Name: Video: Generating a Rabbit Calvarial Defect Model: A Surgical Procedure to Create Round Defects in the Top Portion of the Rabbit Skull to Assess Bone Substitute Materials for Bone Regeneration Capacities - Concept
Uploaded: 2023-04-30T14:55:05.000+00:00
Duration: 1 min 45 s
Description: 1.3K Views. Source: Marger, L. et al., Calvarial Model of Bone Augmentation in Rabbit for Assessment of Bone Growth and Neovascularization in Bone Substitution Materials. J. Vis. Exp. (2019). In this video, we describe the surgical procedure to generate a rabbit calvarial model to evaluate the bone regeneration capacities of bone sample substitution materials of interest.

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All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Surgery
<ol>
	<li>Site preparation
	<ol>
		<li>Place the rabbit on a heated pad (39 &#176;C) covered by a mattress pad (to avoid burns) on the surgery table. Shave the scalp.</li>
		<li>Apply a lubricating gel on the eyes to avoid irritation and dryness. Disinfect the site by scrubbing the skin with povidone iodine (10%). Then drape the rabbit with a sterile surgical drape and cut out an access area for the skull.</li>
		<li>Disinfect the surgical site with povidone iodine (10%) for a second time. Apply a lubricating gel on the eyes to avoid irritation and dryness.</li>
		<li>Prepare a draped table (sterile drape) on which to place the complete surgical tray.</li>
	</ol>
	</li>
	<li>Surgical site opening
	<ol>
		<li>Anesthetize locally with a subcutaneous (SC) injection of lidocaine 2% (1 mL) on the skull.</li>
		<li>Incise through the skin (with a scalpel) along the calvarial sagittal line, from the orbits to the external occipital protuberance (~4 cm in length). Ensure that the periosteum is incised.</li>
		<li>Gently elevate the periosteum (with a periosteal elevator) on both side of the incision. Rinse the site with sterile saline.</li>
	</ol>
	</li>
	<li>Cylinder placement
	<ol>
		<li>Locate the median and coronal sutures on the skull (Figure 2A, B). Note that these anatomical lines form a cross. The cylinders will be placed in each of the quadrants defined by the cross, ensuring that the edge of the cylinder is not over the suture (Figure 2C).</li>
		<li>Place the first cylinder on the left upper quadrant (left frontal bone), and try to lay the device flat. Fix in the position with strong hand pressure and screw a micro-screw, until resistance is felt. Ensure that the screw head is flush with the surface of the cylinder tab.</li>
		<li>Repeat the same procedure on the other tab to fix the cylinder tightly onto the skull. Ensure that the cylinder is hermetically fixed to the bone.</li>
		<li>Repeat the procedure on the right upper quarter (right frontal bone), left lower quarter (left parietal bone) and right lower quarter (right parietal bone).</li>
	</ol>
	</li>
	<li>Bone drilling of 5 intramedullary holes within the area circumscribed by the cylinders (Figure 1)
	<ol>
		<li>Drill an intramedullary hole under saline irrigation (0.8 mm in diameter, ~1 mm in depth) with a round bur on the bone, in the center of the area circumscribed by the cylinder. Ensure that bleeding appears.</li>
		<li>Drill two more intramedullary holes along the axis passing through the two tab screws, at the inner edges of the cylinder. Along the perpendicular axe, drill two more intramedullary holes at the inner edges of the cylinder. Ensure that bleeding appears.</li>
		<li>Repeat the operation within the three other cylinders.</li>
	</ol>
	</li>
	<li>Filling cylinders with material samples and capping (Figure 3)
	<ol>
		<li>Prepare the desired bone substitute material according to manufacturer instructions or material specifications.</li>
		<li>Fill the first cylinder to the brim with the material sample and close the cylinder by fitting the cap. Repeat the process in the 3 other cylinders.</li>
	</ol>
	</li>
	<li>Surgical site closure
	<ol>
		<li>Close the skin above the cylinders with an intermittent non-resorbable suture.</li>
		<li>Apply a sprayable dressing onto the wound.</li>
	</ol>
	</li>
</ol>
2. Post-surgical treatment
<ol>
	<li>Stop analgesia and anesthesia (propofol and fentanyl perfusion arrest) supply and check the recovery of autonomous breathing.</li>
	<li>Stop the ventilation once the animal has recovered autonomous breathing. Maintain the animal under pure oxygen before complete awakening.</li>
	<li>Inject buprenorphine hydrochloride SC (0.02 mg/kg, 0.03 mg/mL, 0.67 mL/kg) and repeat the injection every 6 h for 3 days as post-surgical analgesia.</li>
	<li>Transfer the animal into its usual housing with water and complete feeding.</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Drugs</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fentanyl</td>
			<td>Bischel</td>
			<td></td>
			<td>For analgesia</td>
		</tr>
		<tr>
			<td>Ketalar 50mg/ml</td>
			<td>Pfizer</td>
			<td></td>
			<td>Ketamine for anesthesia</td>
		</tr>
		<tr>
			<td>Lidohex</td>
			<td>Bichsel</td>
			<td></td>
			<td>Lubricating gel for the eyes</td>
		</tr>
		<tr>
			<td>Opsite</td>
			<td>Smith and Nephew</td>
			<td>66004978</td>
			<td>Sprayable dressing</td>
		</tr>
		<tr>
			<td>Povidone iodine 10%, Betadine</td>
			<td>Mundipharma</td>
			<td></td>
			<td>anti-infective agent</td>
		</tr>
		<tr>
			<td>Propofol 2%</td>
			<td>Braun</td>
			<td>3538710</td>
			<td>For anesthesia</td>
		</tr>
		<tr>
			<td>Rapidocain 2%</td>
			<td>sintetica</td>
			<td></td>
			<td>Local anesthesia</td>
		</tr>
		<tr>
			<td>Rompun 2%</td>
			<td>Bayer</td>
			<td></td>
			<td>Xylazin for anesthesia</td>
		</tr>
		<tr>
			<td>Sevoflurane 5%</td>
			<td>Abbvie</td>
			<td></td>
			<td>For anesthesia</td>
		</tr>
		<tr>
			<td>Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fresenius Vial pilot C</td>
			<td>Imexmed</td>
			<td></td>
			<td>Infusion pump</td>
		</tr>
		<tr>
			<td>Heated pad</td>
			<td>Harvard Apparatus</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Suction dominant 50</td>
			<td>Medela</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Suction tubing Optimus</td>
			<td>Promedical</td>
			<td>80342.2</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical motor</td>
			<td>Schick dental</td>
			<td>Qube</td>
			<td>Drilling of intramedullary holes</td>
		</tr>
		<tr>
			<td>Ventilation</td>
			<td>Maquet Servo1</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Material</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cylinders and caps</td>
			<td>Boutyplast</td>
			<td>Customized</td>
			<td>composition: PEEK (poly ether ether ketone)</td>
		</tr>
		<tr>
			<td>Manual self-retaining shaft</td>
			<td>GlobalD</td>
			<td>ACT1K</td>
			<td></td>
		</tr>
		<tr>
			<td>Mobile handle for self-retaining shaft</td>
			<td>GlobalD</td>
			<td>MTM</td>
			<td></td>
		</tr>
		<tr>
			<td>Self- drilling screws</td>
			<td>GlobalD</td>
			<td>VA1.2KL4</td>
			<td>cross-drive screws composed by Titanium grade5, ISO 5832-3</td>
		</tr>
		<tr>
			<td>Surgical tray</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Endotracheal tube Shiley diameter 2,5mm</td>
			<td>Covidien</td>
			<td>86233</td>
			<td>For intubation</td>
		</tr>
		<tr>
			<td>Endotracheal tube Shiley diameter 4,9mm</td>
			<td>Covidien</td>
			<td>107-35G</td>
			<td>For intubation</td>
		</tr>
		<tr>
			<td>Ethicon prolene 4-0</td>
			<td>Ehticon</td>
			<td>8581H</td>
			<td>Non-resorbable suture</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Marcel Blanc</td>
			<td>BD027R</td>
			<td>145 mm</td>
		</tr>
		<tr>
			<td>Intubation catheter</td>
			<td>Cook medical</td>
			<td></td>
			<td>Guide for intubation</td>
		</tr>
		<tr>
			<td>Round surgical burs</td>
			<td>Patterson</td>
			<td>78000</td>
			<td>0.8 mm in diameter, Drilling of intramedullary holes</td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>Swann-Morton</td>
			<td></td>
			<td>n&#176;10 and n&#176;15</td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>Marcel Blanc</td>
			<td>657</td>
			<td>180 mm</td>
		</tr>
		<tr>
			<td>Syringes Omnifix</td>
			<td>Braun</td>
			<td>4616057V</td>
			<td>5ml, 10ml and 50ml</td>
		</tr>
		<tr>
			<td>Venflon G22</td>
			<td>Braun</td>
			<td>42690985-01</td>
			<td>Vasofix safety for the ear iv line</td>
		</tr>
	</tbody>
</table>

generating a rabbit calvarial defect model: a surgical procedure to create round defects in the top portion of the rabbit skull to assess bone substitute materials for bone regeneration capacities

Source: Marger, L. et al., <a href="https://www.jove.com/v/59976">Calvarial Model of Bone Augmentation in Rabbit for Assessment of Bone Growth and Neovascularization in Bone Substitution Materials.</a> J. Vis. Exp. (2019).
In this video, we describe the surgical procedure to generate a rabbit calvarial model to evaluate the bone regeneration capacities of bone sample substitution materials of interest.

<img alt="Figure 1" src="/files/ftp_upload/20930/20930_Figure1.jpg" /> 
Figure 1: Specifications of PEEK cylinders. Two holes (0.8 mm in diameter) were drilled on the lateral stabilizing tabs for screwing. The positions of the 5 intramedullary holes (0.8 mm in diameter) to be drilled on the skull within the area delimited by the cylinder are marked using red circles.
<img alt="Figure 2" src="/files/ftp_uplo...

Generating a Rabbit Calvarial Defect Model: A Surgical Procedure to Create Round Defects in the Top Portion of the Rabbit Skull to Assess Bone Substitute Materials for Bone Regeneration Capacities

Source: Marger, L. et al., Calvarial Model of Bone Augmentation in Rabbit for Assessment of Bone Growth and Neovascularization in Bone Substitution ...

Bone substitutes comprise biocompatible materials implanted into bone defect sites to aid bone regeneration. The bone regenerative capabilities of bone substitutes can be assessed using calvarial defect models.
To generate a calvarial defect rabbit model, prepare an anesthetized rabbit in the prone position on a heating pad with a fixed probe to regulate its body temperature. Shave the scalp and sterilize the surgical site.
Incise along the sagittal line on the top portion of the skull. Incise the periosteum - a dense connective tissue layer - and elevate it to reveal the skull. Identify the sutures on the skull that outline the parietal and frontal bones. Place and secure customized biocompatible cylinders flat on each of the quadrants containing the bones bordered by the sutures, ensuring an airtight seal.
With saline irrigation, carefully drill intramedullary holes into the skull extending into the bone marrow cavity, within the region confined by the cylinders. Successful bleeding confirms penetration of the bone marrow space.&#160; Fill the cylinders fully with the test substitute material and close them. Close the surgical incision and allow the rabbit to recover.
The contact of the bone substitute in the defect site with the bone marrow can facilitate bone regeneration in the area. The established model can help evaluate the bone regenerative ability of bone substitutes.

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1.3K Views. Source: Marger, L. et al., Calvarial Model of Bone Augmentation in Rabbit for Assessment of Bone Growth and Neovascularization in Bone Substitution Materials. J. Vis. Exp. (2019). In this video, we describe the surgical procedure to generate a rabbit calvarial model to evaluate the bone regeneration capacities of bone sample substitution materials of interest. 

Video: Generating a Rabbit Calvarial Defect Model: A Surgical Procedure to Create Round Defects in the Top Portion of the Rabbit Skull to Assess Bone Substitute Materials for Bone Regeneration Capacities - Concept

Adult Zebrafish Injury Models to Study the Effects of Prednisolone in Regenerating Bone Tissue

Zebrafish are able to regenerate various organs, including appendages (fins) after amputation. This involves the regeneration of bone, which regrows within roughly two weeks after injury. Furthermore, zebrafish are able to heal bone rapidly after trepanation of the skull, and repair fractures that can be easily introduced into zebrafish bony fin rays. These injury assays represent feasible experimental paradigms to test the effect of administered drugs on rapidly forming bone. Here, we describe the use of these 3 injury models and their combined use with systemic glucocorticoid treatment, which exerts bone inhibitory and immunosuppressive effects. We provide a workflow on how to prepare for immunosuppressive treatment in adult zebrafish, illustrate how to perform fin amputation, trepanation of calvarial bones, and fin fractures, and describe how the use of glucocorticoids affects both bone forming osteoblasts and cells of the monocyte/macrophage lineage as part of innate immunity in bone tissue.

Here, we describe 3 adult zebrafish injury models and their combined use with immunosuppressive drug treatment. We provide guidance on imaging of regenerating tissues and on detecting bone mineralization therein.

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Adult Mouse Digit Amputation and Regeneration: A Simple Model to Investigate Mammalian Blastema Formation and Intramembranous Ossification

Here, we present a protocol of adult mouse distal terminal phalanx (P3) amputation, a procedurally simple and reproducible mammalian model of epimorphic regeneration, which involves blastema formation and intramembranous ossification analyzed by fluorescence immunohistochemistry and sequential in-vivo microcomputed tomography (&#956;CT). Mammalian regeneration is restricted to amputations transecting the distal region of the terminal phalanx (P3); digits amputated at more proximal levels fail to regenerate and undergo fibrotic healing and scar formation. The regeneration response is mediated by the formation of a proliferative blastema, followed by bone regeneration via intramembranous ossification to restore the amputated skeletal length. P3 amputation is a preclinical model to investigate epimorphic regeneration in mammals, and is a powerful tool for the design of therapeutic strategies to replace fibrotic healing with a successful regenerative response. Our protocol uses fluorescence immunohistochemistry to 1) identify early-and-late blastema cell populations, 2) study revascularization in the context of regeneration, and 3) investigate intramembranous ossification without the need for complex bone stabilization devices. We also demonstrate the use of sequential in vivo &#956;CT to create high resolution images to examine morphological changes after amputation, as well as quantify volume and length changes in the same digit over the course of regeneration. We believe this protocol offers tremendous utility to investigate both epimorphic and tissue regenerative responses in mammals.

Here, we present a protocol of adult mouse terminal phalanx amputation to investigate mammalian blastema formation and intramembranous ossification, analyzed by fluorescent immunohistochemistry and sequential in-vivo microcomputed tomography.

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Construction and Evaluation of a Murine Calvarial Osteolysis Model by Exposure to CoCrMo Particles in Aseptic Loosening

Wear particle-induced osteolysis is a major cause of aseptic loosening in arthroplasty failure, but the underlying mechanism remains unclear. Due to long follow-ups necessary for detection and sporadic occurrence, it is challenging to assess the pathogenesis ofparticle-induced osteolysis in clinical cases. Hence, optimal animal models are required for further studies. The murine model of calvarial osteolysis established by exposure to CoCrMo particles is an effective and valid tool for assessing the interactions between particles and various cells in aseptic loosening. In this model, CoCrMo particles were first obtained by high-vacuum three-electrode direct current and resuspended in phosphate-buffered saline at a concentration of 50 mg/mL. Then, 50 &#181;L of the resulting suspension was applied to the middle of the murine calvaria after separation of the cranial periosteum by sharp dissection. After two weeks, the mice were sacrificed, and calvaria specimens were harvested; qualitative and quantitative evaluations were performed by hematoxylin and eosin staining and micro computed tomography. The strengths of this model include procedure simplicity, quantitative evaluation of bone loss, rapidity of osteolysis development, potential use transgenic or knockout models, and a relatively low cost. However, this model cannot to be used to assess the mechanical force and chronic effects of particles in aseptic loosening. Murine calvarial osteolysis model generated by exposure to CoCrMo particles is an ideal tool for assessing the interactions between wear particles and various cells, e.g., macrophages, fibroblasts, osteoblasts and osteoclasts, in aseptic loosening.

This manuscript describes a murine calvarial osteolysis model by exposure to CoCrMo particles, which constitutes an ideal animal model for assessing the interactions between wear particles and various cells in aseptic loosening.

Wear particle-induced osteolysis is a major cause of aseptic loosening in arthroplasty failure, but the underlying mechanism remains unclear. Due to long ...

Generating a Rabbit Calvarial Defect Model: A Surgical Procedure to Create Round Defects in the Top Portion of the Rabbit Skull to Assess Bone Substitute Materials for Bone Regeneration Capacities

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