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

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Research

JoVE Journal

Medicine

15.1K 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

Use of Human Perivascular Stem Cells for Bone Regeneration

Human perivascular stem cells (PSCs) can be isolated in sufficient numbers from multiple tissues for purposes of skeletal tissue engineering1-3. PSCs are a FACS-sorted population of 'pericytes' (CD146+CD34-CD45-) and 'adventitial cells' (CD146-CD34+CD45-), each of which we have previously reported to have properties of mesenchymal stem cells. PSCs, like MSCs, are able to undergo osteogenic differentiation, as well as secrete pro-osteogenic cytokines1,2. In the present protocol, we demonstrate the osteogenicity of PSCs in several animal models including a muscle pouch implantation in SCID (severe combined immunodeficient) mice, a SCID mouse calvarial defect and a femoral segmental defect (FSD) in athymic rats. The thigh muscle pouch model is used to assess ectopic bone formation. Calvarial defects are centered on the parietal bone and are standardly 4 mm in diameter (critically sized)8. FSDs are bicortical and are stabilized with a polyethylene bar and K-wires4. The FSD described is also a critical size defect, which does not significantly heal on its own4. In contrast, if stem cells or growth factors are added to the defect site, significant bone regeneration can be appreciated. The overall goal of PSC xenografting is to demonstrate the osteogenic capability of this cell type in both ectopic and orthotopic bone regeneration models.

Human perivascular stem cells (PSCs) are a novel stem cell class for skeletal tissue regeneration similar to mesenchymal stem cells (MSCs). PSCs can be isolated by FACS (fluorescence activated cell sorting) from adipose tissue procured during standard liposuction procedures, then combined with an osteoinductive scaffold to achieve bone formation in vivo.

Human perivascular stem cells (PSCs) can be isolated in sufficient numbers from multiple tissues for purposes of skeletal tissue engineering1-3. PSCs are ...

Bioengineering

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 ...

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. ...

A Minimally Invasive Model to Analyze Endochondral Fracture Healing in Mice Under Standardized Biomechanical Conditions

Bone healing models are necessary to analyze the complex mechanisms of fracture healing to improve clinical fracture treatment. During the last decade, an increased use of mouse models in orthopedic research was noted, most probably because mouse models offer a large number of genetically-modified strains and special antibodies for the analysis of molecular mechanisms of fracture healing. To control the biomechanical conditions, well-characterized osteosynthesis techniques are mandatory, also in mice. Here, we report on the design and use of a closed bone healing model to stabilize femur fractures in mice. The intramedullary screw, made of medical-grade stainless steel, provides through fracture compression an axial and rotational stability compared to the mostly used simple intramedullary pins, which show a complete lack of axial and rotational stability. The stability achieved by the intramedullary screw allows the analysis of endochondral healing. A large amount of callus tissue, received after stabilization with the screw, offers ideal conditions to harvest tissue for biochemical and molecular analyses. A further advantage of the use of the screw is the fact that the screw can be inserted into the femur with a minimally invasive technique without inducing damage to the soft tissue. In conclusion, the screw is a unique implant that can ideally be used in closed fracture healing models offering standardized biomechanical conditions.

This protocol describes a minimally invasive osteosynthesis technique using an intramedullary screw for standardized stabilization of femur fractures, which can be used to analyze endochondral bone healing in mice.

Bone healing models are necessary to analyze the complex mechanisms of fracture healing to improve clinical fracture treatment. During the last decade, an ...

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 ...

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

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.

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.

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

Transverse Fracture of the Mouse Femur with Stabilizing Pin

Fracture repair is an essential function of the skeleton that cannot be reliably modeled in vitro. A mouse injury model is an efficient approach to test whether a gene, gene product or drug influences bone repair because murine bones recapitulate the stages observed during human fracture healing. When a mouse or human breaks a bone, an inflammatory response is initiated, and the periosteum, a stem cell niche surrounding the bone itself, is activated and expands. Cells residing in the periosteum then differentiate to form a vascularized soft callus. The transition from the soft callus to a hard callus occurs as the recruited skeletal progenitor cells differentiate into mineralizing cells, and the bridging of the fractured ends results in the bone union. The mineralized callus then undergoes remodeling to restore the original shape and structure of the healed bone. Fracture healing has been studied in mice using various injury models. Still, the best way to recapitulate this entire biological process is to break through the cross-section of a long bone that encompasses both cortices. This protocol describes how a stabilized, transverse femur fracture can be safely performed to assess healing in adult mice. A surgical protocol including detailed harvesting and imaging techniques to characterize the different stages of fracture healing is also provided.

This protocol describes a method to perform fractures on adult mice and monitor the healing process.

Fracture repair is an essential function of the skeleton that cannot be reliably modeled in vitro. A mouse injury model is an efficient approach to test ...

Biology

Precise Growth Plate Injury Study Using Trilineage Fluorescent Mice

Tricolor Transgenic Murine Model for Studying Growth Plate Injury

The cartilage growth plates in the bones of children enable limb lengthening but are weak relative to the bone, making them prone to fracturing when bones are overloaded.&#160;Better treatments for severely fractured growth plates are needed because the response to injury is a bony bridge that prematurely fuses the growth plate, leading to stunted and/or crooked limbs. Murine models of growth plate injury are advantageous for mechanistic studies, but&#160;are challenging because it is difficult to visualize and precisely injure the small growth plates in young mice.&#160;We describe here an improved growth plate injury model using transgenic mice with tri-lineage fluorescent reporters for collagen types I, II, and X.
These mice show native fluorescence associated with the three primary substrata of the growth plate. A growth plate injury similar to a Salter-Harris Type II injury is created reproducibly with a bur using the hypertrophic section of the growth plate as a reference during live imaging under fluorescence stereo microscopy guidance. Frozen histology analysis of the native fluorescence simplifies assessing the cellular response to injury. This methodology represents a substantial leap in growth plate injury research, providing a detailed and reproducible method for investigating pathology and evaluating new therapeutic strategies.

This protocol describes an improved mouse model for adolescent bone growth plate injuries. Using transgenic mice with tri-lineage fluorescent reporters for collagen types I, II, and X, the primary matrices associated with three different substrata of the growth plate, injury placement is guided by native fluorescence under the microscope.

The cartilage growth plates in the bones of children enable limb lengthening but are weak relative to the bone, making them prone to fracturing when bones ...

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 ...