Preparation method of optimized micron-sized cell-matrix co-polymer biological material

Abstract

The invention discloses a preparation method of a micron-sized cell-matrix co-polymer biological material. The preparation method comprises the following steps: firstly, inducing SHED cells in an ultra-low adsorption plate by using an alpha-MEM culture medium containing serum, GlutaMAXI, vitamin C and the like to form micron-sized spheres (SHED spheres); secondly, taking human dentin, and carrying out mechanical crushing, EDTA (Ethylene Diamine Tetraacetic Acid) demineralization, grinding and sterilization to prepare 1-100 microns of dentin matrix particles (DDMPs); and finally, by taking the GelMA hydrogel as a carrier, entrapping SHED spphere and DDMPs, and carrying out 405nm photo-crosslinking for 30 seconds to form the hydrogel. The co-polymer can significantly promote angiogenesis and osteogenesis coupling regeneration in a bone defect model, and is suitable for oral cavity and cranio-maxillofacial bone repair.

Description

Technical Field

[0001] This invention belongs to the field of biomaterials and tissue engineering technology, specifically involving decellularized matrix, three-dimensional culture, multi-site bone regeneration, and cell aggregates. Background Technology

[0002] The ever-growing clinical demand for tissue defect repair continues to drive innovation in regenerative medicine strategies and technologies. In the early stages of organ development, cell self-assembly plays a crucial role in morphogenesis, particularly during tooth and bone formation. Studies have shown that stem cells exhibit similar self-aggregation capabilities in vitro. Based on this, we constructed cell aggregates (CAs) from oral and maxillofacial stem cells—focusing on human exfoliated deciduous tooth stem cells (SHED) with high self-renewal capacity, multi-lineage differentiation potential, and low immunogenicity—to reproduce their developmental potential. These aggregates have shown promising applications in clinical studies such as pulp regeneration and whole-tooth functional regeneration. However, when implanted into pathological microenvironments such as periodontal bone defects, cell aggregates constructed using cell sheet technology exhibited poor repair effects, indicating a need to improve their preparation strategies and treatment methods.

[0003] To reconstruct a regenerative microenvironment, we previously proposed combining cell-associated matrix (CA) materials with specific biomimetic matrix materials to form cell-matrix copolymers. Specifically, our team has established a technical system for preparing human decellularized alveolar bone matrix particles (hDABMPs). hDABMPs are co-cultured with dental follicle stem cells (DFSCs) for 10-14 days to form copolymers, mimicking physiological cell-matrix interactions to promote periodontal bone tissue repair. However, this system has significant limitations in practical applications: 1) Although cell sheet technology preserves the natural cell-extracellular matrix (ECM) contact, the resulting two-dimensional polymers are too large, preparation is time-consuming, and the internal cells often face hypoxia or death due to limited diffusion of oxygen and nutrients; 2) Traditional bone-derived decellularized extracellular matrix (dECM) sources (such as human alveolar bone) are scarce and difficult to obtain, limiting large-scale production. To overcome these challenges, we turned to the more readily available dentin matrix (DM) as an alternative material. DM is highly similar to alveolar bone in mineral composition and biological properties, and can be easily obtained from premolars and third molars clinically extracted for orthodontic or impaction reasons, offering excellent availability and scalability. Using ultra-low adsorption (ULA) culture technology, we successfully prepared three-dimensional spherical cell aggregates. Compared to traditional two-dimensional methods, these spheres require shorter induction times, exhibit tighter cell-cell and cell-ECM connections, and are more cost-effective. However, the direct implantation of micron-sized cell spheres and dECM microparticles into the defect area still faces challenges such as low retention rates and spatial heterogeneity, limiting their efficacy in repairing large-volume bone defects in the skull and periodontal regions. Therefore, achieving stable integration of cells and matrix materials and their sustained retention at the defect site is crucial for advancing clinical translation.

[0004] Oral and craniofacial bone defects have a high incidence rate and present significant repair challenges, severely impacting physiological functions such as chewing, speech, and facial contours. Previous studies have shown that cell-microenvironment copolymers possess regenerative potential in periodontal bone repair. However, their repair efficacy and regulatory mechanisms in craniofacial bones of different developmental origins (such as skulls primarily derived from the mesoderm and partially from neural crest cells) remain unclear. Due to differences in embryonic origin and healing mechanisms among different bone sites, achieving rapid, stable, and widely applicable regenerative regulation remains a key challenge.

[0005] To address the aforementioned issues, this study aims to establish an optimized cell-matrix copolymer strategy using novel cell polymer technology and alternative decellularized dentin matrix particles (DDMPs) to achieve stable retention of cells and matrix in the defect area and synergistic angiogenesis-osteogenic coupling regeneration, thereby enhancing its repair efficacy and adaptability in complex oral and craniofacial bone defects. We hypothesize that this strategy can regulate angiogenesis-osteogenic coupling through specific molecular mechanisms, thereby promoting bone tissue regeneration in multiple sites, providing a new theoretical basis and technical pathway for clinical translation.

[0006] In the fields of tissue engineering and craniofacial bone regeneration, the cell-matrix copolymer strategy has become a research hotspot. Among existing technologies, a developmental biomimetic periodontal regeneration biomaterial and its preparation method and application (CN202410454946.9) proposes a developmental biomimetic periodontal regeneration biomaterial. Although it simulates the natural periodontal development process, it still has significant limitations in practical applications. This approach mainly relies on cell sheet technology to construct two-dimensional cell polymers, resulting in excessively large polymer sizes. Internal cells are prone to hypoxia or death due to limited oxygen and nutrient diffusion, severely affecting cell activity and regeneration performance. Furthermore, the decellularized matrix materials used are mostly derived from scarce tissues such as alveolar bone, making them difficult to obtain and hindering the large-scale preparation and clinical translation of the material. Another similar technical approach, a matrix-cell copolymer biomaterial and its preparation method and application (CN202411457044.7), proposes a matrix-cell copolymer construction concept, but it has shortcomings in terms of carrier systems and structural regulation. The lack of a suitable three-dimensional hydrogel carrier in this approach resulted in poor structural stability of the copolymer after implantation, making it prone to displacement or premature degradation, especially in areas with weight-bearing bone defects where effective retention and spatial distribution were difficult to achieve. Furthermore, the lack of systematic optimization of the particle size and concentration of decellularized matrix particles made the mechanical properties and degradation behavior of the composite uncontrollable, affecting the temporal coordination of the regeneration process and the repair effect. Summary of the Invention

[0007] (1) Technical content To address the shortcomings of the existing technologies mentioned above, this application aims to systematically solve the following key technical problems: First, micron-sized three-dimensional SHED cell spheres were constructed using ultra-low adsorption culture technology to replace the traditional two-dimensional cell sheet structure, thereby improving cell survival rate and enhancing cell-cell and cell-matrix interactions. Secondly, decellularized dentin matrix particles, which are widely available and easy to obtain, are selected as biomimetic matrix materials, breaking through the bottleneck of the scarcity of traditional bone-derived matrix sources and making large-scale production possible. Furthermore, by introducing GelMA hydrogel as a three-dimensional carrier, stable integration and controllable release of cells and matrix in the bone defect area can be achieved, thereby improving the spatial distribution uniformity and retention rate. In addition, by systematically screening the particle size and concentration of DDMPs, the swelling, degradation and mechanical properties of the complex are optimized to better meet the regeneration needs of load-bearing parts such as the periodontium and skull. Finally, the ability of this copolymer to promote the synergistic effect of angiogenesis and bone regeneration was verified at the molecular and functional levels, and its mechanism in regulating the regenerative microenvironment of "vascular-osteogenic coupling" was clarified, so as to provide a novel biomaterial strategy that is efficient, stable and clinically translatable for the repair of complex oral and craniofacial bone defects.

[0008] (2) Technical solution This invention provides an optimized micron-scale cell-matrix copolymer biomaterial, comprising utilizing micron-sized human exfoliated deciduous teeth spheres (SHED spheres) and combining them with decellularized dentin matrix particles (DDMPs) and GelMA hydrogel to develop an optimized cell-matrix copolymer with microenvironment remodeling function.

[0009] The SHEDs described herein are derived from naturally shed deciduous teeth, a relatively non-invasive and abundant source, and possess low immunogenicity. Therefore, SHEDs show broad application prospects in tissue engineering and regenerative medicine. SHEDs are a type of dental mesenchymal stem cell with high self-renewal capacity and multi-lineage differentiation potential. Particularly in craniofacial tissue regeneration research, SHEDs have been shown to differentiate into various cell types, including osteoblasts, odontoblasts, and neuron-like cells, providing a potential cellular source for the repair and regeneration of bone, teeth, and nerve tissues. Therefore, this study selected SHEDs as the basic cells for constructing CAs (corporeal membrane oxygenation).

[0010] The decellularized dentin matrix microparticles are derived from premolars and third molars that need to be extracted for orthodontic reasons or impacted teeth. Previous studies have shown that dECM, as a natural material that retains tissue-specific bioactive components, has unique advantages in constructing biomimetic microenvironments. We have previously established a technical system for processing dECM into microparticle form and co-constructing "cell-matrix copolymers" with stem cells, aiming to simulate cell-matrix interactions under physiological conditions. However, this system still has the following obvious limitations: 1) Although the two-dimensional polymers derived from early cell-sheet technology can maintain the original cell-ECM contact, their large volume and long induction period can easily lead to internal cell hypoxia and death; secondly, traditional bone-derived dECM (such as human alveolar bone) is scarce and difficult to obtain, which restricts its large-scale application. Therefore, we are turning to dentin matrix (DM), which has a wider range of sources, as an alternative material. Its mineral composition and biological characteristics are similar to alveolar bone, and it can be stably obtained from premolars and third molars that need to be extracted clinically for orthodontic reasons or impacted teeth. Its source is convenient and readily available, and it can be collected and applied on a large scale, with good scalability.

[0011] This invention provides an optimized method for developing micron-scale cell-matrix copolymer biomaterials based on GelMA hydrogel, comprising the following steps: (1) Preparation of SHED sphere The isolated and identified SHEDs were placed in an Ultra-Low Attachment (ULA) culture plate, and polymer induction medium was added for induction and culture. The resulting spherical SHED spheres were characterized by scanning electron microscopy (SEM), H&E staining, Masson staining and immunofluorescence staining. The polymer induction medium is prepared according to the following steps: fetal bovine serum, GlutaMAX™ I, penicillin, streptomycin and vitamin C are added to α-MEM medium, and based on 1L of the polymer induction medium, the concentration of GlutaMAX™-I is 0.5-10mM, the concentration of penicillin is 100U / ml, the concentration of streptomycin is 100μg / ml, the concentration of vitamin C is 0-100μg / ml, and the volume concentration of fetal bovine serum is 5-20%, and α-MEM medium is added to make up to 1L.

[0012] (2) Preparation of decellularized dentin matrix particles (DDMPs) After obtaining informed consent from the patient, this invention harvests dentin matrix from healthy, uninfected third molar or premolar tooth structure and prepares DDMPs. The specific steps are as follows: 1) Sample pretreatment: Mechanically remove the crown, pulp, periodontal ligament and other soft tissues, process the remaining tooth matrix into sheet-like DM, and soak it thoroughly in deionized water.

[0013] 2) Ultrasonic cleaning: Use an ultrasonic cleaner to vibrate and clean at a frequency of 30 minutes / time, repeat twice to achieve thorough cleaning.

[0014] 3) Demineralization treatment: Immerse DM in 17% ethylenediaminetetraacetic acid (EDTA) (Proandy, China) solution for 10 minutes, then rinse repeatedly with sterile deionized water until EDTA residue is completely removed.

[0015] 4) Microparticle preparation: After freeze-drying, the particles are ground for 5 minutes at 1200 rpm using a medium flow tissue grinder (DHS, China). The grinding process is repeated 3-6 times according to the particle size requirements to obtain DDMPs.

[0016] 5) Sterilization and aseptic treatment: After DDMPs were sterilized by high temperature and high pressure for 12 hours, 100 g of the particles were resuspended in 200 ml of sterile PBS containing 100 U / mL penicillin and 100 mg / mL streptomycin, and incubated at 37°C for 3 days to verify the sterility.

[0017] 6) Final processing: The cultured suspension was filtered through a 70 μm cell sieve, aliquoted, and stored in an ultra-low temperature freezer at -80℃ for long-term storage. It was thawed and warmed before use.

[0018] (3) Preparation of micron-sized cell-matrix copolymers (GelMA-SHED sphere-DDMPs) Furthermore, using GelMA as a carrier, a composite copolymer containing the previously separated and identified SHED sphere and DDMPs (GelMA-SHED sphere-DDMPs) was constructed.

[0019] The present invention also provides the application of an optimized micron-scale cell-matrix copolymer biomaterial in rat skull and periodontal bone defects.

[0020] Compared with existing technical solutions, this invention demonstrates significant technological advancements and advantages in multiple aspects through systematic technological innovation. Firstly, the most prominent advantage of this invention is the construction of three-dimensional regenerative microunits with higher cell viability and functional activity. This is thanks to our abandonment of traditional two-dimensional cell sheet technology and the adoption of ultra-low adsorption culture technology to induce the self-assembly of human exfoliated deciduous tooth stem cells into micron-sized three-dimensional spheres. This structure effectively overcomes the problems of internal cell hypoxia and death caused by the excessive size and limited diffusion of nutrients and oxygen in two-dimensional aggregates, resulting in tighter cell connections and a more biomimetic cellular microenvironment. Secondly, this invention successfully solves the core bottleneck restricting clinical translation—the scarcity of regenerative material sources. The key lies in the creative use of clinically available dentin matrix as an alternative source, establishing a standardized preparation process for decellularized dentin matrix particles, thereby eliminating dependence on traditional bone-derived decellularized matrix materials. Third, this invention introduces GelMA hydrogel as a three-dimensional carrier to encapsulate and cross-link stem cell spheres with matrix particles to form an integrated copolymer. This technique, acting like "bio-glue," effectively solves the problems of low retention rate and uneven distribution after direct implantation of micron-sized components, ensuring stable anchoring and uniform spatial distribution of repair components at weight-bearing bone defect sites. Furthermore, this invention systematically screens and optimizes the particle size and concentration of decellularized dentin matrix particles, precisely controlling the degradation kinetics and mechanical properties of the composite hydrogel. This allows it to withstand initial compressive stress and adapt to the tissue regeneration process, exhibiting superior physical property compatibility. Finally, this invention goes beyond simple structural repair by actively regulating and enhancing the "angiogenic-osteogenic coupling" regeneration process through the functional integration of each component. In vitro and in vivo experiments have confirmed that this copolymer can significantly upregulate the expression of osteogenic and angiogenic markers, effectively promoting the spatiotemporal synergy of angiogenesis and bone regeneration, providing a reliable technical path for achieving high-quality, multi-site bone tissue regeneration. Attached Figure Description

[0021] Figure 1 This section describes the construction and characterization of SHED spheres. (A) Construction process of SHED spheres. Scale bar: 100 μm. (BC) SEM images of SHED spheres, showing non-section (B) and section (C) views respectively. Left image scale bar: 100 μm; Right image scale bar: 30 μm. (DE) H&E staining (D) and Masson staining (E) of SHED spheres. Scale bar: 100 μm. (FG) Immunofluorescence staining of SHED spheres: co-staining of F-actin (green) and fibronectin (white) (F); co-staining of α-1 type collagen (red) and cell nucleus (blue) (G). Scale bar: 50 μm; Figure 2 This section describes the preparation and characterization of decellularized dentin matrix microparticles (DDMPs) and GelMA-DDMPs complexes. (A) Gross photograph of a clinical dental sample. Scale bar: 3 mm. (B) SEM images of dentin matrix (DM) and decellularized dentin matrix (DDM). Top image scale bar: 50 μm; Bottom image scale bar: 5 μm. (C) Gross image of DDMPs. Scale bar: 3 mm. (D) SEM image of DDMPs. Top image scale bar: 1 mm; Bottom image scale bar: 2 μm. (E) Particle size analysis of DDMPs. (F) X-ray photoelectron spectroscopy (XPS) analysis of DDMPs. (G) Fourier transform infrared (FT-IR) spectrum. (H) Gross photographs and SEM images of GelMA and its complexes with DDMPs of different particle sizes. Top map scale: 1 mm; Middle map scale: 500 μm; Bottom map scale: 100 μm. (I) Swelling capacity assessment. (J) In vitro degradation curves. (K) Quantitative analysis of elastic modulus. n = 3 per group. ns: no statistical significance (p>0.05); *p<0.05; **p<0.01; ***p<0.001; Figure 3 This study aimed to evaluate the biocompatibility, osteogenic and periodontal induction capabilities of DDMPs. (A) Fluorescent staining images of live / dead cells cultured alone and co-cultured with DDMPs on days 1, 3, and 5. Live and dead cells were stained green and red, respectively. Scale bar: 100 μm. (B) Analysis of the expression of osteogenic and periodontal-related genes in GelMA-SHED sphere and GelMA-SHEDsphere-DDMPs complex by qRT-PCR. n = 3 per group. (C) Representative immunofluorescence images of vascular endothelial growth factor (VEGF) (red) and DAPI (blue) in each group. Scale bar: 100 μm. (D) Quantitative analysis of VEGF expression levels in (C). n = 3 per group. ns: no statistical significance (p>0.05); *p<0.05; **p<0.01; ***p<0.001; Figure 4This study evaluated the tissue regeneration of GelMA, GelMA-SHED sphere, and GelMA-SHED sphere-DDMPs in rat models of skull and periodontal bone defects. (A) Schematic diagram of GelMA-SHED sphere-DDMPs implanted in the skull and periodontal bone defect areas. (B) Representative micro-CT images of skull regeneration in each group 10 weeks post-surgery. Scale bar: 2 mm. (C) Representative micro-CT images of periodontal bone regeneration in each group 4 weeks post-surgery. The three-dimensional reconstructed buccal cross-sectional image (top) and the three-dimensional reconstructed images of the defect area (middle and bottom) show the overall healing results of the defect area. Scale bar: 2 mm. (D) H&E stained images at 4 weeks post-surgery showing newly formed bone and periodontal tissue in the periodontal bone defect area. Scale bar: 250 μm. (EF) Quantitative analysis of bone volume fraction (bone volume / tissue volume) in the skull (E) and periodontal bone (F) defect areas. n = 6 per group. (G) Quantitative analysis of alveolar bone area in periodontal bone. n = 3 per group. ns: no statistical significance (p>0.05); *p<0.05; **p<0.01; ***p<0.001; ****p<0.0001; Figure 5 GelMA-SHED sphere-DDMPs promote angiogenesis. (A) Immunofluorescence staining images of CD31 (green) and endothelial mucin (EMCN) (red) in the bone regeneration area, with cell nuclei counterstained by DAPI (blue). (B) Immunofluorescence staining images of CD31 (green) and α-smooth muscle actin (αSMA) (red) in the bone regeneration area, with cell nuclei counterstained by DAPI (blue). Scale bar = 100 μm. (C) Quantitative analysis of CD31⁺EMCN⁺ vessels (%). (D) Quantitative analysis of CD31⁺αSMA⁺ vessels (%). n = 3 per group. Data are expressed as mean ± standard deviation. ns: no statistical significance (p>0.05); **p<0.01; ***p<0.001; ****p<0.0001; Figure 6 GelMA-SHED sphere-DDMPs promote bone regeneration. (A) Immunofluorescence staining images of RUNX2 (green) and DAPI-stained nuclei (blue) in the bone regeneration region. (B) Immunofluorescence staining images of osteocalcin (OCN) (red) and DAPI-stained nuclei (blue) in the bone regeneration region. Scale bar = 100 μm. (C) Quantitative analysis of RUNX2 expression levels. (D) Quantitative analysis of OCN expression levels. n = 3 per group. ns: no statistical significance (p>0.05); *p<0.05; ***p<0.001; ****p<0.0001. Detailed Implementation

[0022] The reagents and consumables used in this invention are all commercially available products that can be purchased on the market. The main reagents and materials were purchased through the following channels: 17% ethylenediaminetetraacetic acid (EDTA) (Maclean; E809070-500g); phosphate-buffered saline (PBS) (ZHHC; PW013); penicillin-streptomycin (Gibco; 15140122); α-MEM medium (Gibco; C12571500bt); fetal bovine serum (Sijiqing; 11011 8611); GlutaMAX™ I (Gibco; 35050079); vitamin C (MP; 100769); GelMA (Engineering For Life, EFL-GM-60); calcein / PI cell viability and cytotoxicity assay kit (Beyotime, China); 30% sucrose solution (Sigma-Aldrich, USA); OCT compounds (Sakura Finetek, USA).

[0023] Example 1: Construction and Characterization of the SHED sphere (1) Construction of SHED sphere With the approval of the hospital's ethics committee, informed consent was obtained from patients and their families before this research project was carried out. Healthy, uninfected deciduous teeth were obtained from the clinic and transferred to the clean bench of the Tissue Engineering Center of the Stomatological Hospital of Air Force Medical University. Under sterile conditions, dental pulp tissue was isolated and primary cell culture was performed. Subsequently, the stem cell characteristics, proliferation capacity, and multiple differentiation potentials were routinely evaluated.

[0024] After the isolated and identified SHED cells were digested with trypsin, the cell suspension was placed in an Ultra-Low Attachment (ULA) culture plate and polymer induction medium was added for induction and culture.

[0025] Preparation of polymer induction medium: Add 25 ml fetal bovine serum, 2.5 ml 100×GlutaMAX™ I, 2.5 ml 100× penicillin-streptomycin and 500 μl 3 mg / ml VC solution to 220 ml α-MEM medium.

[0026] The results are as follows Figure 1 As shown in Figure A, structurally stable SHED spheres can be formed within 24 hours after inoculation with single-cell suspension, and their formation efficiency and morphological characteristics are consistent with the previous experimental results.

[0027] (2) Characterization of the SHED sphere The SHED spheres obtained above were characterized by scanning electron microscopy (SEM), H&E staining, Masson staining, and immunofluorescence staining. The specific steps are as follows: 1) SEM observation of the SHED sphere: The prepared polymer was washed three times with PBS and then fixed in 2.5% glutaraldehyde at room temperature for 5 min. Then, the polymer was dehydrated with different gradients of ethanol solutions (60%, 75%, 85%, 95%, and 100%, 5 min each), and then dried with hexamethyldisilane. Gold was then deposited on the sample using an ion sputtering device, and the morphology of the polymer (both intact and cross-sectional) was observed and photographed under SEM.

[0028] The results are as follows Figure 1 As shown in BC, single cells and collagen fibers intertwine to form a stable and dense three-dimensional spherical structure. Figure 1 B); To further understand the internal structure of the SHED sphere, SEM was used to observe the cross-sectional structure of the SHED sphere. The results show ( Figure 1 C), the stem cells inside the sphere are round or oval, and the cells are physically connected by abundant extracellular matrix (ECM) and tightly connected pseudopodia.

[0029] 2) Histological staining of SHED spheres: The prepared polymers were fixed with 4% PFA at room temperature for 24 h. Subsequently, the polymers were dehydrated in a gradient ethanol solution and then embedded in paraffin. 6 μm thick tissue sections of the polymers were prepared for H&E staining, Masson's staining, and immunofluorescence staining. Images were acquired using a histological analysis workstation.

[0030] The results are as follows Figure 1 As shown in DG, histological staining confirmed that hematoxylin-eosin (H&E) staining showed uniform cell distribution and a moderate nucleoplasmic ratio within the spheroids. Figure 1 D); Masson's staining clearly shows that collagen fibers are densely deposited in a network-like pattern in the ECM (D). Figure 1 E). Immunofluorescence staining results showed that F-actin in SHED spheroids was distributed in a ring around the nucleus, while core ECM components such as fibronectin and α-1 type I collagen (Col1a1) were highly expressed in a network pattern in the matrix. Figure 1 The presence of FG indicates that the three-dimensional polymer possesses a mature cytoskeleton structure and matrix synthesis capabilities.

[0031] Example 2: Preparation and Characterization of DDMPs (1) Preparation method of DDMPs The research protocol was approved by the Medical Ethics Committee of the Stomatological Hospital of the Air Force Medical University, and informed consent was obtained from patients and their families before the research project began. The research involved healthy, uninfected third molars or premolars. Figure 2 A) Dentin matrix (DM) was collected from tooth tissue and transferred to the Tissue Engineering Center of the Stomatological Hospital of Air Force Medical University using culture medium. The specific steps are as follows: 1) Sample pretreatment: Mechanically remove the crown, pulp, periodontal ligament and other soft tissues, process the remaining tooth matrix into sheet-like DM, and soak it thoroughly in deionized water.

[0032] 2) Ultrasonic cleaning: Use an ultrasonic cleaner to vibrate and clean at a frequency of 30 minutes / time, repeat twice to achieve thorough cleaning.

[0033] 3) Demineralization treatment: Immerse DM in 17% EDTA solution for 10 minutes, then rinse repeatedly with sterile deionized water until EDTA residue is completely removed.

[0034] 4) Microparticle preparation: After freeze-drying, the particles are ground at 1200 rpm for 5 minutes using a medium-flow tissue grinder. The grinding process is repeated 3-6 times according to the particle size requirements to obtain DDMPs.

[0035] 5) Sterilization and aseptic treatment: After DDMPs were sterilized by high temperature and high pressure for 12 hours, 100 g of the particles were resuspended in 200 ml of sterile PBS containing 100 U / mL penicillin and 100 mg / mL streptomycin, and incubated at 37°C for 3 days to verify the sterility.

[0036] 6) Final processing: The cultured suspension was filtered through a 70 μm cell sieve, aliquoted, and stored in an ultra-low temperature freezer at -80℃ for long-term storage. It was thawed and warmed before use.

[0037] The results are as follows Figure 2 As shown in C, DM was obtained from healthy third molars or premolars. After decellularization, DM was obtained as DDM. After grinding, DDM was processed into DDMPs, which were pale yellow powdery microparticles.

[0038] (2) Characterization of DDMPs 1) SEM observation of DDMPs The obtained DM and DDM samples were dehydrated in a gradient of ethanol solutions (60%, 75%, 85%, 95%, and 100%, 10 minutes each). Hexamethyldisilane was then added dropwise to the dehydrated samples, which were then dried at room temperature in a fume hood for 8–10 hours. Subsequently, gold was deposited on the dried sample surfaces using an ion sputtering device, and the morphology of DM and DDM was observed and photographed under SEM.

[0039] 2) Particle size analysis of DDMPs The particle size of DDMPs was analyzed using a laser particle size analyzer (Malvern Mastersizer 2000, UK).

[0040] 3) X-ray photoelectron spectroscopy (XPS) analysis The chemical composition of DDMPs was analyzed using an X-ray scanning electron microscopy (EDS) instrument (Octane SDD, USA), according to the manufacturer's instructions.

[0041] 4) Fourier transform-infrared spectroscopy (FTIR) According to the manufacturer's instructions, the chemical properties of DDMPs were analyzed using a Fourier transform infrared spectrometer (Shimadzu, Japan).

[0042] The results are as follows Figure 2 As shown in BG, the SEM results ( Figure 2 (B) shows that, compared to DM, the collagen fibers and dentinal tubules of DDM were fully exposed after decellularization, demonstrating the effectiveness of the decellularization treatment. Subsequently, DDM was ground into pale yellow granules of DDMPs (… Figure 2 C). SEM results ( Figure 2 (D) shows that DDMPs exhibit a non-uniform particle size distribution. The particle sizes of DDMPs after six cycles of milling are: D50: 30.5 μm, D90: 95.484 μm, and the average particle size is 42.999 μm. Figure 2 E). XPS analysis shows ( Figure 2 In addition to essential chemical elements such as calcium and phosphorus, DDMPs also contain elements such as oxygen, carbon, sodium, and magnesium. FT-IR results ( Figure 2 G) indicates that DDMPs contain organic groups such as carbonates and phosphates. These results demonstrate that DDMPs were prepared efficiently.

[0043] (3) Construction and characterization of GelMA-DDMPs complex To systematically investigate the influence of DDMP particle size and concentration on the mechanical properties of GelMA hydrogels, this study first prepared DDMP-GelMA composite hydrogel systems with different particle size gradients (1-100 μm, 100-200 μm, 200-300 μm). The GelMA-DDMP composites were characterized by SEM, swelling experiments, in vitro degradation performance tests, and compression modulus experiments. The specific steps are as follows: 1) Evaluation of swelling performance The swelling properties of GelMA hydrogels with and without DDMPs were determined. Hydrogel samples were prepared into cylinders of equal volume (200 μL, n=3) and exposed to 405 nm UV light for 30 s. The samples were then incubated in PBS at 37 °C for 48 h. After removal from PBS, the wet weight (Ws) of the samples was measured. The samples were then lyophilized for 8 h, and the dry weight (Wd) was measured. The swelling ratio of the hydrogel was calculated using the following formula: Swelling ratio Qs = (Ws−Wd) / Wd × 100%.

[0044] 2) Evaluation of in vitro degradation performance In the in vitro degradation assay, samples were placed in 24-well plates containing 0.5 mg / ml type I collagenase at 37°C, and samples were collected at each predetermined time point (0, 12, 24, 36, and 48 h, n=3). Each sample was then lyophilized for 8 h to obtain the remaining weight (Wr). Using the weight at 0 h as W0, the biodegradation ratio of the hydrogel was calculated using the following formula: Biodegradation ratio Qd = (Wr / W0) × 100%.

[0045] 3) Compression modulus test According to the manufacturer's instructions, the compressive modulus of GelMA hydrogels containing and without DDMPs was tested using a biomechanical tester (EngineeringForLife, China) (n=3).

[0046] The results are as follows Figure 2 As shown in HK, after photocrosslinking to obtain a series of composite hydrogels with varying concentration gradients, macroscopic observation shows that ( Figure 2 H (Top): The composites containing 1-100 μm DDMPs exhibit a uniform milky white color, while the 100-200 μm and 200-300 μm groups show a heterogeneous state with obvious particle sedimentation and hydrogel stratification. We used SEM to further observe the cross-sectional structure of different composite groups ( Figure 2H (Bottom): Compared to the pure GelMA control group, the DDMP-containing composites all exhibited a porous network structure in which DDMPs and GelMA were interwoven and wrapped, with DDMPs embedded in pheromone and interwoven within the GelMA matrix. Among them, the 1-100 μm group showed the best homogeneity—GelMA was uniformly coated on the DDMPs surface, and the pore distribution uniformity was significantly better than other particle size groups. This structural characteristic is consistent with the particle size analysis results (…). Figure 2 E) The particle size distribution was highly consistent, therefore 1-100 μm was selected as the standard particle size for subsequent studies. Based on the particle size screening results, this invention further prepared a series of composite hydrogels with different concentrations of DDMPs and systematically evaluated their swelling properties, degradation kinetics, and compressive mechanical properties. Swelling experiments showed that ( Figure 2 I) The addition of DDMPs did not significantly change the swelling ratio of GelMA (p>0.05). Degradation performance tests showed ( Figure 2 J), the degradation rate of the DDMP-containing complex showed a concentration-dependent increasing trend, with the 0.5 mg / mL group exhibiting a moderate degradation rate, ensuring both initial scaffold stability and meeting the needs of subsequent tissue regeneration. Compression modulus test results ( Figure 2 K) showed that although the addition of DDMPs slightly decreased the compressive modulus of GelMA, the difference between groups was not statistically significant (p>0.05). Notably, the 0.5 mg / mL DDMPs group, while maintaining good compressibility, exhibited excellent swelling adaptability and a moderate degradation rate, making it particularly suitable for tissue engineering applications in load-bearing sites such as periodontal / skull defects—it can both resist compressive stress in the defect area and achieve effective retention and sustained release of the GelMA-SHED sphere-DDMPs copolymer, thereby better exerting its therapeutic effect.

[0047] Example 3: Evaluation of the biocompatibility and osteogenic induction capacity of DDMPs (1) Live and dead staining to assess the biocompatibility of DDMPs After pretreatment with or without DDMPs, GelMA-SHED cell spheres were washed three times with PBS. Subsequently, cell viability in the SHED spheres was assessed using the Calcein / PI Cell Viability and Cytotoxicity Assay Kit, as per the manufacturer's instructions, and observed using a fluorescence microscope.

[0048] (2) In vitro osteogenic and periodontal capacity assessment of GelMA-SHED sphere-DDMPs Total RNA was extracted using the MIsZOL kit. Reverse transcription and qRT-PCR were performed according to the manufacturer's instructions. Reverse transcription and qRT-PCR amplification were performed using the Takara kit. Relative gene expression was quantified using the 2−ΔΔCt method, and mRNA levels were normalized to normal GAPDH levels.

[0049] (3) VEGF immunofluorescence staining After pretreatment with or without DDMPs, GelMA-SHED cell spheres were washed three times with PBS. The resulting SHED spheres were fixed with 4% PFA, then dehydrated in 30% sucrose solution and embedded in OCT. 10 μm frozen sections were prepared using a cryostat. For immunofluorescence staining of the frozen sections, the sections were first thawed at room temperature for 2 hours, then washed three times with PBS for 5 minutes each time. After infiltration with 0.3% Triton X-100, the samples were blocked with goat serum at room temperature for 30 minutes. Diluted primary antibody (VEGF) was then added to the samples, and the samples were incubated overnight at 4°C in a humidified chamber. Excess primary antibody was washed away with PBS, followed by staining with secondary antibody at room temperature for 1 hour and DAPI staining for 15 minutes. Images were acquired and analyzed using confocal microscopy and ImageJ software.

[0050] The results are as follows Figure 3 As shown in AD, SHED spheres induced by ULA culture plates were transferred to GelMA hydrogels and co-cultured with culture systems containing / without DDMPs for 1, 3, and 5 days, respectively. Cell viability was assessed using a live / dead cell staining method. IF staining results showed ( Figure 3 (A) Both groups showed predominantly viable cells labeled with green fluorescent markers at 1, 3, and 5 days of culture, with only a small number of dead cells labeled with red fluorescent markers. Although slight cell death occurred within the SHED sphere in both groups with prolonged culture time, the difference between groups was not statistically significant, confirming that the introduction of DDMPs did not significantly affect the cell viability of SHEDs, indicating that it had no obvious cytological toxicity. To investigate the regulatory role of DDMPs on the osteogenic and periodontal differentiation potential of SHED spheres, this invention further conducted qRT-PCR analysis. The results showed ( Figure 3B), compared with the SHED sphere group alone, the mRNA expression levels of osteogenic markers—alkaline phosphatase (ALP), Runt-related transcription factor 2 (RUNX2), osteopontin (OPN), dentinialophosphoprotein (DSPP), cementum attachment protein (CAP), dentin matrix acidic phosphoprotein 1 (DMP1)—and periodontal markers—periostin and laminin-β1—were significantly increased in the GelMA-SHEDsphere-DDMPs co-aggregates (p<0.05), suggesting that the co-aggregates have stronger osteogenic and periodontal tissue regeneration capabilities. Given that vascularization is a key factor for successful tissue regeneration, this invention examined the secretion dynamics of VEGF in both SHED sphere groups. The results showed ( Figure 3 (C and D) There was no significant difference in VEGF secretion between the two groups on day 1 of culture (p>0.05), but on days 3 and 5, the VEGF secretion in the co-aggregate group increased significantly in a time-dependent manner (p<0.05), indicating that DDMP co-culture can promote VEGF secretion in the SHED sphere. Combined with the expression of osteogenic / periodontal markers and VEGF secretion, this suggests that DDMP co-culture can synergistically enhance the osteogenic, periodontal, and angiogenic differentiation potential of SHED.

[0051] Example 4: Optimized micron-sized cell-matrix copolymer (GelMA-SHED sphere-DDMPs) biomaterial was implanted into the skull and periodontal bone defects of rats, and good tissue regeneration effects were observed. (1) Micro CT detection The aforementioned copolymer was implanted into the skull and periodontal bone defects of rats. Skull and mandibular bone samples were extracted from rats at 10 and 4 weeks post-surgery and fixed in 4% PFA for 24 hours. The mandibular bone samples were scanned using a Quantum GX2 micro-CT imaging system at a resolution of 18 μm, voltage of 50 kV, and current of 80 μA. After scanning and 3D image reconstruction, the original alveolar bone defect area was defined as the region of interest to acquire periodontal bone data, and parameters such as bone volume / total volume (BV / TV) were calculated.

[0052] (2) Histological examination and immunofluorescence staining of the mandible Rat mandibular bone samples were collected after scanning and decalcified with 17% EDTA at room temperature for one month. The tissues were dehydrated in 30% sucrose solution (Sigma-Aldrich, USA) and embedded in an OCT compound (Sakura Finetek, USA). 10 μm frozen sections were obtained using a cryostat (Leica, Germany). H&E staining was performed according to the manufacturer's instructions. Images were obtained under a microscope, and three images were randomly selected for quantification using ImageJ.

[0053] For immunofluorescence staining on frozen sections, the sections were first thawed at room temperature for 2 hours, then washed three times with PBS for 5 minutes each time. After infiltration with 0.3% Triton X-100, the samples were blocked with goat serum at room temperature for 30 minutes. Diluted primary antibodies (CD31, EMCN, αSMA, RUNX2, and OCN) were then added to the samples, and the samples were incubated overnight at 4°C in a humidified chamber. Excess primary antibody was washed away with PBS, followed by staining with secondary antibody at room temperature for 1 hour and DAPI staining for 15 minutes. Images were acquired and analyzed using confocal microscopy and ImageJ software.

[0054] The results are as follows Figure 4-6 As shown, the results of Micro-CT three-dimensional reconstruction and quantitative analysis indicate that in the skull ( Figure 4 B and E) and periodontal bone defect area ( Figure 4 (C and F) All implantation groups showed varying degrees of bone repair. The GelMA-SHEDsphere-DDMPs group exhibited the most significant repair effect, with the defect area almost completely recovering its original bone volume. H&E staining further confirmed that the GelMA-SHEDsphere-DDMPs group had the best bone healing quality, with a large amount of newly formed trabecular bone replacing the defect area ( Figure 4 D and G). To investigate vascular behavior and osteogenic-angiogenic coupling during regeneration, we performed immunofluorescence staining. Results showed that CD31+EMCN+ (neovascularization) was observed in both the early and late stages of implantation in each group. Figure 5 A) and CD31+αSMA+ (mature blood vessels) Figure 5 B) Structure, and in the bone canaliculi and lacunae regions of regenerated bone, RUNX2 (an early osteogenic marker) Figure 6 A) with OCN (late-stage osteogenic marker) Figure 6B) Significant expression around vascular structures suggests a close spatial and functional association between angiogenesis and bone regeneration. Notably, the proportions of CD31+EMCN+ and CD31+αSMA+ vessels were highest in the GelMA-SHED sphere-DDMPs group. Figure 5 C and D), RUNX2 and OCN expression were also significantly upregulated ( Figure 6 (C and D) indicates that this group was most effective in restoring the angiogenic-osteogenic coupling microenvironment. In addition, this group showed high levels of "pro-regenerative" CD31+EMCN+ blood vessels and RUNX2 expression in the early stage of implantation, indicating that DDMPs provided favorable microenvironmental support for stem cell survival, differentiation and mineralization.

[0055] Although embodiments of the present invention have been shown and described above, it is understood that the above embodiments are exemplary and should not be construed as limiting the present invention. Those skilled in the art can make changes, modifications, substitutions and variations to the above embodiments within the scope of the present invention.

Claims

1. An optimized method for preparing micron-sized cell-matrix copolymer biomaterials, characterized in that, Includes the following steps: (1) Preparation of human exfoliated deciduous teeth pulp stem cell sphere (SHED sphere): The isolated and identified SHEDs were placed in a low adhesion (ULA) culture plate, and polymer induction medium was added for induction and culture. The resulting SHED sphere polymers were characterized by SEM, H&E staining, Masson staining and immunofluorescence staining. The polymer induction medium is prepared by adding fetal bovine serum, GlutaMAX™ I, penicillin, streptomycin, and vitamin C to α-MEM medium. The concentration of GlutaMAX™ I is 0.5–10 mM, the concentration of penicillin is 100 U / ml, the concentration of streptomycin is 100 μg / ml, the concentration of vitamin C is 0–100 μg / ml, and the volume concentration of fetal bovine serum is 5–20%, which is then supplemented to 1 L with α-MEM medium. (2) Preparation of decellularized dentin matrix microparticles (DDMPs): Dentin matrix was collected from healthy, uninfected third molar or premolar tooth tissue and DDMPs were prepared, including sample pretreatment, ultrasonic cleaning, demineralization with 17% EDTA for 10 minutes, freeze drying, grinding at 1200 rpm for 3-6 times with a medium flow tissue grinder, high temperature and high pressure sterilization, 100 g particles were resuspended in 200 ml sterile PBS containing 100 U / mL penicillin and 100 mg / mL streptomycin, cultured at 37℃ for 3 days to verify sterility, filtered through a 70 μm cell sieve, and stored at -80℃; (3) Preparation of micron-scale cell-matrix copolymer (GelMA-SHED sphere-DDMPs): Using GelMA hydrogel as a carrier, the SHED sphere obtained in step (1) and the DDMPs obtained in step (2) are encapsulated and crosslinked by 405 nm light for 30 s to form the composite.

2. The preparation method according to claim 1, characterized in that, In step (2), the particle size of DDMPs is 1-100 μm.

3. The preparation method according to claim 1 or 2, characterized in that, In step (3), the concentration of DDMPs in GelMA is 0.5 mg / mL.

4. The preparation method according to any one of claims 1-3, characterized in that, The SHEDs are derived from naturally shed baby teeth.

5. Optimized micron-sized cell-matrix copolymer biomaterials prepared by the preparation method according to any one of claims 1-4.

6. The optimized micron-scale cell-matrix copolymer biomaterial according to claim 5, characterized in that, The biomaterial can promote the synergistic effect of angiogenesis and bone regeneration.

7. The use of the optimized micron-scale cell-matrix copolymer biomaterial according to claim 5 or 6 in the preparation of biomaterials for repairing craniofacial bone defects.

8. The application according to claim 7, characterized in that, The repair is achieved through synergistic regulation of the angiogenesis-osteogenic coupling mechanism.

9. The application according to claim 7 or 8, characterized in that, The craniofacial bone defects include skull defects and periodontal bone defects.

10. A method for promoting bone tissue regeneration at multiple sites, characterized in that, The optimized micron-scale cell-matrix copolymer biomaterial of claim 5 or 6 is implanted into the bone defect site.

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