Method Article

Half-segmental Diaphyseal Bone Defect Model in Rats for Evaluating Bone Substitute Performance in Load-bearing Regions

DOI:

10.3791/69211

⸱

December 30th, 2025

 ,  ,  ,  ,  ,  ,  ,  , 
1State Key Laboratory of Oral Diseases & National Center for Stomatology & National Clinical Research Center for Oral Diseases, West China Hospital of Stomatology, Sichuan University, 2Department of Cariology and Endodontics, West China Hospital of Stomatology, Sichuan University, 3Department of Jinjiang Outpatient, West China Hospital of Stomatology, Sichuan University

In This Article

Summary

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

This study established a rat femoral half-segmental defect model to evaluate the mechanical and osteogenic performance of bone substitute materials under load-bearing conditions without the use of internal or external fixation.

Abstract

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

Accurate preclinical evaluation of bone substitute materials demands models that not only support tissue regeneration but also reflect the mechanical challenges of physiological load-bearing conditions. However, most conventional animal bone defect models fall short in this regard. For example, calvarial defect models, though widely used, are created in non-load-bearing regions. Similarly, drill-hole or partial defects in long bones primarily focus on localized healing, lacking sufficient load transfer to assess mechanical support. This study established a standardized half-segmental diaphyseal bone defect model in the rat femur to assess both the mechanical and biological performance of bone substitute materials in a load-bearing context without the use of fixation devices. Using a high-speed handpiece and a specialized cylindrical dental bur, a semi-cylindrical defect measuring 4 mm in length and 1.5 mm in radius was created in the lateral midshaft of the femur. Two representative materials with distinct properties -- Gelatin methacryloyl (GelMA) hydrogel and 3D-printed polymethyl methacrylate (3DP-PMMA) -- were implanted to validate the model's capacity to differentiate the mechanical properties of materials. Radiographic, histological, and immunofluorescent analyses performed at 4 weeks post-implantation revealed that, due to inadequate mechanical support, GelMA implantation led to malunion with fibrous tissue predominance and limited bone integration. In contrast, 3DP-PMMA facilitated the organization of new bone formation, periosteal continuity, and the recruitment of Piezo type mechanosensitive ion channel component 1-expressing (Piezo1+) and leptin receptor-expressing (LepR+) cells. This model provides a practical and reproducible platform for assessing the load-bearing capacity, osteoinductivity, and osseointegration potential of biomaterials. It can serve as a valuable tool for screening candidate materials for orthopedic applications, such as long bone reconstruction or fracture repair.

Introduction

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

Under physiological conditions, bone plays a central role in maintaining the structural integrity and mechanical function of the body. It supports body weight, enables movement, and protects internal organs. When bone is damaged due to trauma, tumor resection, or infection, especially in load-bearing regions, the mechanical continuity is interrupted. This disruption often leads to complications such as bone deformity, reduced mobility, limb instability, chronic pain, and even life-threatening issues from severe fractures and hemorrhage1,2. Nowadays, bone substitutes such as bone grafting and biomaterials have been clinically applied to replace severely damaged bone tissue or restore large-scale bone defects. Similarly, for the implanted bone substitutes, their compromised mechanical performance can lead to implant failure, resulting in non-union or even catastrophic fractures at the defect site3,4. To restore both structure and function, bone substitute materials must not only promote bone regeneration but also possess sufficient mechanical strength to withstand physiological loading5.

A critical step prior to clinical translation is the realistic assessment of whether a biomaterial, once implanted in vivo, can meet the mechanical demands required for structural support in load-bearing bone regions. Although many biomaterials have shown promise in vitro or in small, non-load-bearing models, their translation to clinical practice is often hindered by the lack of reliable preclinical models that simulate in vivo mechanical conditions. Currently used animal models for bone defect studies include mandibular defects6, calvarial defects7, drill-hole defects in long bones8, post-extraction sockets9, and so on. These models primarily evaluate a material's osteogenic capacity and focus on histological or radiographic regeneration endpoints10,11. However, considering that these sites are either non-load-bearing or mechanically protected, they fail to replicate the dynamic stress conditions encountered in long bones12. Even when mechanical properties are tested by in vitro techniques such as compressive or tensile strength assays, these fail to replicate the dynamic loading and host tissue interactions inherent to in vivo environments13.

A major gap in preclinical research remains the lack of a standardized, reproducible animal model that allows in vivo mechanical assessment of bone substitutes under load-bearing conditions. To address this, the current study established a half-segmental diaphyseal bone defect model in rats. This model provides a stable and reproducible defect site in a load-bearing region, enabling direct evaluation of the mechanical support capacity of bone substitute materials under physiological loading, while concurrently allowing assessment of osteogenic outcomes. Moreover, the model can be applied across various animal species, with defect size and location adjusted according to the specific animal used. Nevertheless, it does not fully replicate the complex biomechanical environment of human bones, and for studies focused solely on non-load-bearing defects, cranial bone healing, or very large structural implants, alternative models may be more appropriate.

Access restricted. Please log in or start a trial to view this content.

Protocol

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

All animal procedures conducted in this study received approval from the Ethical Committee of the West China School of Stomatology, Sichuan University (WCHSIRB-AT-2025-342). Male Sprague-Dawley rats (180 g each) were obtained from a commercial source and randomly assigned to two groups (n = 6 per group) according to the implanted material. The reagents and the equipment used are listed in the Table of Materials.

1. Pre-operative preparation

  1. Surgical items
    1. Prepare surgical instruments referring to Figure 1A, corresponding to the numbered labels in the figure, in the following order: curved forceps, disposable sterile scalpel, straight scissors, periosteal elevator, irrigation needle, a specific dental diamond bur (see the enlarged view in Figure 1B), high-speed handpiece, vernier caliper, surgical motor, monofilament suture (4-0), and needle holder.
    2. Prepare 0.9% sterile saline solution for irrigation.
  2. Sterilization and disinfection
    1. Sterilize surgical instruments that can withstand autoclaving, including curved forceps, scissors, periosteal elevator, dental diamond bur, high-speed handpiece, and needle holder.
      NOTE: Instruments should be wrapped in medical non-woven fabrics and sealed with autoclave tape for steam sterilization at 121-134 °C for 20-25 min.
    2. For heat-sensitive instruments, such as vernier calipers and surgical motor, expose them to ultraviolet light for at least 30 min for disinfection. Disposable items, such as the disposable sterile scalpel, irrigation needle, and sutures, should remain sealed in their original sterile packaging until use.
    3. Ensure the surgical environment is sterile and temperature-controlled. Wipe down the surgical bench and surrounding surfaces with 75% ethanol. Use sterile, medical-grade non-woven fabrics to cover the operating table.
      CAUTION: Ethanol is flammable; handle away from ignition sources and store in closed containers. Collect ethanol waste in designated chemical waste containers for proper disposal. Wear appropriate personal protective equipment, including gloves, a lab coat, and safety goggles.
  3. Anesthesia
    1. Weigh the rat and calculate the anesthetic dosage (100 mg/kg of ketamine and 10 mg/kg of xylazine, intraperitoneally).
    2. Induce anesthesia 20 min before surgery (following institutionally approved protocols) and confirm its depth by checking for the absence of response to toe pinch and the loss of the righting reflex.
    3. Apply ophthalmic ointment to both eyes to prevent corneal drying during the procedure.

2. Surgical procedure

  1. Site preparation
    1. Place the rat in a lateral recumbent position on the sterile surgical bench. Shave the lateral thigh area corresponding to the femur projection and disinfect the skin using 2% iodine tincture, followed by 75% ethanol (Figure 2Ba).
      CAUTION: Iodine tincture is flammable and irritant. Handle in a well-ventilated area while wearing gloves, a lab coat, and safety goggles. Dispose of used wipes and solutions in designated chemical waste containers according to institutional safety regulations.
    2. Change into fresh sterile gloves. Drape the surgical site with the sterile fenestrated sheet to maintain asepsis.
  2. Surgical site opening
    1. Use curved forceps to assist in stabilizing the tissues, holding the muscles, and gently securing the femur throughout the surgical procedure (Figure 2Bb-h).
    2. Make a 2-3 cm longitudinal skin incision over the lateral aspect of the thigh with a sterile scalpel blade to expose the underlying muscles (Figure 2Bb).
    3. Identify the rectus femoris and vastus lateralis muscles, and carefully separate them along the visible white fascia (indicated by the yellow arrow in Figure 2Bb) using a straight scissor (Figure 2Bc).
    4. Locate and expose the muscle attachments on the lateral surface of the femur (Figure 2Bd).
    5. Make a longitudinal incision along the muscle attachments with a disposable sterile scalpel to gain access to the femoral surface (Figure 2Be).
      NOTE: When performing sharp dissection (using a scissor or scalpel) for soft tissue separation, take care not to damage the underlying blood vessels and nerves. If unfamiliar with the anatomical structures, use fine-tipped forceps and a periosteal elevator to perform blunt dissection.
    6. Perform blunt dissection using a periosteal elevator to detach muscle attachments and fully expose the mid-diaphysis of the femur (Figure 2Bf).
    7. Further separate the femur from the surrounding muscles and hold the midshaft of the femur using curved forceps (Figure 2Bg).
  3. Half-segmental diaphyseal bone defect model establishment
    1. Attach the flat-ended cylindrical specific dental diamond (Figure 1B) bur to the high-speed handpiece, powered by the surgical motor. Set the surgical motor to 35,000 rpm.
    2. Under direct visualization, identify the highest point of the femoral midshaft as the anatomical center for defect creation. Drill vertically downward at this point using the bur until a sudden loss of resistance is observed, accompanied by the appearance of blood effusion within the defect, which indicates full cortical penetration and access to the medullary cavity (Figure 2Bh).
      NOTE: Given inter-animal variability in cortical thickness, operators are advised to familiarize themselves with femoral anatomy and to perform pre-experimental training to standardize this step.
    3. Enlarge the defect by applying controlled horizontal push-pull motions centered around the initial penetration point. Extend the defect longitudinally (along the femoral axis) and laterally (along the femoral diameter) to gradually form a standardized semi-cylindrical defect measuring 4 mm in length and 1.5 mm in radius (Figure 2Bi, see Figure 2A for the morphology and position of the defect).
      NOTE: Once the center point is established, use a vernier caliper to assist in stepwise defect enlargement, ensuring uniform dimensions across all specimens. The flat-end cylindrical design of the specialized dental bur matches the semi-cylindrical defect, allowing efficient and reproducible preparation.
    4. Continuously irrigate the surgical site with sterile saline using the irrigation needle to prevent thermal necrosis during bone cutting.
  4. Bone substitute implantation
    1. Gently remove bone debris and irrigate the site to clear residual particles.
    2. Implant non-load-bearing Gelatin methacryloyl (GelMA) hydrogels (see Table of Materials) or load-bearing 3D-printed polymethyl methacrylate (3DP-PMMA) intended for mechanical performance testing into the defect site, ensuring a snug fit (Figure 2Bj).
      NOTE: The two biomaterials used here were deliberately selected to represent contrasting mechanical and biological properties: GelMA hydrogel serves as a soft, non-load-bearing material with limited osteogenic potential, whereas 3DP-PMMA acts as a rigid, load-bearing scaffold capable of effective stress transmission and supporting bone formation14 (Figure 2A). This contrast highlights the model's ability to evaluate implants with a wide range of mechanical strength and biological performance, and in practice, the defect model can be applied to test other candidate materials with mechanical properties.
  5. Surgical site closure
    1. Suture the muscle layers using 4-0 monofilament sutures and a needle holder, taking care to avoid excessive tension (Figure 2Bk).
    2. Perform skin closure with interrupted sutures and disinfect the surgical site using 2% iodophor solution (Figure 2Bl).

3. Post-operative care

  1. Transfer the rats onto a disinfected, constant heating pad (37 °C) to allow post-operative recovery.
  2. Administer appropriate post-operative analgesia (e.g., buprenorphine at 0.03 mg/kg subcutaneously every 12 h for 48-72 h).
  3. Once the rats regain consciousness, place them in clean cages with soft bedding and easy access to food and water.
  4. Monitor the surgical site daily for signs of infection, dehiscence, or abnormal swelling. Observe the rats' behavior, weight, and mobility for at least 7 days post-surgery to ensure recovery and assess welfare.

4. Sample harvesting and data collection

  1. Sample collection
    1. Euthanize the rats with carbon dioxide inhalation followed by secondary confirmation (following institutionally approved protocols).
    2. Harvest the femur specimens at 4 weeks post-implantation with animals initially randomized into two material groups (n = 6 per group) to assess the materials' mechanical performance.
      NOTE: In the GelMA group, three animals developed complete fractures shortly after surgery and were excluded, leaving three specimens available for subsequent analyses.
    3. Fix the femoral specimens in 4% paraformaldehyde for 48 h at 4 °C.
      CAUTION: Paraformaldehyde is toxic and should be handled in a certified fume hood. Wear gloves, a lab coat, and safety goggles or a face shield. Collect waste solutions in labeled hazardous chemical waste containers for disposal.
  2. Micro-computed tomography (µCT) evaluation
    1. Perform µCT scanning on femoral specimens collected at 4 weeks post-implantation with the following scanning parameters: X-ray tube potential, 70 kVp; X-ray intensity, 0.2 mA; filter, AL 0.5 mm; integration time, 1 x 300 ms; and voxel size, 17 µm.
    2. Perform 3D reconstruction using the predefined scripts provided in the µCT machine software. Generate cross-sectional images of the defect site using the reslice function of dedicated image-processing software. Follow the standard workflows as described in the respective software manuals.
  3. Histological staining
    1. Decalcify the samples at 4 °C using 12% ethylenediaminetetraacetic acid (EDTA) (pH 7.2) for 6 weeks on a shaker.
      NOTE: Replace the EDTA solution every 2 days. Verify decalcification progress regularly. Decalcification is complete when the sample is pliable under pressure or passes a needle.
      CAUTION: Dispose of EDTA solutions according to institutional guidelines for aqueous chemical waste. Contaminated biological material should be treated as biohazardous waste.
    2. Dehydrate the samples in a graded series of ethanol dilutions and then embed them in paraffin.
    3. Cut the paraffin-embedded samples sagittally into 6 µm-thick slices.
      NOTE: Ensure sections include the interface between the implanted material and host tissue.
    4. Perform hematoxylin-eosin (H&E) and Masson's trichrome staining using commercially available kits (see Table of Materials). Carry out all procedures strictly according to the manufacturer's standard protocols without any deviations. Observe defect site healing by histopathology.
      CAUTION: Dispose of staining solutions and contaminated consumables in accordance with institutional biosafety and hazardous waste disposal protocols.
  4. Immunofluorescent (IF) staining
    1. Deparaffinize and rehydrate 6 µm-thick paraffin sections using a series of xylene and graded ethanol solutions.
      CAUTION: Xylene is flammable and toxic. Handle xylene in a fume hood while wearing gloves, a lab coat, and safety goggles. Collect waste solvents separately in chemical waste containers for appropriate disposal.
    2. Perform antigen retrieval by incubating the slides in citrate buffer (pH 6.0) at 95-100 °C for 15-20 min. Allow the slides to cool naturally to room temperature.
    3. Wash the sections with phosphate-buffered saline (PBS) for 5 min and treat the sections with 0.1% Triton X-100 in PBS for 20 min to facilitate antibody penetration. Then block non-specific binding sites with 5% bovine serum albumin (BSA) in PBS for 1 h at room temperature.
      CAUTION: Triton X-100 is an irritant; handle with gloves, a lab coat, and safety goggles in a ventilated area. Dispose of Triton X-100 solutions as chemical waste in designated containers. For all procedures, solutions that have come into contact with tissue sections, such as BSA and PBS, should be regarded as biological waste and must be collected and disposed of in accordance with institutional biosafety protocols.
    4. Incubate the sections with primary antibodies (anti-Piezo type mechanosensitive ion channel component 1 (Piezo1), diluted 1:200; anti-leptin receptor (LepR), diluted 1:50) in antibody dilution buffer. Apply 50 µL of diluted antibody solution to each section and incubate for 12-18 h at 4 °C in the dark in a humidified chamber.
    5. Wash the slides with PBS (3 × 5 min), then incubate them with fluorophore-conjugated secondary antibodies (diluted 1:200). Apply 50 µL of diluted antibody solution to each section and incubate for 2 h at room temperature in the dark in a humidified chamber.
    6. Rinse the slides again in PBS (3 × 5 min), then counterstain nuclei with 4',6-diamidino-2-phenylindole (DAPI) (diluted 1:50) for 5 min.
    7. Mount the sections with antifade mounting medium and cover with a coverslip.
    8. Visualize and capture images using a laser scanning confocal microscope with appropriate filter settings.

Access restricted. Please log in or start a trial to view this content.

Results

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

In this study, the half-segmental diaphyseal bone defect model in rats was established through drilling a defect measuring 4 mm in length and 1.5 mm in radius at the femoral midshaft with a flat-ended cylindrical specific dental diamond bur (Figure 2A). To demonstrate the model's capability of testing materials' mechanical and biological performance, we set two comparison groups, with defects implanted with either GelMA hydrogels or 3DP-PMMA (Figure 3,

Access restricted. Please log in or start a trial to view this content.

Discussion

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

Apart from the widely emphasized properties of osteoinductivity, osteointegration, and biocompatibility5, the mechanical responsiveness and load-bearing capacity of bone substitutes are equally vital, particularly for repairing defects in weight-bearing bones22. Conventional assessments of mechanical performance typically rely on in vitro methods, such as universal testing machines, compression testing, or tensile strength analysis23,

Access restricted. Please log in or start a trial to view this content.

Disclosures

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

All the original data and images are included in this paper. The authors declare no conflicts of interest.

Acknowledgements

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,

This study was supported by the Natural Science Foundation of Sichuan Province (2025ZNSFSC0754).

Access restricted. Please log in or start a trial to view this content.

Materials

List of materials used in this article
NameCompanyCatalog NumberComments
2% Iodophor solutionChengdu Jinshan Chemical Reagent Co., Ltd.None
3D-printed polymethyl methacrylate (3DP-PMMA)kindly provided by the College of Polymer Science and Engineering, Sichuan UniversityNone
4% paraformaldehydeBiosharpBL539A
75% EthanolChengdu Jinshan Chemical Reagent Co., Ltd.None
Cotton ballsHaishi Hainuo Group Co.,  Ltd.None
Cotton sticksLakong Medical Devices Co., Ltd.None
DAPI staining solutionBeyotime Cat#C1005
Disposable sterile scalpelHangzhou Huawei Medical Supplies Co., Ltd.20100227
Gelatin methacryloyl (GelMA) hydrogelskindly provided by the College of Polymer Science and Engineering, Sichuan UniversityNone
GelMA 20%CellinkIKG125000005
Goat Anti-Mouse IgG H&L (Alexa Fluor® 594)Abcamab150116
Goat Anti-Rabbit IgG H&L (Alexa Fluor® 488)Abcamab150077
Hematoxylin and Eosin Stain kitBiosharpC1005
High-speed handpieceFoshan SCS Medical Instrument Co., Ltd.9018499000
Irrigation needleSichuan New Century Medical Polymer Products Co., Ltd.None
Masson’s Trichrome Stain KitSolarbioG1340
Medical non-woven fabricsHenan Yadu Industrial Co., Ltd. None
Medical non-woven fabricsHenan Yadu Industrial Co., Ltd. None
Micro-computed tomography (µCT) Scanco Medical AGμCT 45
Mimics 20.0 for cross-sectional images (reslice function)MaterialiseNone
Needle holdersChengdu Shifeng Co., Ltd.None
Ob-R/Leptin Receptor Antibody (B-3)Santa Cruzsc-8391
Olympus Research Grade Whole Slide Scanning System VS200Chengdu Knowledge Technology Co., LtdVS200
Periosteal elevatorChengdu Shifeng Co., Ltd.None
PIEZO1 AntibodyNovusNBP1-78446
Saline solutionSichuan Kelun Pharmaceutical Co., Ltd.None
Scanco medical visualizer software for 3D image reconstruction (predefined scripts)Scanco Medical AGNone
Specific dental diamond bur (TF-22/170-021m/845-021m)Diantong Dental Materials CompanyNone
Sprague-Dawley rats Byrness Weil Biotech LtdNone
Straight ScissorsChengdu Shifeng Co., Ltd.None
Surgical MotorMARATHONN3-140232
Surgical sutures (4-0 monofilament)Hangzhou Huawei Medical Supplies Co., Ltd.None
Vernier caliperDeliDL91150

References

$$\rightleftharpoonup{xx}$$ $$\longleftharp{xx}$$, $$\longrightharp{xx}$$,
  1. Mitchnik, I. Y., et al. Femur fractures and hemorrhagic shock: Implications for point of injury treatment. Injury. 53 (10), 3416-3422 (2022).
  2. Mitchell, S. A. T., Majuta, L. A., Mantyh, P. W. New insights in understanding and treating bone fracture pain. Curr Osteoporos Rep. 16 (4), 325-332 (2018).
  3. Ritchie, R. O. The conflicts between strength and toughness. Nat Mater. 10 (11), 817-822 (2011).
  4. Carson, J. S., Bostrom, M. P. G. Synthetic bone scaffolds and fracture repair. Injury. 38 (1 Suppl ), S33-S37 (2007).
  5. Janicki, P., Schmidmaier, G. What should be the characteristics of the ideal bone graft substitute? Combining scaffolds with growth factors and/or stem cells. Injury. 42, S77-S81 (2011).
  6. Zandi, M., Dehghan, A., Talimkhani, I., Rezaeian, L., Mohammad Gholi Mezerji, N. Histological evaluation of the healing process of autografted mandibular bone defects in rats under treatment with zoledronate. J Cranio Maxillofac Surg. 47 (11), 1779-1786 (2019).
  7. Wu, J., et al. The establishment of calvarial suture-bony composite defects in rats: A standardized model for suture-regenerative therapy investigation. J Vis Exp. (207), e66417(2024).
  8. Chen, Y., Wu, J., Li, F., Ye, L., Wang, H. Creating a box-cavity defect model in the cortical bone of rat femora. J Vis Exp. (201), e66068(2023).
  9. Yu, C., Yu, F., Li, F., Ye, L. The establishment of a murine mandibular molar extraction socket healing model. J Vis Exp. (191), e64855(2023).
  10. Han, L., et al. 3D magnetic nanocomposite scaffolds enhanced the osteogenic capacities of rat bone mesenchymal stem cells in vitro and in a rat calvarial bone defect model by promoting cell adhesion. J Biomed Mater Res A. 109 (9), 1670-1680 (2021).
  11. Han, Z., Bhavsar, M., Leppik, L., Oliveira, K. M. C., Barker, J. H. Histological scoring method to assess bone healing in critical size bone defect models. Tissue Eng Part C Methods. 24 (5), 272-279 (2018).
  12. Gomes, P. S., Fernandes, M. H. Rodent models in bone-related research: the relevance of calvarial defects in the assessment of bone regeneration strategies. Lab Anim. 45 (1), 14-24 (2011).
  13. Tsiklin, I. L., Shabunin, A. V., Kolsanov, A. V., Volova, L. T. In vivo bone tissue engineering strategies: advances and prospects. Polymers. 14 (15), 3222(2022).
  14. Long, S., et al. Architecture mechanics mediated osteogenic progression in bone regeneration of artificial scaffolds. Sci Adv. 11 (29), eadv8804(2025).
  15. Man, K., et al. GelMA hydrogel reinforced with 3D printed PEGT/PBT scaffolds for supporting epigenetically-activated human bone marrow stromal cells for bone repair. J Funct Biomater. 13 (2), 41(2022).
  16. Chen, N. Embedded 3D printing and pressurized thermo-curing of PMMA for medical implants. J Mech Behav Biomed Mater. 146, 106083(2023).
  17. Einhorn, T. A., Gerstenfeld, L. C. Fracture healing: mechanisms and interventions. Nat Rev Rheumatol. 11 (1), 45-54 (2015).
  18. Ikeda, T., et al. Stimulation of osteogenesis in bone defects implanted with biodegradable hydroxyapatite composed of rod-shaped particles under mechanical unloading. Acta Histochem Cytochem. 45 (5), 283-292 (2012).
  19. Du, Y., Xu, B., Li, Q., Peng, C., Yang, K. The role of mechanically sensitive ion channel Piezo1 in bone remodeling. Front Bioeng Biotechnol. 12, 1342149(2024).
  20. Tadge, T., et al. The role of Piezo1 and Piezo2 proteins in tissue engineering: a comprehensive review. Eng Regen. 5 (2), 170-185 (2024).
  21. Prasad, P., Cancelas, J. A. From marrow to bone and fat: exploring the multifaceted roles of Leptin receptor positive bone marrow mesenchymal stromal cells. Cells. 13 (11), 910(2024).
  22. Hart, N. H., et al. Mechanical basis of bone strength: influence of bone material, bone structure and muscle action. J Musculoskelet Neuronal Interact. 17 (3), 114-139 (2017).
  23. Zhong, J., He, D. Combination of universal mechanical testing machine with atomic force microscope for materials research. Sci Rep. 5 (1), 12998(2015).
  24. El Kommos, A., Jackson, A. R., Andreopoulos, F., Travascio, F. Development of improved confined compression testing setups for use in stress relaxation testing of viscoelastic biomaterials. Gels. 10 (5), 329(2024).
  25. Lietaert, K., Cutolo, A., Boey, D., Van Hooreweder, B. Fatigue life of additively manufactured Ti6Al4V scaffolds under tension-tension, tension-compression and compression-compression fatigue load. Sci Rep. 8 (1), 4957(2018).
  26. Bignardi, C., Terzini, M. Novel load systems for in vitro testing of biomaterials and medical devices. Materials. 16 (2), 465(2023).
  27. Teti, A. Bone development: overview of bone cells and signaling. Curr Osteoporos Rep. 9 (4), 264-273 (2011).
  28. McGovern, J. A., Griffin, M., Hutmacher, D. W. Animal models for bone tissue engineering and modelling disease. Dis Model Mech. 11 (4), dmm033084(2018).
  29. Malda, J., et al. 25th anniversary article: engineering hydrogels for biofabrication. Adv Mater. 25 (36), 5011-5028 (2013).
  30. Altmann, B., et al. Promotion of osteoblast differentiation in 3D biomaterial micro-chip arrays comprising fibronectin-coated poly(methyl methacrylate) polycarbonate. Biomaterials. 32 (34), 8947-8956 (2011).
  31. Bastidas-Coral, A. P., et al. Polymethyl methacrylate does not adversely affect the osteogenic potential of human adipose stem cells or primary osteoblasts. J Biomed Mater Res B Appl Biomater. 108 (4), 1536-1545 (2020).
  32. Öztürk, Z., Tosun, B. Comparison of 3D printed and conventional denture base materials in terms of durability and performance characteristics. Sci Rep. 15, 18234(2025).
  33. Wu, J., et al. Transcriptomic and cellular decoding of scaffolds-induced suture mesenchyme regeneration. Int J Oral Sci. 16 (1), 33(2024).
  34. Kan, C., et al. BMP-dependent, injury-induced stem cell niche as a mechanism of heterotopic ossification. Stem Cell Res Ther. 10 (1), 14(2019).
  35. Bao, X., et al. 3D biomimetic artificial bone scaffolds with dual-cytokines spatiotemporal delivery for large weight-bearing bone defect repair. Sci Rep. 7 (1), 7814(2017).

Access restricted. Please log in or start a trial to view this content.

Reprints and Permissions

Request permission to reuse the text or figures of this JoVE article

Request Permission

Tags

Bone Defect ModelLoad Bearing BoneDiaphyseal DefectRat Femur ModelBone Substitute Evaluation3D Printed PMMAGelatin Methacryloyl HydrogelMechanical Performance TestingOsseointegration AssessmentImmunofluorescence Staining

Related Articles