The research reported in this publication was supported by the National Institute of Arthritis and Musculoskeletal and Skin Diseases of the National Institutes of Health under award number F30AR071201 and R01AR066028.

This protocol was developed to ensure that animals are not used needlessly and are spared all unnecessary pain and distress; it adheres to all applicable federal, state, local, and institutional laws and guidelines governing animal research. The protocol was developed under the guidance of a university-wide Laboratory Animal Medicine Program directed by veterinarians specialized in laboratory animal medicine. The protocol was reviewed and approved by the Institutional Animal Care and Use Committee (IACUC).
1. Fracture Tower Construction
Note: All parts are listed in the materials section (Table of Materials). Detailed technical drawings are provided for the machined and 3D-printed parts in Supplementary Figures 1-12. The subassembly technical drawings include fastener details for all mounted parts (Supplementary Figures 1, 2, 7, and 9).
<ol>
	<li>Support subassembly 
	NOTE: For a technical drawing of the support subassembly, see Supplementary Figure 1.
	<ol>
		<li>Attach the Beam Support--Jaw Section at the midpoint of the Beam Support--Horizontal Section.</li>
		<li>Attach the Beam Support--Vertical 1 to the top surface of the Beam Support--Jaw Section, 2 in from the Beam Support--Horizontal Section.</li>
		<li>Attach the Beam Support--Vertical 2 to the top surface of the Beam Support--Horizontal Section at the midpoint (7 in from the end).</li>
		<li>Attach the Beam, Support--Plate Mount to the end of both Beam Support--Vertical 1 and Beam Support--Vertical 2. The end of the plate support should be flush with the back of Beam Support--Vertical 2.</li>
	</ol>
	</li>
	<li>Ram subassembly 
	Note: For a technical drawing of the ram subassembly, see Supplementary Figure 2.
	<ol>
		<li>Machine the Block Stop and the Block Guide (Supplementary Figure 3); the Rod Ram (Supplementary Figure 4); the Screw Alignment (Supplementary Figure 5); and the Plate Mounting (Supplementary Figure 6).</li>
		<li>Attach the Plate Mounting to the Beam Support--Plate Mount of the support subassembly.</li>
		<li>In the following order, slide the first Linear Sleeve Bearing; the Block Guide; the second Linear Sleeve Bearing; and the Block Stop on the Rod Ram. Attach the guides and the blocks to the Plate Mounting.</li>
		<li>Attach three &#8540;-in nuts to the threaded portion of the Rod Ram. One should be flush with the end of the rod to engage with the electromagnet. The other 2 will be used to adjust the fracture depth.</li>
		<li>Align the grove in the Rod Ram to face forward and insert the Alignment Screw into the threaded hole of the Block Guide.</li>
	</ol>
	</li>
	<li>Magnet subassembly 
	Note: For a technical drawing of the magnet subassembly, see Supplementary Figure 7.
	<ol>
		<li>Solder the Electromagnet leads to the wire (polarity is not a factor for the electromagnet operation). Allow enough length to reach the floor, where the fracture device will be positioned. Use zip ties or another form of attachment to stress relieve the wire.</li>
		<li>Strip the Power Supply&#39;s end and connect it to the Foot Pedal. Finally, connect the wire to the Foot Pedal in an &#34;off&#34; (normally open) configuration. Test the circuit to ensure the Electromagnet is on when the Foot Switch is not pressed. This will hold the ram up before the fracture.</li>
		<li>Print the Mount Magnet (Supplementary Figures 8A and 8B) using an additive manufacturing device, or machine the part from aluminum.</li>
		<li>Attach the Electromagnet to the Mount Magnet.</li>
		<li>Attach 2 Corner Brackets to the Beam Support--Magnet.</li>
		<li>In the following order, thread the Rod Magnet through the top Corner Bracket and add one &#188;-in nut; the Mount Magnet; two &#188;-in nuts; and the bottom Corner Bracket. Secure the assembly with two &#188;-in nuts on each end.</li>
	</ol>
	</li>
	<li>Complete assembly 
	Note: For a technical drawing of the complete assembly, see Supplementary Figure 9.
	<ol>
		<li>Attach the Magnet Subassembly to the top surface of the Beam, Support--Plate Mount.</li>
		<li>Adjust the alignment of the Beam Support--Magnet so the magnet engages with the Rod, Ram. 
		NOTE: If the rod does not release when the foot pedal is pressed, reduce the contact area between the electromagnet and the rod by moving the Beam Support--Magnet.</li>
		<li>Machine the Brackets Leg Jaw (Supplementary Figure 10).</li>
		<li>Attach the two Brackets Leg Jaw to the Beam Support--Jaw Section. When dropped, the tip of the ram should be at an equal distance from each jaw.</li>
		<li>Place the Platform Fracture (Supplementary Figures 11A and 11B) above the jaws.</li>
		<li>Print the Jig Positioning Fracture (Supplementary Figures 12A and 12B) and the Jig Pin Gauge (Supplementary Figures 13A and 13B) using an additive manufacturing device, or machine the parts from aluminum. 
		NOTE: The dimensions of the Jigs will be calculated in the optimization steps detailed in step 2.</li>
		<li>Attach the Jig Positioning Fracture to the Platform Fracture.</li>
		<li>Confirm that the depth of the impact can be adjusted using the two stop nuts on the Rod Ram.</li>
		<li>Confirm that the speed of the impact can be adjusted by moving the Mount Magnet up and down.</li>
		<li>Confirm that the width of the fracture can be adjusted by moving the Brackets Leg Jaw closer or further away from the Rod Ram.</li>
	</ol>
	</li>
</ol>
2. Fracture Optimization
<ol>
	<li>Fracture location
	<ol>
		<li>Obtain radiographs of the limb (femur or tibia) to be fractured in a representative sample of 5 euthanized animals. 
		NOTE: The sample should be matched to the specimens, which will be used in the experimental protocol based on age, genotype, and sex. Even if the final protocol calls for only one fractured limb, both sample limbs will be used.</li>
		<li>Position the limb tangential to the x-ray beam to acquire true-lateral and anterior/posterior views to the bone. Place an object of known dimension at the imaging plane to provide a scale for analysis.</li>
		<li>Note: If imaging femurs, ensure the limb is in full extension, where the femur is in the same axial plane as the tibia.</li>
		<li>Mark the desired location of the fracture on the radiograph of the limb to be fractured (Figure 1A - dashed line). Measure from the calcaneal-tibial joint to the level of the marked fracture (Figure 1A). Calculate the mean fracture length (FL) for all trial specimens. Measure from the intercondylar notch for femur fractures.</li>
	</ol>
	</li>
	<li>Fracture-positioning jig
	<ol>
		<li>Measure the distance from the outside surface of one support anvil to the center of the guillotine impact (CGI) (Figure 2). Subtract the CGI from the FL, described in step 2.1.4, to calculate the fracture-positioning jig depth (JD). Machine or 3D-print a U-shaped channel with a height and a width equal to the anvil, and a depth equal to the JD (Figure 3A). A sample technical drawing and CAD file are included in Supplementary Figures 12A and 12B. 
		NOTE: When the limb is placed in the jig, the dorsum of the foot should lie against the surface furthest from the guillotine impact. Modify the U-shaped channel if additional clearance is required for the limb.</li>
		<li>Position the specimen in the fracture apparatus in the prone position for femur fractures or in the supine position for tibia fractures (Figure 4). Press the dorsum of the foot against the end of the fracture-positioning jig. Manually depress the guillotine until the limb fractures. Obtain a radiograph of the fractured limb to confirm the jig size and fracture location (Figure 2B).</li>
		<li>Increase JD if the fracture location is too distal on the bone, or decrease JD if the fracture location is too proximal on the bone.</li>
	</ol>
	</li>
	<li>Stabilization of the pin parameters
	<ol>
		<li>Pin length:&#160;Using the radiographs obtained in step 2.1, measure the limb length (LL) from the tibial plateau to the level of the posterior malleolus for tibia fractures, or the intercondylar notch to the greater trochanter for femur fractures. Multiply the bone length by 0.9 to calculate the pin length (PL) (Figures 1A and 3B).</li>
		<li>Pin Width: Using the radiographs obtained in step 2.1, measure the minimum medullary diameter (MD) in the fractured limb (Figure 1A). Select a needle with a gauge approximately equivalent to the medullary diameter and a length more than 1.5 x PL. 
		NOTE: An approximate pin size for a 14-week-old C57BL/6J mouse is 22 G, 1&#189; in and 27 G, 1&#188; in for femur and tibia, respectively.</li>
	</ol>
	</li>
	<li>Pin cutting gauge
	<ol>
		<li>2.4.1. Machine or 3D-print a gauge with a length equal to PL minus the needle length (CGL) (Figure 3B; Supplementary Figures 13A and 13B). One end should have an overhang to rest against the hub of the needle and the other should indicate where the pin should be cut. A sample technical drawing and CAD file are included in Supplementary Figures 13A and 13B.</li>
	</ol>
	</li>
	<li>Intramedullary pin fracture stabilization
	<ol>
		<li>Using the non-fractured trial specimens from step 2.1, remove hair with an electric clipper or depilatory cream from mid-tibia to mid-femur, exposing the knee joint.</li>
		<li>Tibia pinning: Insert the needle percutaneously lateral to the patellar ligament. Retract the patellar ligament medially and align the tip of the needle to the axis of the tibia. Using a reaming motion, gently breach the tibial plateau and guide the needle down the medullary cavity.</li>
		<li>Femur pinning: Insert the needle percutaneously lateral to the patellar ligament. Retract the patellar ligament medially and align the tip of the needle to the axis of the femur in the intercondylar notch. Using a reaming motion, gently breach the articular surface of the intercondylar notch and guide the needle down the medullary cavity.</li>
		<li>Using the gauge manufactured in step 2.4, ream until the exposed needle is equal to the gauge length. Retract the needle to provide enough room (~3 mm) to cut the needle at the level indicated by the gauge. 
		NOTE: Be sure to hold the proximal (plastic) end of the needle while cutting, so it does not become a hazardous projectile.</li>
		<li>Crimp 0.3 mm of the distal end of the pin using a pin cutter and then cut the pin at the level of the gauge.&#160;Sink the pin to the articular surface using a rod with a diameter 1.5x larger than the diameter of the needle. 
		NOTE: Crimping prevents rotation of the needle and migration by increasing the needle-bone contact.</li>
		<li>Obtain radiographs to confirm the needle extends the length of the medullary canal of the limb and does not protrude from the proximal or distal end (Figure 1C).</li>
	</ol>
	</li>
	<li>Impact depth
	<ol>
		<li>Using the radiographs obtained in step 2.1, measure the diameter of the cortex at the level of the desired fracture (Figure 1A). Calculate the mean cortical diameter (CD) for all trial specimens.</li>
		<li>Position a pinned trial specimen from step 2.5 in the fracture device with the fracture-positioning jig manufactured in step 2.2. Rest the impact ram on the uninjured limb. 
		NOTE: Do not allow the ram to drop; the bone should remain intact during this optimization step.</li>
		<li>Apply enough downward force on the ram to compress soft tissue, but not fracture the bone. Adjust the impact depth (ID) to 0.75 x CD (Figure 2). 
		NOTE: The ideal impact depth is 0.5 x CD when fracturing a bone without any soft tissue. Using 0.75 accounts for the additional soft-tissue compression.</li>
	</ol>
	</li>
	<li>Anvil width
	<ol>
		<li>Set the anvil width (AW) to 0.4 cm for the mouse tibia or femur (Figure 2). 
		NOTE: A wider width is recommended for larger specimens such as rats.</li>
	</ol>
	</li>
	<li>Ram weight
	<ol>
		<li>A minimum weight of 250 g is recommended for murine specimens. 
		NOTE: Additional weight can be threaded on to the ram for larger specimens (Figure 2).</li>
	</ol>
	</li>
	<li>Impact velocity
	<ol>
		<li>Set the drop height (DH) to 2 cm (Figure 2). Position the ram in its starting position by connecting it to the activated electromagnet.</li>
		<li>Position a trial limb in the fracture apparatus. Press the dorsum of the foot against the fracture-positioning jig manufactured in step 2.2. Briefly depress the footswitch to release the ram and then reset it to its starting position.</li>
		<li>Radiograph the impacted trial limb. Analyze the limb for any evidence of a fracture (Figure 1D). 
		NOTE: This can be subtle when using low velocities with a controlled impact depth.</li>
		<li>If no fracture is generated, repeat steps 2.9.1 - 2.9.3 and increase the drop height by 2 cm.</li>
		<li>If a fracture is generated, record the drop height, and multiply it by 1.1. This is the new DH.</li>
		<li>Using the DH from step 2.9.5, fracture the next trial limb.</li>
		<li>If no fracture is generated, repeat steps 2.9.1 - 2.9.6 and increase the drop height by 2 cm.</li>
		<li>If a fracture is generated, repeat steps 2.9.6 - 2.9.7 until all test samples are used. Record the final DH and all parameters (FL, CGI, JD, PL, MD, PS, CGL, CD, ID, AW, and RW) from the optimization. Record the trial specimens&#39; age, sex, genotype, and weight.</li>
	</ol>
	</li>
</ol>
3. Closed-stabilized Fracture Generation
<ol>
	<li>Set-up
	<ol>
		<li>Sterilize all equipment and instruments via autoclave, hot bead immersion, or their equivalent.</li>
		<li>Place a heating element on the surgical table and set it to the optimal temperature. Cover the element with a surgical drape. Prepare 3 x 3 in2 of surgical drape with a 0.75-in circle cut out in the middle.</li>
		<li>Confirm the adjustment of the fracture tower before each trial (Figure 2). Set the ID, AW, RW, and DH to the values derived from the optimization protocol specific to the sex, age, and genotype for the specimen to be studied.</li>
		<li>Weigh and record the weight of the animal.</li>
	</ol>
	</li>
	<li>Surgery
	<ol>
		<li>Adequately sedate the mouse using inhalant anesthetics (isoflurane: 4 - 5% for induction; 1 - 2% for maintenance) or another established laboratory anesthesia protocol. The respiratory rate should be 55 - 100 breaths/min. The animal should not be responsive to a hind-limb toe pinch.</li>
		<li>Administer the first dose of the postoperative analgesia buprenorphine (0.1 mg/kg subcutaneously).</li>
		<li>Apply ocular lubrication to prevent corneal drying.</li>
		<li>Remove the animal&#39;s hair with an electric clipper from mid-tibia to mid-femur, exposing the knee joint. Clean the site of excess hair using non-reactive tape. Prepare the pinning site with a wet swab moistened with 70% EtOH. Repeat as necessary to remove all hair from the incision area.</li>
		<li>Prepare and clean the pinning area with alternative swabs of povidone-iodine and 70% EtOH. Use two alternative swab sequences to ensure sterility.</li>
		<li>A drape is then placed around the surgical site after the skin has been appropriately disinfected.</li>
		<li>Pin the limb to be fractured using the protocol described in step 2.5. Acquire radiographs to confirm the pin extends the length of the medullary canal but does not protrude from the proximal or distal end.</li>
		<li>Turn on the electromagnet and connect the impact ram to place it in the starting position.</li>
		<li>Position the specimen in the fracture apparatus by placing it in a prone position for femur fractures or in a supine position for tibia fractures. The pinned limb should be placed across the anvils and in the fracture-positioning jig with the dorsum of the foot pressed against the outside of the jig.</li>
		<li>While pressing the foot with one hand and ensuring only the limb is in the impact ram target area, briefly depress the footswitch to release the ram. Replace the ram in the starting position.</li>
		<li>Acquire radiographs and confirm the fracture location and type.</li>
	</ol>
	</li>
	<li>Postoperative management
	<ol>
		<li>Monitor the animal every 15 min during its recovery from anesthesia until the animal is conscious, can maintain sternal recumbency, and is ambulatory. Confirm the animal is able to ambulate over a 72-h period.</li>
		<li>House the animal individually until it has completely recovered.</li>
		<li>Maintain analgesia over a 48-h period with buprenorphine (0.1 mg/kg subcutaneously) administered every 12 h.</li>
		<li>Monitor and record the health status of the animal daily for 7 - 10 d or until euthanasia.</li>
	</ol>
	</li>
	<li>Post-fracture analysis
	<ol>
		<li>Measure FL, PL, CD, MD, and the fracture pattern. Record the measurements in a master data file.</li>
	</ol>
	</li>
</ol>

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A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fractures+in+geriatric+mice+show+decreased+callus+expansion+and+bone+volume.">Fractures in geriatric mice show decreased callus expansion and bone volume.</a> Clinical Orthopaedics and Related Research. 472 (11), 3523-3532 (2014).</li><li>Bonnarens, F., Einhorn, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Production+of+a+standard+closed+fracture+in+laboratory+animal+bone.">Production of a standard closed fracture in laboratory animal bone.</a> Journal of orthopaedic research. 2 (1), 97-101 (1984).</li><li>Marturano, J. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+improved+murine+femur+fracture+device+for+bone+healing+studies.">An improved murine femur fracture device for bone healing studies.</a> Journal of Biomechanics. 41 (6), 1222-1228 (2008).</li><li>Jackson, R. W., Reed, C. A., Israel, J. A., Abou-Keer, F. K., Garside, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Production+of+a+standard+experimental+fracture.">Production of a standard experimental fracture.</a> Canadian Journal of Surgery. Journal Canadien De Chirurgie. 13 (4), 415-420 (1970).</li><li>Byrne, M., Cleveland, B., Marturano, J., Wixted, J., Billiar, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Design+of+a+reproducible+murine+femoral+fracture+device.+Conference:+Bioengineering+Conference,+2007.">Design of a reproducible murine femoral fracture device. Conference: Bioengineering Conference, 2007.</a> NEBC '07. IEEE 33rd Annual Northeast. , (2007).</li><li>Carter, D. R., Hayes, W. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Compact+bone+fatigue+damage-I.+Residual+strength+and+stiffness.">Compact bone fatigue damage-I. Residual strength and stiffness.</a> Journal of Biomechanics. 10 (5), 325-337 (1977).</li><li>McGee, A., Qureshi, A., Porter, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Review+of+the+biomechanics+and+patterns+of+limb+fractures.">Review of the biomechanics and patterns of limb fractures.</a> Trauma. 6 (1), 29-40 (2004).</li></ol>

The authors have nothing to disclose.

This fracture optimization and generation protocol provides researchers with an efficient method to derive at fracture parameters and perform a minimally invasive procedure, which produces precise, repeatable, transverse fractures. Additionally, this protocol establishes a common set of fracture generation parameters, which promotes method consistency amongst researchers. These parameters will enable the creation of a common fracture database to establish fracture standards based on a variety of parameters (e.g., age, sex, gender, and genotype). An optimization of fracture variables significantly decreases sample heterogeneity - reducing the amount of wasted time, lost resources, and unusable data.
To generate accurate and precise fractures, it is vital to establish a standardized set of fracture generation parameters that will produce a high degree of specificity and reduce the variability of fracture locations. In addition to fracture generation, adequate stabilization is also required to promote fracture callus formation and to lessen the probability of non-union. Intramedullary pinning is a common fixation method used to stabilize appendicular long bone fractures both experimentally and clinically. Internally fixated fractures tend to heal indirectly - a process involving tissue differentiation, bone resorption at the fracture surface, and the subsequent fracture union via callus formation and remodeling. These processes can be impeded by movement at the fracture junction and migration of the pin within the medullary cavity. This protocol utilizes a fixation method that reduces the degree of displacement at the fracture site following fixation and limits the extent of pin migration without the use of sophisticated surgical equipment and techniques that can cause unnecessary damage to cortical bone tissue. Generating a set of pin parameters that maximize the intramedullary contact per a specific sample type provides the necessary stability for proper callus formation and bone remodeling.
Once the intramedullary pin has been placed, the next critical step is generating a simple transverse fracture. Protocols that generate fractures via externally applied, blunt-force trauma have the potential to produce comminuted fractures and damage fixation hardware. To mitigate these complications, it is important to control the impact depth, which has to be equal to 0.5x the average cortical diameter of each sample set23. Fracture comminution can also result from excessive force applied during external blunt-force trauma procedures. If the impact velocity exceeds a critical threshold, the speed of the crack propagation will generate stress waves resulting in multiple fracture sites24. It is critical to establish a ram weight and drop height that will generate enough kinetic energy to produce a fracture, while also remaining below the impact velocity threshold for stress wave production, reducing the possibility of comminution. A high-impact velocity will cause a rapid loading of the bone, which produces excessive energy absorption before the fracture is generated25. Upon fracture propagation, the excessive energy absorbed during the loading is released non-linearly, which produces comminution. A lower impact velocity and slower loading of the energy has a higher probability of producing a linear fracture compared to high-impact velocities and rapid loading26. To minimize the incidence of comminution, this protocol uses a standard ram weight of 250 g for mice - this can be adjusted to accommodate a larger species. When working with very young animals or with those with a known bone disease (e.g., osteopenia or osteosclerosis), it may be necessary to decrease the ram weight. It is important to use a consistent ram weight when adjusting the drop height so only one variable is being optimized at a time. Calculations for species-specific ideal impact velocities will produce more consistent fractures by accounting for slight variations in the size and soft tissue morphology of the specimen.
The methods described above eliminate many shortcomings of other fracture generation protocols; however, some aspects may require training to efficiently produce desired results. One possible complication of the procedure is an inaccurate pin placement, potentially causing considerable bone or soft-tissue damage. This is due primarily to the limited visibility of the approach and a lack of sufficient bilateral hand dexterity. An internal fixation without an open incision can require a fair amount of skill from the person performing the procedure. Therefore, it is important that he or she has had sufficient training - on cadavers, if necessary - to avoid excess soft-tissue damage that could cause complications throughout the healing process. Recognizing the structures specified in the protocol (the patellar ligament, tibial plateau, and intercondylar notch of the femur) will help produce a consistent, precise pinning with minimal soft-tissue damage. However, the goal of the described study was not to present a detailed procedure for pin placement, but rather to describe methods for generating ideal fractures.
The use of the cutting gauge is highly recommended to avoid any reaming through the proximal end of the femur or distal end of the tibia. Drilling through the proximal end of the femur could cause unnecessary damage to the soft tissue or bone in the hip, causing mobility and injury complications during the healing process. Similarly, reaming through the distal end of the tibia will damage ankle structures, altering the gait mechanics, loading, and callus formation.
To increase the accuracy of the fracture location, a custom limb-positioning jig can be designed to ensure the proper positioning of the limb within the device. A precise and accurate impact placement is essential to consistently generate fractures at the desired location. Our laboratory currently employs two jigs: one for mid-tibial fractures and the other for mid-femoral fractures, but the versatility of a modular design and 3D printing gives researchers the capability to generate fractures at a variety of locations. The addition of a custom jig designed to generate fractures in a particular location increases both the accuracy and precision of fracture generation by limiting the likelihood of operator errors.
The few limitations of this method are similar to those encountered in other existing closed fracture techniques. Excessive soft tissue or fat can impede the generation of fractures, as seen in older or overweight mice. It is important to note that this is normally due to a lack of force and not to a lack of impact depth. This limitation can be overcome by increasing either the ram weight or the velocity to increase the kinetic energy applied to the fracture site. This method also relies upon internal fixation, which can disrupt the endosteal surface of the bone and affect the healing. While endosteal disruption also occurs clinically with intramedullary nailing, if the contribution of endosteum to fracture repair is being studied, external fixation or plates may be a better option. An additional limitation is the required sample of sacrificial animals to establish the initial parameters; however, as the fracture variables for more sample types are established and the database develops, the need for additional sacrificial samples should decrease.
The described protocol increases the precision of induced traumas through the use of standardized parameters specific to sample type, minimizing the fracture heterogeneity typically seen in closed fracture generation procedures. Most current fracture generation protocols are applicable to only murine species and produce moderately consistent fractures. They often require the use of a specific sample type to obtain optimal results or do not account for variations within strains. The protocol presented here accounts for variation in size or bone morphology that may exist between mouse strains and can be adapted to generate consistent fractures in other species. In addition, the widespread application of this protocol will support the adoption of a standardized fracture language between researchers. Using similar protocols with common variables will improve the method consistency and strengthen comparisons between studies. While the parameters discussed above are specific to murine long bones, there is the potential for the fracture optimization protocol to be used in additional fracture models, further increasing the versatility of a collective fracture generation parameter database. Employing this fracture optimization protocol will increase the production of homogeneous, usable samples by improving the consistency of the fracture location and pattern. The higher percent yield of the samples will decrease the waste of laboratory resources, reduce the number of animals needed, and improve study efficiency.

Research on fracture healing is necessary to address a large clinical and economic problem. Each year over 12 million fractures are treated in the United States1, costing $80 billion per year2. The likelihood of a male or female suffering a fracture in their lifetime is 25% and 44%, respectively3. Problems associated with fracture healing are expected to increase with increased comorbidities as the population ages. To study and address this problem, robust models of fracture generation and stabilization are required. Rodent models are ideally suited for this purpose. They provide clinical relevance and can be modified to address specific conditions (i.e., multiple injuries, open, closed, ischemic, and infected fractures). In addition to replicating clinical scenarios, animal fracture models are important for understanding bone biology and developing therapeutics and devices. However, attempts to study differences between interventions may be complicated by the variability introduced by inconsistent fracture generation. Thus, generating reproducible and consistently closed fractures in animal models is essential to the field of musculoskeletal research.
Despite properly controlling for potential subject heterogeneity by ensuring the appropriate genetic background, sex, age, and environmental conditions, the production of clinically-relevant consistent bone injuries is a significant variable affecting reproducibility that must be controlled. Statistical comparisons using inconsistent fractures are plagued with experimental noise and a high variability4; in addition, fracture variability can result in unnecessary animal death because of the need to increase the sample size or the necessity to euthanize animals with comminuted or malpositioned fractures. The purpose of the method described herein is to optimize the fracture generation parameters that are specific to sample type and yield a consistent fracture location and pattern.
Current models of fracture generation fall into two broad categories, each with their own strengths and weaknesses. Open-fracture (osteotomy) models undergo surgery to expose the bone, after which a fracture is induced by cutting the bone or weakening it and then manually breaking it5,6,7,8. The benefits of this method are the direct visualization of the fracture site and a more consistent fracture location and pattern. However, the physiological and clinical relevance of the approach and mechanism of injury is limited. Additionally, open methods of fracture generation require a surgical approach and closure with prolonged periods during which the rodents are exposed to an increased risk of contamination.
Closed techniques address many of the open technique&#39;s limitations. Closed techniques produce fractures using an externally applied blunt force trauma which induces injury to the bone and surrounding tissues, more similar to those seen in human clinical injuries. The most common method was described by Bonnarens and Einhorn in 19849. They described a weighted guillotine being used to impart blunt trauma to break the bone without causing any external skin wounds. This method has been widely adopted to study the effect of genetics10,11, pharmacologic therapy12,13,14,15, mechanics16,17, and physiology18,19,20 on bone healing in mice and rats. While the benefit of closed methods is physiologically relevant fractures, experimental reproducibility and rigor are limited by fracture heterogeneity. The inconsistent fracture generation results in a limited between-group differentiation, lost specimens, and an increase in animals needed to achieve statistical significance.
Controlling the variability in fracture generation and stabilization is essential to produce meaningful results. In order to properly study the biology of fracture repair, a simple yet robust fracture model is needed. The model should be translatable to rodent species, bone types (femur or tibiae, for example), and across variable mouse genetic backgrounds and induced mutations. Furthermore, the ideal procedure should be technically simple and produce consistent results. To address fracture heterogeneity, the method described herein is the construction of a well-controlled fracture device that can then be used to optimize parameters and generate consistently closed fractures regardless of age, sex, or genotype.

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			<td height="20" style="height:20px;width:240px;">Support Subassembly</td>
			<td style="width:209px;"></td>
			<td style="width:147px;"></td>
			<td style="width:315px;">Supplementary Figure 1</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;">Beam, Support--Jaw Section&#160;</td>
			<td style="width:209px;">80/20</td>
			<td>1003 x 9.00</td>
			<td>w/ #7042 at A, C, in Left End</td>
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		<tr height="19">
			<td height="19" style="height:19px;">Beam, Support--Horizontal Section</td>
			<td style="width:209px;">80/20</td>
			<td>1002 x 14.00</td>
			<td></td>
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			<td height="19" style="height:19px;">Beam, Support--Vertical 1</td>
			<td style="width:209px;">80/20</td>
			<td>1050 x 10.50&#160;</td>
			<td>w/ #7042 at A in Left End and at A in Right End</td>
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			<td style="width:209px;">80/20</td>
			<td>1010 x 10.50&#160;</td>
			<td>w/ #7042 at D, B in Left End and at A in Right End</td>
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			<td style="width:209px;">80/20</td>
			<td>1030 x 8.00&#160;</td>
			<td>w/ #7036 at Left End</td>
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			<td height="19" style="height:19px;">Beam, Support--Magnet</td>
			<td style="width:209px;">80/20</td>
			<td>1010 x 13.50&#160;</td>
			<td>w/ #7042 at A, C, in Right End</td>
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			<td height="19" style="height:19px;width:240px;">Anchors (3)</td>
			<td style="width:209px;">80/20</td>
			<td style="width:147px;">3392</td>
			<td></td>
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			<td height="19" style="height:19px;width:240px;">Double Anchor (3)</td>
			<td style="width:209px;">80/20</td>
			<td style="width:147px;">3091</td>
			<td></td>
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		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Bolt Assembly (6)</td>
			<td style="width:209px;">80/20</td>
			<td style="width:147px;">3386</td>
			<td>1/4-20 x 3/8&#34;</td>
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		<tr height="19">
			<td height="19" style="height:19px;">Button Head Socket Cap Screw (6)</td>
			<td style="width:209px;">80/20</td>
			<td style="width:147px;">3604</td>
			<td>1/4-20 x 3/4&#34;</td>
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		<tr height="19">
			<td height="19" style="height:19px;">Ram Subassembly</td>
			<td style="width:209px;"></td>
			<td style="width:147px;"></td>
			<td>Supplementary Figure 2</td>
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		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Block, Stop</td>
			<td style="width:209px;">Custom</td>
			<td style="width:147px;"></td>
			<td>Supplementary Figure 3</td>
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		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Block, Guide</td>
			<td style="width:209px;">Custom</td>
			<td style="width:147px;"></td>
			<td>Supplementary Figure 3</td>
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			<td height="19" style="height:19px;width:240px;">Rod, Ram</td>
			<td style="width:209px;">Custom</td>
			<td style="width:147px;"></td>
			<td>Supplementary Figure 4</td>
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		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Alignment Screw</td>
			<td style="width:209px;">Custom</td>
			<td style="width:147px;"></td>
			<td>Supplementary Figure 5</td>
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			<td height="19" style="height:19px;width:240px;">Plate, Mounting</td>
			<td style="width:209px;">Custom</td>
			<td style="width:147px;"></td>
			<td>Supplementary Figure 6</td>
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		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Linear Sleeve Bearing (2)</td>
			<td style="width:209px;">McMaster-Carr</td>
			<td style="width:147px;">8649T2</td>
			<td></td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Hex Nut (3)</td>
			<td style="width:209px;">McMaster-Carr</td>
			<td>92673A125</td>
			<td>3/8-16 UNC</td>
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		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Socket Cap Screw (8)</td>
			<td style="width:209px;">McMaster-Carr</td>
			<td style="width:147px;">92196A108</td>
			<td>4/40 x 3/8&#34;</td>
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		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Socket Cap Screw (6)</td>
			<td style="width:209px;">McMaster-Carr</td>
			<td style="width:147px;">92196A032</td>
			<td>4/40 x 1 1/8&#34;</td>
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			<td height="19" style="height:19px;width:240px;">Socket Cap Screw (1)</td>
			<td style="width:209px;">McMaster-Carr</td>
			<td style="width:147px;">92196A267&#160;</td>
			<td>10/32 3/8&#34;</td>
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			<td height="19" style="height:19px;width:240px;">Magnet Subassembly</td>
			<td style="width:209px;"></td>
			<td style="width:147px;"></td>
			<td>Supplementary Figure 7</td>
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			<td height="19" style="height:19px;width:240px;">Mount, Magnet</td>
			<td style="width:209px;">Custom</td>
			<td style="width:147px;"></td>
			<td>Supplementary Figure 8</td>
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			<td>70235K23</td>
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			<td style="width:209px;">McMaster-Carr</td>
			<td>7376k2</td>
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			<td style="width:209px;">McMaster-Carr</td>
			<td>5698k111</td>
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			<td style="width:209px;">McMaster-Carr</td>
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			<td height="19" style="height:19px;">Rod, Magnet</td>
			<td style="width:209px;">McMaster-Carr</td>
			<td>95412A566</td>
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			<td height="19" style="height:19px;width:240px;">Corner Bracket (6)</td>
			<td style="width:209px;">80/20</td>
			<td style="width:147px;">4108</td>
			<td></td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Socket Cap Screw (1)</td>
			<td style="width:209px;">McMaster-Carr</td>
			<td style="width:147px;">92196A705</td>
			<td>10/32 1 1/4&#34;</td>
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		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Hex Nut (4)</td>
			<td style="width:209px;">McMaster-Carr</td>
			<td>92673A113</td>
			<td>1/4-20 UNC</td>
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		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Complete Assembly</td>
			<td style="width:209px;"></td>
			<td style="width:147px;"></td>
			<td>Supplementary Figure 9</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Bracket, Leg Jaw (2)</td>
			<td style="width:209px;">Custom</td>
			<td style="width:147px;"></td>
			<td>Supplementary Figure 10</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Platform, Fracture</td>
			<td style="width:209px;">Custom</td>
			<td style="width:147px;"></td>
			<td>Supplementary Figure 11</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Jig, Positioning-Fracture</td>
			<td style="width:209px;">Custom</td>
			<td style="width:147px;"></td>
			<td>Supplementary Figure 12</td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Other</td>
			<td style="width:209px;"></td>
			<td style="width:147px;"></td>
			<td></td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Pin Cutter</td>
			<td style="width:209px;">Medical Supplies and Equipment</td>
			<td style="width:147px;">150S</td>
			<td></td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;width:240px;">Needles</td>
			<td style="width:209px;">Sigma</td>
			<td style="width:147px;">Z192430, Z192376&#160;</td>
			<td>23g x 1.5&#34; - mouse femur, 27g x 1.25&#34; - mouse tibia</td>
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fracture apparatus design and protocol optimization for closed-stabilized fractures in rodents

The reliable generation of consistent stabilized fractures in animal models is essential for understanding the biology of bone regeneration and developing therapeutics and devices. However, available injury models are plagued by inconsistency resulting in wasted animals and resources and imperfect data. To address this problem of fracture heterogeneity, the purpose of the method described herein is to optimize fracture generation parameters specific to each animal and yield a consistent fracture location and pattern. This protocol accounts for variations in bone size and morphology that may exist between mouse strains and can be adapted to generate consistent fractures in other species, such as rat. Additionally, a cost-effective, adjustable fracture apparatus is described. Compared to current stabilized fracture techniques, the optimization protocol and new fracture apparatus demonstrate increased consistency in stabilized fracture patterns and locations. Using optimized parameters specific to the sample type, the described protocol increases the precision of induced traumas, minimizing the fracture heterogeneity typically observed in closed-fracture generation procedures.

Watch this Scientific Journal Video about Fracture Apparatus Design and Protocol Optimization for Closed-stabilized Fractures in Rodents at JoVE.com

Fracture Apparatus Design and Protocol Optimization for Closed-stabilized Fractures in Rodents

The goal of the protocol is to optimize the fracture generation parameters to yield consistent fractures. This protocol accounts for the variations in bone size and morphology that may exist between animals. Additionally, a cost-effective, adjustable fracture apparatus is described.

The reliable generation of consistent stabilized fractures in animal models is essential for understanding the biology of bone regeneration and developing ...

fracture-apparatus-design-protocol-optimization-for-closed-stabilized

Nanyang Technological University

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JoVE Journal

Medicine

13.3K Views.  Michigan State University. The goal of the protocol is to optimize the fracture generation parameters to yield consistent fractures. This protocol accounts for the variations in bone size and morphology that may exist between animals. Additionally, a cost-effective, adjustable fracture apparatus is described.

Video: Fracture Apparatus Design and Protocol Optimization for Closed-stabilized Fractures in Rodents

The Use of Cystometry in Small Rodents: A Study of Bladder Chemosensation

The lower urinary tract (LUT) functions as a dynamic reservoir that is able to store urine and to efficiently expel it at a convenient time. While storing urine, however, the bladder is exposed for prolonged periods to waste products. By acting as a tight barrier, the epithelial lining of the LUT, the urothelium, avoids re-absorption of harmful substances. Moreover, noxious chemicals stimulate the bladder's nociceptive innervation and initiate voiding contractions that expel the bladder's contents. Interestingly, the bladder's sensitivity to noxious chemicals has been used successfully in clinical practice, by intravesically infusing the TRPV1 agonist capsaicin to treat neurogenic bladder overactivity1. This underscores the advantage of viewing the bladder as a chemosensory organ and prompts for further clinical research. However, ethical issues severely limit the possibilities to perform, in human subjects, the invasive measurements that are necessary to unravel the molecular bases of LUT clinical pharmacology. A way to overcome this limitation is the use of several animal models2. Here we describe the implementation of cystometry in mice and rats, a technique that allows measuring the intravesical pressure in conditions of controlled bladder perfusion.
	After laparotomy, a catheter is implanted in the bladder dome and tunneled subcutaneously to the interscapular region. Then the bladder can be filled at a controlled rate, while the urethra is left free for micturition. During the repetitive cycles of filling and voiding, intravesical pressure can be measured via the implanted catheter. As such, the pressure changes can be quantified and analyzed. Moreover, simultaneous measurement of the voided volume allows distinguishing voiding contractions from non-voiding contractions3.
	Importantly, due to the differences in micturition control between rodents and humans, cystometric measurements in these animals have only limited translational value4. Nevertheless, they are quite instrumental in the study of bladder pathophysiology and pharmacology in experimental pre-clinical settings. Recent research using this technique has revealed the key role of novel molecular players in the mechano- and chemo-sensory properties of the bladder.

Cystometry is an efficient technique to measure bladder function of small animals in vivo. The bladder is continuously infused at rates controlled through an intravesical catheter, whereas the urethra is left free for micturition. This allows for repetitive filling and emptying of the bladder, while intravesical pressure and voided volume are recorded.

	The lower urinary tract (LUT) functions as a  dynamic reservoir that is able to store urine and to efficiently expel it at a  convenient time. While ...

Collection Protocol for Human Pancreas

This dissection and sampling procedure was developed for the Network for Pancreatic Organ Donors with Diabetes (nPOD) program to standardize preparation of pancreas recovered from cadaveric organ donors. The pancreas is divided into 3 main regions (head, body, tail) followed by serial transverse sections throughout the medial to lateral axis. Alternating sections are used for fixed paraffin and fresh frozen blocks and remnant samples are minced for snap frozen sample preparations, either with or without RNAse inhibitors, for DNA, RNA, or protein isolation. The overall goal of the pancreas dissection procedure is to sample the entire pancreas while maintaining anatomical orientation.
Endocrine cell heterogeneity in terms of islet composition, size, and numbers is reported for human islets compared to rodent islets 1. The majority of human islets from the pancreas head, body and tail regions are composed of insulin-containing &beta;-cells followed by lower proportions of glucagon-containing &alpha;-cells and somatostatin-containing &delta;-cells. Pancreatic polypeptide-containing PP cells and ghrelin-containing epsilon cells are also present but in small numbers. In contrast, the uncinate region contains islets that are primarily composed of pancreatic polypeptide-containing PP cells 2. These regional islet variations arise from developmental differences. The pancreas develops from the ventral and dorsal pancreatic buds in the foregut and after rotation of the stomach and duodenum, the ventral lobe moves and fuses with the dorsal 3. The ventral lobe forms the posterior portion of the head including the uncinate process while the dorsal lobe gives rise to the rest of the organ. Regional pancreatic variation is also reported with the tail region having higher islet density compared to other regions and the dorsal lobe-derived components undergoing selective atrophy in type 1 diabetes4,5.
Additional organs and tissues are often recovered from the organ donors and include pancreatic lymph nodes, spleen and non-pancreatic lymph nodes. These samples are recovered with similar formats as for the pancreas with the addition of isolation of cryopreserved cells. When the proximal duodenum is included with the pancreas, duodenal mucosa may be collected for paraffin and frozen blocks and minced snap frozen preparations.

This video demonstrates a dissection procedure for processing human pancreas into multiple storage formats. Anatomical orientation is maintained throughout the pancreatic regions to allow definition of regional islet composition and density.

This dissection and sampling procedure was developed for the Network for Pancreatic Organ Donors with Diabetes (nPOD) program to standardize preparation ...

Parabiosis in Mice: A Detailed Protocol

Parabiosis is a surgical union of two organisms allowing sharing of the blood circulation. Attaching the skin of two animals promotes formation of microvasculature at the site of inflammation. Parabiotic partners share their circulating antigens and thus are free of adverse immune reaction. First described by Paul Bert in 18641, the parabiosis surgery was refined by Bunster and Meyer in 1933 to improve animal survival2. In the current protocol, two mice are surgically joined following a modification of the Bunster and Meyer technique. Animals are connected through the elbow and knee joints followed by attachment of the skin allowing firm support that prevents strain on the sutured skin. Herein, we describe in detail the parabiotic joining of a ubiquitous GFP expressing mouse to a wild type (WT) mouse. Two weeks after the procedure, the pair is separated and GFP positive cells can be detected by flow cytometric analysis in the blood circulation of the WT mouse. The blood chimerism allows one to examine the contribution of the circulating cells from one animal in the other.

Parabiotic joining of two organisms leads to the development of a shared circulatory system. In this protocol, we describe the surgical steps to form a parabiotic connection between a wild-type mouse and a constitutive GFP-expressing mouse.

Parabiosis is a surgical union of two organisms allowing sharing of the blood circulation. Attaching the skin of two animals promotes formation of ...

Cell-based Assay Protocol for the Prognostic Prediction of Idiopathic Scoliosis Using Cellular Dielectric Spectroscopy

This protocol details the experimental and analytical procedure for a cell-based assay developed in our laboratory as a functional test to predict the prognosis of idiopathic scoliosis in asymptomatic and affected children. The assay consists of the evaluation of the functional status of Gi and Gs proteins in peripheral blood mononuclear cells (PBMCs) by cellular dielectric spectroscopy (CDS), using an automated CDS-based instrument, and the classification of children into three functional groups (FG1, FG2, FG3) with respect to the profile of imbalance between the degree of response to Gi and Gs proteins stimulation. The classification is further confirmed by the differential effect of osteopontin (OPN) on response to Gi stimulation among groups and the severe progression of disease is referenced by FG2. Approximately, a volume of 10 ml of blood is required to extract PBMCs by Ficoll-gradient and cells are then stored in liquid nitrogen. The adequate number of PBMCs to perform the assay is obtained after two days of cell culture. Essentially, cells are first incubated with phytohemmaglutinin (PHA). After 24 hr incubation, medium is replaced by a PHA-free culture medium for an additional 24 hr prior to cell seeding and OPN treatment. Cells are then spectroscopically screened for their responses to somatostatin and isoproterenol, which respectively activate Gi and Gs proteins through their cognate receptors. Both somatostatin and isoproterenol are simultaneously injected with an integrated fluidics system and the cells&#39; responses are monitored for 15 min. The assay can be performed with fresh or frozen PBMCs and the procedure is completed within 4 days.

A structured protocol is presented for a cell-based assay as a functional test to predict the prognosis of idiopathic scoliosis using cellular dielectric spectroscopy (CDS). The assay can be performed with fresh or frozen peripheral blood mononuclear cells (PBMCs) and the procedure is completed within 4 days.

This protocol details the experimental and analytical procedure for a cell-based assay developed in our laboratory as a functional test to predict the ...

Sequential In vivo Imaging of Osteogenic Stem/Progenitor Cells During Fracture Repair

Bone turns over continuously and is highly regenerative following injury. Osteogenic stem/progenitor cells have long been hypothesized to exist, but in vivo demonstration of such cells has only recently been attained. Here, in vivo imaging techniques to investigate the role of endogenous osteogenic stem/progenitor cells (OSPCs) and their progeny in bone repair are provided. Using osteo-lineage cell tracing models and intravital imaging of induced microfractures in calvarial bone, OSPCs can be directly observed during the first few days after injury, in which critical events in the early repair process occur. Injury sites can be sequentially imaged revealing that OSPCs relocate to the injury, increase in number and differentiate into bone forming osteoblasts. These methods offer a means of investigating the role of stem cell-intrinsic and extrinsic molecular regulators for bone regeneration and repair.

Sequential In vivo Imaging of Osteogenic Stem/Progenitor Cells During Fracture Repair

Quantitative measurement of bone progenitor function in fracture healing requires high resolution serial imaging technology. Here, protocols are provided for using intravital microscopy and osteo-lineage tracking to sequentially image and quantify the migration, proliferation and differentiation of endogenous osteogenic stem/progenitor cells in the process of repairing bone fracture.

Bone turns over continuously and is highly regenerative following injury. Osteogenic stem/progenitor cells have long been hypothesized to exist, but in ...

Catheterization of the Carotid Artery and Jugular Vein to Perform Hemodynamic Measures, Infusions and Blood Sampling in a Conscious Rat Model

The success of a small animal model to study critical illness is, in part, dependent on the ability of the model to simulate the human condition. Intra-tracheal inoculation of a known amount of bacteria has been successfully used to reproduce the pathogenesis of pneumonia which then develops into sepsis. Monitoring hemodynamic parameters and providing standard clinical treatment including infusion of antibiotics, fluids and drugs to maintain blood pressure is critical to simulate routine supportive care in this model but to do so requires both arterial and venous vascular access. The video details the surgical technique for implanting carotid artery and common jugular vein catheters in an anesthetized rat. Following a 72 hr recovery period, the animals will be re-anesthetized and connected to a tether and swivel setup attached to the rodent housing which connects the implanted catheters to the hemodynamic monitoring system. This setup allows free movement of the rat during the study while continuously monitoring pressures, infusing fluids and drugs (antibiotics, vasopressors) and performing blood sampling.

Vascular accesses to measure hemodynamics, provide fluids and perform blood sampling are important to any small animal model study. We present a technique for implanting catheters into the carotid artery and the common jugular vein in an anesthetized rat for connecting to a system to perform monitoring, infusions and sampling.

The success of a small animal model to study critical illness is, in part, dependent on the ability of the model to simulate the human condition. ...

Nerve-sparing Mid-urethral Obstruction (NeMO) in Female Small Rodents

Partial bladder outlet obstruction (pBOO) has a high prevalence, causes significant patient burden, and immense health care costs. The most common animal model to investigate bladder remodeling in pBOO are female rodents undergoing partial obstruction at the proximal urethra. Variability in the degree of obstruction and animal mortality are major concerns with proximal obstruction. Furthermore, dissecting around the proximal urethra and bladder neck jeopardizes bladder innervation.
We developed a nerve-sparing mid-urethral obstruction (NeMO) model for pBOO avoiding the disadvantages of the traditional model. We approached the urethra just inferior to the pubic symphysis, which obviated the need for laparotomy as well as for dissection in this area; also, the striated urethral sphincter remained untouched. We performed NeMO in female Sprague-Dawley rats (12 obstructions, 6 sham animals) as well as in female C57/bl6 mice (20 obstructions, 18 sham animals). After two weeks, we evaluated bladder function, bladder mass, and body mass.
We had no mortalities among obstructed- or sham-operated female rats; as described for the traditional proximal pBOO-method, we tied the suture around the proximal urethra and a temporarily placed 0.9 mm metal rod. NeMO induced an 85% increase in bladder mass after two weeks, average residual urine volume was 0.4 mL in partially obstructed rats while only 0.03 mL in sham animals. In mice, we tested 3 sizes of cannulas that we placed along the urethra when tying the suture. We found that using a 27-gauge cannula resulted in over 50% animal mortality; placing the 25-gauge cannula did not yield the desired response in increasing bladder mass; utilizing a 26-gauge cannula yielded favorable results with minimal animal mortality (1/8) yet a significant 2-fold increase in bladder mass.

Nerve-sparing Mid-urethral Obstruction NeMO in Female Small Rodents

Traditional modeling of partial bladder outlet obstruction in rodents is fraught with animal mortality. A denervation injury from dissection around the proximal urethra and bladder neck is also of major concern. We developed and evaluated a safe and reliable mid-urethral obstruction model, avoiding the shortcomings of the traditional model.

Partial bladder outlet obstruction (pBOO) has a high prevalence, causes significant patient burden, and immense health care costs. The most common animal ...

In Vivo Evaluation of Fracture Callus Development During Bone Healing in Mice Using an MRI-compatible Osteosynthesis Device for the Mouse Femur

Endochondral fracture healing is a complex process involving the development of fibrous, cartilaginous, and osseous tissue in the fracture callus. The amount of the different tissues in the callus provides important information on the fracture healing progress. Available in vivo techniques to longitudinally monitor the callus tissue development in preclinical fracture-healing studies using small animals include digital radiography and &#181;CT imaging. However, both techniques are only able to distinguish between mineralized and non-mineralized tissue. Consequently, it is impossible to discriminate cartilage from fibrous tissue. In contrast, magnetic resonance imaging (MRI) visualizes anatomical structures based on their water content and might therefore be able to noninvasively identify soft tissue and cartilage in the fracture callus. Here, we report the use of an MRI-compatible external fixator for the mouse femur to allow MRI scans during bone regeneration in mice. The experiments demonstrated that the fixator and a custom-made mounting device allow repetitive MRI scans, thus enabling longitudinal analysis of fracture-callus tissue development.

In Vivo Evaluation of Fracture Callus Development During Bone Healing in Mice Using an MRI-compatible Osteosynthesis Device for the Mouse Femur

The evaluation of tissue development in the fracture callus during endochondral bone healing is essential to monitor the healing process. Here, we report the use of a magnetic resonance imaging (MRI)-compatible external fixator for the mouse femur to allow MRI scans during bone regeneration in mice.

Endochondral fracture healing is a complex process involving the development of fibrous, cartilaginous, and osseous tissue in the fracture callus. The ...

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 ...