This study was funded by grants from the National Natural Science Foundation of China 82101000 (H. W.), U21A20368 (L. Y.), and 82100982 (F. L.), and supported by Sichuan Science and Technology Program 2023NSFSC1499 (H. W.).

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
<ol>
	<li>Instrument preparation
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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&#176;C for 20-25 min).</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Anesthesia preparation
	<ol>
		<li>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.</li>
	</ol>
	</li>
</ol>
2. Surgical procedure
<ol>
	<li>Place the rat in a lateral recumbency position on the sterile surgical table and remove the lower limb hairs with an electric shaver.</li>
	<li>Use 5% iodophor solution and 75% alcohol to disinfect the skin tissue in the surgical area.</li>
	<li>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.
	<ol>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Apply a periosteal separator to bluntly separate the femoral surface muscles and expose the middle of the femoral diaphysis.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Use a labeled oral probe parallel to the edge of the defect to determine the defect diameter and morphology during and after preparation.</li>
	<li>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.</li>
</ol>
3. Post-operative care
<ol>
	<li>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.</li>
	<li>Continue analgesia for 24 h and post-operative monitoring for 1 week after surgery.</li>
</ol>
4. Sample collection and analysis
<ol>
	<li>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.</li>
	<li>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 &#181;m; and integration time, 1 &#215; 300 ms. Reconstruct 3D model images using bitmap data.</li>
	<li>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.
	<ol>
		<li>Cut the embedded samples into 5 &#181;m paraffin sections from the sagittal plane.</li>
		<li>Stain sections with both a hematoxylin and eosin (H&#38;E) staining kit and a Masson staining kit. Observe the healing of the defect area by histopathology.</li>
	</ol>
	</li>
</ol>

Establishment and Evaluation of a Rat Femoral Box: Cavity Defect Model

<ol><li>Einhorn, T. A., Gerstenfeld, L. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Fracture+healing%3A+mechanisms+and+interventions%5BTitle%5D)+AND+%22Nat+Rev+Rheumatol%22%5BJournal%5D)">Fracture healing: mechanisms and interventions.</a> Nat Rev Rheumatol. 11 (1), 45-54 (2015).</li>
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<li>Pearce, A. I., Richards, R. G., Milz, S., Schneider, E., Pearce, S. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Animal+models+for+implant+biomaterial+research+in+bone%3A+a+review%5BTitle%5D)+AND+%22Eur+Cell+Mater%22%5BJournal%5D)">Animal models for implant biomaterial research in bone: a review.</a> Eur Cell Mater. 13, 1-10 (2007).</li>
<li>McGovern, J. A., Griffin, M., Hutmacher, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Animal+models+for+bone+tissue+engineering+and+modelling+disease%5BTitle%5D)+AND+%22Dis+Model+Mech%22%5BJournal%5D)">Animal models for bone tissue engineering and modelling disease.</a> Dis Model Mech. 11 (4), dmm033084(2018).</li>
<li>Bigham-Sadegh, A., Oryan, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Selection+of+animal+models+for+preclinical+strategies+in+evaluating+the+fracture+healing%2C+bone+graft+substitutes+and+bone+tissue+regeneration+and+engineering%5BTitle%5D)+AND+%22Connect+Tissue+Res%22%5BJournal%5D)">Selection of animal models for preclinical strategies in evaluating the fracture healing, bone graft substitutes and bone tissue regeneration and engineering.</a> Connect Tissue Res. 56 (3), 175-194 (2015).</li>
<li>Horner, E. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Long+bone+defect+models+for+tissue+engineering+applications%3A+criteria+for+choice%5BTitle%5D)+AND+%22Tissue+Eng+Part+B+Rev%22%5BJournal%5D)">Long bone defect models for tissue engineering applications: criteria for choice.</a> Tissue Eng Part B Rev. 16 (2), 263-271 (2010).</li>
<li>Vajgel, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+systematic+review+on+the+critical+size+defect+model%5BTitle%5D)+AND+%22Clin+Oral+Implants+Res%22%5BJournal%5D)">A systematic review on the critical size defect model.</a> Clin Oral Implants Res. 25 (8), 879-893 (2014).</li>
<li>Samsonraj, R. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+versatile+protocol+for+studying+calvarial+bone+defect+healing+in+a+mouse+model%5BTitle%5D)+AND+%22Tissue+Eng+Part+C+Methods%22%5BJournal%5D)">A versatile protocol for studying calvarial bone defect healing in a mouse model.</a> Tissue Eng Part C Methods. 23 (11), 686-693 (2017).</li>
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<li>Colnot, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Skeletal+cell+fate+decisions+within+periosteum+and+bone+marrow+during+bone+regeneration%5BTitle%5D)+AND+%22J+Bone+Miner+Res%22%5BJournal%5D)">Skeletal cell fate decisions within periosteum and bone marrow during bone regeneration.</a> J Bone Miner Res. 24 (2), 274-282 (2009).</li>
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All the original data and images are included in this paper. Authors declare no conflict of interest

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...

Fractured and defective bone often results from trauma, tumors, inflammation, and congenital malformations1,2. 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 complications5,6. 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 practice9,10.
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-&#224;-vis intramembranous ossification.

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

creating a box-cavity defect model in the cortical bone of rat femora

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&#39;s defect area was evaluated using micro-CT, hematoxylin and eosin (H&#38;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&#39;s healing process provides an effective in vivo method for investigating novel biomaterials and drugs that target intramembranous ossification during bone tissue defect healing.

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,...

Watch this Scientific Journal Video about Creating a Box-Cavity Defect Model in the Cortical Bone of Rat Femora at JoVE.com

Author Spotlight: The Box-Cavity Cortical Approach for Enhanced Evaluation of Biomaterials and Bone Regeneration

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.

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

Severe bone defects or complex fractures can result in serious complications such as nonunion or insufficient bone healing. Tissue engineering, which ...

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700 Views.  West China Hospital of Stomatology, Sichuan University. 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.

Video: Author Spotlight: The Box-Cavity Cortical Approach for Enhanced Evaluation of Biomaterials and Bone Regeneration

Adjustable Stiffness, External Fixator for the Rat Femur Osteotomy and Segmental Bone Defect Models

The mechanical environment around the healing of broken bone is very important as it determines the way the fracture will heal. Over the past decade there has been great clinical interest in improving bone healing by altering the mechanical environment through the fixation stability around the lesion. One constraint of preclinical animal research in this area is the lack of experimental control over the local mechanical environment within a large segmental defect as well as osteotomies as they heal. In this paper we report on the design and use of an external fixator to study the healing of large segmental bone defects or osteotomies. This device not only allows for controlled axial stiffness on the bone lesion as it heals, but it also enables the change of stiffness during the healing process in vivo. The conducted experiments have shown that the fixators were able to maintain a 5 mm femoral defect gap in rats in vivo during unrestricted cage activity for at least 8 weeks. Likewise, we observed no distortion or infections, including pin infections during the entire healing period. These results demonstrate that our newly developed external fixator was able to achieve reproducible and standardized stabilization, and the alteration of the mechanical environment of in vivo rat large bone defects and various size osteotomies. This confirms that the external fixation device is well suited for preclinical research investigations using a rat model in the field of bone regeneration and repair.

One constraint of preclinical research in the field of bone repair is the lack of experimental control over the local mechanical environment within a healing bone lesion. We report the design and use of an external fixator for bone repair with the ability to change fixator stiffness in vivo.

The mechanical environment around the healing of broken bone is very important as it determines the way the fracture will heal. Over the past decade there ...

A Simple Critical-sized Femoral Defect Model in Mice

While bone has a remarkable capacity for regeneration, serious bone trauma often results in damage that does not properly heal. In fact, one tenth of all limb bone fractures fail to heal completely due to the extent of the trauma, disease, or age of the patient. Our ability to improve bone regenerative strategies is critically dependent on the ability to mimic serious bone trauma in test animals, but the generation and stabilization of large bone lesions is technically challenging. In most cases, serious long bone trauma is mimicked experimentally by establishing a defect that will not naturally heal. This is achieved by complete removal of a bone segment that is larger than 1.5 times the diameter of the bone cross-section. The bone is then stabilized with a metal implant to maintain proper orientation of the fracture edges and allow for mobility. 
Due to their small size and the fragility of their long bones, establishment of such lesions in mice are beyond the capabilities of most research groups. As such, long bone defect models are confined to rats and larger animals. Nevertheless, mice afford significant research advantages in that they can be genetically modified and bred as immune-compromised strains that do not reject human cells and tissue.
Herein, we demonstrate a technique that facilitates the generation of a segmental defect in mouse femora using standard laboratory and veterinary equipment. With practice, fabrication of the fixation device and surgical implantation is feasible for the majority of trained veterinarians and animal research personnel. Using example data, we also provide methodologies for the quantitative analysis of bone healing for the model.

Animal models are frequently employed to mimic serious bone injury in biomedical research. Due to their small size, establishment of stabilized bone lesions in mice are beyond the capabilities of most research groups. Herein, we describe a simple method for establishing and analyzing experimental femoral defects in mice.

While bone has a remarkable capacity for regeneration, serious bone trauma often results in damage that does not properly heal. In fact, one tenth of all ...

Semiautomated Longitudinal Microcomputed Tomography-based Quantitative Structural Analysis of a Nude Rat Osteoporosis-related Vertebral Fracture Model

Osteoporosis-related vertebral compression fractures (OVCFs) are a common and clinically unmet need with increasing prevalence as the world population ages. Animal OVCF models are essential to the preclinical development of translational tissue engineering strategies. While a number of models currently exist, this protocol describes an optimized method for inducing multiple highly reproducible vertebral defects in a single nude rat. A novel longitudinal semiautomated microcomputed tomography (&#181;CT)-based quantitative structural analysis of the vertebral defects is also detailed. Briefly, rats were imaged at multiple time points post-op. The day 1 scan was reoriented to a standard position, and a standard volume of interest was defined. Subsequent &#181;CT scans of each rat were automatically registered to the day 1 scan so the same volume of interest was then analyzed to assess for new bone formation. This versatile approach can be adapted to a variety of other models where longitudinal imaging-based analysis could benefit from precise 3D semiautomated alignment. Taken together, this protocol describes a readily quantifiable and easily reproducible system for osteoporosis and bone research. The suggested protocol takes 4 months to induce osteoporosis in nude ovariectomized rats and between 2.7 and 4 h to generate, image, and analyze two vertebral defects, depending on tissue size and equipment.

The goal of this protocol is to generate a nude rat osteoporosis-related vertebral compression fracture model that can be longitudinally evaluated in vivo using a semiautomated microcomputed tomography-based quantitative structural analysis.

Osteoporosis-related vertebral compression fractures (OVCFs) are a common and clinically unmet need with increasing prevalence as the world population ...

Bioengineering

Establishment of a Segmental Femoral Critical-size Defect Model in Mice Stabilized by Plate Osteosynthesis

The use of tissue-engineered bone constructs is an appealing strategy to overcome drawbacks of autografts for the treatment of massive bone defects. As a model organism, the mouse has already been widely used in bone-related research. Large diaphyseal bone defect models in mice, however, are sparse and often use bone fixation which fills the bone marrow cavity and does not provide optimal mechanical stability. The objectives of the current study were to develop a critical-size, segmental, femoral defect in nude mice. A 3.5-mm mid-diaphyseal femoral ostectomy (approximately 25% of the femur length) was performed using a dedicated jig, and was stabilized with an anterior located locking plate and 4 locking screws. The bone defect was subsequently either left empty or filled with a bone substitute (syngenic bone graft or coralline scaffold). Bone healing was monitored noninvasively using radiography and in vivo micro-computed-tomography and was subsequently assessed by ex vivo micro-computed-tomography and undecalcified histology after animal sacrifice, 10 weeks postoperatively. The recovery of all mice was excellent, a full-weight-bearing was observed within one day following the surgical procedure. Furthermore, stable bone fixation and consistent fixation of the implanted materials were achieved in all animals tested throughout the study. When the bone defects were left empty, non-union was consistently obtained. In contrast, when the bone defects were filled with syngenic bone grafts, bone union was always observed. When the bone defects were filled with coralline scaffolds, newly-formed bone was observed in the interface between bone resection edges and the scaffold, as well as within a short distance within the scaffold. 
The present model describes a reproducible critical-size femoral defect stabilized by plate osteosynthesis with low morbidity in mice. The new load-bearing segmental bone defect model could be useful for studying the underlying mechanisms in bone regeneration pertinent to orthopaedic applications.

Although mouse models are invaluable tools for bone tissue engineering, models of long bone defects are sparse. This need motivated development of the present protocol which uses a locking plate with four screws and a dedicated jig to perform and stabilize a reproducible, femoral, critical-size defect with low morbidity.

The use of tissue-engineered bone constructs is an appealing strategy to overcome drawbacks of autografts for the treatment of massive bone defects. As a ...

A Reliable and Reproducible Critical-Sized Segmental Femoral Defect Model in Rats Stabilized with a Custom External Fixator

Orthopedic research relies heavily on animal models to study mechanisms of bone healing in vivo as well as investigate the new treatment techniques. Critical-sized segmental defects are challenging to treat clinically, and research efforts could benefit from a reliable, ambulatory small animal model of a segmental femoral defect. In this study, we present an optimized surgical protocol for the consistent and reproducible creation of a 5 mm critical diaphyseal defect in a rat femur stabilized with an external fixator. The diaphyseal ostectomy was performed using a custom jig to place 4 Kirschner wires bicortically, which were stabilized with an adapted external fixator device. An oscillating bone saw was used to create the defect. Either a collagen sponge alone or a collagen sponge soaked in rhBMP-2 was implanted into the defect, and the bone healing was monitored over 12 weeks using radiographs. After 12 weeks, rats were sacrificed, and histological analysis was performed on the excised control and treated femurs. Bone defects containing only collagen sponge resulted in non-union, while rhBMP-2 treatment yielded the formation of a periosteal callous and new bone remodeling. Animals recovered well after implantation, and external fixation proved successful in stabilizing the femoral defects over 12 weeks. This streamlined surgical model could be readily applied to study bone healing and test new orthopedic biomaterials and regenerative therapies in vivo.

In vivo mammalian models of critical-sized bone defects are essential for researchers studying healing mechanisms and orthopedic therapies. Here, we introduce a protocol for the creation of reproducible, segmental, femoral defects in rats stabilized using external fixation.

Orthopedic research relies heavily on animal models to study mechanisms of bone healing in vivo as well as investigate the new treatment techniques. ...

Calvarial Model of Bone Augmentation in Rabbit for Assessment of Bone Growth and Neovascularization in Bone Substitution Materials

The basic principle of the rabbit calvarial model is to grow new bone tissue vertically on top of the cortical part of the skull. This model allows assessment of bone substitution materials for oral and craniofacial bone regeneration in terms of bone growth and neovascularization support. Once animals are anesthetized and ventilated (endotracheal intubation), four cylinders made of polyether ether ketone (PEEK) are screwed onto the skull, on both sides of the median and coronal sutures. Five intramedullary holes are drilled within the bone area delimited by each cylinder, allowing influx of bone marrow cells. The material samples are placed into the cylinders which are then closed. Finally, the surgical site is sutured, and animals are awaken. Bone growth may be assessed on live animals by using microtomography. Once animals are euthanized, bone growth and neovascularization may be evaluated by using microtomography, immune-histology and immunofluorescence. As the evaluation of a material requires maximum standardization and calibration, the calvarial model appears ideal. Access is very easy, calibration and standardization are facilitated by the use of defined cylinders and four samples may be assessed simultaneously. Furthermore, live tomography may be used and ultimately a large decrease in animals to be euthanized may be anticipated.

Here we present a surgical protocol in rabbits with the aim to assess bone substitution materials in terms of bone regeneration capacities. By using PEEK cylinders fixed onto rabbit skulls, osteoconduction, osteoinduction, osteogenesis and vasculogenesis induced by the materials may be evaluated either on live or euthanized animals.

The basic principle of the rabbit calvarial model is to grow new bone tissue vertically on top of the cortical part of the skull. This model allows ...

Proximal Cadaveric Femur Preparation for Fracture Strength Testing and Quantitative CT-based Finite Element Analysis

Cadaveric fracture testing is routinely used to understand factors that affect proximal femur strength. Because ex vivo biological tissues are prone to lose their mechanical properties over time, specimen preparation for experimental testing must be performed carefully to obtain reliable results that represent in vivo conditions. For that reason, we designed a protocol and a set of fixtures to prepare the femoral specimens such that their mechanical properties experienced minimal changes. The femora were kept in a frozen state except during preparation steps and mechanical testing. The relevant clinical measures of total hip and femoral neck bone mineral density (BMD) were obtained with a clinical dual X-ray absorptiometry (DXA) bone densitometer, and the 3D geometry and distribution of bone mineral were obtained using CT with a calibration phantom for quantitative estimations based on the greyscale values. Any possible bone disease, fracture, or the presence of implants or artifacts affecting the bone structure, was ruled out with X-ray scans. For preparation, all bones were carefully cleaned of excess soft tissue, and were cut and potted at the internal rotation angle of interest. A cutting fixture allowed the distal end of the bone to be cut off leaving the proximal femur at a desired length. To allow positioning of the femoral neck at prescribed angles during later CT scanning and mechanical testing, the proximal femoral shafts were potted in polymethylmethacrylate (PMMA) using a fixture designed specifically for desired orientations. The data collected from our experiments were then used for validation of quantitative computed tomography (QCT)-based finite element analysis (FEA), as described in a different protocol. In this manuscript, we present the protocol for the precise bone preparation for mechanical testing and subsequent QCT/FEA modeling. The current protocol was successfully applied to prepare about 200 cadaveric femora over a 6-year time period.

We present a robust protocol on how to carefully preserve and prepare cadaveric femora for fracture testing and quantitative computed tomography imaging. The method provides precise control over input conditions for the purpose of determining relationships between bone mineral density, fracture strength, and defining finite element model geometry and properties.

Cadaveric fracture testing is routinely used to understand factors that affect proximal femur strength.  Because ex vivo biological tissues are prone to ...

Development and Evaluation of a Rat Model of Full-Thickness Cartilage Defects

Cartilage defects of the knee joint caused by trauma are a common sports joint injury in the clinic, and these defects result in joint pain, impaired movement, and eventually, knee osteoarthritis (kOA). However, there is little effective treatment for cartilage defects or even kOA. Animal models are important for developing therapeutic drugs, but the existing models for cartilage defects are unsatisfactory. This work established a full-thickness cartilage defects (FTCD) model by drilling holes in the femoral trochlear groove of rats, and the subsequent pain behavior and histopathological changes were used as readout experiments. After surgery, the mechanical withdrawal threshold was decreased, chondrocytes at the injured site were lost, matrix metalloproteinase MMP13 expression was increased, and type II collagen expression decreased, consistent with the pathological changes observed in human cartilage defects. This methodology is easy and simple to perform and enables gross observation immediately after the injury. Furthermore, this model can successfully mimic clinical cartilage defects, thus providing a platform for studying the pathological process of cartilage defects and developing corresponding therapeutic drugs.

This protocol establishes a full-thickness cartilage defects (FTCD) model by drilling holes in the femoral trochlear groove of rats and measuring the subsequent pain behavior and histopathological changes.

Cartilage defects of the knee joint caused by trauma are a common sports joint injury in the clinic, and these defects result in joint pain, impaired ...

Standardizing Rat Calvarial Suture-Bony Defect Model for Regenerative Studies

The Establishment of Calvarial Suture-Bony Composite Defects in Rats: A Standardized Model for Suture-Regenerative Therapy Investigation

Large-scale calvarial defects often coincide with cranial suture disruption, leading to impairments in calvarial defect restoration and skull development (the latter occurs in the developing cranium). However, the lack of a standardized model hinders progress in investigating suture-regenerative therapies and poses challenges for conducting comparative analyses across distinct studies. To address this issue, the current protocol describes the detailed modeling process of calvarial suture-bony composite defects in rats.
The model was generated by drilling full-thickness rectangular holes measuring 4.5 mm &#215; 2 mm across the coronal sutures. The rats were euthanized, and the cranium samples were harvested postoperatively at day 0, week 2, week 6, and week 12. &#181;CT results from samples collected immediately post-surgery confirmed the successful establishment of the suture-bony composite defect, involving the removal of the coronal suture and the adjacent bone tissues.
Data from the 6th and 12th postoperative weeks demonstrated a natural healing tendency for the defect to close. Histological staining further validated this trend by showing increased mineralized fibers and new bone at the defect center. These findings indicate progressive suture fusion over time following calvarial defects, underscoring the significance of therapeutic interventions for suture regeneration. We anticipate that this protocol will facilitate the development of suture-regenerative therapies, offering fresh insights into the functional restoration of calvarial defects and reducing adverse outcomes associated with suture loss.

Author Spotlight: Development and Evaluation of a Standardized Rat Model for Calvarial Suture-Bony Composite Defects

We describe the detailed surgical procedures of calvarial suture-bony composite defects in rats, alongside the investigations into the short-term and long-term prognoses of the model. We aim to construct a standardized model for developing suture-regenerative therapies.

Large-scale calvarial defects often coincide with cranial suture disruption, leading to impairments in calvarial defect restoration and skull development ...

Establishing a Diaphyseal Femur Fracture Model in Mice

Bones have a significant regenerative capacity. However, fracture healing is a complex process, and depending on the severity of the lesions and the age and overall health status of the patient, failures can occur, leading to delayed union or nonunion. Due to the increasing number of fractures resulting from high-energy trauma and aging, the development of innovative therapeutic strategies to improve bone repair based on the combination of skeletal/mesenchymal stem/stromal cells and biomimetic biomaterials is urgently needed. To this end, the use of reliable animal models is fundamental to better understanding the key cellular and molecular mechanisms that determine the healing outcomes. Of all the models, the mouse is the preferred research model because it offers a wide variety of transgenic strains and reagents for experimental analysis. However, the establishment of fractures in mice may be technically challenging due to their small size. Therefore, this article aims to demonstrate the procedures for the surgical establishment of a diaphyseal femur fracture in mice, which is stabilized with an intramedullary wire and resembles the most common bone repair process, through cartilaginous callus formation.

This protocol describes a surgical procedure for the establishment of a diaphyseal fracture in the femur of mice, which is stabilized with an intramedullary wire, for fracture healing studies.

Bones have a significant regenerative capacity. However, fracture healing is a complex process, and depending on the severity of the lesions and the age ...