Creating a Box-Cavity Defect Model in the Cortical Bone of Rat Femora

Summary

Here, we present a protocol for creating a box-cavity defect in rat femoral diaphysis tissue. This model can assess biomaterial performance under biomechanical stress and explore mechanisms of bone regeneration related to intramembranous osteogenesis.

Abstract

Severe bone defects or complex fractures can result in serious complications such as nonunion or insufficient bone healing. Tissue engineering, which involves the application of cells, scaffolds, and cytokines, is considered a promising solution for bone regeneration. Consequently, various animal models that simulate bone defects play a crucial role in exploring the therapeutic potential of tissue engineering for bone healing. In this study, we established a box-shaped cortical bone defect model in the mid-femur of rats, which could serve as an ideal model for assessing the function of biomaterials in promoting bone healing. This box-shaped cortical bone defect was drilled using an oral low-speed handpiece and shaped by a lathe needle. Post-operative micro-CT analysis was immediately conducted to confirm the successful establishment of the box-cavity cortical bone defect. The femurs on the operated side of the rats were then harvested at multiple time points post-surgery (0 days, 2 weeks, 4 weeks, and 6 weeks). The healing process of each sample's defect area was evaluated using micro-CT, hematoxylin and eosin (H&E) staining, and Masson trichrome staining. These results demonstrated a healing pattern consistent with intramembranous ossification, with healing essentially complete by 6 weeks. The categorization of this animal model's healing process provides an effective in vivo method for investigating novel biomaterials and drugs that target intramembranous ossification during bone tissue defect healing.

Introduction

Fractured and defective bone often results from trauma, tumors, inflammation, and congenital malformations^1,2. Although bone tissue in young healthy individuals typically possesses robust regenerative abilities³, defects exceeding a critical size or healing impediments due to systemic diseases (e.g., diabetes, osteoporosis, and infections) may still lead to complications such as bone discontinuity or impaired healing⁴. To address this clinical challenge, bone grafting or biomaterials are commonly used to replace severely defective bone or to reconstruct large bone segments. However, these treatments have limitations. For instance, although considered the gold standard, autologous bone grafting suffers from restricted donor supply and potential donor site complications^5,6. Allografts also present certain risks, such as immune-mediated rejection, potential transmission of diseases, and negative impacts on the biomechanical and biological properties of the graft⁷.

Recent years have witnessed a surge in research focusing on bone defect healing mechanisms. The use of alternative biomaterials and advancements in tissue engineering have emerged as prominent topics within the domain of bone regeneration⁸. Before these biomaterials can be applied to human therapy, they must be tested in vitro and in vivo to ensure their efficacy and safety. However, the reduced complexity of in vitro environments and the absence of immune and inflammatory responses limit the evaluation of various biomaterials in vitro. Consequently, the establishment of animal models for various types of bone tissue defects is needed⁹. Animal models allow the evaluation of biomaterials under different loading conditions, facilitate understanding of species-specific bone characteristics, and provide insight into the similarity between animal models and human clinical situations. These advantages are essential for studying bone-scaffold interactions and translating research findings into clinical practice^9,10.

Currently, mechanical bone defect animal models are widely used to validate the performance of biomaterials, with cranial bone defect models and segmental bone defect models being the most commonly applied methods¹¹. Segmental bone defect models, often utilized to mimic severe long bone or tibial trauma ending in bone nonunion, are advantageous due to their uniform dimensions and defined anatomical positions, simplifying radiological or histological evaluations of new bone formation and revascularization. However, these models require metal implants to stabilize bilateral fracture segments and necessitate a complex healing process involving both endochondral and intramembranous ossification¹². On the other hand, calvarial bone defect models have become a primary screening tool for evaluating biomaterials due to their standardized defect diameters, convenient surgical access, and the supportive function of dura mater and soft tissue¹³. Although they are widely used for modeling intramembranous bone formation in clinically relevant scenarios, they are unsuitable for evaluating bone healing under biomechanical loading conditions due to their non-load-bearing nature during the healing process¹⁴.

To address these limitations, we established a box-cavity cortical bone defect model in the femoral diaphysis tissue of rats. Utilizing micro-computed tomography (CT) three-dimensional (3D) reconstruction, and histopathological staining (Hematoxylin and eosin [HE] and Masson), we analyzed the healing process of this model under hemostasis conditions. We aim to offer fresh insights for evaluating biomaterial performance under biomechanical loading conditions and for studying the bioengineering and mechanism of bone regeneration vis-à-vis intramembranous ossification.

Protocol

All animal procedures in this study were reviewed and approved by the Ethical Committee of the West China School of Stomatology, Sichuan University (WCHSIRB-D-2021-597). Sprague-Dawley rats (male, body weight 300 g) were used for the present study.

1. Presurgical preparation

1. Instrument preparation
   1. Refer to Figure 1A for the surgical instruments used in this study: electric shaver, tissue scissors, ophthalmic scissors, ophthalmic forceps, disposable scalpel, periosteal separator, oral low-speed handpiece, oral probe, disposable irrigation vac, needle holder, 3.0 suture.
   2. Prepare an oral probe and mark the large curved end of the probe with a marker pen according to the diameter of the defect (Figure 1B). Use this to determine the size of the defect during the surgery.
   3. Sterilize all surgical materials and instruments used to perform the procedure before use. Pack the desired materials in folded cloth or wrapping paper and seal them with autoclave tape for steam sterilization (125-135°C for 20-25 min).
   4. Surgical area preparation: Disinfect the operating table and the environment around the table by spraying with 75% alcohol. Create an approximately 60 cm x 90 cm sterile area with autoclaved drapes on the operating table.
2. Anesthesia preparation
   1. Anesthetize the rats intraperitoneally with 10% ketamine hydrochloride (50-100 mg/kg) and 2% xylazine 2 mg/kg). Use subcutaneous injection of carprofen (5 mg/kg) for preoperative and intraoperative analgesia. Examine the depth of anesthesia by toe pinch test. After anesthesia, apply sterile eye ointment to the eyes to prevent dry eyes and corneal injury.

2. Surgical procedure

1. Place the rat in a lateral recumbency position on the sterile surgical table and remove the lower limb hairs with an electric shaver.
2. Use 5% iodophor solution and 75% alcohol to disinfect the skin tissue in the surgical area.
3. Locate the proximal and distal femur and make a 2.5 cm incision along the long axis of the femur to cut through the skin tissue of the rat.
   1. Separate the skin layer from the fascia with ophthalmic forceps and tissue scissors, and expose the lateral approach to the femur through the biceps femoris and lateral femoral muscles.
   2. Locate the intersection of the two muscle septa (a white tissue line) and carefully separate with a disposable surgical blade along the muscle border until the femoral surface is reached.
      NOTE: When using a disposable blade to separate the muscle, it is important to separate along the muscle septum and be careful of causing vascular injury within the soft tissue. Beginners and those unfamiliar with the anatomy must use ophthalmic forceps and periosteal separators in blunt dissection between the two muscle bulks.
4. Apply a periosteal separator to bluntly separate the femoral surface muscles and expose the middle of the femoral diaphysis.
5. Use a sterile marker pen to mark the area of the defect site on the mid-surface of the femoral diaphysis, located at the top of the lateral 1/3 oblique crest of the femoral head.
6. Use the oral low-speed handpiece with a 1.2 mm diameter slow-motion ball drill to drill a small hole perpendicular to the bone surface at the marked site, destroying the periosteum and the bone cortex with a depth deep enough to reach the bone marrow cavity. At this drilling depth level, expand the size of the hole parallel to this depth in all directions, trimming to achieve a box-cavity shape.
7. Use a labeled oral probe parallel to the edge of the defect to determine the defect diameter and morphology during and after preparation.
8. Close the muscle and skin layers with 3-0 monofilament absorbable sutures, respectively, and disinfect the surgical area with 5% iodophor from the inside out.

3. Post-operative care

1. After surgery, administer carprofen (5 mg/kg) subcutaneously and put the rat on a constant temperature heating pad until recovery from anesthesia. When the rat regains consciousness, gently move it to a cage that contains dry, autoclaved bedding.
2. Continue analgesia for 24 h and post-operative monitoring for 1 week after surgery.

4. Sample collection and analysis

1. Humanly euthanize the rat after surgery by injecting pentobarbital sodium 100-200 mg/kg intraperitoneally. Carefully separate the muscle and fascial tissue on the surface of the femur and remove the femur on the operated side completely. Collect specimens at 0 days (Figure 2A, B), 2 weeks, 4 weeks, and 6 weeks postoperatively.
2. Fix the femoral specimens in 4% paraformaldehyde for 24 h. Analyze the structure of the femora by using micro-computed tomography. Set the scanning parameters as follows: X-ray tube potential, 70 kVp; filter, AL 0.5 mm; X-ray intensity, 0.2 mA; voxel size, 17 µm; and integration time, 1 × 300 ms. Reconstruct 3D model images using bitmap data.
3. Decalcify the specimens in 10% EDTA for 8 weeks before dehydrating the femoral specimens in a graded series of ethanol dilutions. Then, embed the samples in paraffin wax¹⁵.
   1. Cut the embedded samples into 5 µm paraffin sections from the sagittal plane.
   2. Stain sections with both a hematoxylin and eosin (H&E) staining kit and a Masson staining kit. Observe the healing of the defect area by histopathology.

Results

In this protocol, we successfully establish a rat femoral box-cavity defect model with dimensions of 4.5 mm x 1.5 mm by drilling. In order to analyze the healing process, we collected the femoral tissue on the operated side at 0 days, 2 weeks, 4 weeks, and 6 weeks after surgery, which are the key time points of endochondral ossification, intramembranous ossification, and bone remodeling during the healing process of femoral trauma in rats². On post-operative day 0,...

Discussion

Preclinical animal models are vital for examining bone healing and the influence of biomaterials on bone regeneration. This protocol illustrates a femoral box-cavity defect model replicating the intramembranous bone formation process associated with clinical bone regeneration. The defect area was intraoperatively standardized using a pre-marked oral probe. Micro-CT and histopathological staining results showed progressive healing over 6 weeks, with thickened periosteum and new trabecular bone formation, followed by dense...

Materials

Name	Company	Catalog Number	Comments
1.2 mm slow speed ball drill	Dreybird Medical Equipment Co., Ltd.	RA3-012	For preparation of box cavity defects
3.0 suture	Chengdu Shifeng Co., Ltd.	None	For suturing wounds
4% paraformaldehyde	Biosharp	BL539A	For fix the femoral specimens
Cotton balls	Haishi Hainuo Group Co.,  Ltd.	20120047	For skin sterilization and cleaning of surgical field
Cotton sticks	Lakong Medical Devices Co., Ltd.	M6500R	For skin disinfection
Dental technician grinding machine	Marathon	N3-140232	For preparation of box cavity defects
Disposable scalpel	Hangzhou Huawei Medical Supplies Co., Ltd.	20100227	For creating skin incisions as well as to sharply separate muscle tissue
Electric shaver	JASE	BM320210	Removal of hair tissue from the surgical area
Hematoxylin and Eosin Stain kit	Biosharp	C1005	For the histological analysis of the specimens
Masson’s Trichrome Stain Kit	Solarbio	G1340	For the histological analysis of the specimens
Micro CT	Scanco medical ag	µCT 45	For analyzing the healing of defects in femoral samples
Needle holder	Chengdu Shifeng Co., Ltd.	None	For suture-holding needles
Olympus Research Grade Whole Slide Scanning System VS200	Chengdu Knowledge Technology Co.	VS200	For analyzing the results of HE staining and Masson staining
Ophthalmic forceps	Chengdu Shifeng Co., Ltd.	None	For clamping skin, muscle tissue
Ophthalmic scissors	Chengdu Shifeng Co., Ltd.	None	For forming a skin incision approach
Oral low-speed handpiece	Marathon	Y221101003	For preparation of box cavity defects
Oral probe	Shanghai Sangda Medical Insurance Co., Ltd.	20000143	For measuring the diameter of defects
Periosteal separator	Chengdu Shifeng Co., Ltd.	None	For blunt separation of muscle tissue
Sprague–Dawley rats	Byrness Weil Biotech Ltd	None	For the establishment of femoral bone boxy cavitary defect
Tissue scissors	Chengdu Shifeng Co., Ltd.	None	For forming a skin incision approach