A Reliable and Reproducible Critical-Sized Segmental Femoral Defect Model in Rats Stabilized with a Custom External Fixator

Summary

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.

Abstract

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.

Introduction

Orthopedic trauma surgery focuses on treating a wide range of complex fractures. Critical diaphyseal segmental bone defects have proven difficult to treat clinically due to the decreased regenerative ability of the surrounding muscle and periosteum as well as the failure of localized angiogenesis¹. Modern treatment techniques include operative fixation with bone grafting, delayed bone (Masquelet) grafting, bone transport, fusion, or amputation^2,3,4. In most patients who have ambulatory function preserved after their trauma, with well-functioning distal limbs, limb salvage is clearly a better treatment option⁵. These salvage treatments often require staged surgical interventions over a long treatment course. Some authors have suggested that external fixation is superior as compared to the internal fixation for these applications due to the decreased tissue damage during implantation, decreased implanted surface area, and increased postoperative adjustability of the fixator⁶. However, a prospective randomized controlled trial is currently underway to help clarify this controversy of internal versus external fixation in severe open fractures of the tibia⁷. Unfortunately, with either treatment selected, significant complication and failure rates persist^8,9. With either treatment method, with respect to the segmental bone loss, the surgeon must contend with segmental diaphyseal defects that present significant challenges. Corrections of segmental defects must maximize bone stabilization and simultaneously enhance the osteogenic process^10,11.

Due to the clinical importance, yet the lower volume, of critical-sized diaphyseal segmental defects, an effective, reproducible animal model is necessary to enable research teams to advance treatment techniques and ultimately improve clinical outcomes. Researchers need to study in vivo physiologic healing mechanisms in a mammalian animal model. While such models of external fixation already exist^12,13,14,15, we hope to provide a more reliable method for non-unions in the untreated animals, decrease costs through the choice of affordable fixator materials, and outline a straightforward surgical protocol for the easy application to future studies. The primary goal of this protocol is to establish a reliable and reproducible model of a critical diaphyseal defect in rats. The procedure was evaluated by assessing the stabilization and bone healing in rat femurs over 12 weeks. The secondary goals included: making an affordable model as a cost effective as possible, simplifying the surgical approach and stabilization, and ensuring ethical care of the animals. The authors and research team conducted preliminary experiments with a range of different biomaterials and potential regenerative therapies to improve healing in this segmental defect.

Protocol

The rats used in this study received daily care in accordance with the AVMA Guidelines for the Euthanasia of Animals: 2013 Edition¹⁶. The Institutional Animal Care and Use Committee at the University of Wisconsin-Madison evaluated and approved this experimental protocol before the project began.

1. Animals

1. Use the outbred Sprague-Dawley male rats weighing approximately 350 g.

2. Preparation of Bone Morphogenetic Protein-2 (rhBMP-2) soaked sponge scaffolds

NOTE: Scaffold preparation should occur just before implantation in the femur (see Step 6.14).

1. Follow manufacturer instructions for the use of an established rhBMP-2 bone graft kit containing a collagen sponge, lyophilized rhBMP-2, and sterile water for reconstitution¹⁷. Maintaining sterility, reconstitute the rhBMP-2 with the sterile water to a concentration of 1.5 mg/mL.
2. Using sterile scissors and a sterile ruler, trim the rhBMP-2 soaked collagen sponge to reshape to fit a 5 mm x 3 mm x 3 mm defect.
3. Using a syringe, distribute the rhBMP-2 solution evenly over the collagen sponge so that it is absorbed.

3. Preparation of custom external fixation device

NOTE: See Figure 1A for the more complete listing of dimensions.

1. Cut aluminum sheet stock (type 6061, 0.088” thickness) to two pieces (1.4” x 6”) using a jigsaw or other appropriate tool.
2. Mount one piece in the milling machine and, using a 1/8” 90°-point carbide drill mill, cut four 'V' grooves (0.035” deep) lengthwise. Leave the other piece free of cuts.
3. Cut individual plates of 0.3” width from the two pieces (Figure 1B). Measure and drill screw holes for 4-40 thread. Tap plate with ‘V’ grooves with the 4-40 thread. Drill the plate without grooves for a #4 screw body drill.
4. Sand both pieces to round corners and reduce weight (Figure 1C).
5. Screw pieces together with 4-40, 18-8 stainless steel button head cap screws (0.25”) so that grooves are flush against the plain plate (Figure 1D)

4. Anesthetic procedure and analgesia

1. Induce anesthesia by placing the rat in induction chamber delivering 4 L O₂/min with 4% isoflurane.
   CAUTION: Research personnel must avoid inhalation of anesthetic gas and maintain proper hood and ventilation in the laboratory.
2. Remove the rat from the chamber after the rat loses righting reflex, attach a nose cone and place at the maintenance dose of anesthesia through the nose (O₂ delivery rate to 2-3 L/min and 0.8% isoflurane).
3. Place the rat on the heating pad or under the warming light to prevent hypothermia.
4. Confirm the adequate depth of anesthesia by pinching the toe or testing the palpebral reflex.
5. Apply lubrication to eyes to prevent drying out of the cornea.
6. Deliver a subcutaneous injection of extended-release buprenorphine (1 mg/kg) on the trunk/dorsum of the rat, far from the surgical site, to provide analgesia for up to 3 days following surgery.

5. Aseptic preparation and antibiotic preventatives

1. Shave area around hindleg using the 13^th rib, the foot, the dorsal midline, and the ventral midline as margins.
2. Scrub shaved area using sterile 2 x 2 gauze soaked with 10% povidone-iodine followed by 70% EtOH (4 times each, alternating).
3. Administer an intramuscular injection of cefazolin (20 mg/kg) into the operative quadriceps.
4. Administer enrofloxacin (0.25 mg/ml) in drinking water for 7 days postoperatively for continued antibiotic protection.
5. Place rats on medicated feed (e.g., Uniprim) for the duration of the study to prevent pin tract infections.
6. Apply double antibiotic ointment to the skin-pin interface once daily for 3 days postoperatively.
   NOTE: Avoid any external fixation pin or clamp loosening which can contribute to the development of an infection.

6. Surgical procedure

NOTE: Make a concerted to effort to maintain a sterile field and workspace and follow sterile technique throughout the entirety of the case.

1. Extend shaved leg through fenestrated, clear sticky drape and cover surgical bench in sterile towels to create a sterile field.
2. Palpate the femur and use a #15 blade to create an anterolateral incision through the skin extending from the patella to the greater trochanter at the proximal femur.
3. Carefully incise the lateral leg fascia along the intermuscular septum to separate the vastus lateralis muscle of the quadriceps anteriorly from the hamstrings posteriorly until the lateral femur is exposed. Preserve the abductor gluteal tendon insertion on the greater trochanter.
4. Perform a careful, atraumatic circumferential soft tissue dissection and expose the femur at its mid-diaphysis starting on the lateral surface. To do this, use a #15 blade to gently cut the muscle away from the underlying bone by keeping the blade parallel against the contour of the bone surface. Use a periosteal elevator to lift the muscle away from the exposed bone as it is dissected and proceed around the femoral shaft until 7-10 mm of central diaphysis has been cleared of soft tissue on all sides to prepare for ostectomy.
   NOTE: Avoid injury to the medial femoral neurovascular bundle.
5. Insert four 1.0 mm Kirschner (k) wires: 2 proximal and 2 distal in the femur perpendicular to the lateral femur, directed straight lateral to medial. Ensure all pins engage both cortices (bicortical) for adequate stability (Figure 2A).
6. Start with the distal-most pin first, just at the level of the lateral epicondyle. Place jig flush to the lateral distal femur and insert a 1.0 threaded tip k-wire.
7. Maintaining the position of the jig on the bone, identify where the most proximal pin will enter the bone based on the jig holes. Once the position is determined, carefully incise parallel to the fibers of the gluteal tendon as needed to create a small gap in the tissue for the proximal pin to pass through, thus minimizing iatrogenic damage to the tendon. Drill a 1.0 mm non-threaded k-wire in this gap, again ensuring the pin engages both cortices (Figure 2B).
8. Maintain the jig’s position in contact with the bone and drill two 1.0 mm threaded k-wires, one on either side of the future defect site. Ensure pins engage both cortices (Figure 2C).
9. Place the external fixator bar level 1 cm above skin and screw tightly, locking the bar in place. Clip the excess pin lengths (Figure 2D).
10. Prepare for the ostectomy (defect creation) by placing a small, curved retractor around the anterior and posterior femur to protect the surrounding soft tissue, muscle, and neurovascular bundle. Utilizing a ~5 mm sagittal oscillating saw blade, very cautiously create a 5 mm segmental defect through the mid-diaphysis. Apply a light, even pressure with the saw to avoid unnecessary fracture (Figure 2E).
11. Apply small amounts of irrigation (room temperature 0.9% sterile normal saline (NS)) as needed while creating defect to avoid thermal necrosis of the bone.
12. Flush the wound using 10 mL of NS after creating the defect.
13. Administer 0.1 mL of a 0.25% bupivacaine with epinephrine (1:200,000) to the wound as an analgesic and vasoconstrictor.
14. Insert the scaffold (5 mm x 3 mm x 3 mm) of collagen sponge or rhBMP-2 soaked sponge (from Step 2) into the defect. Each scaffold should be sized appropriately to span the length and volume of the defect, helping the sponge stay in position.
    NOTE: At this point, mRNA complexes may be prepared and injected as outlined in Steps 7.1-7.3 below if performing bioluminescence imaging.
15. Close the muscle plane using the simple interrupted pattern with 4-0 absorbable suture. Close the skin layer using a running subcuticular pattern with 4-0 absorbable suture and skin glue to close gaps around the protruding pins.
16. Remove the rat from the nose cone, remaining on the heating pad, and monitor continuously until the rat is able to consistently maintain an upright posture. At this point, place in a clean cage to recover.

7. Preparation of complexed mRNA and bioluminescence imaging

NOTE: Transfection with mRNA complexes should be performed during surgery 1 day before luminescence imaging. Use sterile techniques when handling mRNA.

1. Mix 10 µL of mRNA encoding for Gaussia luciferase (stock concentration of 1 µg/µL) with 30 µL of the lipidic transfecting agent.
2. Allow for the mRNA-lipid complexes to form by incubating for at least 5 min at room temperature. The lipidic transfecting agent will condense the mRNA molecules, stabilizing them and enhancing transfection efficiency.
   NOTE: If the complexes are not used immediately, store them in ice for a maximum of 1 h.
3. Using a 20 µL pipette equipped with filtered tips, inject half of the volume of mRNA complexes to the distal and proximal ends of the defect, respectively.
4. The following day, 3 min before imaging, anesthetize the rat using inhaled isoflurane as previously described in Step 4.1.
5. Position the rat in an in vivo imaging chamber equipped with a nose cone delivering maintenance isoflurane (0.8% isoflurane, O₂ delivery rate of 2-3 L/min).
6. Inject coelenterazine resuspended in saline at a dose of 4 mg/kg body weight in the proximity of the defect.
7. Acquire bioluminescence images with the in vivo imaging system (IVIS) according to the manufacturer’s instructions¹⁸.

8. Imaging Protocol

1. After calibrating the plain radiographic machine, an X-ray system¹⁹, anesthetize rat using inhaled isoflurane as previously described (see Step 4.1) and position the rat in a nose cone with inhaled isoflurane (0.8% isoflurane, O₂ delivery rate of 2-3 L/min) for an anteroposterior (AP) femur radiograph.
   1. While the rat is in sternal recumbency, advance the surgical hindlimb forward, flexing at the hip and stifle joint. Flex the stifle joint to approximately 90°. Tape the paw plantar side down, close to the body wall. Position the tibia forward from the femur to eliminate the possibility of superimposing the bones. To provide slight abduction of the hip, place a translucent sponge (approximately 15 mm thick) in the groin region. Then obtain an anterior-posterior (cranial-caudal) image of the femur.
2. Repeat this AP femur radiographic view immediately following surgery, 4 weeks, and 12 weeks. Use tape and gauze to appropriately position the animal’s extremity for quality and consistent imaging.
3. Remove the rat from the nose cone and monitor continuously until the rat is able to consistently maintain an upright posture. Then, place back into the cage.

9. Histological Procedure

1. Euthanize rats in a chamber with inhaled CO₂ according to AVMA ethical standards¹⁶.
2. Following euthanasia, shave the hindlimb, remove the skin from the operative extremity and disarticulate femur at the hip. Carefully remove all soft tissue from the operative femur (including all muscles, tendons, and ligaments). Leave only a thin layer of muscle surrounding the defect site to protect the healing region from inadvertent damage during dissection.
3. Place the femur in 10% Neutral Buffered Formalin at room temperature for 3-4 days to allow for fixation. Keep a 15:1 formalin to tissue volume ratio. Change the solution once halfway through the fixation process.
4. Decalcify the femur in a 15% Ethylenediaminetetraacetic acid (EDTA) pH 6.5 solution for 3-4 weeks. Collect serial radiographs to determine decalcification endpoint.
5. Bisect the femur longitudinally with a cut from anterior to posterior in the mid-sagittal plane. Submit tissue for standard paraffin embedding and hematoxylin and eosin (H&E) staining.
6. Send H&E slides to a pathologist for histological assessment.

Results

Surgeries were performed in approximately one hour by one surgeon with the help of one assistant. After surgical optimization, intra- and postoperative complications were greatly minimized and use of the jig apparatus ensured consistent size (5 x 3 x 3 mm) and localization of femoral defects. Rats were ambulatory immediately following recovery from anesthesia and did not appear to have any altered behavioral patterns; their gait was not antalgic, and they did not appear to be disturbed by...

Discussion

Small animal models of orthopedic injuries such as complete bone fractures enable research that explores the mechanisms of osteogenesis and assessing the therapeutic potential of biomaterials²⁰. This study introduces a rat segmental defect model stabilized by a custom external fixator that a lab and biomedical engineering team can readily reproduce for further studies of load-bearing osteosynthetic bone repair.

Previous studies using critical-sized defects in rat models...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.9% Sterile Saline	Baxter	2F7124	Used for irrigating wound and rehydration
10% Iodine/Povidone	Carefusion	1215016	Used to prep skin
10% Neutral Buffered Formalin	VWR	89370094	Used as fixative
1mm non-threaded kirschner wire	DePuy Synthes	VW1003.15	Sterilized, used for the most proximal pin
1mm threaded kirschner wire	DePuy Synthes	VW1005.15	Sterilized, used for the 3 most distal pin slots
2x2 gauze	Covidien	4006130	Sterilized, used to prep skin and absorb blood
4-0 Vicryl Suture	Ethicon	4015304	Used to close muscle and skin layers
4-40 x 0.25",18-8 stainless steel button head cap screws	Generic		External fixator assembly
4200 Cordless Driver	Stryker	OR-S-4200	Used to drill kirschner wires
4x4 gauze	Covidien	1219158	Sterilized, used to absorb blood
70 % Ethanol			Used to prep skin
Baytril	Bayer Healthcare LLC, Animal health division	312.10010.3	Added to water as an antibiotic
Cefazolin	Hikma Pharmaceuticals	8917156	Pre-op antibiotic
CleanCap Gaussia Luciferase mRNA (5moU)	TriLink Biotechnologies	L-7205	Modified mRNA encoding for Gaussia Luciferase, keep on ice during use
Coelenterazine native	NanoLight Technology	303	Substrate for Guassia Luciferase, used to assess luciferase activity in vivo
Double antibiotic ointment	Johnson & Johnson consumer Inc	8975432	Applied to pin sites post-op as wound care
Dual Cut Microblade	Stryker	5400-003-410	Used to create 5mm defect in femur
Ethylenediamine Tetraacetic Acid (EDTA)	Fisher	BP120-500	Used to decalcify bone to prep for histology
Extended Release Buprenorphine	ZooPharm		Used as 3 day pain relief
Fenestrated drapes	3M	1204025	Used to establish sterile field
Handpiece cord for TPS	Stryker	OR-S-5100-4N	Used to create 5mm defect in femur
Heating pad	K&H Pet Products	121239	Rat body temperature maintenance
Hexagonal head screwdriver	Wiha	263/1/16 " X 50	External fixator tightening
Induction chamber	Generic		Anesthesia for rats
Infuse collagen sponge with recombinant human Bone Morphogenic Protein-2	Medtronic	7510200	Clinically relevant treatment used as positive control
Isoflurane	Clipper	10250	Anesthesia for rats
IVIS	Perkin Elmer	124262	Bioluminescence imaging modality
Jig	Custom		Used to place bicortical pins
Lipofectamine MessengerMAX	Fisher Scientific	LMRNA003	mRNA complexing agent that enables mRNA delivery
Sensorcaine-MPF (Bupivicane (0.25%) and Epinephrine (1:200,000))	APP Pharmaceuticals, LLC	NDC 63323-468-37	Applied to surgical site for pain relief and vasoconstriction
Sterile water	Hospira	8904653	Used as solvent for cefazolin powder
Titanium external fixator plates	Custom		Prepared in house with scrap titanium and milling machine
Total Performance System (TPS) Console	Stryker	OR-S-5100-1	Used to create 5mm defect in femur
TPS MicroSaggital Saw	Stryker	OR-S-5100-34	Used to create 5mm defect in femur
Ultrafocus Faxitron with DXA	Faxitron		High resolution radiographic imaging modality
Uniprim rat diet	Envigo	TD.06596	Medicated rat diet
Universal Handswitch for TPS	Stryker	OR-S-5100-9	Used to create 5mm defect in femur
Vetbond Tissue Adhesive	3M	1469	Skin closure