We thank the staff and veterinarians at the Scott &#38; White Hospital Department of Comparative Medicine, Temple, Texas, for their invaluable advice and assistance during the development of this technique. This work was funded in part by The Institute for Regenerative Medicine Program Funds, Scott &#38; White RGP grant #90172, NIH 2P40RR017447-07 and NIH R01AR066033-01 (NIAMS). We thank Dr Suzanne Zeitouni for proofing the manuscript.

NOTE: This protocol is designed for female nude (Nu/J) mice (18-25 g, 6 weeks) acquired from Jackson Laboratories. Since strains of mice vary slightly in terms of anatomy and growth rate, we advise that the fabrication of pins is optimized to the strain, gender and age of the recipients prior to implantation into live subjects. If the strains are carefully matched, the interference fit between the pin and marrow cavity is highly reproducible. Procedures for housing, diet and general animal husbandry are beyond the scope of this protocol, but all mice were housed in accordance with the Guide for the Care and Use of Laboratory Animals (8th Edition) and institutional policies set by the Institutional Animal Care and Use Committee (IACUC) and Department of Comparative Medicine (DCM) at Scott and White Hospital.
1. Fabrication of Pins
<img alt="Figure 2" src="/files/ftp_upload/52368/52368fig2highres.jpg" /> 
Figure 2: Pin assembly. (A) Photographs of the surgical steel tubing at various stages of assembly and a 4 mm diameter cylinder cut from 5 mm thick Gelfoam sheet. (B) After polishing the assembled pin (above), the Gelfoam cylinder is trimmed, positioned over the steel collar (middle), then autoclaved, resulting in a sterile pin coated with dried Gelfoam. This improves cell attachment at the site of injury and maintains the direction of healing (below). (C) An excised femur fitted with the medullary pin. (D) Examples of pin assemblies at various thicknesses and lengths demonstrating the flexibility of this approach. (E) &#181;CT reconstruction of a pin-stabilized femoral defect illustrating the interaction of the medullary pin with the trabecular bone in the proximal and distal ends of the femur.
<ol>
	<li>Cut a 9 mm length of 22 G stainless hypodermic tubing, and a 3 mm length of 19 G tubing using a 23.8 mm diameter fiberglass reinforced heavy duty cut-off wheel fitted to a rotary tool. Use small fine-tipped pliers to immobilize the tubing (Figure 2A, above). Wear appropriate eye protection and handle rotary tool with care.</li>
	<li>Place a small amount (approximately 10 &#181;l) of cyanoacrylate adhesive to the middle of the 9 mm shaft. Pass the 3 mm collar over the 9 mm shaft until in the center and twist to evenly distribute the glue between the collar and shaft. Measure dimensions with a pair of digital calipers and compare with Figure 1. Allow to set for at least 15 hr (Figure 2A, center).</li>
	<li>Use a 220 grit emery-impregnated disc to remove burrs and excessive glue.</li>
	<li>Using the rotary tool, polish the pin using a felt polishing disc.</li>
	<li>Rinse with deionized water and dry with compressed air.</li>
	<li>Test integrity by placing a 19 G hypodermic blunt needle over the shaft of the newly made pin and push against the collar. Ensure that the collar can resist approximately 25 g of weight.</li>
	<li>Rinse the pin in sterile-filtered phosphate buffered saline (Figure 2B, top). Ensure that the shaft of the pin fits snugly into the femoral medullary cavity with the collar flush against the edges of the cut bone (Figure 2C). Prototype various configurations for the specific purposes of the study and the recipient (Figure 2D). Make sure that the ends of the pin penetrate into the trabecular bone of the diaphysis so as to maximize fixation and improve the interference fit (Figure 2E). 
	NOTE: It is recommended that the fit is tested on bone specimens before implantation in live mice (e.g. Figure 2C).</li>
	<li>Use a 4 mm diameter punch-biopsy cutter to cut a cylinder from a 5 mm thick sheet of surgical gelatin sponge. Use a scalpel to trim the cylinder to 3 mm length and pass a 20 G hypodermic needle along the length of the cylinder to generate a hole (Figure 2A, below). 
	NOTE: We have found that the positioning of a gelatin or collagen scaffold at the site of the defect improves cell retention and induces longitudinal growth along the axis of the bone.</li>
	<li>Pass the gelatin foam cylinder over the pin and align with the collar (Figure 2B, center) and autoclave on a dry cycle at 120 &#176;C at 15 psi. The gelatin foam will dry and darken in color (Figure 2B, below).</li>
</ol>
2. Surgical Technique
NOTE: The following procedure is written observing all guidelines set by the Guide for the Care and Use of Laboratory Animals (8th Edition). Please observe all additional policies set by the local IACUC and ensure an IACUC-approved animal use protocol (or equivalent) is in place before proceeding with the following sections.
<img alt="Figure 3" src="/files/ftp_upload/52368/52368fig3highres.jpg" /> 
Figure 3: Surgical instruments. (A) Surgical kit. Micro-drill (1), cutting wheel (2), absorbable suture (3), outer suture (4), scalpel blades (5), scalpel handle (6), sterile medullary pins wrapped in foil (7), medullary depth gauge made form hypodermic tubing (8), fine nosed forceps (9), rat-teeth forceps (10), blunt reaming needles (11), needle drivers (12), small hemostats (13), fine scissors (14), periosteal elevator (15), modified Kern forceps (16). (B): Modifications required to produce Kern-style bone-grips for mice.
<ol>
	<li>Select a surgical procedure room or clean laboratory with a closable door and no through-traffic. Sanitize a suitable work surface with a clinical grade quaternary-based disinfectant cleaner.</li>
	<li>While wearing disposable sterile gloves and robe, set up a sterile field of approximately 60 x 90 cm2 on the sanitized work surface with autoclaved cloth drapes. 
	NOTE: It is advisable to have an assistant remove the outer packaging and present the sterile materials to the investigator setting up the field.</li>
	<li>Place a sanitized heating pad fitted to a warm water re-circulator on the drape and cover with sterile disposable drapes. Warm a bead sterilizer to operating temperature (250-265 &#176;C, this takes up to 30 min).</li>
	<li>On the sterile field, arrange the surgical kit (Figure 3A) to provide convenient access to all components. Also, provide sterile cotton gauze (2 x 2 inch2), sterile Q-tips, a sterile steel bowl (500 ml) containing sterile saline (0.9% w/v), and chlorhexidine/isopropyl alcohol surgical disinfectant applicators.</li>
	<li>Assemble a small animal anesthesia unit next to the sterile field with an induction chamber and nose-cone assembly in accordance with veterinary guidance and the user manual. Use clinical grade oxygen as the gas supply and isoflurane, USP.</li>
</ol>
<img alt="Figure 4" src="/files/ftp_upload/52368/52368fig4highres.jpg" /> 
Figure 4: Surgical procedure. (A-C) Approach and exposure of the femur. (D-E) Elevation and cuts. (F-K) Reaming and sizing of medullary canals.
<img alt="Figure 5" src="/files/ftp_upload/52368/52368fig5highres.jpg" /> 
Figure 5: Surgical procedure. (A-B) Installation of pin. (C-D) Closure. (E) Typical image of properly positioned pin 24 hr after surgery.
<ol start="6">
	<li>Place a mouse in the induction chamber of the anesthesia system and set the output to 2 L min-1 O2 and the isoflurane concentration to 3% (v/v).
	<ol>
		<li>Check that the mouse is unconscious within 1 min; raise the concentration to 4% (v/v) if necessary. Remove the mouse, place on sterile field with the heating pad positioned underneath&#160;&#160;and apply the nose cone. Transfer flow to the cone, and reduce the isoflurane to 2.5%, wait for a further 20 sec, and test for adequate surgical plane in accordance with institutional policy. Typically, lack of a hind limb reflex when gently squeezed is adequate for determining adequate anesthesia.</li>
		<li>During the entire process, have an assistant monitor for strong regular breathing and pink coloration of the extremities and mouth region to ensure an appropriate level of oxygenation while unconscious. Adjust anesthesia in accordance with veterinary guidance and institutional policy if necessary. Apply sterile artificial tears lubricant ointment (15% (v/v) mineral oil, 83% (v/v) white petrolatum) to the eyes.</li>
	</ol>
	</li>
	<li>Turn the mouse onto one side with the hind-limb facing upwards on a new disposable drape. If necessary, remove fur with an electric razor or hair removal cream. Wipe the site with sterile saline and remove the additional drape with excess fur. Place a new fenestrated drape over the mouse so as to cover all parts but the entire hind-limb (Figure 4A) but maintain view of the face and paw to monitor coloration and breathing.</li>
	<li>Locate the proximal and distal ends of the femur and incise the skin for 5-10 mm along the longitudinal axis (Figure 4B). Separate the skin layer from the fascia with a #15 scalpel, exposing a lateral approach to the femur via the biceps femoris and vastus lateralis. Locate where the septa of both muscles meet (it is a line of white tissue against the pink coloration of the muscle). With a scalpel, carefully dissect along the intermuscular boundary until the femur is visible (Figure 4B).</li>
	<li>Develop the incision with a blunt periosteal elevator so as to expose the entire diaphysis (Figure 4C). Use the elevator to further expose the central two thirds of the femur while taking care to preserve the posterior neurovascular bundle on the medial side (Figure 4D). Gently scrape soft tissue off of the bone with a scalpel, and dry with a sterile Q-tip or equivalent.</li>
	<li>Locate the center of the femur with calipers if necessary and mark with a sterile scalpel or marker, then mark 1.5 mm proximally and distally from the center. Gently grasp the femur using a pair of fine-nosed forceps previously fashioned in the Kern style to prevent excessive pressure on the bone. 
	NOTE: Exact dimensions for modifications are provided in Figure 3B. It is advisable to test the forceps on a euthanized specimen prior to use to ensure the bone will not break under pressure required for immobilization during cutting.</li>
	<li>Using a fine drill fitted with a fine diamond-grit coated cutting wheel (8 mm diameter x 0.1 mm width), make the first cut with the elevator in place to protect tissue below (Figure 4E). Raising the cut femur to 45&#176; while firmly holding the extremity of the diaphysis, make the second cut, removing a 3 mm segment. 
	NOTE: Face protection is recommended at this stage.</li>
	<li>With the bone immobilized by forceps, carefully ream the medullary cavities of each end with a blunt 23 G hypodermic needle. Using a pre-made depth-gauge made from a length of 22 G tubing placed in 19 G tubing (Figure 3A, item 8), assess the depth of the reamed medullary cavity and ensure it is 3 mm (Figure 4F-K). If the medullary cavity resists the 22 G depth gauge, ream again with a blunt 22 G hypodermic needle.</li>
	<li>Carefully insert the medullary pin into the proximal then distal medullary cavities to bring the femur back out to its original length and establish a stable 3 mm gap (Figure 5A, B). If needed, apply a small amount of manual stress to achieve a good interference fit of the rod with the cortical bone of the femur. Ensure that the pin fits snugly into the medullary cavities of both sides and the edges of the cut bone are flush with the collar. If there is a gap, ream once again with a 22 G blunt needle.</li>
	<li>Reposition the muscle and peripheral tissue over the pin and close with a continuous absorbable 5-0 suture (Figure 5C). Close skin incision with 5-7 square knot nylon 5-0 sutures and seal with surgical adhesive (Figure 5D).</li>
</ol>
3. Postoperative Procedures
<ol>
	<li>After closing the skin incision, withdraw anesthesia, but allow O2 to remain flowing until the mouse begins to move; this should take less than 1 min. If the mouse remains motionless after 5 min of O2 administration, refer to institutional policies on veterinary intervention.</li>
	<li>When the mouse begins to move, gently transfer it to a cage preferably containing dry, autoclaved bedding.
	<ol>
		<li>Place an igloo-type nest in each cage and the mouse will retreat into it, reducing movement. Check that the mouse recovers hind-limb mobility 5-10 min after recovery of consciousness.</li>
		<li>Perform daily postoperative monitoring in accordance with institutional policies. Provide gelatinized hydration and food on the floor of the cage for the first 5-7 days and administer analgesia and postoperative monitoring in accordance with institutionally approved policies. 
		NOTE: We administer 0.05-0.1 mg kg-1 buprenorphine24 with 0.25 ml saline subcutaneously, twice daily for the first 3 days and thereafter if the mouse does not regain normal utility.</li>
	</ol>
	</li>
	<li>After the first 24 hr of recovery, perform live-animal X-ray imaging under anesthesia to visualize pin placement (Figure 4E). If the pin is dislocated, consider immediate revision surgery.</li>
	<li>At day 7 post-surgery, remove sutures under anesthesia and house in groups in accordance with institutional animal husbandry policies.</li>
	<li>After 2-5 weeks, humanely euthanize animals in accordance with the American Veterinary Medical Association approved methods for euthanasia25. 
	NOTE: An intraperitoneal injection of commercially available barbiturate-combination euthanasia cocktail is effective and humane, but local institutional policies on euthanasia should be followed.</li>
	<li>Carefully dissect away the hind-limb by exposing the proximal femur and the pelvis from the medial side. Gently press the joint inwards from the lateral side of the limb while detaching the femoral head from the acetabulum (the hip socket) with a scalpel. Cut remaining muscle and skin with a sharp scalpel or micro-scissors, releasing the entire limb form the pelvis. With a sharp pair of rongeurs or heavy scissors, cut the lower hind-limb (tibia/fibula) approximately 5 mm below the knee joint. 
	NOTE: It is recommended that all specimens are stored in an identical manner prior to analysis.</li>
	<li>Remove skin, but leave muscle in place. Fix the tissue in 10% buffered formalin supplemented with 10 mM CaCl2 fixative for 1 week followed by storage in phosphate buffered saline supplemented with 10 mM CaCl2 for up to 1 month prior to imaging. Perform fixation and storage at 4 &#176;C. Alternatively, scan specimens immediately without fixation.</li>
</ol>
4. Analysis of Specimens
NOTE: Bone healing can be assessed by a wide variety of methodologies that are beyond the scope of this protocol. The following is a method that we have successfully employed using a specimen microCT (&#181;CT) imager. It is recommended that the following parameters are tested initially, then optimized for the specific needs of the project.
<ol>
	<li>Wrap specimen in a thin layer of plastic sealing film and position it vertically in the instrument chamber. Make sure that the femur is perpendicular to the sample stage during the entire scan. Set the scan to the following parameters; voltage = 29 kV, current = 661 &#181;A, power = 19 W, image pixel size (&#956;m) = 21.00; 360 degree rotation = yes; frame averaging = on (5); rotation step (deg) = 1.00, random movement = on. Save images as JPEG files in a single destination folder per scan (e.g. Figure 6A). 
	NOTE: The scan should be complete in 57 min.</li>
	<li>Utilize the reconstruction software to generate axial images based on the following parameters; smoothing = on (4), misalignment compensation = on, ring artifact reduction = on (5), beam-hardening correction (40%), CS rotation (deg) = 0.00. Set output to 2,000-15,000 Hounsfield units. Save images as JPEG files in a single destination folder per scan.</li>
	<li>Using the axial images and analytical software, define the region of interest (ROI) by first setting the proximal and distal edges of the original defect. Achieve this by selecting the sections that encompass the collar only. Because the collar is thicker than the medullary pin, these sections are easily defined (Figure 6B, left). Exclude the collar from calculations by drawing an exclusion zone around it (with a 100 &#181;m margin) and transferring the zone to each of the sections in the ROI (Figure 6B, right). 
	NOTE: Polar moments of inertia, 3D reconstructions and calculations such as the volume of new bone (Figure 6C, D) can readily be achieved using the software.</li>
</ol>

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S., Yao, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Overview:+animal+models+of+osteopenia+and+osteoporosis.">Overview: animal models of osteopenia and osteoporosis.</a> J Musculoskelet Neuronal Interact. 1, 193-207 (2001).</li></ol>

The authors have no competing financial interests.

Herein, we describe a simple method to generate a critical-sized pin-stabilized defect of the murine femur using standard laboratory and veterinary equipment. While the assembly of the pins and the surgical procedure itself requires practice, it is well within the capabilities of a well-trained biomedical research scientist or veterinarian.
The pin is positioned into the medullary canal without additional fixation, making the procedure technically more feasible than more complicated approaches...

It is estimated that half of the US population experience a fracture by the age of 651. For those patients with fractures treated surgically, 500,000 procedures involve the use of a bone graft2 and this number is expected to rise with an increasingly aging population3. Although bone is one of the few organs that has the capacity to completely heal without scarring, there are instances where the process fails3,4. Depending on the circumstances and quality of treatment, 2-30% of long bone fractures fail, resulting in non-union3,5. While there remains some debate on the definition, pseudoarthrosis, critical-sized or non-union bone injuries generally refers to an injury that does not heal over the natural lifetime of the subject6. For experimental purposes, this duration is shortened to the average time required for complete healing of an average-sized bone injury. Non-union bone lesions occur for numerous reasons, but major factors include extreme trauma resulting in a critically-sized gap, infection, poor angiogenesis, tobacco use, or inhibited osteoregenerative capacity due to disease or age7. Even if non-unions are successfully treated, it can cost in excess of $60,000 per procedure, depending on the type of injury and the approaches employed8.
In moderate cases, autologous bone grafting is employed. This strategy involves recovery of bone from a donor site and implantation at the site of injury. While this approach is extremely effective, the volume of available donor-derived bone is limited and the procedure involves an additional surgery, which results in persistent pain in many patients9,10. In addition, the efficacy of the autologous bone graft is dependent of the health of the patient. Bone substitutes made from synthetic materials or processed cadaveric bone are abundantly available11-13, but they have significant limitations, including poor host-cell adhesion properties, reduced osteoconductivity, and the potential for immune rejection14. There is therefore an urgent need for bone regeneration technologies that are safe, effective and widely available.
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. Although it can vary with species15, this is achieved by complete removal of a bone segment that is larger than 1.5 times the diameter of the bone cross-section16. 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.
For human cell-based applications, immune-compromised mice are attractive to work with because they are physiologically well-characterized, easy to house, cost effective, and easily analyzed radiologically and histologically. Of paramount importance is that immune-compromised mice do not reject cells from different species including humans. Their small size also permits the testing of very small numbers of cells or volumes of experimental scaffolds in orthopedic applications. Several murine orthopedic models have been reported that afford various degrees of bone stability17,18. Those systems that result in very high levels of stability, such as external fixators and locking plates predominantly heal by intramembranous ossification although endochondral healing has been reported19. In contrast, those that permit some micro- and/or macro-motion, such as those employing unfixed or partially-fixed medullary pins, generally heal with a predominance of endochondral ossification20,21. Delayed union or non-union defects of long bone are particularly difficult to achieve in mice due to the extra level of stabilization required. However, a number of approaches have been reported, including medullary pins with interlocking nails, locking plates and external fixators22. These systems generally work well, but given their complicated design they can be technically challenging to install. For example, Garcia et al.23 devised an elegant interlocking pin system for use in mice, but the procedure involves incisions at two separate sites and extensive modification of the femur to accommodate the pins. These procedures were performed under a dissecting microscope.
Herein, we describe a simple femoral medullary pin with a central collar designed to prevent closing of a 3 mm bone deficit and also delineate the original edges of the defect. While the pin was not fixed to the bone itself, precise sizing of the pin diameter and reaming of the medullary cavity results in sufficient interference to minimize torsional motion (Figure 1). With careful selection of inbred age, gender and strain-matched mice, the result is a highly reproducible hypertrophic non-uniondefect22 which can be easily evaluated radiologically. Moreover regions of interest can be reproducibly defined after micro-computed tomography (&#181;CT) for measurement of de novo bone formation and histomorphological parameters. The pins were prototyped in our laboratory using readily available tools.
<img alt="Figure 1" src="/files/ftp_upload/52368/52368fig1highres.jpg" /> 
Figure 1: Experimental principle. Diagrammatic summary of the segmental defect model. The central 3 mm segment of a 9-10 mm murine femur is excised surgically (left). A 3 mm long, 19 gauge surgical steel tube is passed over a 9 mm long, 22 G stainless steel tube and fixed with adhesive at the exact center (right). The resultant pin is fitted into the medullary canals of the remaining proximal and distal portions of the femur with the 19 G collar replacing the 3 mm segment of bone (below, center).

<table border="0" cellpadding="0" cellspacing="0" width="799">
	<tbody>
		<tr height="20">
			<td height="20" style="height:20px;width:235px;">Name of Equipment/Material*</td>
			<td style="width:207px;">Company</td>
			<td style="width:180px;">Catalog or model</td>
			<td style="width:179px;">Notes</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Pin Assembly</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Dremel rotary tool</td>
			<td>Dremel</td>
			<td>8220</td>
			<td>or equivalent</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Heavy duty cut off wheel</td>
			<td>Dremel</td>
			<td>420</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Surgical tubing 19G</td>
			<td>Small Parts (Amazon)</td>
			<td>B000FMZ8LY</td>
			<td>OD 1.07mm, ID 0.889mm</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Surgical tubing 21G</td>
			<td>Small Parts (Amazon)</td>
			<td>B000FMZ8YQ</td>
			<td>OD 0.82mm, ID 0.635mm</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Surgical tubing 22G</td>
			<td>Small Parts (Amazon)</td>
			<td>B000FMYLZS</td>
			<td>OD 0.719mm, ID 0.502mm</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Surgical tubing 23G</td>
			<td>Small Parts (Amazon)</td>
			<td>B000FN0SY0</td>
			<td>OD 0.643mm, ID 0.444mm</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Cyanoacrylate adhesive</td>
			<td>Loctite</td>
			<td>1365882</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Emery disc</td>
			<td>Dremel</td>
			<td>413</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Rubber polishing point</td>
			<td>Dremel</td>
			<td>462</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Felt polishing disc</td>
			<td>Dremel</td>
			<td>414</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Gelatin sponge</td>
			<td>Surgifoam/Ethicon</td>
			<td>1974</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Punch biopsy cutter</td>
			<td>Miltex</td>
			<td>33-34</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Surgery/post-operative</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Warm pad and circulator pump</td>
			<td>Stryker/Thermocare</td>
			<td>TP700, TP700C, TPP722</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Coverage quaternary spray</td>
			<td>Steris</td>
			<td>1429-77</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Bead sterilizer</td>
			<td>Germinator/CellPoint Scentific</td>
			<td>Germinator 500</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Anesthesia system</td>
			<td>VetEquip Inc</td>
			<td>901806 or 901807/901809</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Isofluorane anesthetic</td>
			<td>VETone/MWI</td>
			<td>501017, 502017</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Surgical disinfectant</td>
			<td>Chloraprep/CareFusion</td>
			<td>260449</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Surgical tools</td>
			<td>Fine Science Tools</td>
			<td>various</td>
			<td>recommend German made</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Face protection</td>
			<td>Splash Shield</td>
			<td>4505</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Rechargable high speed drill</td>
			<td>Fine Science Tools</td>
			<td>18000-17</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Diamond cutting wheel</td>
			<td>Strauss Diaiond</td>
			<td>361.514.080HP</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Absorbable sutures&#160;</td>
			<td>Covidien</td>
			<td>UM-213</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Outer sutures</td>
			<td>Ethicon</td>
			<td>668G</td>
			<td>or equivalent</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Vetbond</td>
			<td>3M</td>
			<td>1469SB</td>
			<td>or equivalent</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Hydration gel</td>
			<td>Clear H2O</td>
			<td>70-01-1082</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Diet gel</td>
			<td>Clear H2O</td>
			<td>72-01-1062</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Buprenorphine</td>
			<td>Reckitt and Benckser</td>
			<td>12496-0757-01</td>
			<td>controlled substance</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Mouse igloos</td>
			<td>Bio Serv</td>
			<td>K3328, 3570,3327</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Euthanasia cocktail</td>
			<td>Euthasol/Virbac</td>
			<td>710101</td>
			<td>controlled substance</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Analysis</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Live animal imager&#160;</td>
			<td>Orthoscan</td>
			<td>FD Pulse</td>
			<td>or equivalent</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Micro-CT unit and software</td>
			<td>Bruker</td>
			<td>Skyscan1174</td>
			<td>or equivalent</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Sealing film/Parafilm M</td>
			<td>VWR or Fisher</td>
			<td>100501-338, S37441</td>
			<td></td>
		</tr>
		<tr height="20">
			<td colspan="4" height="20" style="height:20px;">*Generic sources are suitable for all other items such as gause, drapes, protective clothing, animal care equipment.</td>
		</tr>
	</tbody>
</table>

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.

Mice typically recover consciousness and hind-limb mobility 5-10 minutes after withdrawal of anesthesia. During the first 5 days, it is advisable to house mice individually and introduce environmental enrichment to prevent excess use of the limb. For this purpose, igloo-type nests reduce the need for nest building and encourage resting. We have also observed that provision of food and hydrogel on the floor of the cage reduces the probability of pin displacement. During the 5-day period, analgesia should be administered a...

Watch this Scientific Journal Video about A Simple Critical-sized Femoral Defect Model in Mice at JoVE.com

A Simple Critical-sized Femoral Defect Model in Mice

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 ...

a-simple-critical-sized-femoral-defect-model-in-mice

Research

JoVE Journal

Medicine

15.0K Views.  Texas A&M Health Science Center. 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. 

Video: A Simple Critical-sized Femoral Defect Model in Mice

A Simple Guide Screw Method for Intracranial Xenograft Studies in Mice

The grafting of human tumor cells into the brain of immunosuppressed mice is an established method for the study of brain cancers including glioblastoma (glioma) and medulloblastoma. The widely used stereotactic approach only allows for the injection of a single animal at a time, is labor intensive and requires highly specialized equipment. The guide screw method, initially developed by Lal et al.,1 was developed to eliminate cumbersome stereotactic procedures. We now describe a modified guide screw approach that is rapid and exceptionally safe; both of which are critical ethical considerations. Notably, our procedure now incorporates an infusion pump that allows up to 10 animals to be simultaneously injected with tumor cells.
To demonstrate the utility of this procedure, we established human U87MG glioma cells as intracranial xenografts in mice, which were then treated with AMG102; a fully human antibody directed to HGF/scatter factor currently undergoing clinical evaluation2-5. Systemic injection of AMG102 significantly prolonged the survival of all mice with intracranial U87MG xenografts and resulted in a number of complete cures. 
This study demonstrates that the guide screw method is an inexpensive, highly reproducible approach for establishing intracranial xenografts. Furthermore, it provides a relevant physiological model for validating novel therapeutic strategies for the treatment of brain cancers.

In order to evaluate novel therapeutic paradigms for the treatment of glioma, physiological relevant models are essential. We utilize an implantable guide screw procedure for establishment of intracranial xenograft models that is more rapid and safer than stereotactic approaches.

The grafting of human tumor cells into the brain of immunosuppressed mice is an established method for the study of brain cancers including glioblastoma ...

Creating Rigidly Stabilized Fractures for Assessing Intramembranous Ossification, Distraction Osteogenesis, or Healing of Critical Sized Defects

Assessing modes of skeletal repair is essential for developing therapies to be used clinically to treat fractures. Mechanical stability plays a large role in healing of bone injuries. In the worst-case scenario mechanical instability can lead to delayed or non-union in humans. However, motion can also stimulate the healing process. In fractures that have motion cartilage forms to stabilize the fracture bone ends, and this cartilage is gradually replaced by bone through recapitulation of the developmental process of endochondral ossification. In contrast, if a bone fracture is rigidly stabilized bone forms directly via intramembranous ossification. Clinically, both endochondral and intramembranous ossification occur simultaneously. To effectively replicate this process investigators insert a pin into the medullary canal of the fractured bone as described by Bonnarens4. This experimental method provides excellent lateral stability while allowing rotational instability to persist. However, our understanding of the mechanisms that regulate these two distinct processes can also be enhanced by experimentally isolating each of these processes. We have developed a stabilization protocol that provides rotational and lateral stabilization. In this model, intramembranous ossification is the only mode of healing that is observed, and healing parameters can be compared among different strains of genetically modified mice 5-7, after application of bioactive molecules 8,9, after altering physiological parameters of healing 10, after modifying the amount or time of stabilization 11, after distraction osteogenesis 12, after creation of a non-union 13, or after creation of a critical sized defect. Here, we illustrate how to apply the modified Ilizarov fixators for studying tibial fracture healing and distraction osteogenesis in mice.

This article describes a method for stabilizing long bone fractures that is based on the application of modified Ilizarov external fixators 1-3. After application of the fixators and creation of the bone injury, healing can be assessed, distraction osteogenesis can be performed, or non-union or critical sized defect can be created and used to study therapeutic interventions.

Assessing modes of skeletal repair is essential for developing therapies to be used clinically to treat fractures. Mechanical stability plays a large role ...

Repair of a Critical-sized Calvarial Defect Model Using Adipose-derived Stromal Cells Harvested from Lipoaspirate

Craniofacial skeletal repair and regeneration offers the promise of de novo tissue formation through a cell-based approach utilizing stem cells. Adipose-derived stromal cells (ASCs) have proven to be an abundant source of multipotent stem cells capable of undergoing osteogenic, chondrogenic, adipogenic, and myogenic differentiation. Many studies have explored the osteogenic potential of these cells in vivo with the use of various scaffolding biomaterials for cellular delivery. It has been demonstrated that by utilizing an osteoconductive, hydroxyapatite-coated poly(lactic-co-glycolic acid) (HA-PLGA) scaffold seeded with ASCs, a critical-sized calvarial defect, a defect that is defined by its inability to undergo spontaneous healing over the lifetime of the animal, can be effectively show robust osseous regeneration. This in vivo model demonstrates the basis of translational approaches aimed to regenerate the bone tissue - the cellular component and biological matrix. This method serves as a model for the ultimate clinical application of a progenitor cell towards the repair of a specific tissue defect.

This protocol describes the isolation of adipose-derived stromal cells from lipoaspirate and the creation of a 4 mm critical-sized calvarial defect to evaluate skeletal regeneration.

Craniofacial skeletal repair and regeneration offers the promise of de novo tissue formation through a cell-based approach utilizing stem cells. ...

A Simple Method of Mouse Lung Intubation

A simple procedure to intubate mice for pulmonary function measurements would have several advantages in longitudinal studies with limited numbers or expensive animal. One of the reasons that this is not done more routinely is that it is relatively difficult, despite there being several published studies that describe ways to achieve it. In this paper we demonstrate a procedure that eliminates one of the major hurdles associated with this intubation, that of visualizing the trachea during the entire time of intubation. The approach uses a 0.5 mm fiberoptic light source that serves as an introducer to direct the intubation cannula into the mouse trachea. We show that it is possible to use this procedure to measure lung mechanics in individual mice over a time course of at least several weeks. The technique can be set up with relatively little expense and expertise, and it can be routinely accomplished with relatively little training. This should make it possible for any laboratory to routinely carry out this intubation, thereby allowing longitudinal studies in individual mice, thereby minimizing the number of mice needed and increasing the statistical power by using each mouse as its own control.

This paper describes a striaghforward and efficient method of intubating mice for pulmonary function measurements or pulmonary instillation, that allows the mice to recover and be studied at later times. The procedure involves an inexpensive fiberoptic light source that directly illuminates the trachea.

A  simple procedure to intubate mice for pulmonary function measurements would  have several advantages in longitudinal studies with limited numbers or  ...

Femoral Bone Marrow Aspiration in Live Mice

Serial sampling of the cellular composition of bone marrow (BM) is a routine procedure critical to clinical hematology. This protocol describes a detailed step-by-step technical procedure for an analogous procedure in live mice which allows for serial characterization of cells present in the BM. This procedure facilitates studies aimed to detect the presence of exogenously administered cells within the BM of mice as would be done in xenograft studies for instance. Moreover, this procedure allows for the retrieval and characterization of cells enriched in the BM such as hematopoietic stem and progenitor cells (HSPCs) without sacrifice of mice. Given that the cellular composition of peripheral blood is not necessarily reflective of proportions and types of stem and progenitor cells present in the marrow, procedures which provide access to this compartment without requiring termination of the mice are very helpful. The use of femoral bone marrow aspiration is illustrated here for cytological analysis of marrow cells, flow cytometric characterization of the hematopoietic stem/progenitor compartment, and culture of sorted HSPCs obtained by femoral BM aspiration compared with conventional marrow harvest.

This protocol describes a procedure for serial sampling of femoral bone marrow (BM) without requiring the sacrifice of mice. This procedure facilitates longitudinal studies of the BM composition of mice over time and provides serial access to cells within the BM for ex vivo and transplantation studies.

Serial sampling of the cellular composition of bone marrow (BM) is a routine procedure critical to clinical hematology. This protocol describes a detailed ...

A Murine Model of Arterial Restenosis: Technical Aspects of Femoral Wire Injury

Cardiovascular disease caused by atherosclerosis is the leading cause of death in the developed world. Narrowing of the vessel lumen, due to atherosclerotic plaque development or the rupturing of established plaques, interrupts normal blood flow leading to various morbidities such as myocardial infarction and stroke. In the clinic endovascular procedures such as angioplasty are commonly performed to reopen the lumen. However, these treatments inevitably damage the vessel wall as well as the vascular endothelium, triggering an excessive healing response and the development of a neointimal plaque that extends into the lumen causing vessel restenosis (re-narrowing). Restenosis remains a major cause of failure of endovascular treatments for atherosclerosis. Thus, preclinical animal models of restenosis are vitally important for investigating the pathophysiological mechanisms as well as translational approaches to vascular interventions. Among several murine experimental models, femoral artery wire injury is widely accepted as the most suitable for studies of post-angioplasty restenosis because it closely resembles the angioplasty procedure that injures both endothelium and vessel wall. However, many researchers have difficulty utilizing this model due to its high degree of technical difficulty. This is primarily because a metal wire needs to be inserted into the femoral artery, which is approximately three times thinner than the wire, to generate sufficient injury to induce prominent neointima. Here, we describe the essential surgical details to effectively overcome the major technical difficulties of this model. By following the presented procedures, performing the mouse femoral artery wire injury becomes easier. Once familiarized, the whole procedure can be completed within 20 min.

The mouse femoral artery wire injury model of restenosis is technically challenging. In this protocol we show the key technical details essential for successfully performing wire injury to induce consistent&#160;neointima for studies of restenosis.

Cardiovascular disease caused by atherosclerosis is the leading cause of death in the developed world. Narrowing of the vessel lumen, due to ...

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 Repetitive Concussive Head Injury Model in Mice

Despite the concussion/ mild traumatic brain injury (mTBI) being the most frequent occurrence of traumatic brain injury, there is still a lack of knowledge on the injury and its effects. To develop a better understanding of concussions, animals are often used because they provide a controlled, rigorous, and efficient model. Studies have adapted traditional animal models to perform mTBI to stimulate mild injury severity by changing the injury parameters. These models have been used because they can produce morphologically similar brain injuries to the clinical condition and provide a spectrum of injury severities. However, they are limited in their ability to present the identical features of injuries in patients. Using a traditional impact system, a repetitive concussive injury (rCHI) model can induce mild to moderate human-like concussion. The injury degree can be determined by measuring the period of loss of consciousness (LOC) with a sign of a transient termination of breathing. The rCHI model is beneficial to use for its accuracy and simplicity in determining mTBI effects and potential treatments.

Concussion presents the most common type of traumatic brain injury. Therefore, a repetitive concussive animal model, which replicates the important features of an injury in patients, may provide a means to study concussion in a rigorous, controlled, and efficient manner.

Despite the concussion/ mild traumatic brain injury (mTBI) being the most frequent occurrence of traumatic brain injury, there is still a lack of ...

The Generation of Closed Femoral Fractures in Mice: A Model to Study Bone Healing

Bone fractures impose a tremendous socio-economic burden on patients, in addition to significantly affecting their quality of life. Therapeutic strategies that promote efficient bone healing are non-existent and in high demand. Effective and reproducible animal models of fractures healing are needed to understand the complex biological processes associated with bone regeneration. Many animal models of fracture healing have been generated over the years; however, murine fracture models have recently emerged as powerful tools to study bone healing. A variety of open and closed models have been developed, but the closed femoral fracture model stands out as a simple method for generating rapid and reproducible results in a physiologically relevant manner. The goal of this surgical protocol is to generate unilateral closed femoral fractures in mice and facilitate a post-fracture stabilization of the femur by inserting an intramedullary steel rod. Although devices such as a nail or a screw offer greater axial and rotational stability, the use of an intramedullary rod provides a sufficient stabilization for consistent healing outcomes without producing new defects in the bone tissue or damaging nearby soft tissue. Radiographic imaging is used to monitor the progression of callus formation, bony union, and subsequent remodeling of the bony callus. Bone healing outcomes are typically associated with the strength of the healed bone and measured with torsional testing. Still, understanding the early cellular and molecular events associated with fracture repair is critical in the study of bone tissue regeneration. The closed femoral fracture model in mice with intramedullary fixation serves as an attractive platform to study bone fracture healing and evaluate therapeutic strategies to accelerate healing.

The murine closed femoral fracture model is a powerful platform to study fracture healing and novel therapeutic strategies to accelerate bone regeneration. The goal of this surgical protocol is to generate unilateral closed femoral fractures in mice using an intramedullary steel rod to stabilize the femur.

Bone fractures impose a tremendous socio-economic burden on patients, in addition to significantly affecting their quality of life. Therapeutic strategies ...

A Modified Simple Method for Induction of Myocardial Infarction in Mice

Myocardial infarction (MI) represents one of the leading causes of death. MI models are widely used for investigating the pathomechanisms of post-MI remodeling and evaluation of novel therapeutics. Different methods (e.g., isoproterenol treatment, cryoinjury, coronary artery ligation, etc.) have been used to induce MI. Compared with isoproterenol treatment and cryoinjury, coronary artery ligation may better reflect the ischemic response and chronic remodeling after MI. However, traditional methods for coronary ligation in mice are technically challenging. The current study describes a simple and efficient process for induction of MI in mice with readily available materials. The mouse chest skin was cut open under stable anesthesia. The heart was immediately externalized through the intercostal space after blunt separation of the pectoralis major and pectoralis minor. The left anterior descending branch (LAD) was ligated with a 6-0 suture 3 mm from its origin. Following LAD ligation, staining with 2,3,5-Triphenyltetrazolium chloride (TTC) indicated successful induction of MI and temporal changes of post-MI scar size. Meanwhile, survival analysis results showed overt mortality within 7 days after MI, mainly due to cardiac rupture. Moreover, post-MI echocardiographic assessment demonstrated successful induction of contractile dysfunction and ventricular remodeling. Once mastered, an MI model can be established in mice within 2-3 min with readily available materials.

Under adequate anesthesia, the mouse heart was externalized through the intercostal space, and myocardial infarction was successfully induced by ligating the left anterior descending artery (LAD) using materials readily available in most laboratories.

Myocardial infarction (MI) represents one of the leading causes of death. MI models are widely used for investigating the pathomechanisms of post-MI ...