shape, with diameters primarily around 550µm (Fig. 3 G). Additionally, with varying experimental conditions, the porosity and pore size of the microspheres changed. We selected group G from the factorial experiment, which had higher porosity and pore sizes primarily between 30–50µm (Fig. 3 H), for subsequent experiments. To verify the stability and degradation capability of the prepared microspheres, they were stored in PBS, samples at different time intervals, lyophilized, and SEM characterization was performed to observe changes in microsphere morphology (Fig. 3 A-E). The results showed that in the first stage of degradation, the spheres largely retained their shape, with finer structures on the surface beginning to degrade. In the second stage, the spherical structure of the microspheres gradually collapsed, with fragments breaking off. By the third stage, the spherical structure of the microspheres had completely disappeared, resulting in a large number of fragments. The pH changes in the solution during the degradation process were also characterized (Fig. 3 F), revealing no significant change in pH over the 8-month degradation period, maintaining a neutral environment around 7.2–7.4. Additionally, the degradation of the microspheres under static and dynamic conditions was characterized (Figs. 3 I-J), showing a faster degradation rate under dynamic conditions, with most of the material degrading by the 8th month. (2) Cell experiments (Fig. 4 A) DPSCs migrated out from the tissue block and were in good condition. (Fig. 4 B) The fixed third-generation cells were in good condition. Compared to (Fig. 4 C) cells positive for stro-1, more than 95% were stro-1 positive. (Fig. 4 D) DPSCs stained for CD90 showed positive responses, confirming the cells are mesenchymal stem cells. Flow cytometry detected about 9.66% CD34-positive cells, indicating low expression and excluding hematopoietic tissue origin. These experiments confirmed the cultivation of DPSCs. (Fig. 4 F) The cell proliferation curve was "S"-shaped, typical of fibroblast growth patterns. (Figs. 4 G and J) MTT assay results for 0-1.0mg/mL PLGA HOPMs concentrations showed no difference between the experimental and control groups, indicating the material is non-toxic; (Figs. 4 H and K) Different concentrations (0-1.5mg/mL) of PLGA microspheres promoted DPSC mineralization, suggesting PLGA HOPMs can promote osteogenic differentiation of dental pulp cells, with a certain positive correlation; (Figs. 4 I and L) Different concentrations of PLGA HOPMs had no impact on cell migration. (Figs. 4 M and N) PLGA HOPMs promoted the transcription of inflammatory genes IL-6 and IL-10 mRNA. (Figs. 4 O and P) PLGA HOPMs enhanced the transcription of mineralization genes ALP and OPG mRNA. (3) Subcutaneous implantation experiment in animals (Fig. 5 A-D) Dental pulp stem cells co-cultured with PLGA HOPMs demonstrated good integration; (Fig. 5 F) Dental pulp stem cells transfected with a lentivirus carrying GFP, maintained cell integrity and carried green fluorescent markers for tracking; (Fig. 5 G, H) Subcutaneous implantation of co-cultured PLGA HOPMs and cells showed no significant inflammation or necrosis in subcutaneous tissues; (Fig. 5 I, J) GFP expression in subcutaneous tissue sections after one week indicated survival of lentivirus-marked dental pulp cells within the microspheres; (Fig. 5 K) Over time, cells progressively infiltrated the interior of the PLGA HOPMs implanted subcutaneously, showing good biocompatibility; (Fig. 5 L, M) Masson staining revealed that PLGA HOPMs loaded with dental pulp stem cells promoted fibrous tissue formation; (Fig. 5 N) Silver staining around the edges of cell-loaded microspheres indicated more pronounced nerve tissue formation; Figures O, P, Q, R show immunohistochemistry and quantification of IL-6 and TNF-α, demonstrating that cell-loaded PLGA HOPMs can reduce TNF-α expression and inflammation in tissues. (4) Bone tissue implantation experiment in animals Figure 6 shows PLGA HOPMs co-cultured with DPSCs implanted into rat alveolar bone. (Figs. 6 A-D) Micro-CT images demonstrate that in the NC group, bone formation was mainly at the defect edges and less abundant, while in the PLGA group, internal bone formation was more pronounced than in the NC group. Cell-loaded specimens showed further enhanced internal bone formation. Gross observation indicated better healing in cell-loaded alveolar bone defects, as indicated by arrows. (Fig. 6 E) HE staining results showed more extensive internal tissue formation and cellular proliferation in the PLGA group compared to the NC group, with increasing vascular formation in cell-loaded specimens, as indicated by arrows. (Fig. 6 F) TRAP staining for osteoclasts showed no significant difference between groups. (Fig. 6 G) Hemolysis reactions were similar across all groups. (Figs. 6 H, I) Bone mineral density (BMD) and bone volume fraction (BV/TV) results indicated superior bone formation in the PLGA HOPMs group compared to the NC group, with cell loading further enhancing the osteogenic response of PLGA HOPMs. Figure 7 PLGA HOPMs and DPSCs co-cultured and implanted into rat alveolar bone. CD34 and VEGF are important markers for vascularization. (Figs. 7 A, E) PLGA HOPMs loaded with DPSCs promoted CD34 expression and vascular formation, with fluorescent localization indicating newly formed vessels inside the microspheres, as shown by arrows; (Figs. 7 B, F) VEGF staining results were consistent with CD34, showing more pronounced angiogenic response in microspheres loaded with a double concentration of DPSCs, as indicated by arrows; (Fig. 7 G) There was no difference in TNF-α expression among groups. (Figs. 7 C, H, D, I) Immunohistochemistry for OC and OPG showed enhanced mineralization response in the PLGA group, but cell loading did not further promote this effect; (Fig. 7 J) Weight change graphs indicated no significant difference between groups, demonstrating that the material implantation had no noticeable harm to the animals' health. Discussion Periodontitis, orthodontic treatment, and maxillofacial trauma can all lead to alveolar bone defects or atrophy. These defects are challenging to restore to their original height and volume after treatment. The implantation of tissue engineering materials aids in the ingress, differentiation, and reformation of bone tissue by stem cells, but this process is time-consuming. Inducing differentiation of stem cells within scaffold materials and forming a substantial amount of new tissue before implantation into the defect area can reduce this time. In this experiment, we prepared PLGA HOPMs with adjustable diameters and pore sizes (Figs. 2 , 3 G, H). The degradation rate in vitro can last over six months (Figs. 3 A-E, I-J), aligning with the requirements of oral clinical treatment. DPSCs were discovered relatively late and are often overlooked in dental medicine, not being isolated and cultured until 2000 by Gronthos et al. [10] . Identification of DPSCs primarily involves morphological characteristics and surface markers. As mesenchymal in origin, DPSCs exhibit mesenchymal stem cell markers without hematopoietic stem cell markers. In this study, the cultured cells appeared as spindle-shaped fibroblast-like cells (Figs. 1 A, B), with central nuclei, abundant cytoplasm, and large cell bodies, consistent with previous reports [10, 11] . Stro-1 and CD90, mesenchymal stem cell markers, and CD34, a hematopoietic stem cell marker, were assessed via immunofluorescence staining. The cultured cells showed positive expression of Stro-1 and CD90, while CD34 positivity was below 10%, indicating a low expression rate (Figs. 1AC, D, E). This suggests that the cultured cells are mesenchymal, not hematopoietic. The cell proliferation curve exhibited a typical 'S' shape (Fig. 1 F), indicating healthy proliferation. When DPSCs were co-cultured with PLGA HOPMS, cell cytotoxicity was assessed at 12–24 hours, and cell proliferation response was observed from day 1 to 7. It was found that PLGA HOPMS did not exhibit cytotoxicity towards DPSCs nor affect their proliferation, demonstrating the material's safety (Figs. 1 G, J). Mineralization assays showed that PLGA HOPMs could promote the formation of mineralized nodules by DPSCs, with a positive correlation with concentration (Figs. 1 H, K). These results align with our previous findings using PLGA nanoparticles to test the proliferation and mineralization capabilities of bone marrow mesenchymal stem cells, indicating non-toxicity and the ability to promote cell mineralization [12] . We also analyzed mineralization gene transcription, observing increased transcription of ALP and OPGmRNA, suggesting that the mechanism by which the material promotes mineralization might be related to this pathway. The material did not affect cell migration (Figs. 1 I, L). We assessed the material's impact on cellular immune responses by measuring IL-6 and IL-8mRNA expression (Figs. 1 M, N), finding increased expression of both, indicating the material's potential impact on cellular immune balance. We then co-cultured DPSCs with PLGA HOPMS to form microscale cell carriers and conducted subcutaneous experiments to assess the material's tissue compatibility and tissue formation capabilities. DPSCs were first transfected with lentivirus and marked with green fluorescence for future use (Figs. 2 E, F). Scanning electron microscopy and H&E staining revealed a tight integration and healthy growth of cells on the PLGA HOPMS (Figs. 3 A-E). When these microscale cell carriers were implanted subcutaneously in rats, no significant necrosis or inflammatory response was observed around the implants (Figs. 3 G, H), demonstrating that the PLGA scaffolds had excellent formability and were highly compatible as tissue engineering materials [13] . Upon retrieval of the implanted microscale cell carriers, DAPI staining showed robust green fluorescence expression of DPSCs, primarily concentrated within the microscale cell carriers, with no significant GFP + cells in the surrounding tissue, confirming that PLGA HOPMS can confine DPSCs to the damaged area (Figs. 3 I, J). H&E results revealed that over time, surrounding tissue grew into the microscale cell carriers, eventually integrating with them (Fig. 3 K). Masson staining indicated that the more DPSCs present, the more fibers formed in the microscale cell carriers, demonstrating DPSCs' ability to promote fiber formation (Figs. 3 L, M). Silver staining results showed that the more DPSCs, the more neurons formed in the microscale cell carriers, primarily at the interface between the microscale cell carriers and surrounding tissue, as indicated by the black areas in Fig. 3 N with arrows. This confirms that DPSC transplantation can enhance the tissue formation capabilities of the scaffold material [14] . Further study of DPSCs' immunomodulatory abilities revealed that microscale cell carriers with DPSCs significantly reduced TNF-α expression in the implants compared to the PLGA HOPMS group alone (Figs. 3 P, O), while the decrease in IL-6 expression was not statistically significant (Fig. 3 Q.R). We implanted microscale cell carriers into the alveolar bone of SD rats to explore their osteogenic and vasogenic effects. (Figs. 4 A, B, C, D) show that the group with DPSC-enriched microscale cell carriers exhibited superior bone defect healing compared to other groups, with a positive correlation observed with the concentration of DPSCs. MicroCT images revealed a higher density of material in the microscale cell carriers group, and results of bone mineral density (BMD) and bone volume/total volume (BV/TV) were consistent with the imaging, confirming DPSCs' role in promoting bone tissue formation. This aligns with other researchers' findings that DPSCs, as tissue engineering materials, can facilitate bone tissue repair [15] . H&E staining results were in agreement with MicroCT, showing cellular tissue within the PLGA HOPMS group and more extensive tissue formation, including vascular structures, within the microscale cell carriers group (Fig. 5 E). TRAP staining (Fig. 5 F) indicated no significant difference in osteoclasts between the experimental and control groups, possibly because the tissue samples were extracted one month post-implantation, a stage where bone tissue is in repair rather than osteoclast absorption. Hemolysis tests (Fig. 5 G) demonstrated that the materials implanted in each group did not significantly affect hemolysis. Further molecular-level investigations were conducted to elucidate the mechanisms behind osteogenesis and angiogenesis. Research by Evandro Piva et al. suggests that DPSCs promote tissue vascularization by secreting various angiogenic factors [16] . We performed VEGF and CD34 staining on bone tissue; immunofluorescence (Figs. 5 A, B) showed stronger VEGF and CD34 expression in the microscale cell carriers group. The expression and localization of VEGF and CD34 indicated that higher DPSC content correlated with more evident internal vascular formation. TNF-α immunohistochemistry revealed no significant difference in expression between the experimental and control groups, inconsistent with subcutaneous experiment results. This discrepancy may be due to the subcutaneous tissue being analyzed at one week and bone tissue at one month, by which time the material might have been tolerated by the body. OC and OPG are important molecules in the osteogenic pathway and indicators of bone formation [17, 18] . Immunohistochemical staining for OC and OPG revealed higher expression in both PLGA HOPMS and microscale cell carriers groups compared to the NC group, but no significant difference between the PLGA HOPMS and microscale cell carriers groups. This suggests that PLGA indeed facilitates osteogenesis. However, compared to the PLGA HOPMS group, the osteogenic effect of the DPSCs in the microscale cell carriers group may be more related to their early promotion of vascular formation rather than OC and OPG expression. Conclusion The prepared PLGA HOPMS are stable in form with an appropriate degradation rate; DPSCs proliferate rapidly with strong mineralization capabilities. PLGA HOPMS does not impede DPSC proliferation or migration and promotes their mineralization. PLGA HOPMS loaded with DPSCs to form microscale cell carriers enhance tissue fiber and neural tissue formation. The microscale cell carriers accelerate bone tissue and vascular formation promoted by PLGA HOPMS, without significant immune rejection. DPSCs, often regarded as medical waste, are excellent tool cells for forming microscale cell carriers, promoting vascular and bone tissue formation, and are safe for use. Declarations Authors’ contribution ， competing interest All authors read and approved the final manuscript. Availability of data and materials. All data generated or analysed during this study are included in this published article. Declaration of conflicting interests. The author(s) declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article. All authors read and approved the final manuscript. Renze Shen and Aizheng Chen contributed to conception and design of the study. YiCheng Wang contributed to data acquisition. Zhanchao Ye contributed to data analysis. Others are contributed to Interpretation、drafting and critical revision of the article, final approval of the version to be published, and agreed to be accountable for all aspects of the work. All authors contributed to data analysis, drafting or revising the article, gave final approval of the version to be published, and agree to be accountable for all aspects of the work. Ethical approval, consent to participate and publication The submission reported data collected from animals and human, and all studies were conducted according to the regulations for animal experimentation issued by the State Committee of Science and Technology of the People’s Republic of China. For any patient under the age of 18, a parent or legal guardian provided consent. Ethical approval for this study was obtained from Medical Ethics Committee of Zhangshan Hospital of Xiamen University (No xmzsyyky-2021019). The submission has reported data collected from animals and humans, and all studies were conducted according to the regulations for animal experimentation issued by the State Committee of Science and Technology of the People’s Republic of China. Funding The author(s) disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: This study was supported by Science and Technology Planning Project of Fujian province (2022j011347) and Fujian Key Laboratory of Oral Diseases , School and Hospital of Stomatology，Fujian Medical University (2021kq002). Acknowledge financial support from the National Natural Science Foundation of China (NSFC, 32071323, 81971734, and U1605225) 。 Data sharing statement All data generated or analyzed during this study are included in this published article. References YUHE J, PING Z, XIAO Z, et al. Advances in mesenchymal stem cell transplantation for the treatment of osteoporosis [J]. Cell proliferation, 2020, 54(1): e12956-e. EL-SAYED M E, ATWA A, SOFY A R, et al. Mesenchymal stem cell transplantation in burn wound healing: uncovering the mechanisms of local regeneration and tissue repair [J]. Histochem Cell Biol, 2023. 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A Systematic Review of the Circadian Rhythm of Bone Markers in Blood [J]. Calcif Tissue Int, 2023, 112(2): 126-47. Additional Declarations No competing interests reported. Cite Share Download PDF Status: Posted Version 1 posted 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. As a division of Research Square Company, we’re committed to making research communication faster, fairer, and more useful. We do this by developing innovative software and high quality services for the global research community. Our growing team is made up of researchers and industry professionals working together to solve the most critical problems facing scientific publishing. Also discoverable on Platform About Our Team In Review Editorial Policies 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-3921403","acceptedTermsAndConditions":true,"allowDirectSubmit":true,"archivedVersions":[],"articleType":"Research Article","associatedPublications":[],"authors":[{"id":270974409,"identity":"19a8dcfa-f6e7-4e33-ac17-abf937382428","order_by":0,"name":"RenZe Shen","email":"","orcid":"","institution":"Zhongshan Hospital of Xiamen University, Fujian Medical University","correspondingAuthor":false,"submittingAuthor":false,"prefix":"","firstName":"RenZe","middleName":"","lastName":"Shen","suffix":""},{"id":270974410,"identity":"2d9d3101-595e-41e0-aeea-6fa1e0c8fd62","order_by":1,"name":"YiCheng Wang","email":"","orcid":"","institution":"Huaqiao University","correspondingAuthor":false,"submittingAuthor":false,"prefix":"","firstName":"YiCheng","middleName":"","lastName":"Wang","suffix":""},{"id":270974411,"identity":"d1574895-b8b3-4ec2-88a6-13a9e2d75381","order_by":2,"name":"JiaCheng Huang","email":"","orcid":"","institution":"Guangdong Hospital of Traditional Chinese Medicine.guangdong","correspondingAuthor":false,"submittingAuthor":false,"prefix":"","firstName":"JiaCheng","middleName":"","lastName":"Huang","suffix":""},{"id":270974412,"identity":"f561b7d9-dd66-4b08-bc05-3a00883c19a8","order_by":3,"name":"YongMei Tan","email":"","orcid":"","institution":"Guangdong Hospital of Traditional Chinese Medicine.guangdong","correspondingAuthor":false,"submittingAuthor":false,"prefix":"","firstName":"YongMei","middleName":"","lastName":"Tan","suffix":""},{"id":270974413,"identity":"f1603ca3-48e6-42ac-a9de-f638eac3bcba","order_by":4,"name":"SongXia Ke","email":"","orcid":"","institution":"Zhongshan Hospital of Xiamen University, Fujian Medical University","correspondingAuthor":false,"submittingAuthor":false,"prefix":"","firstName":"SongXia","middleName":"","lastName":"Ke","suffix":""},{"id":270974414,"identity":"858395b6-c2b6-4a77-82f6-31ebf220c49b","order_by":5,"name":"ZiQing Zhou","email":"","orcid":"","institution":"Zhongshan Hospital of Xiamen University, Fujian Medical University","correspondingAuthor":false,"submittingAuthor":false,"prefix":"","firstName":"ZiQing","middleName":"","lastName":"Zhou","suffix":""},{"id":270974415,"identity":"d3242b4c-a66e-4fce-8acd-8c5cda7f58da","order_by":6,"name":"WeiWen Zheng","email":"data:image/png;base64,iVBORw0KGgoAAAANSUhEUgAAAZAAAAAyAQMAAABI0h/eAAAABlBMVEX///8AAABVwtN+AAAACXBIWXMAAA7EAAAOxAGVKw4bAAAA0UlEQVRIiWNgGAWjYBACNmbm4z8/VPznkWdvPkCcFj72tgRpiTPMcoY9xxKI0yLHc0ZBgreN2ZjhRo4BkQ6TyGEwkDjDltg4I+fjjTcMdnK6DQS15B5IKKjgSWznebvZcg5DsrHZAYJa8hIOSJyRSGxsz90mzcNwIHEbYS05hg28bQaJDQdynhGpheeMMQNvW4Ixw4kcNiK1sLelMUucOQAKZGPLOQZE+EW+mfkY44eKA6CofHjjTYWdHEEtKECCh8ioQdZCqo5RMApGwSgYEQAAGvBB8zvrJbwAAAAASUVORK5CYII=","orcid":"","institution":"Zhongshan Hospital of Xiamen University, Fujian Medical University","correspondingAuthor":true,"submittingAuthor":false,"prefix":"","firstName":"WeiWen","middleName":"","lastName":"Zheng","suffix":""},{"id":270974416,"identity":"16db79b5-81fb-420f-8976-1d8babdca6a0","order_by":7,"name":"AiZheng Chen","email":"","orcid":"","institution":"Huaqiao University","correspondingAuthor":false,"submittingAuthor":false,"prefix":"","firstName":"AiZheng","middleName":"","lastName":"Chen","suffix":""},{"id":270974417,"identity":"63485248-6739-46bc-8394-0163e9af3f41","order_by":8,"name":"ZhanChao Ye","email":"","orcid":"","institution":"Zhongshan Hospital of Xiamen University, Fujian Medical University","correspondingAuthor":false,"submittingAuthor":false,"prefix":"","firstName":"ZhanChao","middleName":"","lastName":"Ye","suffix":""}],"badges":[],"createdAt":"2024-02-02 15:29:43","currentVersionCode":1,"declarations":"","doi":"10.21203/rs.3.rs-3921403/v1","doiUrl":"https://doi.org/10.21203/rs.3.rs-3921403/v1","draftVersion":[],"editorialEvents":[],"editorialNote":"","failedWorkflow":false,"files":[{"id":50726662,"identity":"6dc65321-b1bf-441c-8656-aec9de6347cc","added_by":"auto","created_at":"2024-02-06 11:36:13","extension":"png","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":399615,"visible":true,"origin":"","legend":"\u003cp\u003eA:Material preparation diagram; B: Bone tissue implantation diagram; C: Experimental flow chart.\u003c/p\u003e","description":"","filename":"floatimage1.png","url":"https://assets-eu.researchsquare.com/files/rs-3921403/v1/b2871a861fa2e4805e781a99.png"},{"id":50726663,"identity":"125549bd-bf06-4bc8-882c-c967843a754b","added_by":"auto","created_at":"2024-02-06 11:36:13","extension":"png","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":1515547,"visible":true,"origin":"","legend":"\u003cp\u003eScanning electron microscopy images\u003c/p\u003e","description":"","filename":"floatimage2.png","url":"https://assets-eu.researchsquare.com/files/rs-3921403/v1/43e01b466a22ecdd0f3c11b0.png"},{"id":50726666,"identity":"0301de0a-e1c8-4528-9d38-f84b2b09cd8e","added_by":"auto","created_at":"2024-02-06 11:36:14","extension":"png","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":1035768,"visible":true,"origin":"","legend":"\u003cp\u003ePhysicochemical Performance Testing; A-E: SEM Images of Lyophilized Microspheres at Different Degradation Times (0.5, 2, 4, 6, 8 months), Scale: 200μm; F: pH Value Changes During Degradation; G: Particle Size Distribution of the Prepared Microspheres; H: Pore Size Distribution of Microspheres Prepared under Group G Experimental Conditions; I-J: Weight Changes of Microspheres Under Static and Dynamic Degradation Conditions.\u003c/p\u003e","description":"","filename":"floatimage3.png","url":"https://assets-eu.researchsquare.com/files/rs-3921403/v1/5adac2448a72de35cf0515f1.png"},{"id":50726664,"identity":"f80c9681-550b-4bbe-a127-0842f9ab009d","added_by":"auto","created_at":"2024-02-06 11:36:14","extension":"png","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":2221146,"visible":true,"origin":"","legend":"\u003cp\u003eA: Primary culture of rat dental pulp stem cells; B: Third-generation dental pulp stem cells; C Stro-1 fluorescence staining of dental pulp stem cells; D: CD-90 fluorescent staining of dental pulp stem cells; E: Flow cytometry detection of CD34; F: Growth curve of dental pulp stem cells; G: MTT assay for cell cytotoxicity; H, K: Mineralization of dental pulp stem cells induced by microspheres and corresponding quantification; I, L: Impact of PLGA microspheres on dental pulp stem cell migration and corresponding quantification; J: MTT assay for cell proliferation at different microsphere concentrations; M-P: RT-PCR detection of mRNA expression.\u003c/p\u003e","description":"","filename":"floatimage4.png","url":"https://assets-eu.researchsquare.com/files/rs-3921403/v1/80fec350855405c505a8b0d6.png"},{"id":50727777,"identity":"4d629709-1d19-4528-9cff-90cab4656b6d","added_by":"auto","created_at":"2024-02-06 11:52:14","extension":"png","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":3479604,"visible":true,"origin":"","legend":"\u003cp\u003eA: Panoramic HE staining of dental pulp stem cells co-cultured with PLGA HOPMs ×10; B: Zoomed-in HE staining of co-cultured dental pulp stem cells and PLGA HOPMs ×40; C: Scanning electron microscopy of co-cultured cells and PLGA HOPMs after 3 days; D: Scanning electron microscopy after 7 days; E: Third-generation dental pulp stem cells; F: Third-generation dental pulp stem cells transduced with GFP lentivirus; G: Subcutaneous implantation in rats of co-cultured PLGA HOPMs and virus-transduced cells; H: Subcutaneous tissue block 7 days post-implantation; I: GFP expression in subcutaneous tissue sections under a fluorescence microscope; J: GFP and DAPI staining in subcutaneous tissue sections; K: HE staining of PLGA HOPMs implanted subcutaneously at different time points; L, M: Masson staining and quantification of cell concentration and PLGA HOPMs post-subcutaneous implantation; N: Silver staining of cell-loaded PLGA HOPMs subcutaneously implanted; O, P, Q, R: Immunohistochemistry and quantification for IL-6 and TNF-α.\u003c/p\u003e","description":"","filename":"floatimage5.png","url":"https://assets-eu.researchsquare.com/files/rs-3921403/v1/9ae848f47fcc63f6c5f50fb0.png"},{"id":50726665,"identity":"9f6e40e0-9523-4e5e-87d3-a0e8117cb794","added_by":"auto","created_at":"2024-02-06 11:36:14","extension":"png","order_by":6,"title":"Figure 6","display":"","copyAsset":false,"role":"figure","size":2104647,"visible":true,"origin":"","legend":"\u003cp\u003ePLGA HOPMs and DPSCs co-cultured and implanted into rat alveolar bone. A: NC group; B: PLGA HOPMs; C: PLGA HOPMs+Cells; D: PLGA Microspheres+2Cells; E: HE staining; F: TRAP staining; G: Hemolysis reaction monitoring; H: Bone mineral density (BMD); I: Bone volume fraction (BV/TV)..\u003c/p\u003e","description":"","filename":"floatimage6.png","url":"https://assets-eu.researchsquare.com/files/rs-3921403/v1/ccbb1c654aaa2a1cdcdd391c.png"},{"id":50727170,"identity":"8bec2aeb-4196-42d8-a5a5-19e58881d756","added_by":"auto","created_at":"2024-02-06 11:44:14","extension":"png","order_by":7,"title":"Figure 7","display":"","copyAsset":false,"role":"figure","size":2333341,"visible":true,"origin":"","legend":"\u003cp\u003epresents histological sections of PLGA+DPSCs bone tissue; A, E CD34 immunofluorescence staining and corresponding quantitative analysis; B, F, G VEGF (red) and TNF-α (green) immunofluorescence staining with quantitative analysis; C, H osteocalcin (OC) immunohistochemical staining and its quantification; D, I osteoprotegerin (OPG) immunohistochemical staining and quantification; and J body weight changes.\u003c/p\u003e","description":"","filename":"floatimage7.png","url":"https://assets-eu.researchsquare.com/files/rs-3921403/v1/781c88a09cc726260c1e7c1d.png"},{"id":51400211,"identity":"82d460b1-139c-423a-881b-44aa105f0f23","added_by":"auto","created_at":"2024-02-20 22:22:28","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":10051463,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-3921403/v1/0b2aa53f-5fdb-45cf-bf77-7c38535601e5.pdf"}],"financialInterests":"No competing interests reported.","formattedTitle":"The Role and Mechanism of Dental Pulp Stem Cells in Highly Porous Microcarriers Forming New Micro-tissues to Accelerate Mandibular Bone Defect Repair","fulltext":[{"header":"Introduction","content":"\u003cp\u003eInadequate alveolar bone is a significant factor affecting the prognosis of periodontal therapy, implant surgery outcomes, and the stability of orthodontically treated teeth. However, once lost, the alveolar bone cannot naturally regenerate its original height and shape. Under natural conditions, bone marrow mesenchymal stem cells (BMSCs) are recruited to the defect area and differentiate into osteoblasts to form bone tissue. Nevertheless, in such case, due to the lack of structural support in the defect area, the restored height of the alveolar bone is challenging to achieve. Scaffold-supported stem cell transplantation techniques are key to resolving this issue.\u003c/p\u003e \u003cp\u003eMesenchymal stem cell transplantation has several advantages over other cell transplantation methods. First, these cells can migrate to the injury site and modulate the immune response, enhancing immunity when it is low and downregulating it when it is high. Second, mesenchymal stem cell transplantation is expected to increase osteoblast differentiation, inhibit osteoclast activation, and rebalance bone formation and absorption\u003csup\u003e[1]\u003c/sup\u003e. Third, mesenchymal stem cells can differentiate in multiple directions. For instance, in burn treatment, transplanting these cells onto burned skin significantly reduces local inflammation and promotes angiogenesis\u003csup\u003e[2]\u003c/sup\u003e. In treating liver cirrhosis, mesenchymal stem cell transplantation can foster liver tissue regeneration\u003csup\u003e[3]\u003c/sup\u003e. BMSCs are also crucial for new bone tissue formation and are often the preferred choice for repairing bone tissue defects.\u003c/p\u003e \u003cp\u003eDental pulp stem cells (DPSCs), as a type of mesenchymal stem cell, share embryonic homology with bone marrow mesenchymal stem cells. In current stem cell transplantation practices, DPSCs are often overlooked. In reality, throughout a person's life, numerous teeth are extracted, ranging from deciduous teeth to wisdom teeth and teeth removed due to orthodontic treatment or periodontitis in adults. Unfortunately, these extracted teeth are typically discarded. A single tooth can contain enough DPSCs for 2\u0026ndash;3 stem cell transplantations, and these cells exhibit potent proliferation and differentiation abilities\u003csup\u003e[4]\u003c/sup\u003e. Their strong paracrine effects also play a significant role in injury repair\u003csup\u003e[5]\u003c/sup\u003e. Autologous DPSCs, used as tool cells, avoid immune rejection and eliminate the need for additional surgeries required for harvesting other stem cells. Research indicates that DPSC transplantation can promote the recovery of hematopoietic function in aplastic anemia, enhancing treatment for hematopoietic failure and providing blood protection\u003csup\u003e[6]\u003c/sup\u003e. Additionally, when co-cultured with PLCL scaffolds to form micro-tissues, DPSCs can be used in nerve injury repair, leading to the formation of new neurons\u003csup\u003e[7]\u003c/sup\u003e. These applications are based on the multi-directional differentiation capabilities and immunomodulatory functions of DPSCs. Extensive experiments have also verified the effectiveness and safety of DPSCs as tool cells.\u003c/p\u003e \u003cp\u003eMicroscale cell carriers have demonstrated significant advantages in tissue injury repair. Comprising scaffold structures and stem cells, these carriers provide a 3D environment conducive to localized cell proliferation and differentiation. The differentiation and paracrine actions of stem cells create a microenvironment favorable for tissue repair. Our preliminary work involved preparing PLGA highly open porous microspheres (HOPMS) and co-culturing them with cells to form micro-tissues. We observed that their implantation promotes tissue vascularization and myofibrosis\u003csup\u003e[8, 9]\u003c/sup\u003e. The structure of PLGA HOPMS is loosely porous with large interstitial diameters, avoiding the drawbacks of insufficient internal nutrient supply and poor cell migration or internal formation in the scaffold. The spherical 3D structure also provides ample growth space for stem cells.\u003c/p\u003e \u003cp\u003eBased on this theories, we propose using PLGA HOPMS co-cultured with dental pulp stem cells (DPSCs) to create microscale cell carriers for repairing alveolar bone defects (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003e). Figure\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eA involves preparing PLGA HOPMS; culturing Sprague Dawley (SD) rat DPSCs and performing their identification; and forming microscale cell carriers by co-culturing DPSCs with PLGA HOPMS. Figure\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eB illustrates the implantation of these carriers into SD rat alveolar bone defects, utilizing the scaffold function of the carriers for tissue growth and the tissue repair properties of dental pulp stem cells to promote local calcification and vascular formation for bone tissue repair. Figure\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eC is a schematic of the experimental process, involving the physicochemical characterization of prepared PLGA HOPMS, such as degradation performance and characterization. This is followed by cell experiments to verify the material's cytotoxicity, mineralization ability, and cell immunity. Finally, subcutaneous and bone tissue implantation in animals are used to validate the material's osteogenic and angiogenic properties and immune response, exploring its function and mechanism.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e"},{"header":"Materials and Methods","content":"\u003cdiv id=\"Sec3\" class=\"Section2\"\u003e \u003ch2\u003eMaterials\u003c/h2\u003e \u003cp\u003eGelatin (from porcine skin, Type A), DMEM (HyClone), FBS (Thermo Fisher Scientific)。PEGDA (Mw: 6\u0026ndash;7 kDa), polyvinyl alcohol (PVA), SYBR Premix DimerEraser kit (Takara)。sodium alginate (SA, low viscosity),DMEM (HyClone). PrimeScript\u0026reg; RT reagent kit (Takara), PLGA (66\u0026ndash;107 kDa,lactide:glycolide 75:25) were purchased from SigmaAldrich (St. Louis, USA). RGD was obtained from the \u003cb\u003eChinese Peptide Company (Hangzhou, China).\u003c/b\u003e\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec4\" class=\"Section2\"\u003e \u003ch2\u003eMethods\u003c/h2\u003e \u003cdiv id=\"Sec5\" class=\"Section3\"\u003e \u003ch2\u003e(1) Preparation of Materials\u003c/h2\u003e \u003cp\u003e PLGA HOPMs were prepared by initially dissolving gelatin in 925\u0026micro;L of 1% PVA solution, followed by heating and stirring for 6 hours. 0.06g of PLGA was dissolved in 2220\u0026micro;L of dichloromethane. Once fully dissolved, it was mixed with the gelatin solution and processed with an ultrasonic disruptor for 90 seconds to form a uniform emulsion. This emulsion and 1% PVA solution were loaded into a microfluidic device and injected into a beaker containing 1% PVA under ice bath conditions. After microsphere formation, the mixture was left undisturbed for 1 hour, then the upper solution was gently stirred for another hour, and finally left in a refrigerator at 4\u0026deg;C overnight. The PVA was removed multiple times slowly, followed by 4\u0026ndash;5 washes with ultrapure water and heating in a water bath for 1 hour. After several more washes, the microspheres were transferred into a tube and frozen at -20℃ overnight, then lyophilized in a vacuum freeze dryer for 24 hours.A factorial experiment was designed using Minitab to investigate the effects of ultrasonic power, gelatin content, and flow rate on the morphology of the prepared microspheres (Table\u0026nbsp;\u003cspan refid=\"Tab1\" class=\"InternalRef\"\u003e1\u003c/span\u003e).\u003c/p\u003e \u003cp\u003e \u003cdiv class=\"gridtable\"\u003e\u003ctable float=\"Yes\" id=\"Tab1\" border=\"1\"\u003e \u003ccaption language=\"En\"\u003e \u003cdiv class=\"CaptionNumber\"\u003eTable 1\u003c/div\u003e \u003cdiv class=\"CaptionContent\"\u003e \u003cp\u003eMinitab factorial experimental design\u003c/p\u003e \u003c/div\u003e \u003c/caption\u003e \u003ccolgroup cols=\"4\"\u003e \u003cdiv align=\"left\" class=\"colspec\" colname=\"c1\" colnum=\"1\"\u003e\u003c/div\u003e \u003cdiv align=\"char\" char=\".\" class=\"colspec\" colname=\"c2\" colnum=\"2\"\u003e\u003c/div\u003e \u003cdiv align=\"char\" char=\".\" class=\"colspec\" colname=\"c3\" colnum=\"3\"\u003e\u003c/div\u003e \u003cdiv align=\"char\" char=\".\" class=\"colspec\" colname=\"c4\" colnum=\"4\"\u003e\u003c/div\u003e \u003cthead\u003e \u003ctr\u003e \u003cth align=\"left\" colname=\"c1\"\u003e\u0026nbsp;\u003c/th\u003e \u003cth align=\"left\" colname=\"c2\"\u003e \u003cp\u003eultrasonic power/W\u003c/p\u003e \u003c/th\u003e \u003cth align=\"left\" colname=\"c3\"\u003e \u003cp\u003egelatin/g\u003c/p\u003e \u003c/th\u003e \u003cth align=\"left\" colname=\"c4\"\u003e \u003cp\u003eflow velocity/(mL/min)\u003c/p\u003e \u003c/th\u003e \u003c/tr\u003e \u003c/thead\u003e \u003ctbody\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eA\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c2\"\u003e \u003cp\u003e200\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c3\"\u003e \u003cp\u003e0.05\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c4\"\u003e \u003cp\u003e0.1\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eB\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c2\"\u003e \u003cp\u003e200\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c3\"\u003e \u003cp\u003e0.05\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c4\"\u003e \u003cp\u003e0.5\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eC\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c2\"\u003e \u003cp\u003e200\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c3\"\u003e \u003cp\u003e0.10\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c4\"\u003e \u003cp\u003e0.5\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eD\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c2\"\u003e \u003cp\u003e200\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c3\"\u003e \u003cp\u003e0.10\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c4\"\u003e \u003cp\u003e0.1\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eE\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c2\"\u003e \u003cp\u003e400\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c3\"\u003e \u003cp\u003e0.05\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c4\"\u003e \u003cp\u003e0.1\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eF\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c2\"\u003e \u003cp\u003e400\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c3\"\u003e \u003cp\u003e0.05\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c4\"\u003e \u003cp\u003e0.5\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eG\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c2\"\u003e \u003cp\u003e300\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c3\"\u003e \u003cp\u003e0.075\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c4\"\u003e \u003cp\u003e0.3\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eH\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c2\"\u003e \u003cp\u003e400\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c3\"\u003e \u003cp\u003e0.10\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c4\"\u003e \u003cp\u003e0.1\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003ctr\u003e \u003ctd align=\"left\" colname=\"c1\"\u003e \u003cp\u003eI\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c2\"\u003e \u003cp\u003e400\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c3\"\u003e \u003cp\u003e0.10\u003c/p\u003e \u003c/td\u003e \u003ctd align=\"char\" char=\".\" colname=\"c4\"\u003e \u003cp\u003e0.5\u003c/p\u003e \u003c/td\u003e \u003c/tr\u003e \u003c/tbody\u003e \u003c/colgroup\u003e \u003c/table\u003e\u003c/div\u003e \u003c/p\u003e \u003c/div\u003e \u003c/div\u003e \u003cdiv id=\"Sec6\" class=\"Section2\"\u003e \u003ch2\u003e(2) Cell experiments:\u003c/h2\u003e \u003cp\u003eCell culture and identification: Dental pulps were extracted from 5-week-old Sprague Dawley rats in a sterile environment, minced and washed in PBS, then digested with Type I collagenase for 3 minutes. Digestion was stopped with 10% fetal bovine serum\u0026thinsp;+\u0026thinsp;culture medium, followed by washing and culturing in dishes. Cells were observed for migration. At the third passage, cells were stained for stro-1 and CD90, and CD34-positive cells were detected using flow cytometry. Cell proliferation was assessed on days 1, 3, 5, and 7 using the MTT assay.\u003c/p\u003e \u003cp\u003eCytotoxicity, proliferation, mineralization response, and migration experiments: Third-generation dental pulp stem cells were co-cultured with 0mg/mL, 0.25mg/mL, 0.50mg/mL, and 1.0mg/mL PLGA HOPMs. Cytotoxic responses were assessed at 0.5 and 1 day using the MTT assay, and effects on cell proliferation were evaluated on days 1, 3, 5, and 7. Cell migration was assessed using a Transwell chamber with DPSCs in the upper chamber and 0-0.15mg/mL PLGA HOPMs in the lower chamber, followed by crystal violet staining after 24 hours. Cells were co-cultured with PLGA HOPMs for 14 days, fixed with paraformaldehyde, and stained with Alizarin Red to assess mineralized nodule formation. RT-PCR was performed at set times to assess mRNA transcription in the material-cell co-cultures.\u003c/p\u003e \u003cdiv id=\"Sec7\" class=\"Section3\"\u003e \u003ch2\u003e(3) Subcutaneous implantation in animals:\u003c/h2\u003e \u003cp\u003eCells were transfected with a GFP-labeled lentivirus provided by Shanghai Jiama Pharmaceutical Technology Co., Ltd. The lentiviral expression vector was pGLV3/Hl/GFP\u0026thinsp;+\u0026thinsp;Puro with the sequence 5 '-ACTACCGTTGTTATAGGTG-3'. The packaging system included pGag/Pol, pRev, and pVSV-G, with 293T cells as the packaging cells. After transduction for one day, GFP\u0026thinsp;+\u0026thinsp;DPSCs were imaged using a fluorescence microscope. GFP\u0026thinsp;+\u0026thinsp;DPSCs were co-cultured with PLGA HOPMs and examined with HE staining after 7 days and scanning electron microscopy on days 3 and 7 for cell growth.\u003c/p\u003e \u003cp\u003eSD rats weighing 200-250g were purchased and acclimatized for two weeks. Under general anesthesia, microscale cell carriers formed from co-cultured PLGA HOPMs and GFP\u0026thinsp;+\u0026thinsp;dental pulp stem cells, as well as pure PLGA HOPMs, were implanted subcutaneously. The PLGA\u0026thinsp;+\u0026thinsp;cell group included a cell concentration of 3\u0026times;10\u003csup\u003e6\u003c/sup\u003e/mL, and the PLGA\u0026thinsp;+\u0026thinsp;2cell group had a concentration of 6\u0026times;10\u003csup\u003e6\u003c/sup\u003e/mL. Samples were removed after 7 days; sections were stained with DAPI to observe the growth of GFP\u0026thinsp;+\u0026thinsp;cells inside the PLGA HOPMs and the DAPI staining pattern. Tissues were removed on days 3, 7, 10, and 14 for HE staining to observe the growth of the implanted tissue. At one week post-implantation, samples were taken for Masson staining, silver staining, and immunohistochemical staining to assess collagen formation, nerve tissue development, and the expression of TNF-α and IL-6.\u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec8\" class=\"Section3\"\u003e \u003ch2\u003e(4) Animal bone tissue implantation experiment:\u003c/h2\u003e \u003cp\u003eCells were co-cultured with PLGA HOPMs for 7 days, with cell concentrations as previously described. Sprague Dawley rats weighing 200-250g were purchased and acclimatized for two weeks. Under general anesthesia, a dental bur was used to create a bone defect in the mandible, approximately 2 millimeters deep and 2.5 millimeters in diameter. The NC group did not receive any implantation, while the other groups were implanted with either PLGA HOPMs or microscale cell carriers co-cultured with cells. One month post-implantation, specimens were removed for gross observation, micro-CT analysis to calculate bone mineral density (BMD), and bone volume fraction (BV/TV); HE staining to observe tissue healing; TRAP staining to examine osteoclast activity; and to check for hemolysis in the animals. Immunofluorescence was used to observe the expression of CD34 and VEGF for angiogenic reaction and vascular formation location; immunohistochemistry to explore osteogenesis mechanisms through OC and OPG expression; and to assess bone tissue inflammatory response by measuring TNF-α expression. Animal weights were monitored to evaluate the implant's impact on their growth.\u003c/p\u003e \u003c/div\u003e \u003c/div\u003e"},{"header":"Results","content":"\u003cp\u003e(1) \u003cb\u003eMaterial preparation and physicochemical properties\u003c/b\u003e\u003c/p\u003e \u003cp\u003eFigure \u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003e shows SEM images of microspheres prepared under different experimental conditions, demonstrating that the microspheres had a uniform spherical shape, with diameters primarily around 550\u0026micro;m (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eG). Additionally, with varying experimental conditions, the porosity and pore size of the microspheres changed. We selected group G from the factorial experiment, which had higher porosity and pore sizes primarily between 30\u0026ndash;50\u0026micro;m (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eH), for subsequent experiments.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eTo verify the stability and degradation capability of the prepared microspheres, they were stored in PBS, samples at different time intervals, lyophilized, and SEM characterization was performed to observe changes in microsphere morphology (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eA-E). The results showed that in the first stage of degradation, the spheres largely retained their shape, with finer structures on the surface beginning to degrade. In the second stage, the spherical structure of the microspheres gradually collapsed, with fragments breaking off. By the third stage, the spherical structure of the microspheres had completely disappeared, resulting in a large number of fragments. The pH changes in the solution during the degradation process were also characterized (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eF), revealing no significant change in pH over the 8-month degradation period, maintaining a neutral environment around 7.2\u0026ndash;7.4. Additionally, the degradation of the microspheres under static and dynamic conditions was characterized (Figs.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eI-J), showing a faster degradation rate under dynamic conditions, with most of the material degrading by the 8th month.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cdiv id=\"Sec10\" class=\"Section2\"\u003e \u003ch2\u003e(2) Cell experiments\u003c/h2\u003e \u003cp\u003e(Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eA) DPSCs migrated out from the tissue block and were in good condition. (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eB) The fixed third-generation cells were in good condition. Compared to (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eC) cells positive for stro-1, more than 95% were stro-1 positive. (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eD) DPSCs stained for CD90 showed positive responses, confirming the cells are mesenchymal stem cells. Flow cytometry detected about 9.66% CD34-positive cells, indicating low expression and excluding hematopoietic tissue origin. These experiments confirmed the cultivation of DPSCs. (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eF) The cell proliferation curve was \"S\"-shaped, typical of fibroblast growth patterns. (Figs.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eG and J) MTT assay results for 0-1.0mg/mL PLGA HOPMs concentrations showed no difference between the experimental and control groups, indicating the material is non-toxic; (Figs.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eH and K) Different concentrations (0-1.5mg/mL) of PLGA microspheres promoted DPSC mineralization, suggesting PLGA HOPMs can promote osteogenic differentiation of dental pulp cells, with a certain positive correlation; (Figs.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eI and L) Different concentrations of PLGA HOPMs had no impact on cell migration. (Figs.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eM and N) PLGA HOPMs promoted the transcription of inflammatory genes IL-6 and IL-10 mRNA. (Figs.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eO and P) PLGA HOPMs enhanced the transcription of mineralization genes ALP and OPG mRNA.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec11\" class=\"Section2\"\u003e \u003ch2\u003e(3) Subcutaneous implantation experiment in animals\u003c/h2\u003e \u003cp\u003e(Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eA-D) Dental pulp stem cells co-cultured with PLGA HOPMs demonstrated good integration; (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eF) Dental pulp stem cells transfected with a lentivirus carrying GFP, maintained cell integrity and carried green fluorescent markers for tracking; (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eG, H) Subcutaneous implantation of co-cultured PLGA HOPMs and cells showed no significant inflammation or necrosis in subcutaneous tissues; (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eI, J) GFP expression in subcutaneous tissue sections after one week indicated survival of lentivirus-marked dental pulp cells within the microspheres; (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eK) Over time, cells progressively infiltrated the interior of the PLGA HOPMs implanted subcutaneously, showing good biocompatibility; (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eL, M) Masson staining revealed that PLGA HOPMs loaded with dental pulp stem cells promoted fibrous tissue formation; (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eN) Silver staining around the edges of cell-loaded microspheres indicated more pronounced nerve tissue formation; Figures O, P, Q, R show immunohistochemistry and quantification of IL-6 and TNF-α, demonstrating that cell-loaded PLGA HOPMs can reduce TNF-α expression and inflammation in tissues.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e \u003cdiv id=\"Sec12\" class=\"Section2\"\u003e \u003ch2\u003e(4) Bone tissue implantation experiment in animals\u003c/h2\u003e \u003cp\u003eFigure \u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003e shows PLGA HOPMs co-cultured with DPSCs implanted into rat alveolar bone. (Figs.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eA-D) Micro-CT images demonstrate that in the NC group, bone formation was mainly at the defect edges and less abundant, while in the PLGA group, internal bone formation was more pronounced than in the NC group. Cell-loaded specimens showed further enhanced internal bone formation. Gross observation indicated better healing in cell-loaded alveolar bone defects, as indicated by arrows. (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eE) HE staining results showed more extensive internal tissue formation and cellular proliferation in the PLGA group compared to the NC group, with increasing vascular formation in cell-loaded specimens, as indicated by arrows. (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eF) TRAP staining for osteoclasts showed no significant difference between groups. (Fig.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eG) Hemolysis reactions were similar across all groups. (Figs.\u0026nbsp;\u003cspan refid=\"Fig6\" class=\"InternalRef\"\u003e6\u003c/span\u003eH, I) Bone mineral density (BMD) and bone volume fraction (BV/TV) results indicated superior bone formation in the PLGA HOPMs group compared to the NC group, with cell loading further enhancing the osteogenic response of PLGA HOPMs.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003cp\u003eFigure \u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003e PLGA HOPMs and DPSCs co-cultured and implanted into rat alveolar bone. CD34 and VEGF are important markers for vascularization. (Figs.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eA, E) PLGA HOPMs loaded with DPSCs promoted CD34 expression and vascular formation, with fluorescent localization indicating newly formed vessels inside the microspheres, as shown by arrows; (Figs.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eB, F) VEGF staining results were consistent with CD34, showing more pronounced angiogenic response in microspheres loaded with a double concentration of DPSCs, as indicated by arrows; (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eG) There was no difference in TNF-α expression among groups. (Figs.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eC, H, D, I) Immunohistochemistry for OC and OPG showed enhanced mineralization response in the PLGA group, but cell loading did not further promote this effect; (Fig.\u0026nbsp;\u003cspan refid=\"Fig7\" class=\"InternalRef\"\u003e7\u003c/span\u003eJ) Weight change graphs indicated no significant difference between groups, demonstrating that the material implantation had no noticeable harm to the animals' health.\u003c/p\u003e \u003cp\u003e \u003c/p\u003e \u003c/div\u003e"},{"header":"Discussion","content":"\u003cp\u003ePeriodontitis, orthodontic treatment, and maxillofacial trauma can all lead to alveolar bone defects or atrophy. These defects are challenging to restore to their original height and volume after treatment. The implantation of tissue engineering materials aids in the ingress, differentiation, and reformation of bone tissue by stem cells, but this process is time-consuming. Inducing differentiation of stem cells within scaffold materials and forming a substantial amount of new tissue before implantation into the defect area can reduce this time. In this experiment, we prepared PLGA HOPMs with adjustable diameters and pore sizes (Figs.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003e, \u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eG, H). The degradation rate in vitro can last over six months (Figs.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eA-E, I-J), aligning with the requirements of oral clinical treatment.\u003c/p\u003e \u003cp\u003eDPSCs were discovered relatively late and are often overlooked in dental medicine, not being isolated and cultured until 2000 by Gronthos et al.\u003csup\u003e[10]\u003c/sup\u003e. Identification of DPSCs primarily involves morphological characteristics and surface markers. As mesenchymal in origin, DPSCs exhibit mesenchymal stem cell markers without hematopoietic stem cell markers. In this study, the cultured cells appeared as spindle-shaped fibroblast-like cells (Figs.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eA, B), with central nuclei, abundant cytoplasm, and large cell bodies, consistent with previous reports\u003csup\u003e[10, 11]\u003c/sup\u003e. Stro-1 and CD90, mesenchymal stem cell markers, and CD34, a hematopoietic stem cell marker, were assessed via immunofluorescence staining. The cultured cells showed positive expression of Stro-1 and CD90, while CD34 positivity was below 10%, indicating a low expression rate (Figs.\u0026nbsp;1AC, D, E). This suggests that the cultured cells are mesenchymal, not hematopoietic. The cell proliferation curve exhibited a typical 'S' shape (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eF), indicating healthy proliferation.\u003c/p\u003e \u003cp\u003eWhen DPSCs were co-cultured with PLGA HOPMS, cell cytotoxicity was assessed at 12–24 hours, and cell proliferation response was observed from day 1 to 7. It was found that PLGA HOPMS did not exhibit cytotoxicity towards DPSCs nor affect their proliferation, demonstrating the material's safety (Figs.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eG, J). Mineralization assays showed that PLGA HOPMs could promote the formation of mineralized nodules by DPSCs, with a positive correlation with concentration (Figs.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eH, K). These results align with our previous findings using PLGA nanoparticles to test the proliferation and mineralization capabilities of bone marrow mesenchymal stem cells, indicating non-toxicity and the ability to promote cell mineralization\u003csup\u003e[12]\u003c/sup\u003e. We also analyzed mineralization gene transcription, observing increased transcription of ALP and OPGmRNA, suggesting that the mechanism by which the material promotes mineralization might be related to this pathway. The material did not affect cell migration (Figs.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eI, L). We assessed the material's impact on cellular immune responses by measuring IL-6 and IL-8mRNA expression (Figs.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003eM, N), finding increased expression of both, indicating the material's potential impact on cellular immune balance.\u003c/p\u003e \u003cp\u003eWe then co-cultured DPSCs with PLGA HOPMS to form microscale cell carriers and conducted subcutaneous experiments to assess the material's tissue compatibility and tissue formation capabilities. DPSCs were first transfected with lentivirus and marked with green fluorescence for future use (Figs.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eE, F). Scanning electron microscopy and H\u0026amp;E staining revealed a tight integration and healthy growth of cells on the PLGA HOPMS (Figs.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eA-E). When these microscale cell carriers were implanted subcutaneously in rats, no significant necrosis or inflammatory response was observed around the implants (Figs.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eG, H), demonstrating that the PLGA scaffolds had excellent formability and were highly compatible as tissue engineering materials\u003csup\u003e[13]\u003c/sup\u003e. Upon retrieval of the implanted microscale cell carriers, DAPI staining showed robust green fluorescence expression of DPSCs, primarily concentrated within the microscale cell carriers, with no significant GFP + cells in the surrounding tissue, confirming that PLGA HOPMS can confine DPSCs to the damaged area (Figs.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eI, J). H\u0026amp;E results revealed that over time, surrounding tissue grew into the microscale cell carriers, eventually integrating with them (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eK). Masson staining indicated that the more DPSCs present, the more fibers formed in the microscale cell carriers, demonstrating DPSCs' ability to promote fiber formation (Figs.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eL, M). Silver staining results showed that the more DPSCs, the more neurons formed in the microscale cell carriers, primarily at the interface between the microscale cell carriers and surrounding tissue, as indicated by the black areas in Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eN with arrows. This confirms that DPSC transplantation can enhance the tissue formation capabilities of the scaffold material\u003csup\u003e[14]\u003c/sup\u003e. Further study of DPSCs' immunomodulatory abilities revealed that microscale cell carriers with DPSCs significantly reduced TNF-α expression in the implants compared to the PLGA HOPMS group alone (Figs.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eP, O), while the decrease in IL-6 expression was not statistically significant (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eQ.R).\u003c/p\u003e \u003cp\u003eWe implanted microscale cell carriers into the alveolar bone of SD rats to explore their osteogenic and vasogenic effects. (Figs.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003eA, B, C, D) show that the group with DPSC-enriched microscale cell carriers exhibited superior bone defect healing compared to other groups, with a positive correlation observed with the concentration of DPSCs. MicroCT images revealed a higher density of material in the microscale cell carriers group, and results of bone mineral density (BMD) and bone volume/total volume (BV/TV) were consistent with the imaging, confirming DPSCs' role in promoting bone tissue formation. This aligns with other researchers' findings that DPSCs, as tissue engineering materials, can facilitate bone tissue repair\u003csup\u003e[15]\u003c/sup\u003e. H\u0026amp;E staining results were in agreement with MicroCT, showing cellular tissue within the PLGA HOPMS group and more extensive tissue formation, including vascular structures, within the microscale cell carriers group (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eE). TRAP staining (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eF) indicated no significant difference in osteoclasts between the experimental and control groups, possibly because the tissue samples were extracted one month post-implantation, a stage where bone tissue is in repair rather than osteoclast absorption. Hemolysis tests (Fig.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eG) demonstrated that the materials implanted in each group did not significantly affect hemolysis.\u003c/p\u003e \u003cp\u003eFurther molecular-level investigations were conducted to elucidate the mechanisms behind osteogenesis and angiogenesis. Research by Evandro Piva et al. suggests that DPSCs promote tissue vascularization by secreting various angiogenic factors\u003csup\u003e[16]\u003c/sup\u003e. We performed VEGF and CD34 staining on bone tissue; immunofluorescence (Figs.\u0026nbsp;\u003cspan refid=\"Fig5\" class=\"InternalRef\"\u003e5\u003c/span\u003eA, B) showed stronger VEGF and CD34 expression in the microscale cell carriers group. The expression and localization of VEGF and CD34 indicated that higher DPSC content correlated with more evident internal vascular formation. TNF-α immunohistochemistry revealed no significant difference in expression between the experimental and control groups, inconsistent with subcutaneous experiment results. This discrepancy may be due to the subcutaneous tissue being analyzed at one week and bone tissue at one month, by which time the material might have been tolerated by the body. OC and OPG are important molecules in the osteogenic pathway and indicators of bone formation\u003csup\u003e[17, 18]\u003c/sup\u003e. Immunohistochemical staining for OC and OPG revealed higher expression in both PLGA HOPMS and microscale cell carriers groups compared to the NC group, but no significant difference between the PLGA HOPMS and microscale cell carriers groups. This suggests that PLGA indeed facilitates osteogenesis. However, compared to the PLGA HOPMS group, the osteogenic effect of the DPSCs in the microscale cell carriers group may be more related to their early promotion of vascular formation rather than OC and OPG expression.\u003c/p\u003e \u003cp\u003e\u003c/p\u003e \u003cdiv id=\"Sec14\" class=\"Section2\"\u003e \u003cp\u003e\u003c/p\u003e \u003c/div\u003e"},{"header":"Conclusion","content":"\u003cp\u003eThe prepared PLGA HOPMS are stable in form with an appropriate degradation rate; DPSCs proliferate rapidly with strong mineralization capabilities. PLGA HOPMS does not impede DPSC proliferation or migration and promotes their mineralization. PLGA HOPMS loaded with DPSCs to form microscale cell carriers enhance tissue fiber and neural tissue formation. The microscale cell carriers accelerate bone tissue and vascular formation promoted by PLGA HOPMS, without significant immune rejection. DPSCs, often regarded as medical waste, are excellent tool cells for forming microscale cell carriers, promoting vascular and bone tissue formation, and are safe for use.\u003c/p\u003e"},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eAuthors\u0026rsquo; contribution\u003c/strong\u003e\u003cstrong\u003e，\u003c/strong\u003e\u003cstrong\u003ecompeting interest\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAll authors read and approved the final manuscript. Availability of data and materials. All data generated or analysed during this study are included in this published article. Declaration of conflicting interests. The author(s) declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eAll authors read and approved the final manuscript. Renze Shen and Aizheng Chen contributed to conception and design of the study. YiCheng Wang contributed to data acquisition. Zhanchao Ye contributed to data analysis. Others are contributed to Interpretation、drafting and critical revision of the article, final approval of the version to be published, and agreed to be accountable for all aspects of the work. All authors contributed to data analysis, drafting or revising the article, gave final approval of the version to be published, and agree to be accountable for all aspects of the work.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eEthical approval, consent to participate and publication\u0026nbsp;\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe submission reported data collected from animals and human, and all studies were conducted according to the regulations for animal experimentation issued by the State Committee of Science and Technology of the People\u0026rsquo;s\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eRepublic of China. For any patient under the age of 18, a parent or legal guardian provided consent. Ethical approval for this study was obtained from Medical Ethics Committee of Zhangshan Hospital of Xiamen University (No xmzsyyky-2021019). The submission has reported data collected from animals and humans, and all studies were conducted according to the regulations for animal experimentation issued by the \u003cstrong\u003eState Committee of Science and Technology of the People\u0026rsquo;s Republic of China.\u0026nbsp;\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eFunding\u0026nbsp;\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe author(s) disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: This study was supported by Science and Technology Planning Project of Fujian province (2022j011347) and Fujian Key Laboratory of Oral Diseases , School and Hospital of Stomatology，Fujian Medical University (2021kq002).\u0026nbsp;Acknowledge financial support from the National Natural Science Foundation of China (NSFC, 32071323, 81971734, and U1605225)\u0026nbsp;。\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eData sharing statement\u003c/strong\u003e\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eAll data generated or analyzed during this study are included in this published article.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\n\u003cli\u003eYUHE J, PING Z, XIAO Z, et al. Advances in mesenchymal stem cell transplantation for the treatment of osteoporosis [J]. Cell proliferation, 2020, 54(1): e12956-e.\u003c/li\u003e\n\u003cli\u003eEL-SAYED M E, ATWA A, SOFY A R, et al. Mesenchymal stem cell transplantation in burn wound healing: uncovering the mechanisms of local regeneration and tissue repair [J]. Histochem Cell Biol, 2023.\u003c/li\u003e\n\u003cli\u003eWATANABE Y, TSUCHIYA A, TERAI S. The development of mesenchymal stem cell therapy in the present, and the perspective of cell-free therapy in the future [J]. Clin Mol Hepatol, 2021, 27(1): 70-80.\u003c/li\u003e\n\u003cli\u003eZHANG X M, SUN Y, ZHOU Y L, et al. Therapeutic effects of dental pulp stem cells on vascular dementia in rat models [J]. Neural Regen Res, 2021, 16(8): 1645-51.\u003c/li\u003e\n\u003cli\u003eBAR J K, LIS-NAWARA A, GRELEWSKI P G. Dental Pulp Stem Cell-Derived Secretome and Its Regenerative Potential [J]. Int J Mol Sci, 2021, 22(21).\u003c/li\u003e\n\u003cli\u003eGONZAGA V F, WENCESLAU C V, VIEIRA D P, et al. Therapeutic Potential of Human Immature Dental Pulp Stem Cells Observed in Mouse Model for Acquired Aplastic Anemia [J]. Cells, 2022, 11(14).\u003c/li\u003e\n\u003cli\u003ePINEDA J R, POLO Y, PARDO-RODR\u0026iacute;GUEZ B, et al. In vitro preparation of human Dental Pulp Stem Cell grafts with biodegradable polymer scaffolds for nerve tissue engineering [J]. Methods Cell Biol, 2022, 170: 147-67.\u003c/li\u003e\n\u003cli\u003eKANKALA R K, ZHAO J, LIU C G, et al. Highly Porous Microcarriers for Minimally Invasive In Situ Skeletal Muscle Cell Delivery [J]. Small, 2019, 15(25): e1901397.\u003c/li\u003e\n\u003cli\u003eWANG Y, KANKALA R K, CAI Y Y, et al. Minimally invasive co-injection of modular micro-muscular and micro-vascular tissues improves in situ skeletal muscle regeneration [J]. Biomaterials, 2021, 277: 121072.\u003c/li\u003e\n\u003cli\u003eGRONTHOS S, MANKANI M, BRAHIM J, et al. Postnatal human dental pulp stem cells (DPSCs) in vitro and in vivo [J]. Proc Natl Acad Sci U S A, 2000, 97(25): 13625-30.\u003c/li\u003e\n\u003cli\u003eMA D, GAO J, YUE J, et al. Changes in proliferation and osteogenic differentiation of stem cells from deep caries in vitro [J]. J Endod, 2012, 38(6): 796-802.\u003c/li\u003e\n\u003cli\u003eXUE Y, HONG X, GAO J, et al. Preparation and biological characterization of the mixture of poly(lactic-co-glycolic acid)/chitosan/Ag nanoparticles for periodontal tissue engineering [J]. Int J Nanomedicine, 2019, 14: 483-98.\u003c/li\u003e\n\u003cli\u003eBAZGIR M, ZHANG W, ZHANG X, et al. Degradation and Characterisation of Electrospun Polycaprolactone (PCL) and Poly(lactic-co-glycolic acid) (PLGA) Scaffolds for Vascular Tissue Engineering [J]. Materials (Basel), 2021, 14(17).\u003c/li\u003e\n\u003cli\u003eCARTER K, LEE H J, NA K S, et al. Characterizing the impact of 2D and 3D culture conditions on the therapeutic effects of human mesenchymal stem cell secretome on corneal wound healing in vitro and ex vivo [J]. Acta Biomater, 2019, 99: 247-57.\u003c/li\u003e\n\u003cli\u003eLORUSSO F, INCHINGOLO F, DIPALMA G, et al. Synthetic Scaffold/Dental Pulp Stem Cell (DPSC) Tissue Engineering Constructs for Bone Defect Treatment: An Animal Studies Literature Review [J]. Int J Mol Sci, 2020, 21(24).\u003c/li\u003e\n\u003cli\u003ePIVA E, TARL\u0026eacute; S A, N\u0026ouml;R J E, et al. Dental Pulp Tissue Regeneration Using Dental Pulp Stem Cells Isolated and Expanded in Human Serum [J]. J Endod, 2017, 43(4): 568-74.\u003c/li\u003e\n\u003cli\u003eYAO Z, GETTING S J, LOCKE I C. Regulation of TNF-Induced Osteoclast Differentiation [J]. Cells, 2021, 11(1).\u003c/li\u003e\n\u003cli\u003eDIEMAR S S, DAHL S S, WEST A S, et al. A Systematic Review of the Circadian Rhythm of Bone Markers in Blood [J]. Calcif Tissue Int, 2023, 112(2): 126-47.\u003c/li\u003e\n\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":true,"highlight":"","institution":"","isAcceptedByJournal":false,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"researchsquare","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":true,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"/submission","title":"Research Square","twitterHandle":"researchsquare","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"","reportingPortfolio":"","inReviewEnabled":false,"inReviewRevisionsEnabled":true},"keywords":"PLGA, Dental pulp stem cells, Osteogenesis, GBR","lastPublishedDoi":"10.21203/rs.3.rs-3921403/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-3921403/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003ch2\u003eObjective\u003c/h2\u003e \u003cp\u003eDental pulp stem cells (DPSCs) exhibit strong differentiation and proliferation capabilities but are often discarded. This experiment aimed to prepare PLGA (poly lactic-co-glycolic acid) highly open porous microspheres (HOPMS) and form microscale cell carriers with DPSCs, exploring their efficacy and mechanism in promoting alveolar bone defect repair.\u003c/p\u003e\u003ch2\u003eMethods\u003c/h2\u003e \u003cp\u003eRat dental pulp stem cells were cultured and identified. PLGA HOPMS were prepared and their porosity, pore size, degradation rate, and PH change were evaluated. Cytological experiments examined the effects of PLGA HOPMS on cell proliferation, mineralized nodule formation, mineralization gene, and inflammatory factor expression. Cells were transduced with lentivirus for fluorescence marking and co-cultured with PLGA HOPMS to form microscale cell carriers. Subcutaneous and intraosseous implantations were performed to investigate their biocompatibility, angiogenesis, and inflammatory response.\u003c/p\u003e\u003ch2\u003eResults\u003c/h2\u003e \u003cp\u003eThe size and pore diameter of PLGA HOPM were adjustable. PLGA HOPMS promoted the formation of mineralized nodules, mineralization gene expression, and exhibited no cytotoxicity. Subcutaneous experiments demonstrated that microscale cell carriers had excellent biocompatibility with no significant immune rejection. Intraosseous experiments confirmed that this material promoted osteogenesis and angiogenesis. Conclusion: PLGA HOPMS loaded with DPSCs effectively promoted bone and vascular formation in alveolar bone defect repair, proving to be an excellent stem cell transplantation carrier.\u003c/p\u003e","manuscriptTitle":"The Role and Mechanism of Dental Pulp Stem Cells in Highly Porous Microcarriers Forming New Micro-tissues to Accelerate Mandibular Bone Defect Repair","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2024-02-06 11:36:09","doi":"10.21203/rs.3.rs-3921403/v1","editorialEvents":[{"type":"communityComments","content":0}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"researchsquare","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":true,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"/submission","title":"Research Square","twitterHandle":"researchsquare","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"","reportingPortfolio":"","inReviewEnabled":false,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"c762bb82-b7a1-4ddb-afdb-abde1deda093","owner":[],"postedDate":"February 6th, 2024","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"posted","subjectAreas":[],"tags":[],"updatedAt":"2024-02-20T22:14:19+00:00","versionOfRecord":[],"versionCreatedAt":"2024-02-06 11:36:09","video":"","vorDoi":"","vorDoiUrl":"","workflowStages":[]},"version":"v1","identity":"rs-3921403","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-3921403","identity":"rs-3921403","version":["v1"]},"buildId":"zQwnuV7TCBrMSSSToR1PI","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}