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. Author manuscript; available in PMC: 2022 Jan 1.
Published in final edited form as: Methods Mol Biol. 2021;2230:75–89. doi: 10.1007/978-1-0716-1028-2_5

A Mouse Femoral Ostectomy Model to Assess Bone Graft Substitutes

Ryan Trombetta 1,#, Emma K Knapp 2, Hani A Awad 2,3
PMCID: PMC8299532  NIHMSID: NIHMS1720744  PMID: 33197009

Abstract

The shortcomings of autografts and allografts in bone defect healing have prompted researchers to develop suitable alternatives. Numerous biomaterials have been developed as bone graft substitutes each with their own advantages and disadvantages. However, in order to test if these biomaterials provide an adequate replacement of the clinical standard, a clinically representative animal model is needed to test their efficacy. In this chapter, we describe a mouse model that establishes a critical sized defect in the mid-diaphysis of the femur to evaluate the performance of bone graft substitutes. This is achieved by performing a femoral ostectomy and stabilization utilizing a femoral plate and titanium screws. The resulting defect enables the bone regenerative potential of bone graft substitutes to be investigated. Lastly, we provide instruction on assessing the torsional strength of the healed femurs to quantitatively evaluate the degree of healing as a primary outcome measure.

Keywords: bone graft substitute, critical sized defect, mouse model, scaffold, healing, bone, biomechanics, torsional testing

1. INTRODUCTION

Bone grafting is a common and necessary procedure for healing bone defects that result from trauma, disease, cancer, or congenital anomalies. Due to the high demand for bone grafts, bone is the second most transplanted tissue [1]. In the United States, over half a million patients receive bone defect repairs with an estimated cost exceeding 2.5 billion US dollars [2]. The current gold standard for bone grafting procedures are autografts harvested from the patient’s own tissue typically from the iliac crest. Although autografts provide the osteoinduction, osteoconduction, and osteogenesis needed for bone regeneration, only a limited amount of tissue can be harvested from the patient, which results in donor site morbidity and severe blood loss [3], [4]. Because of these caveats, allografts are the clinical standard for large bone defects. However, prior to being implanted allografts are decellularized to abate the host’s immune rejection of the graft and gamma irradiated to remove viruses, bacteria, and fungi [5,6]. This renders the allograft a nonviable scaffold incapable of remodeling the damage accumulated during daily activities as living bone normally does. Thus, allogenic failure rates reported as high as 37% are attributed to the rigorous decellularization and sterilization process of allografts, which result in loss of osteogenic properties that limits the bone healing potential. Due to the various shortcomings of allogenic and autogenic bone transplantations, researchers have thoroughly investigated alternative biomaterials that could be used as bone graft substitutes.

Various biomaterials have been utilized as bone graft substitutes. The primary objective of all these substitutes is to enable osseous healing with complete bone union restoring the biomechanical strength and function. The most commonly investigated bone graft alternatives are synthetic bone grafts. These consist of calcium salt derivatives (i.e. calcium sulfate [7] and calcium phosphate [8]), bioactive glass [9], synthetic (and natural) polymers [10], and composite grafts [11]. These, materials have been produced in multiple physical forms such as powders [12], pastes [13], cements [14], pellets [15], spacers [16], putties [17], and coatings [18]. These biomaterial formulations also share the key characteristic of being bioresorbable. The design of these scaffolds generally aims to have resorption rates comparable to new bone formation rates [19]. However, the major limiting factor is that synthetic bone grafts are solely osteoconductive, lacking osteoinductive and osteogenic properties. Hence the host tissue is required to initiate and orchestrate the bone healing and regeneration process. To circumvent this limitation, orthobiologics are added to the synthetic scaffolds in order to incorporate osteoinductive components to initiate and enhance bone regeneration [20]. These include growth factors such as bone morphogenetic proteins [21], vascular endothelial growth factors [22], insulin-like growth factors [23], fibroblast growth factors [24], and platelet-derived growth factors [25]. Demineralized Bone Matrix (DBM) [26] and platelet rich plasma (PRP) [27] can also be added to these biomaterials, which contain a multitude of undefined growth factors. Bone graft substitutes can also be seeded with bone forming cells, such as osteoblasts [28], mesenchymal stem cells [29], and osteoprogenitor cells [30] in order to enhance bone formation. An emerging technique currently being investigated is incorporation of engineered periosteum around bone graft substitutes in order to augment osteogenesis [31]. Stem cell sheet technology is utilized to create the thin outermost portion of bone consisting of MSCs and osteoprogenitor cells, which has been shown to be critical for bone healing [32]. Despite the various grafting materials and combinations of orthobiologics, there is minimum level 1 evidence supporting the use of bone graft substitutes over conventional grafting standards [20]. Therefore, the field of tissue engineering must continue to push the scientific development and evaluation of bone graft substitutes that promise to replace the clinical standards of allografts in preclinical models. Equally as important as the grafting material itself are the preclinical models used to investigate and assess the regeneration of bone in these critical defects.

Clinically-relevant animal models are essential for the preclinical evaluation of promising bone graft substitutes. Bone healing models encompass various modalities ranging from fracture to large critical sized defect reconstruction. Numerous species have been implemented, and each has their advantages and disadvantages. This variety of models provides researchers with numerous options for investigating the efficacy of their bone graft substitutes. Ultimately the researcher must choose a model that will provide the appropriate answers for their underlying research questions. With regard to bone graft substitutes, a critical sized defect (does not heal spontaneously) is ideal for testing the regenerative properties of bone grafts and their biomaterial substitutes. Critical sized defects are associated with high rates of complications and poor functional outcomes in the clinic [33]. In animal models, critical sized defects are achieved by one of three ways, 1) cylindrical metaphyseal defects (tibial plateau, femoral plug), 2) segmental mid-diaphysis defects, or 3) circular calvaria defects. Metaphyseal and calvaria defects provide inexpensive and easy surgical approaches due to the lack of required stabilization hardware. Metaphyseal and calvaria defects are not fully load bearing, while segmental mid-diaphysis defects are load bearing. Studies have shown that controlled loading positively affect bone healing [34]. Segmental mid-diaphysis defects are commonly associated with trauma and infection scenarios and require stabilization hardware that replicates clinical practice. The degree of load bearing is determined by both the choice of the stabilization hardware and biomaterial scaffold used as a bone graft substitute. These models are often performed in larger animals, such as canines or sheep, which can be expensive. However, due to advances in micro-machining, small fracture-fixation hardware has been developed for use in rodents, including mice. Mice are easily handled, housed, and offer a cost-effective animal model allowing for high sample sizes. Furthermore, the mouse genome is easily manipulated enabling an abundance of precise disease models for investigation. However, due to their small size and limited bone volume researchers face challenges to rigor and reproducibility when performing invasive surgical procedures that are associated with segmental bone defects. This concern is eliminated with mouse-specific stabilization hardware, specialized surgical tools, and an established and repeatable surgical approach that we present herein to enable reproducible reconstruction of a critical-size femoral defect in a mouse.

In this chapter, we describe the methods of establishing a critical-size segmental defect by performing an ostectomy in a murine femur stabilized by using a polyether ether ketone (PEEK) femoral plate and titanium screws. This model enables the study of bone graft substitutes made from various biomaterials to induce and orchestrate bone healing and regeneration in the defect as determined by torsional mechanical testing as the definitive standard outcome of healing in bone.

2. MATERIAL

2.1. Presurgical Prep

  1. Hair clipper

  2. Povidone prep solution and 70% isopropyl alcohol

  3. Ophthalmic ointment

  4. Surgical pad

  5. General surgical disposables consisting of face masks, sterile drapes, sterile gloves, and sterile gauze

2.2. Anesthetic and Analgesic Drugs

  • 2.

    Anesthesia drugs: 130 mg/kg Ketamine-HCl and 12 mg/kg xylazine. Drugs are combined and administered as a single intraperitoneal (IP) injection. The following regimen will produce a surgical level of anesthesia lasting 45–60 min and sedation 2–3 h: combine 1.0 ml of 130 mg/mL ketamine HCl with 1.0 ml of 12 mg/ml xylazine and 8.0 ml 1xPBS. The combined drugs are to be administered at 0.1 ml/10 g body weight via an IP injection using a 1 ml syringe with 25G 5/8 in. needle.

  • 3.

    Analgesic drugs: 0.3 mg/ml Buprenorphine HCl (Buprenex®). 0.1 mg/kg Buprenex® is administered subcutaneously preoperatively and every 12 h up to 3 days postoperatively (see Note 1). Subcutaneous injections are given using a 1 ml syringe with a 25–30 G 5/8 in. needle.

2.3. Femoral Ostectomy

  1. RISystems MouseFix plate 6-hole, polyether ether ketone (PEEK) (see Note 2)

  2. RISystems MouseFix screw, L=2.00 mm

  3. RISystems MouseFix saw guide (2–3 mm) (see Note 3)

  4. RISystems drill bit 0.30 mm

  5. RISystems Hand Drill

  6. Saw guide mount (Fig. 1a–b, see Note 4)

  7. Dremel

  8. Forceps

  9. Surgical scissors

  10. Curved forceps

  11. Hemostatic forceps

  12. Gigli wire saw 0.22 mm

  13. 5–0 monofilament sutures

  14. Sterile drapes

  15. No. 10 scalpel

  16. LX-60 X-ray cabinet (Faxitron Bioptics© or equivalent)

Figure 1.

Figure 1.

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Photograph of the saw guide mount.

2.4. Sample Prep for Biomechanical Torsion Testing

  1. Harvested femurs, cleaned of all soft tissue

  2. Scalpel

  3. Forceps

2.5. Set-up Prep for Biomechanical Torsion Testing

  1. Square aluminum tubing (outer dimension of ¼”, cut to ¾” lengths)

  2. Plumber’s putty

  3. Plastic alignment jig – with 6mm gauge length opening

  4. Alignment release bars

  5. Aluminum compacting bar

2.6. Alignment fixation for Biomechanical Torsion Testing

  1. 5mL syringes

  2. Autopolymerizable bone cement mix (acrylic powder and liquid)

  3. Wooden tongue depressors

  4. Graduated cylinder

2.7. Sample rehydration for Biomechanical Torsion Testing

  1. 1X Phosphate buffer solution

  2. Petri dish

2.8. Biomechanical Torsion Testing

  1. EnduraTec Test Bench™, with 200N.mm torque cell (Bose Corp. Minnetonka, MN)

  2. WinTest7 software

3. METHODS

Please note that all personnel performing this protocol should be familiar with their institutional animal welfare policies as well as the anatomy of the mouse’s hind limb. Adherence to guidelines such as the ARRIVE (Animal Research: Reporting In Vivo Experiments) Guidelines Checklist is highly recommended.

3.1. Surgical Preparation of Mouse Hindlimb

  1. Anesthetize the mouse as describe in Subheading 2.2, item 1. This will provide a surgical plane of anesthesia that will last for approximately 45–60 minutes, which is enough time for an experienced rodent surgeon to complete this procedure. The average time for this procedure is 25–30 minutes.

  2. Administer pain management drugs preoperatively as per Subheading 2.2, item 2, with a subcutaneous injection of buprenorphine.

  3. Place the mouse on the surgical pad. Using the clipper, shave the hair on the posterior surface of the hindlimb spanning from the distal end of the tibia to halfway up the spine and just beyond the medial plane of the spine. Shave any remaining hair on the anterior surface of the hindlimb (Fig. 2a).

  4. Using a gauze pad or Q-tip applicator, prep the posterior surface on the hind leg with alternating scrubs of povidone iodine and 70% isopropanol. Repeat this procedure three times.

  5. Prepare a sterile drape and organize sterile instruments.

  6. Position the mouse in the prone position (Fig. 2a).

Figure 2.

Figure 2.

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Procedure of the femoral ostectomy.

3.2. Femoral Ostectomy

  1. Using sterile gloves and aseptic technique, create a longitudinal incision in line with the femur spanning from the hip joint to the knee using surgical scissors. To initiate the cut, pinch excess skin at the hip with teethed forceps to make initial cut. Use minimum number of cuts to establish surgical wound.

  2. Perform a blunt dissection to separate the vastus lateralis and biceps femoris muscles to expose the full length of the femur while preserving the sciatic nerve. To perform the blunt dissection, stretch exposed tissue with forceps downward to look for a gap between the lateralis and biceps femoris muscles (Fig. 2b). Insert closed surgical scissors and open in order to separate the two muscles (Fig. 2c). Repeat this action multiple times if necessary to expose the femur.

  3. Uncover and skeletonize the femur using forceps. Then use curved forceps to perform a circular preparation of the femur at the transition from the middle third to the distal third of the femur (Fig. 2d).

  4. Use the forceps with teeth to remove any soft tissue attached to the femur by lightly grasping the soft tissue and pulling away from the center of the bone. This must be performed very carefully as to not damage the periosteum (outermost vascular connective tissue of the bone), which plays a critical role in bone healing. Further expose the length of the femur by using the surgical scissors to cut any attached soft tissue at the proximal and distal ends of the femur.

  5. Place the six-hole PEEK plate to the anterolateral surface of the femur (see Note 5). Temporarily suture the PEEK plate to femur via the middle notch (Fig. 2e). As a visual reference to align the plate at the center of the femur, the second most proximal hole is adjacent to the third trochanter.

  6. Ensure that the plate is centered and use the hemostatic forceps in the nondominant hand to grip the plate at the middle and rotate away from the mouse (toward the surgeon).

  7. Use the drill bit inserted into the Dremel to drill the most distal hole (Fig. 2f). During the act of drilling, the surgeon should feel two gives corresponding to the drill bit penetrating through each cortex of the femur. When drilling it is important to make sure the drill is perpendicular to the bone in order to ensure stability and avoid fracture.

  8. Use the hand drill to insert a Titanium screw perpendicular to the bone. Tighten slowly until the screw head is flush with the plate (this is the part that sits right on top of the threads for the plate). When finished, shear off the pin by rotating away from the center of the plate in a clockwise manner (Fig. 2g).

  9. Repeat this process for the most proximal and then the second most proximal screws.

  10. Insert the final screw into the second most distal hole. Do not shear off the pin, leave intact for the saw guide (Fig 2h).

  11. Remove the sutures with either surgical scissors or a No. 10 scalpel.

  12. Install the saw guide by aligning with PEEK plate and insert the intact screw pin into the distal most screw hole of the saw guide. Grasp the intact screw pin with the forceps and push the saw guide down until it is flush with the plate (see Note 6).

  13. Clamp the saw guide in place with the hemostatic forceps (Fig. 2i). Use only the first locking point to prevent bone fracture.

  14. Move the mouse by gripping the curved forceps and saw guide to the saw guide mount and lock into place (Fig. 2j).

  15. Use approximately 8 inches of the Gigli saw wire to perform the osteotomy. Run the Gigli saw wire underneath the bone and align to saw guide’s slots using two forceps. Using a narrow grip, slowly initiate the cut with short strokes. Gradually widen your grip and perform longer strokes until the bone is completely cut (see Note 7).

  16. Unlatch the clamp and remove the mouse. Unclamp the hemostats and remove the saw guide. Check to ensure the bone was cleanly cut at both ends through both cortices. Remove the bone with forceps (Fig. 2k).

  17. Shear off the intact screw pin by reinserting hand drill and grasping the exposed plate with the hemostatic forceps.

  18. Suture proximal and distal ends of the femur using a 5–0 nylon monofilament suture in order to prevent screw pull-out (see Note 8).

  19. Place gauze pad underneath the mouse and spray the defect area with sterile 1xPBS from a syringe in order to irrigate the surgical wound and remove any bone debris.

  20. Insert any grafting or scaffolding material into the defect using forceps. Secure the graft of scaffold in a cerclage fashion using a 6–0 nylon braided suture and cut carefully with surgical scissors (Fig. 2l).

  21. Remove the curved forceps.

  22. Suture the muscle with two to four 5–0 sutures ensuring that surgical wound is closed.

  23. Close the skin with 5–0 sutures.

  24. Perform a radiographic evaluation of the mouse’s hindlimb to assess the placement and success of the surgical hardware and bone graft substitute (Fig. 3). Acquire planar x-ray images using a LX-60 X-ray cabinet or similar device. Scans are performed with an energy of 55 kV, intensity of 145 μA, and 300 ms integration time (see Note 9).

  25. Place mouse on a heated pad for anesthesia recovery.

Figure 3.

Figure 3.

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Planar X-ray image post-surgery of an implanted calcium phosphate bone graft substitute in a 3 mm defect.

3.3. Sample Prep for Biomechanical Torsion Testing

  1. At the end of the study (see Note 10), euthanize the mice and harvest the femur ensuring the proximal and distal ends are intact.

  2. Remove all soft tissue and surgical hardware from the harvested femur. If necessary, some soft tissue can be kept around the implanted graft/callus to ensure that it remains stable (see Note 11).

3.4. Set-up for Biomechanical Torsion Testing

  1. For each femur, 2 square aluminum tubes, one plastic alignment jig and one alignment release bar will be used. Label each tube (Fig. 4a).

  2. Fill one tube completely full of plumbers putty. Fill the other tube half way with plumbers putty.

  3. Insert the release bar into the alignment jig (Fig. 4b).

  4. Insert the proximal end of the bone into the tubing that is filled with putty. Using forceps, orient the bone, so the mid-diaphysis is centered and in line with the tubing (Fig 4c).

  5. Place the tube with the bone into the plastic alignment jig, so the edge of the tube with the bone protruding is flush with the inner edge of the alignment jig.

Figure 4.

Figure 4.

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Schematic depicting the alignment jig used to line up the harvested femur in the aluminum tubing with putty and bone cement.

3.5. Alignment Fixation for Biomechanical Torsion Testing

  1. Mix a small batch of autopolymerizable bone cement (see Note 12).

  2. Pour the bone cement into the tube that is half filled with plumbers putty. Make sure that bone cement is filled to the top of the tube.

  3. Insert the bottom tube, so that the distal end of the bone is submerged in wet cement. Slide the tube up, so the edge of the tube is flush with the inner edge of the alignment jig. Set the jig upright, to allow the bone cement to harden (Fig 4d). This will create an exposed bone gauge length of 6mm. If any excess bone cement gets on the exposed gauge length, gently scrap off the bone cement before it hardens, being careful to keep the bone aligned.

  4. Allow the bone cement to harden for 15–20 minutes.

  5. Using the alignment release bar, remove the specimen and tubes from the alignment jig. The distal end of the bone should be fixed within the tube at this point.

  6. Remove the proximal end of the bone from the tube and use the compacting bar to remove half of the putty, leaving the tube half full.

  7. Reinsert the alignment release bar into the alignment jig, and insert the specimen, so the tube with the distal end cemented is now flush with the inner edge of the alignment jig.

  8. Repeat steps 1–5 to cement the proximal end of the bone in the tube.

  9. Once the sample has both the proximal and distal end firmly fixed in bone cement, place the sample in a petri dish with 1X phosphate buffer solution for at least 2 hours, to rehydrate the sample.

3.6. Biomechanical Torsion Testing

  1. Turn on the EnduraTec Test Bench™ and open the WinTest7 software.

  2. The samples are inserted into the clamps, and 4 screws are used to secure the clamps. Custom fabricated clamps firmly grip and anchor the square tubes (Fig 5a–c).

  3. Initialize the program, with rotation and torque being recorded. Continue the test until the femur is completely fractured.

  4. Test the samples at a rotation speed of 1 deg/sec and a data collection rate of 20 samples/sec.

  5. The WinTest7 software produces a .txt file for each sample tested. This .txt file is then imported into Excel to determine the torsional rigidity, maximum torque and rotation at maximum torque of each sample.

Figure 5.

Figure 5.

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Custom machined clamps fabricated to clasp the square tubes containing the potted femur.

ACKOWLEDGEMENTS

This manuscript was supported by the NIAMS/NIH grants P30AR069655 and P50AR072000, and the AOTrauma Clinical Priority Program.

Footnotes

1.

As a substitution for 12 h doses of 0.1 mg/kg buprenorphine, extended release buprenorphine can be adminstered once preopertatively at a dose of 3.25 mg/kg subcutaneously.

2.

A PEEK plate is radiolucent and will not be visible on radiographic imaging, such as X-rays and microcomputed tomography (μCT). This was chosen for instances where μCT is performed as an added outcome measure to quantify bone regeneration because metal plates will produce metallic artifiacts and noise making quantification difficult.

3.

Various saw guide sizes can be purchased or fabricated. For a mouse critical-sized femoral defect, a defect size of 2 mm is require.

4.

The saw guide mount was created in-house using a low-profile hold-down toggle clamp seated at the end of a cantilever arm attached to a weighted base

5.

Installing the first screw is crucial because at this moment the alignment of the plate is determined. Pay extra attention to aligning the surface prior to implantation. Adjust the oritentation of the long axis so that the plate is sitting flush on the anterolateral surface of the femur and is centered.

6.

Rotating the bone can help align the saw guide, if the plate is sitting perfectly in the middle of the bone the saw guide will fit flushly with the plate. Otherwise, attempt to adjust orientation of the saw guide so that is fits over the plate and bone.

7.

It is important to ensure the bone is cut all the way through for both slots. The surgeon should be able to feel the difference in friction between cutting the bone verse cutting plate indicating the bone is completely cut. If bone was not completely cut and the ostectomy was incomplete, the bone cannot be removed. If this occurs install the saw guide and continue cutting with the gigli saw until the osteoctomy is complete. On the contrary, cutting too much can damage the plate and this must be avoided to ensure stabilization of the femur.

8.

This step is not required, but is performed as a safety precaution.

9.

When assessing quality of surgery, examine the position of the defect and screws to ensure the defect is centered in the femur. Confirm that the bone graft subsitute is sucessfully implanted in the defect. Also examine if any bone fragments or fractures resulted from the installation of the titanium screws and ostectomy.

10.

Study duration is determined by the time needed to heal the defect. Larger defects will require a longer healing time. A minimum of 12 weeks is suggested as the time required to heal a 2 mm critical sized femoral defect.

11.

When potting, the bone cement will only adhere to exposed bone, and excess soft tissue can cause an air pocket to form around the ends of the bone when curing, comprising the structural integrity of the fixation.

12.

The brand used was Bosworth Fastray. Approximately 10g of powder and 3mL of liquid were used to make the bone cement.

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