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.

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 ...

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, cl...

Materials

List of materials used in this article

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.