The study was supported by the NIH P30AR069655 and R01AR070613 (H. A. A.).

Animal studies were approved by The University of Rochester Committee of Animal Resources. The mice used in this study were C57BL/6 males and females ranging from age 24-29 weeks of age. Mice were housed in standard conditions with food and water ad libitum. Upon euthanasia via carbon dioxide inhalation, followed by cervical dislocation, 20 right femurs (10 male and 10 female) were harvested and frozen at -20 &#176;C until tested.
1. Creation of custom 3D printed mounting guides
NOTE: This step might be needed because different strains and genetic phenotypes might have different anatomical geometries.
<ol>
	<li>Obtain &#181;CT scans of the representative samples.
	<ol>
		<li>Scan representative samples on a &#181;CT scanner with the following settings: 55 kV, 145 &#181;A for 300 ms integration times, and resolution of 10.5 &#181;m voxels.</li>
		<li>Ensure that the captured region covers the proximal end of the femur and continues down through the midshaft. 
		NOTE: If a &#181;CT scanner is unavailable, 2D planar X-rays of the representative samples can be used.</li>
	</ol>
	</li>
	<li>Analyze the &#181;CT scans.
	<ol>
		<li>Using the representative set of &#181;CT scans, obtain a 2D rendering of the anterior view of the proximal femur.
		<ol>
			<li>Obtain &#181;CT images with a resolution of 10.5 &#181;m voxels from the midshaft to the proximal end of the femur. Compile these slices using software (see Table of Materials) into a 3D rendering of the sample.</li>
			<li>Determine a threshold to distinguish bone from the surrounding tissue and apply a Gaussian filter for noise reduction.</li>
			<li>Orient the 3D renderings to eliminate off-axis tilt and ensure that the femur&#39;s anterior surface is viewed.</li>
			<li>Export this 2D view of the 3D rendering as an image file, such as .jpg or .png.</li>
		</ol>
		</li>
		<li>Using an image analysis software (see Table of Materials), measure the femoral shaft angle by drawing a line perpendicular to the femoral shaft 7 mm distally and a second line through the peak of the greater trochanter to the midpoint of the aforementioned perpendicular line (Figure 1).</li>
		<li>Along the 7 mm distal perpendicular line, measure the femoral shaft diameter below the third trochanter.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63394/63394fig01.jpg" /> 
Figure 1: &#181;CT analysis. &#181;CT images of femurs of C57Bl/6 mice are used to calculate the average shaft angle, measured from the top of the greater trochanter through the center of the midshaft, ~7 mm distally. The midshaft diameter was also measured at this position. The 3D renderings of the proximal femur were oriented in an anterior view to display the profile of the third trochanter. The average shaft angle was 93.13&#176; (SD = 1.19&#176;), and the average midshaft diameter was 1.53 mm (SD = 0.14 mm) (n = 20). Scale bar = 1 mm. <a href="https://www.jove.com/files/ftp_upload/63394/63394fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="3">
	<li>Create the mounting guides using a 3D modeling software program (see Table of Materials) (Figure 2, Supplementary File 1). 
	NOTE: The guides are rectangular cuboids measuring 6.25 mm x 3.25 mm x 7mm with an angled slot, slightly larger than the average shaft diameter determined in step 1.1.2. The angle of the slot will create a consistent angle of 20&#176; from vertical. The guides should be consistent in length, height and width, but can be made with various slot diameters to accommodate anatomical differences among the bone samples.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63394/63394fig02.jpg" /> 
Figure 2: Designing the guides. (A) 3D sketch and (B) visualization of midshaft angling fixture before 3D printing. Based on previous literature, a midshaft angle between 20&#176; maximizes the stiffness. It minimizes the maximal bending moment in the femoral shaft to ensure fractures occur in the neck and variability in mechanical outcomes5. To compensate for the 3.13&#176; deviation from perpendicular in the midshaft average angles, the fixture angle was set to 73.13&#176; to produce an angle of 20&#176;. Alignment fixtures were printed with diameters ranging from 1.9-2.2 mm to ensure a proper fit for varying midshaft diameters. <a href="https://www.jove.com/files/ftp_upload/63394/63394fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="4">
	<li>Using a 3D printer, print the guides. The guides can remain on during the testing process, so printing multiple replicates of the guides can be beneficial for preparing multiple samples at once.</li>
</ol>
2. Sample preparation
<ol>
	<li>Harvest the mouse femurs by making a transverse incision entirely around the mouse abdomen and removing the tissue from the incision to the ankles. Following this, locate the hip socket and carefully use the tip of a pair of fine forceps to dislocate the hip. Cut the additional soft tissue to remove the leg from the mouse.</li>
	<li>Once the leg is harvested, use a scalpel to dislocate and cut through the knee joint. Manually clean the femurs of all soft tissue using forceps, scalpels, and paper towels.</li>
	<li>Test the harvested samples immediately or store them at -20 &#176;C for up to 6 months. If samples are frozen, allow them to come to room temperature and hydrate in PBS for 2 h before prepping.</li>
	<li>Using &#188;&#34; x &#188;&#34; square aluminum tubing (see Table of Materials), cut tubing sections &#189;&#34; to 1&#34; in length. Using an etching tool, label each aluminum segment with the sample IDs.</li>
	<li>Fill half of the tubing segments with putty. Place these tubing segments into a fixture to hold them upright.</li>
	<li>Place the cleaned femurs into the 3D printed guides. To do this, place the samples flat on the benchtop so the anterior surface is facing up. Place the guide directly below the third trochanter, where the shaft diameter becomes more consistent. 
	NOTE: This will leave ~7mm of the proximal femur above the guide.</li>
	<li>To prevent the femur from rotating to the lateral or medial side while placing on the guide, hold the proximal and distal ends with one hand when applying the guides, firmly press the femur onto the workbench and using your other hand, place the 3D printed guide on the midshaft of the femur. Ensure to apply the appropriate diameter guide gently, as the midshaft of the femur can snap if forced into a guide too small.</li>
	<li>Once the guides are on the femurs, place them in front of the corresponding aluminum segments. Using bone cement or other hardening agents, fill the aluminum segments until just full, leaving a little room for displacement.</li>
	<li>Place the femurs with guides on in the correct aluminum segment. 
	NOTE: The guides will not be centered on the aluminum segments, seated slightly to one side to allow the distal end of the femur to sit in the center of the aluminum pot.</li>
	<li>Allow the hardening agent to set. Once set, place the samples in a Petri dish with room temperature phosphate-buffered saline (PBS) and allow to rehydrate for 2 h (Figure 3).</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63394/63394fig03.jpg" /> 
Figure 3: Sample preparation using custom jigs and angling fixtures. (A) Samples in aluminum pots with the proper alignment are maintained using the 3D printed guides while the bone cement is drying. (B) X-ray before testing shows the shadow of angling fixtures and complete coverage of bone cement surrounding the distal end of femurs. The saturated white area at the bottom of the aluminum pots is putty, used to keep bone cement in pots when hardening. Scale bar (Panel B) = 5 mm. <a href="https://www.jove.com/files/ftp_upload/63394/63394fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Hardware set-up
<ol>
	<li>Using a mechanical testing system (MTS), attach and calibrate a load cell with resolution &#60;1 N (see Table of Materials) (Figure 4A). 
	NOTE: The load cell can be mounted on the stage or, preferably, the actuator when possible.</li>
	<li>Attach a fixture with a square slot that will firmly hold the aluminum segments with the samples. Attach set screws to the two sides of the holding fixture to firmly hold samples in place. (Figure 4B). 
	NOTE: This fixture can be 3D printed or machined and then tapped with threaded screw holes to mount to the testing frame.</li>
	<li>Attach a loading platen to the actuator. This can be simply a tapered screw with a flattened tip (Figure 4C).</li>
	<li>Place a stereomicroscope on a table or surface directly in front of the MTS. If additional lighting is needed to see the set-up through the microscope, place these around the system.</li>
</ol>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63394/63394fig04.jpg" /> 
Figure 4: Hardware set-up. (A) Set-up of testing on mechanical testing system, with 1 kN load cell (resolution &#60; 1 N) and black biaxial stage to ensure proper sample positioning. (B) Close up of the 3D printed mounting fixture attached to the load cell with an M10 threaded rod and two M4 bolts used to hold the aluminum pot in place. (C) View of the sample through a stereo microscope with a tapered loading fixture. Scale bar (Panel C) = 5 mm. <a href="https://www.jove.com/files/ftp_upload/63394/63394fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Software set-up
<ol>
	<li>In the MTS software, begin the creation of a new flexural (bending) protocol. Ensure that the protocol will operate in displacement control.</li>
	<li>Set the loading rate of the protocol to 0.5 mm/s.</li>
	<li>If the software has a setting for soft keys, add the soft keys &#34;Balance&#34; and &#34;Zero Extension&#34; to the protocol. 
	NOTE: This will quickly set the load and actuator position to 0 before testing each sample.</li>
	<li>Ensure that the software program will record the time in seconds, load in Newtons, and extension or displacement in millimeters at a minimum sampling rate of 100 Hz.</li>
	<li>Save the new protocol and return to the main screen of the software program to begin testing a new sample set.</li>
</ol>
5. Testing set-up
<ol>
	<li>Before mounting the specimens on the MTS, obtain an X-ray image of the samples in the aluminum pots. Multiple samples can be imaged at once. Ensure that the anterior view of samples is captured to allow for verification measurements of potting angle (Figure 5).</li>
</ol>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63394/63394fig05.jpg" /> 
Figure 5. Assessment of sample alignment. (A) The shaft angle from vertical is measured from planar digital x-rays. (B) Representative potted femoral shaft angles ranged from 18.11&#176; to 23.99&#176;, with a coefficient of variation (COV) of 7.1% (n = 20). Sex differences due to anatomical variations were not statistically significant, as determined using a one-tailed unpaired t-test (p &#60; 0.05). Scale bar (Panel A) = 1 mm. <a href="https://www.jove.com/files/ftp_upload/63394/63394fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="2">
	<li>Place the aluminum segment with sample into the holding fixture and tighten set screws.</li>
	<li>Lower actuator/loading platen until it is within a few millimeters of the femoral head. 
	NOTE: Do not preload the sample with any force and be careful not to lower the actuator too quickly, as it is very easy to damage samples.</li>
	<li>Using the stereomicroscope, adjust the biaxial stage to align the position of the femoral head directly underneath the loading platen. Lock biaxial stage in place.</li>
	<li>In the MTS software, zero the position of the actuator and balance the load cell using the soft keys added in step 4.3.</li>
	<li>Begin the loading protocol. Depending on how much space was left between the loading platen and sample, testing will only take 10-30 s.</li>
	<li>After testing, capture another anterior X-ray of the sample. This will be used to discern and document the mode of fracture (Figure 6).</li>
</ol>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/63394/63394fig06.jpg" /> 
Figure 6: X-ray image of samples after testing. All samples fractured in a bifurcated line through the femoral neck and along the femoral neck-shaft attachment (highlighted by the orange circle). Scale bar = 1 mm. <a href="https://www.jove.com/files/ftp_upload/63394/63394fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
6. Data analysis
<ol>
	<li>After data collection, export force and displacement data into software (see Table of Materials) that allows for graphing and mathematical calculations.</li>
	<li>Plot the load vs. displacement for each sample (Figure 7A). Fit a linear approximation to the linear segment of the load-displacement curve. The slope of this linear fit will define the stiffness, a measure of the elasticity of the sample.</li>
	<li>Calculate the additional outcomes such as maximum load, maximum displacement, yield load, displacement at yield point, energy to maximum load, and energy to yield point. 
	NOTE: The yield point can be determined by off-setting the linear approximation determined in step 6.2 by 0.2%6. The point at which the off-set line and the load vs. displacement curve intersect will determine the yield point. In the case of very brittle samples that show little yield, the yield point may be the same as the maximum point.</li>
</ol>

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	<li>Reports of the Surgeon General. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Health+and+Osteoporosis:+A+Report+of+the+Surgeon+General.">Health and Osteoporosis: A Report of the Surgeon General.</a> Reports of the Surgeon General. , (2004).</li><li>Veronese, N., Maggi, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Epidemiology+and+social+costs+of+hip+fracture.">Epidemiology and social costs of hip fracture.</a> Injury. 49 (8), 1458-1460 (2018).</li><li>Gurumurthy, C. B., Lloyd, K. C. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generating+mouse+models+for+biomedical+research:+Technological+advances.">Generating mouse models for biomedical research: Technological advances.</a> Disease Models and Mechanisms. 12 (1), 029462 (2019).</li><li>Boymans, T. A. E. J., Veldman, H. D., Noble, P. C., Heyligers, I. C., Grimm, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+femoral+head+center+shifts+in+a+mediocaudal+direction+during+aging.">The femoral head center shifts in a mediocaudal direction during aging.</a> Journal of Arthroplasty. 2 (32), 581-586 (2017).</li><li>Voide, R., van Lenthe, G. H., Muller, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Femoral+stiffness+and+strength+critically+depend+on+loading+angle:+A+parametric+study+in+a+mouse-inbred+strain.">Femoral stiffness and strength critically depend on loading angle: A parametric study in a mouse-inbred strain.</a> Biomedical Engineering. 53 (3), 122-129 (2008).</li><li>CRC Press. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Bone Mechanics Handbook. Second end. , (2001).</li><li>Middleton, K. M., 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=The+relative+importance+of+genetics+and+phenotypic+plasticity+in+dictating+bone+morphology+and+mechanics+in+aged+mice:+evidence+from+an+artificial+selection+experiment.">The relative importance of genetics and phenotypic plasticity in dictating bone morphology and mechanics in aged mice: evidence from an artificial selection experiment.</a> Zoology (Jena). 111 (2), 135-147 (2008).</li><li>Jamsa, T., Koivukangas, A., Ryhanen, J., Jalovaara, P., Tuukkanen, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Femoral+neck+is+a+sensitive+indicator+of+bone+loss+in+immobilized+hind+limb+of+mouse.">Femoral neck is a sensitive indicator of bone loss in immobilized hind limb of mouse.</a> Journal of Bone and Mineral Research. 14 (10), 1708-1713 (1999).</li><li>Kamal, B., 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=Biomechanical+properties+of+bone+in+a+mouse+model+of+Rett+syndrome.">Biomechanical properties of bone in a mouse model of Rett syndrome.</a> Bone. 71, 106-114 (2015).</li><li>Jamsa, T., Tuukkanen, J., Jalovaara, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Femoral+neck+strength+of+mouse+in+two+loading+configurations:+Methods+evaluation+and+fracture+characteristics.">Femoral neck strength of mouse in two loading configurations: Methods evaluation and fracture characteristics.</a> Journal of Biomechanics. 31 (8), 723-729 (1998).</li><li>Brent, M. B., Bruel, A., Thomsen, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=PTH+(1-34)+and+growth+hormone+in+prevention+of+disuse+osteopenia+andsarcopenia+in+rats.">PTH (1-34) and growth hormone in prevention of disuse osteopenia andsarcopenia in rats.</a> Bone. 110, 244-253 (2018).</li><li>Bromer, F. D., Brent, M. B., Pedersen, M., Thomsen, J. S., Bruel, A., Foldager, C. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effect+of+normobaric+intermittent+hypoxia+therapy+on+bone+in+normal+and+disuse+osteopenic+mice.">The effect of normobaric intermittent hypoxia therapy on bone in normal and disuse osteopenic mice.</a> High Altitude Medicine and Biology. 22 (2), 225-234 (2021).</li><li>Vegger, J. B., Bruel, A., Brent, M. B., Thomsen, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Disuse+osteopenia+induced+by+botulinum+toxin+is+similar+in+skeletally+mature+young+and+aged+female+C57BL/6J+mice.">Disuse osteopenia induced by botulinum toxin is similar in skeletally mature young and aged female C57BL/6J mice.</a> Journal of Bone and Mineral Metabolism. 36, 170-179 (2018).</li><li>Lodberg, A., Vegger, J. B., Jensen, M. V., Larsen, C. M., Thomsen, J. S., Bruel, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immobilization+induced+osteopenia+is+strain+specific+in+mice.">Immobilization induced osteopenia is strain specific in mice.</a> Bone Reports. 2, 59-67 (2015).</li><li>Varacallo, M. A., Fox, E. 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The authors have nothing to disclose.

This protocol outlines a reliable cantilever bending test for murine femoral necks. The natural cantilever flexure scenario that occurs at the femoral neck is typically not represented in standard 3- and 4-point bending tests5. This testing method is better and more reliably replicates the type of femoral neck fractures experienced by bone fragility patients. The main focus when performing this protocol is eliminating the variability due to inconsistent potting of the femoral shaft. Critically, cl...

Fracture risk is a serious medical concern associated with osteoporosis. Over 1.5 million fragility fractures are reported each year in the United States alone, with fractures occurring in the hip, specifically the femoral neck, as the leading fracture type1. It is estimated that 18% of women and 6% of men will experience a femoral neck fracture in their lifetime2, and the mortality rate at 1 year following the fracture is greater than 20%1. Therefore, mouse models that allow biomechanical testing of the femoral neck can be suitable for studying fragility fractures. Mouse models also offer powerful tools to elucidate translatable cellular and molecular events involved in osteoporosis potentially. This is due to the availability of genetic reporters, gain and loss of function models, and the expansive library of molecular techniques and reagents. Mechanical testing of mouse bones can provide necessary outcome measures to determine bone health, genotypic and phenotypic variations that could explain the etiology of the disease, and assess therapies based on outcome measures of the quality of the bone and the risk of fracture3.
The anatomy of the femoral neck creates unique mechanical loading scenarios, which typically lead to flexural (bending) fractures. The femoral head is loaded in the acetabular socket at the proximal end of the femur. This creates a cantilever bending scenario on the femoral neck, which is rigidly attached to the femoral shaft distally4. This differs from traditional 3- or 4-point bending tests on the femoral mid-diaphysis. While these tests are helpful, they do not replicate the loading that typically leads to fragility fractures in osteopenic and osteoporotic individuals in terms of fracture location or the loading scenario.
To better assess fragility fracture risk in mice, it was sought to improve the reproducibility of cantilever bending tests of murine femoral necks. As theoretically predicted, the loading angle on the femoral head relative to the femoral shaft has been shown to significantly affect the outcome measures5, thereby creating a challenge for reliability and reproducibility of reported outcomes. To ensure proper and consistent alignment of the femurs during sample preparation, guides were designed, and 3D printed based on anatomic measurements made on &#181;CT scans of C57BL/6 mouse femurs. The guides were designed to aid in consistently potting the samples such that the femoral shaft is maintained at ~20&#176; from the vertical loading direction. This angle was chosen because it maximizes the stiffness while minimizing the maximal bending moment along the femoral shaft, which increases the likelihood of femoral neck fractures and leads to more consistent and reproducible testing5. Guides were 3D printed in various sizes to accommodate anatomical differences between samples and used to hold samples in a stable position while potting in acrylic bone cement. The stiffness, maximum force, yield force, and maximum energy were calculated from the force-displacement graphs. This testing method showed consistent results for the aforementioned biomechanical outcome. With practice and the aid of the 3D printed guide, measurement errors due to misalignment can be minimized, resulting in reliable outcome measures.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>&#188;&#8221; x &#188;&#8221; square aluminum tubing</td>
			<td>Grainger</td>
			<td>48KU67</td>
			<td>Cut to lengths of 1/2&#34; to 1&#34; lengths</td>
		</tr>
		<tr>
			<td>1 kN load cell</td>
			<td>Instron</td>
			<td>2527-130</td>
			<td>Any load cell with sub 1 N resolution can be used.</td>
		</tr>
		<tr>
			<td>3.5x-45x Zoom Stereo Boom Microscope</td>
			<td>Omano</td>
			<td>OM2300S-GX4</td>
			<td>Microscope used to precisely line up samples with loading platen.</td>
		</tr>
		<tr>
			<td>3D printed guides</td>
			<td>Custom made</td>
			<td></td>
			<td>Angled slots at 73.13&#176;, with diameters between 1.9 mm and 2.2 mm</td>
		</tr>
		<tr>
			<td>3D printed mount</td>
			<td>Custom made</td>
			<td></td>
			<td>Tapped with M10 threads to fit the mount attachment and with 2 M4 threaded holes adjacent sides to hold the aluminum tubing with sample in place.</td>
		</tr>
		<tr>
			<td>Acrylic Base Plate Material Kit</td>
			<td>Keystone Industries</td>
			<td>921392</td>
			<td>Mix 3.5 g of powder with 2 mL of liquid. This will be enough for approximately 8 samples, and will begin to harden quickly.</td>
		</tr>
		<tr>
			<td>Amira</td>
			<td>ThermoFisher Scientific</td>
			<td></td>
			<td>Used to compile &#181;CT scans</td>
		</tr>
		<tr>
			<td>Biaxial stage</td>
			<td>Custom made</td>
			<td></td>
			<td>Used to center femoral head of sample under the loading platen.</td>
		</tr>
		<tr>
			<td>BioMed Amber Resin</td>
			<td>formlabs</td>
			<td>RS-F2-BMAM-01</td>
			<td>Any resin from formlabs could be used for this project.</td>
		</tr>
		<tr>
			<td>Bluehill 3</td>
			<td>Instron</td>
			<td>V3.66</td>
			<td>Software used to set up loading protocol and collect load, displacement and time data.</td>
		</tr>
		<tr>
			<td>ElectroPuls 10000</td>
			<td>Instron</td>
			<td>E10000</td>
			<td>Mechanical testing system</td>
		</tr>
		<tr>
			<td>Faxitron UltraFocus</td>
			<td>Faxitron BioOptics</td>
			<td>2327A40311</td>
			<td>X-ray imaging system</td>
		</tr>
		<tr>
			<td>Form 2</td>
			<td>formlabs</td>
			<td>F2</td>
			<td>Used to print the mount and guides</td>
		</tr>
		<tr>
			<td>Form 2 Resin Tank LT</td>
			<td>formlabs</td>
			<td>RT-F2-02</td>
			<td>LT Tank was used to be compatible with the BioMed Resin</td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td>National Institutes of Health</td>
			<td>ImageJ</td>
			<td>Used to assess &#181;CT and X-ray images</td>
		</tr>
		<tr>
			<td>Laxco iLED Series LED Light Source</td>
			<td>ThermoFisher Scientific</td>
			<td>AMPSILED30W</td>
			<td>Light source used in conjugtion with microscope.</td>
		</tr>
		<tr>
			<td>Loading platen</td>
			<td>Custom made</td>
			<td></td>
			<td>This can be any metal rod that is tapered to a diameter of approximately 2.5 mm. We used an M6 screw that was tapered on a lathe.</td>
		</tr>
		<tr>
			<td>Mount attachment</td>
			<td>Custom made</td>
			<td></td>
			<td>To secure the 3D printed mount to the load cell. We used a M10/M6 threaded rod</td>
		</tr>
		<tr>
			<td>Phosphate Buffer Saline (PBS)</td>
			<td>ThermoFisher Scientific</td>
			<td>10010031</td>
			<td>Need to rehydrate the samples once acrylic base plate material has set.</td>
		</tr>
		<tr>
			<td>Plumber&#39;s putty</td>
			<td>Oatey</td>
			<td>31174</td>
			<td>Used to seal the end of the aluminum tubing when pouring acrylic base plate material in. Any clay or putty could be used.</td>
		</tr>
		<tr>
			<td>PreForm</td>
			<td>formlabs</td>
			<td>Preform 3.15.2</td>
			<td>Formlabs software</td>
		</tr>
		<tr>
			<td>Tissue Culture Dish</td>
			<td>Corning</td>
			<td>353003</td>
			<td>Samples can be laid flat in culture dish and covered in PBS to rehydrate.</td>
		</tr>
		<tr>
			<td>vivaCT 40</td>
			<td>Scanco</td>
			<td>&#181;CT 40</td>
			<td>Representative set or actual samples can be scanned prior to printing of guides to calculate femoral shaft angle and diameter.</td>
		</tr>
	</tbody>
</table>

cantilever bending of murine femoral necks

Fractures in the femoral neck are a common occurrence in individuals with osteoporosis. Many mouse models have been developed to assess disease states and therapies, with biomechanical testing as a primary outcome measure. However, traditional biomechanical testing focuses on torsion or bending tests applied to the midshaft of the long bones. This is not typically the site of high-risk fractures in osteoporotic individuals. Therefore, a biomechanical testing protocol was developed that tests the femoral necks of murine femurs in cantilever bending loading to replicate better the types of fractures experienced by osteoporosis patients. Since the biomechanical outcomes are highly dependent on the flexural loading direction relative to the femoral neck, 3D printed guides were created to maintain a femoral shaft at an angle of 20&#176; relative to the loading direction. The new protocol streamlined the testing by reducing variability in alignment (21.6&#176; &plusmn; 1.5&#176;, COV = 7.1%, n = 20) and improved reproducibility in the measured biomechanical outcomes (average COV = 26.7%). The new approach using the 3D printed guides for reliable specimen alignment improves rigor and reproducibility by reducing the measurement errors due to specimen misalignment, which should minimize sample sizes in mouse studies of osteoporosis.

When potted with the aid of the guide, the femoral shafts were aligned at 21.6&#176; &#177; 1.5&#176;. While this represents &#60;10% deviation from the intended angle of 20&#176;, the coefficients of variation (COV) of the potting angle across all samples tested were 7.6% and 6.5% for male and female mice, respectively (n = 10 per group) as verified by pre-test planar x-rays (Figure 5). Additionally, the post-testing X-rays should be used to assess the mode in which the samples failed. Fail...

Watch this Scientific Journal Video about Cantilever Bending of Murine Femoral Necks at JoVE.com

Cantilever Bending of Murine Femoral Necks

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

Fractures in the femoral neck are a common occurrence in individuals with osteoporosis. Many mouse models have been developed to assess disease states and ...

cantilever-bending-of-murine-femoral-necks

Research

JoVE Journal

Bioengineering

2.0K Views.  University of Rochester Medical Center. The present protocol describes the development of a reproducible testing platform for murine femoral necks in a cantilever bending set-up. Custom 3D printed guides were used to consistently and rigidly fix the femurs in optimal alignment. 

Video: Cantilever Bending of Murine Femoral Necks

Tissue Engineering of the Intestine in a Murine Model

Tissue-engineered small intestine (TESI) has successfully been used to rescue Lewis rats after massive small bowel resection, resulting in return to preoperative weights within 40 days.1 In humans, massive small bowel resection can result in short bowel syndrome, a functional malabsorptive state that confers significant morbidity, mortality, and healthcare costs including parenteral nutrition dependence, liver failure and cirrhosis, and the need for multivisceral organ transplantation.2 In this paper, we describe and document our protocol for creating tissue-engineered intestine in a mouse model with a multicellular organoid units-on-scaffold approach. Organoid units are multicellular aggregates derived from the intestine that contain both mucosal and mesenchymal elements,3 the relationship between which preserves the intestinal stem cell niche.4 In ongoing and future research, the transition of our technique into the mouse will allow for investigation of the processes involved during TESI formation by utilizing the transgenic tools available in this species.5The availability of immunocompromised mouse strains will also permit us to apply the technique to human intestinal tissue and optimize the formation of human TESI as a mouse xenograft before its transition into humans. Our method employs good manufacturing practice (GMP) reagents and materials that have already been approved for use in human patients, and therefore offers a significant advantage over approaches that rely upon decellularized animal tissues. The ultimate goal of this method is its translation to humans as a regenerative medicine therapeutic strategy for short bowel syndrome.

This article and the accompanying video present our protocol for generating tissue-engineered intestine in the mouse, using an organoid units-on-scaffold approach.

Tissue-engineered small intestine (TESI) has successfully been used to rescue Lewis rats after massive small bowel resection, resulting in return to ...

Method and Instrumented Fixture for Femoral Fracture Testing in a Sideways Fall-on-the-Hip Position

Mechanical testing of femora brings valuable insights into understanding the contribution of clinically-measureable variables such as bone mineral density distribution and geometry on the femoral mechanical properties. Currently, there is no standard protocol for mechanical testing of such geometrically complex bones to measure strength, and stiffness. To address this gap we have developed a protocol to test cadaveric femora to fracture and to measure their biomechanical parameters. This protocol describes a set of adaptable fixtures to accommodate the various load magnitudes and directions accounting for possible bone orientations in a fall on the hip configuration, test speed, bone size, and left leg-right leg variations. The femora were prepared for testing by cleaning, cutting, scanning, and potting the distal end and greater trochanter contact surfaces in poly(methyl methacrylate) (PMMA) as presented in a different protocol. The prepared specimens were placed in the testing fixture in a position mimicking a sideways fall on the hip and loaded to fracture. During testing, two load cells measured vertical forces applied to the femoral head and greater trochanter, a six-axis load cell measured forces and moments at the distal femoral shaft, and a displacement sensor measured differential displacement between the femoral head and trochanter contact supports. High speed video cameras were used to synchronously record the sequence of fracture events during testing. The reduction of this data allowed us to characterize the strength, stiffness, and fracture energy for nearly 200 osteoporotic, osteopenic, and normal cadaveric femora for further development of engineering-based diagnostic tools for osteoporosis research.

In this manuscript, we present a protocol to fracture test cadaveric proximal femora in a sideways fall on the hip configuration using instrumented fixtures mounted on a standard servo hydraulic frame. Nine digitized signals comprising forces, moments, and displacement along with two high speed video streams are acquired during testing.

Mechanical testing of femora brings valuable insights into understanding the contribution of clinically-measureable variables such as bone mineral density ...

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and dampen loads in the spine. The IVD consists of a proteoglycan-rich nucleus pulposus (NP) and a collagen-rich annulus fibrosis (AF) sandwiched by cartilaginous end-plates. Together with the adjacent vertebrae, the vertebrae-IVD structure forms a functional spine unit (FSU). These microstructures contain unique cell types as well as unique extracellular matrices. Whole organ culture of the FSU preserves the native extracellular matrix, cell differentiation phenotypes, and cellular-matrix interactions. Thus, organ culture techniques are particularly useful for investigating the complex biological mechanisms of the IVD. Here, we describe a high-throughput approach for culturing whole lumbar mouse FSUs that provides an ideal platform for studying disease mechanisms and therapies for the IVD. Furthermore, we describe several applications that utilize this organ culture method to conduct further studies including contrast-enhanced microCT imaging and three-dimensional high-resolution finite element modeling of the IVD.

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Whole organ culture of the intervertebral disc (IVD) preserves the native extracellular matrix, cell phenotypes, and cellular-matrix interactions. Here we describe an IVD culture system using mouse lumbar and caudal IVDs in their functional spinal units and several applications utilizing this system.

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and ...

Isolation of Primary Murine Retinal Ganglion Cells (RGCs) by Flow Cytometry

Neurodegenerative diseases often have a devastating impact on those affected. Retinal ganglion cell (RGC) loss is implicated in an array of diseases, including diabetic retinopathy and glaucoma, in addition to normal aging. Despite their importance, RGCs have been extremely difficult to study until now due in part to the fact that they comprise only a small percentage of the wide variety of cells in the retina. In addition, current isolation methods use intracellular markers to identify RGCs, which produce non-viable cells. These techniques also involve lengthy isolation protocols, so there is a lack of practical, standardized, and dependable methods to obtain and isolate RGCs. This work describes an efficient, comprehensive, and reliable method to isolate primary RGCs from mice retinae using a protocol based on both positive and negative selection criteria. The presented methods allow for the future study of RGCs, with the goal of better understanding the major decline in visual acuity that results from the loss of functional RGCs in neurodegenerative diseases.

Isolation of Primary Murine Retinal Ganglion Cells RGCs by Flow Cytometry

Millions of people suffer from retinal degenerative diseases that result in irreversible blindness. A common element of many of these diseases is the loss of retinal ganglion cells (RGCs). This detailed protocol describes the isolation of primary murine RGCs by positive and negative selection with flow cytometry.

Neurodegenerative diseases often have a devastating impact on those affected. Retinal ganglion cell (RGC) loss is implicated in an array of diseases, ...

Retroductal Nanoparticle Injection to the Murine Submandibular Gland

Two common goals of salivary gland therapeutics are prevention and cure of tissue dysfunction following either autoimmune or radiation injury. By locally delivering bioactive compounds to the salivary glands, greater tissue concentrations can be safely achieved versus systemic administration. Furthermore, off target tissue effects from extra-glandular accumulation of material can be dramatically reduced. In this regard, retroductal injection is a widely used method for investigating both salivary gland biology and pathophysiology. Retroductal administration of growth factors, primary cells, adenoviral vectors, and small molecule drugs has been shown to support gland function in the setting of injury. We have previously shown the efficacy of a retroductally injected nanoparticle-siRNA strategy to maintain gland function following irradiation. Here, a highly effective and reproducible method to administer nanomaterials to the murine submandibular gland through Wharton's duct is detailed (Figure 1). We describe accessing the oral cavity and outline the steps necessary to cannulate Wharton's duct, with further observations serving as quality checks throughout the procedure.

Local drug delivery to the submandibular glands is of interest in understanding salivary gland biology and for the development of novel therapeutics. We present an updated and detailed retroductal injection protocol, designed to improve delivery accuracy and experimental reproducibility. The application presented herein is the delivery of polymeric nanoparticles.

Two common goals of salivary gland therapeutics are prevention and cure of tissue dysfunction following either autoimmune or radiation injury. By locally ...

Light-sheet Fluorescence Microscopy for the Study of the Murine Heart

Light-sheet fluorescence microscopy has been widely used for rapid image acquisition with a high axial resolution from micrometer to millimeter scale. Traditional light-sheet techniques involve the use of a single illumination beam directed orthogonally at sample tissue. Images of large samples that are produced using a single illumination beam contain stripes or artifacts and suffer from a reduced resolution due to the scattering and absorption of light by the tissue. This study uses a dual-sided illumination beam and a simplified CLARITY optical clearing technique for the murine heart. These techniques allow for deeper imaging by removing lipids from the heart and produce a large field of imaging, greater than 10 x 10 x 10 mm3. As a result, this strategy enables us to quantify the ventricular dimensions, track the cardiac lineage, and localize the spatial distribution of cardiac-specific proteins and ion-channels from the post-natal to adult mouse hearts with sufficient contrast and resolution.

This study uses a dual-sided illumination light-sheet fluorescence microscopy (LSFM) technique combined with optical clearing to study the murine heart.

Light-sheet fluorescence microscopy has been widely used for rapid image acquisition with a high axial resolution from micrometer to millimeter scale. ...

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

Orthopedic research relies heavily on animal models to study mechanisms of bone healing in vivo as well as investigate the new treatment techniques. Critical-sized segmental defects are challenging to treat clinically, and research efforts could benefit from a reliable, ambulatory small animal model of a segmental femoral defect. In this study, we present an optimized surgical protocol for the consistent and reproducible creation of a 5 mm critical diaphyseal defect in a rat femur stabilized with an external fixator. The diaphyseal ostectomy was performed using a custom jig to place 4 Kirschner wires bicortically, which were stabilized with an adapted external fixator device. An oscillating bone saw was used to create the defect. Either a collagen sponge alone or a collagen sponge soaked in rhBMP-2 was implanted into the defect, and the bone healing was monitored over 12 weeks using radiographs. After 12 weeks, rats were sacrificed, and histological analysis was performed on the excised control and treated femurs. Bone defects containing only collagen sponge resulted in non-union, while rhBMP-2 treatment yielded the formation of a periosteal callous and new bone remodeling. Animals recovered well after implantation, and external fixation proved successful in stabilizing the femoral defects over 12 weeks. This streamlined surgical model could be readily applied to study bone healing and test new orthopedic biomaterials and regenerative therapies in vivo.

In vivo mammalian models of critical-sized bone defects are essential for researchers studying healing mechanisms and orthopedic therapies. Here, we introduce a protocol for the creation of reproducible, segmental, femoral defects in rats stabilized using external fixation.

Orthopedic research relies heavily on animal models to study mechanisms of bone healing in vivo as well as investigate the new treatment techniques. ...

Optical Imaging of Isolated Murine Ventricular Myocytes

The ability to isolate adult cardiac myocytes has permitted researchers to study a variety of cardiac pathologies at the single cell level. While advances in calcium sensitive dyes have permitted the robust optical recording of single cell calcium dynamics, recording of robust transmembrane optical voltage signals has remained difficult. Arguably, this is because of the low single to noise ratio, phototoxicity, and photobleaching of traditional potentiometric dyes. Therefore, single cell voltage measurements have long been confined to the patch clamp technique which while the gold standard, is technically demanding and low throughput. However, with the development of novel potentiometric dyes, large, fast optical responses to changes in voltage can be obtained with little to no phototoxicity and photobleaching. This protocol describes in detail how to isolate adult murine myocytes which can be used for cellular shortening, calcium, and optical voltage measurements. Specifically, the protocol describes how to use a ratiometric calcium dye, a single-excitation calcium dye, and a single excitation voltage dye. This approach can be used to assess the cardiotoxicity and arrhythmogenicity of various chemical agents. While phototoxicity is still an issue at the single cell level, methodology is discussed on how to reduce it.

We present the methodology for the isolation of murine myocytes and how to obtain voltage or calcium traces simultaneously with sarcomere shortening traces using fluorescence photometry with simultaneous digital cell geometry measurements.

The ability to isolate adult cardiac myocytes has permitted researchers to study a variety of cardiac pathologies at the single cell level. While advances ...

Biomechanical Testing of Murine Tendons

Tendon disorders are common, affect people of all ages, and are often debilitating. Standard treatments, such as anti-inflammatory drugs, rehabilitation, and surgical repair, often fail. In order to define tendon function and demonstrate efficacy of new treatments, the mechanical properties of tendons from animal models must be accurately determined. Murine animal models are now widely used to study tendon disorders and evaluate novel treatments for tendinopathies; however, determining the mechanical properties of mouse tendons has been challenging. In this study, a new system was developed for tendon mechanical testing that includes 3D-printed fixtures that exactly match the anatomies of the humerus and calcaneus to mechanically test supraspinatus tendons and Achilles tendons, respectively. These fixtures were developed using 3D reconstructions of native bone anatomy, solid modeling, and additive manufacturing. The new approach eliminated artifactual gripping failures (e.g., failure at the growth plate failure rather than in the tendon), decreased overall testing time, and increased reproducibility. Furthermore, this new method is readily adaptable for testing other murine tendons and tendons from other animals.

The protocol describes efficient and reproducible tensile biomechanical testing methods for murine tendons through the use of custom-fit 3D printed fixtures.

Tendon disorders are common, affect people of all ages, and are often debilitating. Standard treatments, such as anti-inflammatory drugs, rehabilitation, ...

Transplantation of a 3D Bioprinted Patch in a Murine Model of Myocardial Infarction

Testing regenerative properties of 3D bioprinted cardiac patches in vivo using murine models of heart failure via permanent left anterior descending (LAD) ligation is a challenging procedure and has a high mortality rate due to its nature. We developed a method to consistently transplant bioprinted patches of cells and hydrogels onto the epicardium of an infarcted mouse heart to test their regenerative properties in a robust and feasible way. First, a deeply anesthetized mouse is carefully intubated and ventilated. Following left lateral thoracotomy (surgical opening of the chest), the exposed LAD is permanently ligated and the bioprinted patch transplanted onto the epicardium. The mouse quickly recovers from the procedure after chest closure. The advantages of this robust and quick approach include a predicted 28-day mortality rate of up to 30% (lower than the 44% reported by other studies using a similar model of permanent LAD ligation in mice). Moreover, the approach described in this protocol is versatile and could be adapted to test bioprinted patches using different cell types or hydrogels where high numbers of animals are needed to optimally power studies. Overall, we present this as an advantageous approach which may change preclinical testing in future studies for the field of cardiac regeneration and tissue engineering.

This protocol aims to transplant a 3D bioprinted patch onto the epicardium of infarcted mice modeling heart failure. It includes details regarding anesthesia, the surgical chest opening, permanent ligation of the left anterior descending (LAD) coronary artery and application of a bioprinted patch onto the infarcted area of the heart.

Testing regenerative properties of 3D bioprinted cardiac patches in vivo using murine models of heart failure via permanent left anterior descending (LAD) ...