Name: Establishing a Diaphyseal Femur Fracture Model in Mice (Video) | JoVE Journal
Uploaded: 2022-12-09T14:30:00.000+00:00
Duration: 5 min 46 s
Description: Learn an open surgical technique to establish diaphyseal femur fractures in mice for studying long-bone healing. This low-cost model enables local testing of stem cells, drugs, and biomaterials in vivo.

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5 minutes

This work was funded by the Carlos Chagas Filho Foundation for Research Support of the State of Rio de Janeiro (FAPERJ).

All the experiments were approved by the Animal Use and Care Committee of the Center for Health Sciences of the Federal University of Rio de Janeiro (Protocol Number 101/21). Male Balb/c mice at 10-12 weeks of age (25-30 g body weight) were used in this study. The surgical procedure takes approximately 15-20 min per mouse. Before each procedure, the required instruments (listed in the Table of Materials) must be organized over a sterile surgical field covering the operating table (Figure 1A). The metallic surgical instruments must be autoclaved in self-sealing envelopes at 123 &#176;C for 30 min. Disposable items, such as needles and gauze pads, must be procured sterile.
1. Animal preparation
<ol>
	<li>Anesthetize the mouse and perform analgesia in accordance with the veterinary-recommended regimen approved by the&#160;institutional animal care and use program. 
	NOTE: If available, inhalation anesthesia should preferably be performed. A description of the protocol for inhalation anesthesia can be found in the report by Ewald et al.18. However, if the fracture is produced for osteoimmunology studies, this type of anesthesia must be avoided, as evidence shows that several volatile anesthetics, including isoflurane, affect the activity of both innate and adaptive immune cells19,20.</li>
	<li>Once the mouse is immobile, shave the left leg, and then transfer it to the surgical table on a warm heating pad (see Table of Materials) at 37 &#176;C covered with a sterile surgical drape.</li>
	<li>Perform antiseptic washing of the incision area by rubbing the skin with a 10% povidone-iodine sponge. Disinfection should begin along the incision line and extend outward in a circular pattern.&#160;Dry the rubbed area with sterile gauze pads, wash with 70% ethanol, and dry again with a sterile gauze pad. Repeat this procedure three times.</li>
	<li>Place the mouse in the right lateral decubitus position, and immobilize the paws with surgical tape (Figure 1C).</li>
	<li>Drape the mouse so that only the incision region is visible (Figure 1D).</li>
</ol>
2. Surgical procedure
<ol>
	<li>During the surgical procedure, constantly check that the mouse is breathing and provide eye drops in its eyes to avoid dryness and prevent the mouse from becoming blind. 
	NOTE: The whole surgical procedure usually takes ~15-20 min when performed by a trained surgeon. Therefore, applying the eye drops once at the beginning of the procedure should suffice. If the procedure starts to become longer, additional applications can be performed whenever it is identified that the eyes are starting to dry.</li>
	<li>Before proceeding to the incision, evaluate the anesthetic depth by pinching the tail to check the pain response reflex and visually inspecting the rate of breathing (counting the number of thoracic movements per minute)21. Under optimal anesthesia, the mouse should not respond to a tail pinch, and the rate of breathing must be around 55-65 breaths/min21.</li>
	<li>Make a 1 cm cutaneal lateral parapatellar incision with a scalpel blade (Number 11, see Table of Materials), beginning at the level of the tibial tuberosity and extending to the level of the patella and then, for an equal distance, toward the distal femur (Figure 1E).</li>
	<li>With blunt-end scissors, dissect the subcutaneous fascia around the incision line to expose the fascia lata, the lateral vastus, and the femoral biceps muscles22.</li>
	<li>With the scalpel blade Number 11, make another incision in the fascia lata similar to the one made in the skin, beginning at the level of the tibial tuberosity and running along the biceps femoris aponeurosis until the level of the distal femur, to open the articular capsule and access the knee joint (Figure 1F, G).</li>
	<li>Perform a medial luxation of the patella by placing the tip of a straight serrated precision tip tweezer (see Table of Materials) under it and pushing it to the side together with the patellar and quadriceps ligaments, thus exposing the condyles of the femur (Figure 1H).</li>
	<li>Holding the femur with a serrated tip tweezer, flex the knee at 90&#176;, and manually perforate the intramedullary canal of the femur through the intercondylar fossa with a 26 G hypodermic needle (Figure 1I, J).</li>
	<li>Maintaining the knee flexed at 90&#176;, insert a segment of 1.0 cm of a 0.016 in (0.40 mm) stainless steel rod wire (Figure 1K, insert) (see Table of Materials) through the opening into the medullary canal of the femur toward the great trochanter (Figure 1K). 
	NOTE: Maintaining the knee flexed at 90&#176; is crucial for the proper insertion of the wire into the medullary canal. Not doing so will result in the extravasation of the wire out of the bone and surrounding soft tissue lesions.</li>
	<li>Adjust the pre-bent distal extremity of the wire with a straight serrated tip tweezer to tightly fix it in the lateral condyle (Figure 1L). In addition to fixing the wire in place, the bent extremity will facilitate the postmortem removal of the wire.</li>
	<li>Separate the lateral vastus and femoral biceps muscles through blunt end dissection with a serrated tip tweezer to access the distal diaphysis of the femur (Figure 1M).</li>
	<li>Insert a dissecting scissor around the femur diaphysis at an angle of approximately 90&#176;, and gently perform a complete cortical osteotomy (Figure 1N). 
	NOTE: Mice femurs are easily cut. Refrain from applying excessive force during osteotomy to avoid the bending of the intramedullary wire and extensive fracture comminution.</li>
	<li>Reposition the muscles and the patella by pushing the tip of a straight serrated precision tip tweezer over the condyle region.</li>
	<li>Close the muscle fascia with a 6-0 resorbable suture and then the skin using a 6-0 nylon suture (see Table of Materials), both in a simple interrupted fashion (Figure 1O).</li>
	<li>Transfer the mouse to an individual clean cage for recovery. Once awake, the mouse must be able to move freely with unrestricted weight bearing.</li>
	<li>In the following days after surgery, perform analgesia in accordance with the veterinary-recommended regimen approved by the&#160;institutional animal care and use program.</li>
</ol>
3. X-ray imaging
<ol>
	<li>Anesthetize the mouse as described in step 1.1. 
	NOTE: If the radiography is performed right after the surgical procedure and the mouse is still under optimal anesthesia (step 2.2), it is not necessary to perform this step.</li>
	<li>For a clean lateral view of the fractured femur, place the mouse in the dorsal decubitus position, and slightly pull the operated hindlimb to the side.</li>
	<li>Immobilize the paws with surgical tape.</li>
	<li>Perform radiographic imaging according to the available equipment protocol. 
	&#8203;NOTE: For this study, a digital dental X-ray generator was used with the following parameters: 70 kVp voltage, 7 mA current, and 0.2 s exposure time.</li>
</ol>
4. Histology processing and H&#38;E staining
<ol>
	<li>Euthanize the mice with an intraperitoneal overdose of anesthetics (please refer to the veterinary-recommended regimen approved by the&#160;institutional animal care and use program). After checking the depth of anesthesia with a tail pinch, perform cervical dislocation. Next, collect the fractured bone, remove excess surrounding muscle tissue23, and fix the bone in 10% buffered formalin solution (pH 7.4) for 3 days.</li>
	<li>Place the bone samples in labeled histology cassettes (see Table of Materials), and immerse them in 10% EDTA in phosphate-buffered saline (PBS), pH 7.4, for 14 days for decalcification. Change the decalcification solution twice per week.</li>
	<li>Dehydrate the samples in a series of solutions of increasing ethanol concentrations (70%, 80%, 90%, 100%, 100%) for 1 h each.</li>
	<li>Clear the samples in two sequential baths of xylene for 30 min each.</li>
	<li>For wax infiltration, immerse the samples in two sequential paraffin baths at 60 &#176;C for 30 min. Next, embed the samples into blocks for sectioning24. 
	NOTE: To better view the callus, embed the bone with its long axis in the horizontal position to allow for longitudinal cuts.</li>
	<li>Cut the tissue into 4 &#181;m thick sections with a microtome (see Table of Materials).</li>
	<li>Float the sections in a 56 &#176;C water bath, and mount the sections on histological slides (see Table of Materials).</li>
	<li>For H&#38;E staining, deparaffinize the slides in three sequential baths of xylene for 5 min, and rehydrate the tissue in a series of solutions of decreasing ethanol concentrations (95%, 80%, and 70%) for 5 min.</li>
	<li>Rinse the slides in tap water for 30 s, stain the slides with Harris hematoxylin (see Table of Materials) for 6 min, and rinse them in tap water for another 30 s.</li>
	<li>Immerse the slides in 1% hydrochloric acid in ethanol for 30 s and then in 70% ethanol for 30 s.</li>
	<li>Stain with eosin (see Table of Materials) for 2 min, and wash with tap water for 30 s.</li>
	<li>Dehydrate the slides with ethanol (70%, 80%, and 95% for 5 min), and clarify with two baths of xylene for 5 min each.</li>
	<li>For mounting, drip one to two drops of mounting medium (see Table of Materials) onto each slide, and cover the slide with a clean coverslip.</li>
</ol>

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The authors have no conflicting financial interests.

As the number of fractures increases worldwide9,10,25, innovative treatments for nonunion are becoming increasingly urgent. As fracture healing involves a complex and tightly orchestrated summation of events that occur over a long timescale3, the use of valid animal models is central to improving our understanding of the mechanisms that determine the success of bone repair and to selecting effective drugs...

The skeleton is a vital and functionally versatile organ. The bones of the skeleton enable body posture and movement, protect the internal organs, produce hormones that integrate physiological responses, and are the site of hematopoiesis and mineral storage1. If fractured, bones have a remarkable capacity to regenerate and fully restore their pre-injury form and function. The healing process begins with the formation of a hematoma and an inflammatory response, which induces the activation and condensation of skeletal stem/progenitor cells from the periosteum, endosteum, and bone marrow and their subsequent differentiation to form the soft cartilaginous callus. The bridging of the fractured ends then occurs through a process that resembles endochondral bone formation, in which the cartilaginous scaffold expands and then mineralizes, forming the hard osseous callus. Finally, the hard callus is gradually remodeled by osteoclasts and osteoblasts to restore the original bone structure2,3.
Although the fracture healing process is fairly robust, it involves an intricate summation of events and is significantly influenced by several individual factors, including the general health status, age, and sex of the patient, as well as injury factors, such as the mode of mechanical stabilization of the fractured bone, the occurrence of infection, and the severity of the surrounding soft tissue injury4,5,6. Therefore, failures are common, leading to the development of nonunion, which greatly impacts patient rehabilitation and quality of life7,8. Due to the increasing number of fractures as a result of high-energy trauma and aging, as well as the high costs of treatments, nonunion fractures have become a burden for health systems worldwide9,10. This increasing burden highlights the urgent need for innovative therapeutic strategies to improve bone repair11,12 based on the combination of skeletal/mesenchymal stem/stromal cells and biomimetic biomaterials13,14.
In pursuit of this goal, animal models have been widely used in studies aiming to understand the fundamental biology of fracture healing mechanisms and in proof-of-concept preclinical studies aiming to devise new therapeutic strategies to promote bone repair15,16,17. Small-animal models, such as the mouse, are excellent for fracture healing studies because of the wide availability of genetically modified strains and reagents for experimental analyses and their low maintenance costs. Additionally, mice have a rapid healing time course, which allows for the temporal analysis of all stages of the repair process15. However, the small size of the animal can pose challenges for the surgical production of fractures with fixation modes similar to those applied in humans. This protocol describes a simple and low-cost model of fracture healing in mice using an open femoral osteotomy stabilized with an intramedullary wire, which resembles the most common bone repair process, through cartilaginous callus formation, and can be used both in basic and translational investigations in which access to the fracture site is required.

<table><tbody><tr><td>Alcohol 70&amp;ordm;</td><td>Merck</td><td>109-56-8</td><td>Or any general available supplier</td></tr><tr><td>Canada balsam (mounting medium)</td><td>Merck</td><td>C1795</td><td>Or any general available supplier</td></tr><tr><td>Cefazoline</td><td>ABL</td><td>Not applicable</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Coverslip</td><td>Merck</td><td>CSL284525</td><td>Or any general available supplier</td></tr><tr><td>Dental X-Ray Generator</td><td>Focus</td><td>-</td><td>Sold by Instrumentarium Dental Inc.&amp;nbsp;</td></tr><tr><td>DEPC water</td><td>Merck</td><td>W4502</td><td>Or any general available supplier</td></tr><tr><td>Dissecting Scissor</td><td>ABC Instrumentos</td><td>0327</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>EDTA</td><td>Vetec</td><td>60REAVET014340</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Eosin solution</td><td>Laborclin</td><td>EA-65</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Ethanol P.A</td><td>Vetec</td><td>60REAVET012053</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Gauze&amp;nbsp;pads</td><td>Cremer</td><td>Not applicable</td><td>Or any general available supplier</td></tr><tr><td>Harris Hematoxylin Solution</td><td>Laborclin</td><td>620503</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Heating pad</td><td>Tonkey Electrical Technology</td><td>E114273</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Histological slides</td><td>Merck</td><td>CSL294875X25</td><td>Or any general available supplier</td></tr><tr><td>Histology cassettes</td><td>Merck</td><td>H0542-1CS</td><td>Or any general available supplier</td></tr><tr><td>Hydrochloric acid - 37%</td><td>Merck</td><td>258148</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Insulin syringe</td><td>BD</td><td>324918</td><td>Or any general available supplier</td></tr><tr><td>Iodopovidone&amp;nbsp;sponge</td><td>Rioqu&amp;iacute;mica</td><td>372106</td><td>Or any general available supplier</td></tr><tr><td>Ketamine hydrochloride</td><td>Ceva</td><td>Not applicable</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Lacribel collyrium</td><td>Cristalia</td><td>Not applicable</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Microtome</td><td>Leica</td><td>149AUTO00C1</td><td></td></tr><tr><td>Mouse Tooth Forceps Tweezer</td><td>ABC Instrumentos</td><td>0164</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Needle 26 G</td><td>BD</td><td>2239</td><td>Or any general available supplier</td></tr><tr><td>Needle Holder&amp;nbsp;</td><td>Golgran</td><td>135-18</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Nonresorbable Nylon Suture thread n&amp;ordm; 6</td><td>Atramat</td><td>C1546-NT</td><td>Or any general available supplier</td></tr><tr><td>Paraffin</td><td>Exodo</td><td>8002 - 74 - 2</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Paraformaldehyde</td><td>Sigma</td><td>30525-89-4</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>PBS 1x&amp;nbsp;</td><td>Lonza</td><td>&amp;nbsp;BE17-516F</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Resorbable Nylon Suture thread n&amp;ordm; 6</td><td>Atramat</td><td>C1596-45B</td><td>Or any general available supplier</td></tr><tr><td>Rod Wire SS CrNi 0.016&quot;</td><td>Orthometric</td><td>56.50.2016</td><td></td></tr><tr><td>Scalpel n&amp;ordm; 11</td><td>Descarpak</td><td>15782</td><td>Or any general available supplier</td></tr><tr><td>Serrated Tip Tweezer</td><td>Quinelato</td><td>QC.404.12</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Shaver</td><td>Phillips</td><td>Not applicable</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Surgical tape</td><td>3M</td><td>2734</td><td>Or any general available supplier</td></tr><tr><td>Surgical tnt field</td><td>Polarfix</td><td>6153</td><td>Or any general available supplier</td></tr><tr><td>Tramadol hydrochloride</td><td>Teuto&amp;nbsp;</td><td>Not applicable</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Water bath for histology</td><td>Leica</td><td>HI1210</td><td></td></tr><tr><td>Xylazine hydrochloride</td><td>Ceva</td><td>Not applicable</td><td>Similar brands of the item may be used according to local availability</td></tr><tr><td>Xylene</td><td>Dinamica</td><td>60READIN001105</td><td>Similar brands of the item may be used according to local availability</td></tr></tbody></table>

establishing a diaphyseal femur fracture model in mice

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.

The most simple and immediate way to evaluate the success of the surgical procedure in producing the fracture is X-ray imaging. Radiographs can be performed immediately after surgery, with the mouse still under anesthesia, and subsequently 7 days, 14 days, and 21 days after the fracture to evaluate the callus formation and progression. Acceptable fracture patterns are those in which the cortices are fully ruptured, the wires are correctly placed within the medullary canal, and the fracture lines are transverse (with an a...

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 ...

establishing-a-diaphyseal-femur-fracture-model-in-mice

Research

JoVE Journal

Biology

Pseudofracture, a reproducible murine model of sterile musculoskeletal trauma, allows for evaluation of late term post-traumatic immune responses. This article describes the procedural execution of the model step by step, including the potential for experimental model combinations to permit study of multiple trauma.

Pseudofracture: An Acute Peripheral Tissue Trauma Model

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

A Simple Critical-sized Femoral Defect Model in Mice

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.

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

This protocol describes an osteosynthesis technique using an intramedullary locking nail for standardized fixation of femur osteotomies, which can be used to analyze normal and defective bone healing in mice.

An Intramedullary Locking Nail for Standardized Fixation of Femur Osteotomies to Analyze Normal and Defective Bone Healing in Mice

This protocol describes a mouse model of orthopedic surgery that has been used to study mechanisms of postoperative neuroinflammation and behavioral changes, and when combined with parabiosis, to study tissue regeneration during aging.

A Mouse Model of Orthopedic Surgery to Study Postoperative Cognitive Dysfunction and Tissue Regeneration

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.

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

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.

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

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

Transverse Fracture of the Mouse Femur with Stabilizing Pin

The present protocol describes the development of a reproducible testing platform for murine femoral necks in a cantilever bending set-up. Custom 3D printed guides were used to consistently and rigidly fix the femurs in optimal alignment.

Cantilever Bending of Murine Femoral Necks

Here, we present a protocol to iatrogenically fracture the shaft of the femur of Wistar albino rats and follow up on the development of the callus. This femur osteotomy model can help researchers evaluate the process of fracture healing and to study how a drug could influence fracture healing.

Protocol for Developing a Femur Osteotomy Model in Wistar Albino Rats

Learn an open surgical technique to establish diaphyseal femur fractures in mice for studying long-bone healing. This low-cost model enables local testing of stem cells, drugs, and biomaterials in vivo.

Establishing a Diaphyseal Femur Fracture Model in Mice

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