Cantilever Bending of Murine Femoral Necks

Summary

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.

Abstract

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° relative to the loading direction. The new protocol streamlined the testing by reducing variability in alignment (21.6° ± 1.5°, 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.

Introduction

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 lifetime^2, 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 µCT 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° 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.

Protocol

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 °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.

1. Obtain µCT scans of the representative samples.
   1. Scan representative samples on a µCT scanner with the following settings: 55 kV, 145 µA for 300 ms integration times, and resolution of 10.5 µm voxels.
   2. Ensure that the captured region covers the proximal end of the femur and continues down through the midshaft.
      NOTE: If a µCT scanner is unavailable, 2D planar X-rays of the representative samples can be used.
2. Analyze the µCT scans.
   1. Using the representative set of µCT scans, obtain a 2D rendering of the anterior view of the proximal femur.
      1. Obtain µCT images with a resolution of 10.5 µm 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.
      2. Determine a threshold to distinguish bone from the surrounding tissue and apply a Gaussian filter for noise reduction.
      3. Orient the 3D renderings to eliminate off-axis tilt and ensure that the femur's anterior surface is viewed.
      4. Export this 2D view of the 3D rendering as an image file, such as .jpg or .png.
   2. 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).
   3. Along the 7 mm distal perpendicular line, measure the femoral shaft diameter below the third trochanter.

Figure 1: µCT analysis. µCT 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° (SD = 1.19°), and the average midshaft diameter was 1.53 mm (SD = 0.14 mm) (n = 20). Scale bar = 1 mm. Please click here to view a larger version of this figure.

3. 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° 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.

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° 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° deviation from perpendicular in the midshaft average angles, the fixture angle was set to 73.13° to produce an angle of 20°. Alignment fixtures were printed with diameters ranging from 1.9-2.2 mm to ensure a proper fit for varying midshaft diameters. Please click here to view a larger version of this figure.

4. 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.

2. Sample preparation

1. 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.
2. 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.
3. Test the harvested samples immediately or store them at -20 °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.
4. Using ¼" x ¼" square aluminum tubing (see Table of Materials), cut tubing sections ½" to 1" in length. Using an etching tool, label each aluminum segment with the sample IDs.
5. Fill half of the tubing segments with putty. Place these tubing segments into a fixture to hold them upright.
6. 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.
7. 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.
8. 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.
9. 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.
10. 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).

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. Please click here to view a larger version of this figure.

3. Hardware set-up

1. Using a mechanical testing system (MTS), attach and calibrate a load cell with resolution <1 N (see Table of Materials) (Figure 4A).
   NOTE: The load cell can be mounted on the stage or, preferably, the actuator when possible.
2. 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.
3. Attach a loading platen to the actuator. This can be simply a tapered screw with a flattened tip (Figure 4C).
4. 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.

Figure 4: Hardware set-up. (A) Set-up of testing on mechanical testing system, with 1 kN load cell (resolution < 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. Please click here to view a larger version of this figure.

4. Software set-up

1. In the MTS software, begin the creation of a new flexural (bending) protocol. Ensure that the protocol will operate in displacement control.
2. Set the loading rate of the protocol to 0.5 mm/s.
3. If the software has a setting for soft keys, add the soft keys "Balance" and "Zero Extension" to the protocol.
   NOTE: This will quickly set the load and actuator position to 0 before testing each sample.
4. 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.
5. Save the new protocol and return to the main screen of the software program to begin testing a new sample set.

5. Testing set-up

1. 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).

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° to 23.99°, 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 < 0.05). Scale bar (Panel A) = 1 mm. Please click here to view a larger version of this figure.

2. Place the aluminum segment with sample into the holding fixture and tighten set screws.
3. 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.
4. 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.
5. In the MTS software, zero the position of the actuator and balance the load cell using the soft keys added in step 4.3.
6. Begin the loading protocol. Depending on how much space was left between the loading platen and sample, testing will only take 10-30 s.
7. After testing, capture another anterior X-ray of the sample. This will be used to discern and document the mode of fracture (Figure 6).

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. Please click here to view a larger version of this figure.

6. Data analysis

1. After data collection, export force and displacement data into software (see Table of Materials) that allows for graphing and mathematical calculations.
2. 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.
3. 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.

Representative Results

When potted with the aid of the guide, the femoral shafts were aligned at 21.6° ± 1.5°. While this represents <10% deviation from the intended angle of 20°, 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. Failure was consistently observed in the femoral necks, as intended, in a bifurcated manner, with one fracture line parallel to the femoral shaft and the other line perpendicular to the femoral neck (Figure 6). If significant variations were to occur in the breakage pattern amongst samples, then the bone quality of samples could be further assessed via µCT by measuring outcomes such as volumetric bone mineral density, trabecular and cortical thickness, spacing, and mineralization. If failure is not consistently induced in the femoral neck, the 3D printed guides may be adjusted.

The biomechanical outcome measures reported herein are consistent with values reported in similar axial bending of femoral neck configurations^{7,8,9,10,11,12,13,14}. However, the consistent alignment attained using the 3D printed guides generally improved the COV of the maximum load in particular (Table 1).

Current study	Sex	Midshaft Angle	Max Load	Stiffness	Work to failure
	Male	8%	10%	20%	24%
	Female	7%	9%	35%	38%
Jämsä et al10	Male	NR	22%	NR	NR
Jämsä et al8	Male	NR	19%	NR	NR
Kamal et al9	Female	NR	16%-25%	11%-28%	NR
Middleton et al7	Female	NR	24%-27%	NR	NR
Brent et al11	Female - rats	NR	18%-24%	NR	NR
Bromer et al12*	Female	NR	11%-27%	NR	NR
Vegger et al13*	Female	NR	16%-32%	NR	NR
Lodberg14*	Female	NR	11%-45%	NR	NR
NR: Not Reported
*: Data extrapolated from published figures

Table 1: Coefficients of variation for measured flexural properties of mouse femoral necks. The coefficients of variation represent a ratio of the standard deviation and mean of a data set. As COV decreases, this indicates a tighter grouping of the individual data points around the mean. This protocol decreased COV for maximum load compared to other publications performing similar testing.

As expected, sex differences were observed in the measured mechanical properties. Statistical analyses were performed using a one-tailed unpaired t-test. Femoral necks from male mice were significantly stronger and stiffer than specimens from female mice (p = 0.009 and p = 0.0006, respectively). In addition, the female femoral necks experienced more significant deformations (p = 0.014) and worked to failure (p = 0.024) compared to specimens from male mice (Figure 7). This is consistent with the lower bone mineral density in females and underscores the sensitivity of the test to detect physiologically relevant differences. In the male and female mice cohorts used in this study, the female mice bone mineral density was significantly lower than their male counterparts, as determined by a dual-energy X-ray absorptiometry scan (DEXA) and a one-tailed unpaired t-test (p = 0.036).

Figure 7: Biomechanical outcomes. (A) A Representative force-displacement curve, displaying a 0.2% offset linear fit, is used to derive the stiffness and yield point. Selected outcome measures are scatter plotted displaying mean and standard deviation, including (B) maximum load (at failure), (C) stiffness, (D) maximum displacement (at failure), and (E) work to failure (area under the curve up to the failure point). Asterisks indicate significant differences determined using a one-tailed unpaired t-test (*p < 0.05, **p < 0.01, ***p < 0.001, n = 10 per sex cohort). Please click here to view a larger version of this figure.

To confirm that the slight variations in the potting angle did not contribute to the experimental variability, each biomechanical outcome measure was plotted against the potting angle and performed a simple linear regression for the male cohort, female cohort, and all samples grouped together (Figure 8). The hypothesis was then tested that the linear regression slope is not zero. The regression analysis demonstrated that except for the stiffness, the slight variations in the potting angle (range 18° to 24°) did not affect the biomechanical outcome measures. For stiffness, there was a significant linear correlation with the potting angle (R² = 0.29, p < 0.05).

Figure 8: Effect of the potting angle on biomechanical outcomes. Biomechanical outcome measures including (A) maximum load, (B) stiffness, (C) maximum displacement, and (D) work to failure were plotted against the potting angle and correlated using a simple linear regression for the male cohort, female cohort, and all samples grouped together. Solid black lines show linear regression of grouped samples, with dotted lines indicating confidence intervals. Variability in the potting angle did not significantly affect the maximum load, maximum displacement, or failure work. However, as the potting angle increased, the stiffness increased, as determined by a Pearson's test (p = 0.0126, n = 20). Please click here to view a larger version of this figure.

Supplementary File 1: Standard Triangle Language (.STL) file of the guides. This file can be used to print the guides described in the protocol. Please click here to download this File.

Discussion

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, closely following the steps outlined in the first and fourth sections of the protocol will ensure that the creation of the guides and the loading protocol will replicate what is reported in this publication. As can be theoretically predicted and shown experimentally, the midshaft angle relative to the loading axis can affect the stresses experienced by the femoral neck and the probability that the fracture occurs at the femoral neck. Previous groups have demonstrated that the midshaft angle significantly influences the femur's compressive stiffness and strength when loaded through the femoral head. Parametric analysis of the effects of midshaft inclination angles showed that the maximal bending moments in the femoral shafts experience a minimum at midshaft angles between 15° and 25°, which also maximizes the shaft stiffness⁵. This angle, therefore, minimizes the likelihood of compression fractures in the shaft and increases the probability of flexural fractures in the femoral neck.

Several parameters could affect the outcomes of any biomechanical test and confound the ability to detect significant differences due to physiologically relevant experimental variables. This variability is compounded by the small size of long mouse bones. Among the parameters that require attention in this test, in particular, are the number of freeze-thaw cycles and hydration state of the bone, the loading rate, and the alignment of the femoral shaft relative to the axis of loading. The protocol stipulates that all samples go through the same number of freeze-thaw cycles and a 2 h window for hydration in PBS at room temperature. The loading rate is also set to a uniform value of 0.5 mm/s^3,4. Furthermore, 3D printed guides were designed to consistently position the femur at a midshaft angle of ~20° during the potting step. This resulted in consistent midshafts angles in the range of 18° to 24°, with no significant sex effects due to anatomic differences and coefficients of variation of 7.6% and 6.5% for male and female mice, respectively. These guides are accessible, easily modified using standard solid modeling software, and reproduced on demand using an inexpensive desktop 3D printer.

The representative results demonstrated that the testing protocol is sensitive to subtle physiological differences, such as sex, with a reasonable sample size of n = 10. A retrospective power analysis accounting for experimentally determined size effects (δ = Δmean/SD) at n = 10 suggested that the power was estimated to be 57% for the maximum load at failure (δ = 0.8), >95% for the stiffness (δ = 1.77) and the maximum displacement (δ = 1.9), and 83% for the work to failure (δ = 1.77), respectively. Along with the small coefficients of variation (Table 1), this power analysis confirms that the variation in the potting angle did adversely affect the sensitivity and reliability of the protocol.

The subjective analysis of the mode of failure also demonstrated that 100% of the samples tested failed in the femoral neck, as all of them showed a bifurcated fracture, with one fracture line running parallel to the shaft at the site it meets the neck and another fracture line perpendicular to femoral neck at the apex of the bifurcation. This encompasses features from two clinically relevant modes of femoral neck fractures; the intertrochanteric and transcervical neck fractures¹⁵. Femoral neck cantilever bending tests are not as commonly used and described in the literature as standard torsion or flexure testing of femoral and tibial midshafts in rodent models of osteoporosis. Only a handful of studies were identified to describe such protocols using mouse and rat models^{5,7,8,9,16,17}. The angle at which the femurs were positioned during testing is not always reported. Some with detailed descriptions use an excessive amount of custom fixtures and software to align their samples⁵ but still resort to potting by hand, introducing the same human error in other protocols.

This protocol is designed for murine samples and is specified for C57Bl/6 mice but could easily be adapted to large animal models or other murine strains with different femoral geometry. Future investigators using this protocol may need to modify the amount of exposed bone, as the third trochanter may not be precisely 7 mm distally from the femoral head. Additional modifications to the protocol include using a hardening agent that can be softened post-testing to release the sample if further testing is desired. This could be done with a bismuth alloy that could be melted in a hot water bath after testing to release the sample⁷. The final modification users could make to this protocol is eluded to in step 3.1, being the type and positioning of the load cell. An axial load cell should be used with sub 1 N resolution. A 50 N load cell would be appropriate based on the maximum loads observed. Furthermore, a load cell that only measures tension or compression should be used to avoid any compounding bending moment that the load cell may experience from eccentric loading relative to the load cell. Another way to avoid compounding force measurements would be to fix the load cell to the actuator to ensure the loading force is in line with the load cell.

This protocol simplifies the need for custom fixtures, describes how guides can be printed on any commercially available 3D printer, and utilizes common laboratory equipment to thoroughly and reproducibly test samples, as demonstrated by the lower coefficients of variation reported in the current study (Table 1). However, this protocol is not restricted by the need for a 3D printer. Commercially available solutions exist, where the 3D rendering files can be sent to printing companies, and the parts can be shipped back. Additionally, this mode of bending loading on the femoral neck simulates the location and types of fractures clinically encountered. With the number of people at high risk for fragility fractures, it is predicted that there will be upwards of 21.3 million hip fractures each year by 2050¹⁸. The immense societal, financial, and medical burden this poses, reliable testing in rodent models can improve the rigor and reproducibility of research geared towards understanding the etiology of osteoporosis and therapeutics to treat it effectively.

Materials

Name	Company	Catalog Number	Comments
¼” x ¼” square aluminum tubing	Grainger	48KU67	Cut to lengths of 1/2" to 1" lengths
1 kN load cell	Instron	2527-130	Any load cell with sub 1 N resolution can be used.
3.5x-45x Zoom Stereo Boom Microscope	Omano	OM2300S-GX4	Microscope used to precisely line up samples with loading platen.
3D printed guides	Custom made		Angled slots at 73.13°, with diameters between 1.9 mm and 2.2 mm
3D printed mount	Custom made		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.
Acrylic Base Plate Material Kit	Keystone Industries	921392	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.
Amira	ThermoFisher Scientific		Used to compile µCT scans
Biaxial stage	Custom made		Used to center femoral head of sample under the loading platen.
BioMed Amber Resin	formlabs	RS-F2-BMAM-01	Any resin from formlabs could be used for this project.
Bluehill 3	Instron	V3.66	Software used to set up loading protocol and collect load, displacement and time data.
ElectroPuls 10000	Instron	E10000	Mechanical testing system
Faxitron UltraFocus	Faxitron BioOptics	2327A40311	X-ray imaging system
Form 2	formlabs	F2	Used to print the mount and guides
Form 2 Resin Tank LT	formlabs	RT-F2-02	LT Tank was used to be compatible with the BioMed Resin
ImageJ	National Institutes of Health	ImageJ	Used to assess µCT and X-ray images
Laxco iLED Series LED Light Source	ThermoFisher Scientific	AMPSILED30W	Light source used in conjugtion with microscope.
Loading platen	Custom made		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.
Mount attachment	Custom made		To secure the 3D printed mount to the load cell. We used a M10/M6 threaded rod
Phosphate Buffer Saline (PBS)	ThermoFisher Scientific	10010031	Need to rehydrate the samples once acrylic base plate material has set.
Plumber's putty	Oatey	31174	Used to seal the end of the aluminum tubing when pouring acrylic base plate material in. Any clay or putty could be used.
PreForm	formlabs	Preform 3.15.2	Formlabs software
Tissue Culture Dish	Corning	353003	Samples can be laid flat in culture dish and covered in PBS to rehydrate.
vivaCT 40	Scanco	µCT 40	Representative set or actual samples can be scanned prior to printing of guides to calculate femoral shaft angle and diameter.