The study was supported by the NIH / NIAMS (R01 AR055580, R01 AR057836).

Animal studies were approved by Columbia University Institutional Animal Care and Use Committee. Mice used in this study were of a C57BL/6J background and were purchased from The Jackson Laboratory (Bar Harbor, ME, USA). They were housed in pathogen-free barrier conditions and were provided food and water ad libitum.
1. Development of custom-fit 3D printed fixtures for gripping bone
<ol>
	<li>Bone image acquisition and 3D bone model construction
	<ol>
		<li>Dissect the bone of interest in preparation for 3D model creation and 3D bone grip printing; the humerus and the calcaneus are used as examples in the current protocol. 
		NOTE: Detailed instructions to dissect bone-tendon-muscle specimens for mechanical testing are provided in step 2.1.1. The following steps should be followed to isolate bones for the purpose of creating 3D-printed bone grips.
		<ol>
			<li>Dissection of the humerus: Euthanize a mouse per IACUC-approved procedure. Remove upper extremity skin, remove all muscles over the humerus, disarticulate the elbow and glenohumeral joints, and carefully remove all connective tissues attached to the humerus.</li>
			<li>Dissection of the calcaneus: Euthanize a mouse per IACUC-approved procedure. Remove lower extremity skin, disarticulate the Achilles tendon-calcaneus joint and joints between calcaneus and other foot bones, and carefully remove all connective tissues attached to the calcaneus.</li>
		</ol>
		</li>
		<li>Perform a microcomputed tomography scan of the entire bone, e.g., scan the humerus and calcaneus samples. 
		NOTE: Depending on the scanner used, the settings will be different. For the scanner used in the current study (Table of Materials), the recommended settings are: scan at an energy of 55 kVP, Al 0.25 filter, at a resolution of 6 &#956;m.
		<ol>
			<li>Mix agarose powder in ultrapure water and microwave for 1-3 min until the agarose is completely dissolved. It is helpful to microwave for 30-45 s, stop and swirl, and then continue towards a boil. Fill cryotubes up to three-quarters full with agarose. Let the agarose cool for about 5-10 min.</li>
			<li>Insert bone into the agarose gel (this will prevent movement artifacts during scanning). Insert a cryotube with bone into the scanner. 
			NOTE: For the scanner used in the current study, a 16-position automatic sample changer was used for all scans. This scanner can automatically select magnification according to a sample&#8217;s size and shape.&#160;</li>
		</ol>
		</li>
		<li>Reconstruct microcomputed tomography scan projection images into cross-section images. Use recommended parameters for the experimenter&#8217;s scanner/software combination. 
		NOTE: For the program used in the current study (Table of Materials) it is recommended to use the following reconstruction parameters: Smoothing: 0-2, Beam Hardening Correction: 45, Ring Artefact Reduction: 4-9 and to reconstruct slices in 16-bit TIFF format.</li>
		<li>Create a 3D&#160;model and save into a standard STL format compatible with most 3D printers and rapid prototyping. For the program used in the current study (Table of Materials), do the following:
		<ol>
			<li>Select the command File &#62; Open to open the file dataset. Open the dialog File &#62; Preferences and select the Advanced tab.</li>
			<li>Use the adaptive rendering algorithm to construct the 3D models. This algorithm minimizes the number of facet triangles and provides smoother surface detail. Use 10 as the locality parameter; this parameter defines the distance in pixels to the neighboring point used for finding the object border. Minimize tolerance to 0.1 to decrease file size. 
			NOTE: After opening the dataset, the images are shown in the &#8220;Raw Images&#8221; page.</li>
			<li>To specify the volume of interest (VOI), manually select two images to set as the top and bottom of the selected VOI range.</li>
			<li>Move to the second page, Region of Interest. Manually select the region of interest on a single cross section image. 
			NOTE: The selected region will be highlighted in red (i.e., the humerus cross-sectional area).</li>
			<li>Repeat the previous step every 10&#8211;15 cross-section images.</li>
			<li>Move to the third page Binary Selection. On the histogram menu, click From Dataset. The histogram distribution of brightness from all images of the dataset will be shown. Also on the histogram menu, click the Create a 3D Model file menu.</li>
		</ol>
		</li>
		<li>Save a 3D model of the bone in STL file format.</li>
		<li>Refine the mesh: Manipulate the mesh to reduce the size of the STL file and make it compatible with any solid modeling computer-aided design program. For the program used in the current study (Table of Materials), follow the steps below:
		<ol>
			<li>Import mesh and select all to edit. Choose Reduce&#160;from the toolset Edit. Then, select&#160;Triangle Budget from the toolset Reduce Target. Reduce the Tri Count and accept changes.&#160;Resave the newly reduced file in STL format by choosing Export as&#8230;</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Design of custom-fit bone fixtures
	<ol>
		<li>Supraspinatus tendon-humeral bone
		<ol>
			<li>Use a solid modeling computer-aided design program to create a custom-fit model of humerus gripping fixture (Figure 1, Supplemental Files). 
			NOTE: The program used in the current study is listed in the Table of Materials.</li>
			<li>Open the STL format file of the humerus bone in a solid modeling program and save as a part file. 
			NOTE: For the software used in the current study (Table of Materials), the 3D bone object was saved in SLDPRT format.</li>
			<li>Open the part file and manually create three anatomically relevant planes (i.e., sagittal, coronal, transverse).
			<ol>
				<li>Manually define the sagittal plane to cut through the supraspinatus tendon attachment at the greater tuberosity. Ensure that the 3D block contains the sagittal plane as a plane of symmetry. To achieve this, add or cut material from the block if needed. 
				NOTE: This plane of symmetry ensures that when the specimen is inserted into the fixtures the tendon and tendon attachment are located in the central axis of the fixture.</li>
			</ol>
			</li>
			<li>Measure dimensions of the bone along each of the three planes (i.e., height, width, length).</li>
			<li>Measure the dimensions of the mechanical testing grips where the 3D printed fixture will be attached.</li>
			<li>Begin by designing a solid block part (e.g., a solid cylinder).
			<ol>
				<li>Ensure that each dimension of the block is at least 5 mm greater than the dimensions of the humerus.</li>
				<li>Account for design constraints from mechanical testing grips (i.e., ensure that the 3D printed fixture can be assembled and disassembled freely in the mechanical testing grips).</li>
			</ol>
			</li>
			<li>Create an assembly model with two components: the solid block and either the right or left humerus bone. Define the orientation of the bone within the block (i.e., the angle between the tendon and bone). Ensure that the entire bone volume fits inside the block.</li>
			<li>Create a cavity in the block using the humerus bone as the mold. If using the software specified in the Table of Materials, follow the following steps:
			<ol>
				<li>Insert the design part (humerus) and the mold base (cylinder block) into an interim assembly. In the assembly window, select the block, and click Edit Component from the Assembly toolbar.</li>
				<li>Click Insert &#62; Features &#62; Cavity. Select Uniform Scaling and enter 0% as the value to scale in all directions.</li>
			</ol>
			</li>
			<li>Suppress the bone part and save the assembly as a part.</li>
			<li>Open part (cylinder with cavity). Cut the part along the sagittal plane to create two symmetrical components that fit the bone anteriorly and posteriorly (e.g., two half cylinders, as seen in Figure 1). 
			NOTE: Two components are designed that fit the bone anteriorly and posteriorly. The anterior component includes a half spherical-shaped cavity extended from the anterior side of the humeral head up to the supraspinatus tendon attachment. The posterior component cavity is shaped like the posterior part of the humerus (i.e., posterior side of the humeral head, deltoid tuberosity, and medial and lateral epicondyle).</li>
			<li>Save each component as a separate file part.</li>
			<li>For the anterior component, ensure that the humeral head is embedded in the cavity of the part by defining appropriate tolerances. 
			NOTE: In the current study, using the software specified in the Table of Materials, it is suggested to follow the steps below:
			<ol>
				<li>Create a revolved cut to smooth the mesh geometry of the cavity. Create a sketch for the cut by emulating the cavity geometry and adding a locational clearance. 
				NOTE: The clearance allows for free assembly and disassembly between the bone and the anterior component.</li>
			</ol>
			</li>
			<li>Modify the posterior component to imitate the cavity geometry to create a cut that adds clearance, as described above for the anterior component.</li>
			<li>Make a cut in the transverse plane starting from the top of the posterior component up to the crest of the greater/lesser tubercle. 
			NOTE: As seen in Figure 1 and Figure 2, the posterior component includes a cut that creates an opening at the tendon attachment.</li>
			<li>Create a snug fit between the two components to allow for free assembly and disassembly. 
			NOTE: A hole-shaft fit with a loose running clearance was created for the fixtures in the current study.</li>
			<li>Create 3D mirror models for each component of the fixture for the opposite limb (i.e., left or right).</li>
			<li>Add an etch on the bottom of the fixtures to distinguish between the left and right sides.</li>
			<li>Save all fixture parts in STL standard file format in preparation for 3D printing.</li>
		</ol>
		</li>
		<li>Achilles tendon-calcaneus bone
		<ol>
			<li>Follow the same steps as described above for supraspinatus-humeral head fixture. 
			NOTE: Only one set of fixtures is necessary for the Achilles-calcaneal, since the anatomy of the left and right calcaneus bones is nearly symmetrical.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Biomechanical testing of murine tendons
<ol>
	<li>Specimen preparation and cross-sectional area measurement
	<ol>
		<li>Dissect the muscle-tendon-bone of interest in preparation for tensile mechanical testing. In the current study, supraspinatus muscle - tendon - humerus bone specimens (N=10, 5 male, 5 female) and gastrocnemius muscle - Achilles tendon-calcaneus bone specimens (N=12, 6 male, 6 female) were isolated from 8 week old C57BL/6J mice.
		<ol>
			<li>Dissection of the supraspinatus muscle - tendon - humerus bone specimen
			<ol>
				<li>Euthanize a mouse per IACUC-approved procedure. Position the mouse in a prone position. Make an incision in the skin from above the elbow of the forepaw towards the shoulder.</li>
				<li>Carefully remove the skin with blunt dissection so that the musculature of the shoulder is visible. Remove the tissue surrounding the humerus until the bone is exposed and can be held securely with forceps.</li>
				<li>Hold the humerus with forceps and carefully remove the deltoid and trapezius muscles to expose the coracoacromial arch. Identify the acromioclavicular joint and carefully separate the clavicle from the acromion with a scalpel blade.</li>
				<li>Taking care not to damage the supraspinatus tendon and its bony attachment, remove the muscle from its scapular attachment using a scalpel blade. Taking care not to damage the supraspinatus tendon and its bony attachment, detach the humeral head from the glenoid; using a scalpel blade, lacerate the joint capsule and the infraspinatus, subscapularis, and teres minor tendons.</li>
				<li>Disarticulate the elbow joint to separate the humerus from the ulna and radius. Isolate the humerus - supraspinatus tendon - muscle specimen and clean off excess soft tissues on the humerus and humeral head.</li>
			</ol>
			</li>
			<li>Dissection of the Achilles tendon - calcaneus bone sample
			<ol>
				<li>Euthanize a mouse per IACUC-approved procedure. Position the mouse in a prone position. Taking care not to damage the Achilles tendon and its bony attachment, remove the skin with blunt dissection so that the musculature around the ankle and knee joints is exposed.</li>
				<li>Using a scalpel blade, starting at the Achilles tendon - calcaneus attachment, carefully detach the gastrocnemius muscle from its proximal attachments.</li>
				<li>Carefully disarticulate the calcaneus from the various adjacent bones. Isolate the Achilles tendon - calcaneus specimen and clean off excess soft tissues.</li>
			</ol>
			</li>
		</ol>
		</li>
		<li>Determine the cross-sectional area of the tendon using microcomputed tomography. 
		NOTE: For the scanner used in the current study (Table of Materials), the recommended settings are: scan at an energy of 55 kVP, Al 0.25 filter, at a resolution of 5 &#956;m.
		<ol>
			<li>Mix agarose powder in ultrapure water and microwave for 1-3 min until the agarose is completely dissolved. It is helpful to microwave for 30-45 s, stop and swirl, and then continue towards a boil. Fill cryotubes up to three-quarters full with agarose. Let the agarose cool for about 5-10 min.</li>
			<li>Suspend the specimen in the cryotube by inserting the bone upside down. 
			NOTE: Only the bone should be in the agarose gel. The tendon and muscle should be suspended outside.</li>
		</ol>
		</li>
		<li>After the scan, gently remove muscle from tendon using scalpel blade. Insert the specimen into the 3D-printed fixture. 
		NOTE: The grips are reusable for each test. Do not use glue or epoxy in the fixture; the bone is held in a press fit.</li>
		<li>Insert and glue the tendon between a folded thin tissue paper (2 cm x 1 cm) and clamp the construct using thin film grips. Attach the 3D printed fixture with the specimen into the testing grips.</li>
		<li>Insert the sample and the grips into a testing bath of phosphate buffered saline (PBS) at 37 &#176;C (i.e., mouse body temperature23).</li>
	</ol>
	</li>
	<li>Tensile testing
	<ol>
		<li>Perform tensile mechanical test on a material testing frame. 
		NOTE: For the testing frame used in the current study (Table of Materials), the recommended protocol is:
		<ol>
			<li>Define the gauge length as the distance from the tendon attachment to the upper grip.</li>
			<li>Precondition with 5 cycles between 0.05 N and 0.2 N.</li>
			<li>Hold for 120 s.</li>
			<li>Use a tension to failure of 0.2%/s.</li>
		</ol>
		</li>
		<li>Collect load-deformation data.</li>
		<li>Calculate the strain as the displacement relative to the initial gauge length of the tendon.</li>
		<li>Calculate the stress as the force divided by the initial tendon cross-sectional area (as measured from microCT).</li>
		<li>If interested in viscoelastic behavior, perform a stress relaxation prior to the tension test to failure and use the data to calculate parameters such as A, B, C, tau1, and tau2 from the quasilinear viscoelastic model24.</li>
		<li>From the load deformation curve, calculate the stiffness (slope of linear portion of curve), the maximum force, and the work to yield (the area under the curve up to yield force).
		<ol>
			<li>Identify the linear portion by choosing a window of points in the load-deformation curve that maximizes the R2 value for a linear least squares regression25.</li>
			<li>Determine the stiffness as the slope of the linear portion of the load-displacement curve25,26.</li>
		</ol>
		</li>
		<li>From the stress strain curve, calculate the modulus (slope of linear portion of curve), the strength (maximum stress), and the resilience (area under the curve up to yield stress). 
		NOTE: Using the RANSAC algorithm, the yield strain (x-value) is defined as the first point when the y-fit has deviated more than 0.5% of the expected stress value (y-value). Yield stress is the corresponding y-value of the yield strain. 
		NOTE: In addition to the monotonic tensile loading to failure described in the current study, cyclic loading can provide important information about tendon fatigue and/or viscoelastic properties. For example, Freedman et al. reported fatigue properties of the murine Achilles tendons27.</li>
		<li>After completion of tensile testing, perform a microcomputed tomography scan of the entire bone, e.g., scan the humerus and calcaneus samples. 
		NOTE: For the scanner used in the current study (Table of Materials), the recommended settings are: scan at an energy of 55 kVP, Al 0.25 filter, at a resolution of 6 &#956;m.
		<ol>
			<li>Repeat steps 1.1.2.1&#8211;1.1.2.2.</li>
		</ol>
		</li>
		<li>Repeat step 1.1.3.</li>
		<li>Use a 3D visualization program compatible with the scanner to create a volume-rendered 3D model of the scanned object. 
		NOTE: The program used in the current study is listed in the Table of Materials.</li>
		<li>Determine the failure mode and failure site area by inspecting the 3D object.</li>
	</ol>
	</li>
	<li>Statistical analysis: Show all sample results as mean &#177; standard deviation (SD). Make comparisons between groups using student&#8217;s t-tests (two-tailed and unpaired). Set significance as p &#60; 0.05. 
	NOTE: The statistical software used in the current study is listed in the Table of Materials.</li>
</ol>

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanical,+Compositional,+and+Structural+Properties+of+the+Post-natal+Mouse+Achilles+Tendon.">Mechanical, Compositional, and Structural Properties of the Post-natal Mouse Achilles Tendon.</a> Annals of Biomedical Engineering. 39, 1904-1913 (2011).</li><li>Shu, C. C., Smith, M. M., Appleyard, R. C., Little, C. B., Melrose, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Achilles+and+tail+tendons+of+perlecan+exon+3+null+heparan+sulphate+deficient+mice+display+surprising+improvement+in+tendon+tensile+properties+and+altered+collagen+fibril+organisation+compared+to+C57BL/6+wild+type+mice.">Achilles and tail tendons of perlecan exon 3 null heparan sulphate deficient mice display surprising improvement in tendon tensile properties and altered collagen fibril organisation compared to C57BL/6 wild type mice.</a> PeerJ. 6, 5120 (2018).</li><li>Probst, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+clamping+technique+for+biomechanical+testing+of+tendons+in+small+animals.">A new clamping technique for biomechanical testing of tendons in small animals.</a> Journal of Investigative Surgery. 13, 313-318 (2000).</li><li>Talan, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Body+temperature+of+C57BL/6J+mice+with+age.">Body temperature of C57BL/6J mice with age.</a> Experimental Gerontology. 19, 25-29 (1984).</li><li>Newton, M. D., 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+influence+of+testing+angle+on+the+biomechanical+properties+of+the+rat+supraspinatus+tendon.">The influence of testing angle on the biomechanical properties of the rat supraspinatus tendon.</a> Journal of Biomechanics. 49, 4159-4163 (2016).</li><li>Schwartz, A. G., Lipner, J. H., Pasteris, J. D., Genin, G. M., Thomopoulos, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Muscle+loading+is+necessary+for+the+formation+of+a+functional+tendon+enthesis.">Muscle loading is necessary for the formation of a functional tendon enthesis.</a> Bone. 55, 44-51 (2014).</li><li>Gimbel, J. A., Van Kleunen, J. P., Williams, G. R., Thomopoulos, S., Soslowsky, L. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long+durations+of+immobilization+in+the+rat+result+in+enhanced+mechanical+properties+of+the+healing+supraspinatus+tendon.">Long durations of immobilization in the rat result in enhanced mechanical properties of the healing supraspinatus tendon.</a> Journal of Biomechanical Engineering. 129, 400-404 (2006).</li><li>Freedman, B. R., Sarver, J. J., Buckley, M. R., Voleti, P. B., Soslowsky, L. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biomechanical+and+structural+response+of+healing+Achilles+tendon+to+fatigue+loading+following+acute+injury.">Biomechanical and structural response of healing Achilles tendon to fatigue loading following acute injury.</a> Journal of Biomechanics. 47, 2028-2034 (2014).</li><li>Deymier, A. C., 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+multiscale+structural+and+mechanical+effects+of+mouse+supraspinatus+muscle+unloading+on+the+mature+enthesis.">The multiscale structural and mechanical effects of mouse supraspinatus muscle unloading on the mature enthesis.</a> Acta Biomaterialia. 83, 302-313 (2019).</li><li>Killian, M. L., Thomopoulos, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Scleraxis+is+required+for+the+development+of+a+functional+tendon+enthesis.">Scleraxis is required for the development of a functional tendon enthesis.</a> FASEB Journal. 30, 301-311 (2016).</li><li>Schwartz, A. G., Long, F., Thomopoulos, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Enthesis+fibrocartilage+cells+originate+from+a+population+of+Hedgehog-responsive+cells+modulated+by+the+loading+environment.">Enthesis fibrocartilage cells originate from a population of Hedgehog-responsive cells modulated by the loading environment.</a> Development. 142, 196-206 (2015).</li><li>Bell, R., Taub, P., Cagle, P., Flatow, E. L., Andarawis-Puri, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+a+mouse+model+of+supraspinatus+tendon+insertion+site+healing.">Development of a mouse model of supraspinatus tendon insertion site healing.</a> Journal of Orthopaedic Research. 33, 25-32 (2014).</li><li>Connizzo, B. K., Bhatt, P. R., Liechty, K. W., Soslowsky, L. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diabetes+Alters+Mechanical+Properties+and+Collagen+Fiber+Re-Alignment+in+Multiple+Mouse+Tendons.">Diabetes Alters Mechanical Properties and Collagen Fiber Re-Alignment in Multiple Mouse Tendons.</a> Annals of Biomedical Engineering. 42, 1880-1888 (2014).</li><li>Eekhoff, J. D., 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=Functionally+Distinct+Tendons+From+Elastin+Haploinsufficient+Mice+Exhibit+Mild+Stiffening+and+Tendon-Specific+Structural+Alteration.">Functionally Distinct Tendons From Elastin Haploinsufficient Mice Exhibit Mild Stiffening and Tendon-Specific Structural Alteration.</a> Journal of Biomechanical Engineering. 139, 111003 (2017).</li><li>Mikic, B., Bierwert, L., Tsou, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Achilles+tendon+characterization+in+GDF-7+deficient+mice.">Achilles tendon characterization in GDF-7 deficient mice.</a> Journal of Orthopaedic Research. 24, 831-841 (2006).</li><li>Sikes, K. J., 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=Knockout+of+hyaluronan+synthase+1,+but+not+3,+impairs+formation+of+the+retrocalcaneal+bursa.">Knockout of hyaluronan synthase 1, but not 3, impairs formation of the retrocalcaneal bursa.</a> Journal of Orthopaedic Research. 36, 2622-2632 (2018).</li><li>Wang, V. M., Banack, T. M., Tsai, C. W., Flatow, E. L., Jepsen, K. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Variability+in+tendon+and+knee+joint+biomechanics+among+inbred+mouse+strains.">Variability in tendon and knee joint biomechanics among inbred mouse strains.</a> Journal of Orthopaedic Research. 24, 1200-1207 (2006).</li><li>Wang, V. 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=Murine+tendon+function+is+adversely+affected+by+aggrecan+accumulation+due+to+the+knockout+of+ADAMTS5.">Murine tendon function is adversely affected by aggrecan accumulation due to the knockout of ADAMTS5.</a> Journal of Orthopaedic Research. 30, 620-626 (2011).</li><li>Zhang, K., 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=Tendon+mineralization+is+progressive+and+associated+with+deterioration+of+tendon+biomechanical+properties,+and+requires+BMP-Smad+signaling+in+the+mouse+Achilles+tendon+injury+model.">Tendon mineralization is progressive and associated with deterioration of tendon biomechanical properties, and requires BMP-Smad signaling in the mouse Achilles tendon injury model.</a> Matrix Biology. 52-54, 315-324 (2016).</li><li>Rooney, S. I., 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=Ibuprofen+differentially+affects+supraspinatus+muscle+and+tendon+adaptations+to+exercise+in+a+rat+model.">Ibuprofen differentially affects supraspinatus muscle and tendon adaptations to exercise in a rat model.</a> American Journal of Sports Medicine. 44, 2237-2245 (2016).</li><li>Galasso, O., 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=Quality+of+Life+and+Functional+Results+of+Arthroscopic+Partial+Repair+of+Irreparable+Rotator+Cuff+Tears.">Quality of Life and Functional Results of Arthroscopic Partial Repair of Irreparable Rotator Cuff Tears.</a> Arthroscopy - Journal of Arthroscopic and Related Surgery. 33, 261-268 (2017).</li><li>Sarver, D. C., 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=Sex+differences+in+tendon+structure+and+function.">Sex differences in tendon structure and function.</a> Journal of Orthopaedic Research. 35, 2117-2126 (2017).</li><li>Razmjou, H., 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=Disability+and+satisfaction+after+Rotator+Cuff+decompression+or+repair:+A+sex+and+gender+analysis.">Disability and satisfaction after Rotator Cuff decompression or repair: A sex and gender analysis.</a> BMC Musculoskeletal Disorders. 12, 66 (2011).</li></ol>

The authors have nothing to disclose.

Murine animal models are commonly used to study tendon disorders, but characterization of their mechanical properties is challenging and uncommon in the literature. The purpose of this protocol is to describe a time efficient and reproducible method for tensile testing of murine tendons. The new methods reduced the time required to test a sample from hours to minutes and eliminated a major gripping artifact that was a common problem in previous methods.
Several steps described in this protocol...

Tendon disorders are common and highly prevalent among the aging, athletic, and active populations1,2,3. In the United States, 16.4 million connective tissue injuries are reported each year4 and account for 30% of all injury-related physician office visits3,5,6,7,8. The most commonly affected sites include the rotator cuff, Achilles tendon, and patellar tendon9. Although a variety of non-operative and operative treatments have been explored, including anti-inflammatory drugs, rehabilitation, and surgical repair, outcomes remain poor, with limited return to function and high rates of failure5,6. These poor clinical outcomes have motivated basic and translational studies seeking to understand tendinopathy and to develop novel treatment approaches.
Tensile biomechanical properties are the primary quantitative outcomes defining tendon function. Therefore, laboratory characterization of tendinopathy and treatment efficacy must include a rigorous testing of tendon tensile properties. Numerous studies have described methods to determine the biomechanical properties of tendons from animal models such as rats, sheep, dogs, and rabbits10,11,12. However, few studies have tested the biomechanical properties of murine tendons, primarily due to the difficulties in gripping the small tissues for tensile testing. As murine models have numerous advantages for mechanistically studying tendinopathy, including genetic manipulation, extensive reagent options, and low cost, development of accurate and efficient methods to biomechanically test murine tissues is needed.
In order to properly test the mechanical properties of tendons, the tissue must be gripped effectively, without slipping or artifactual tearing at the grip interface or fracturing of the growth plate. In many cases, particularly for short tendons, the bone is gripped on one end and the tendon is gripped on the other end. Bones are typically secured by embedding them in materials such as epoxy resin13 and polymethylmethacrylate14,15. Tendons are often placed between two layers of sandpaper, glued with cyanoacrylate, and secured using compression clamps (if the cross section is flat) or in a frozen medium (if the cross section is large)15,16,17. These methods have been applied to biomechanically test murine tendons, but challenges arise due to the small size of the specimens and the compliance of the growth plate, which never ossifies18. For example, the diameter of the murine humeral head is only a few millimeters, thus making gripping of the bone difficult. Specifically, tensile testing of murine supraspinatus tendon-to-bone samples often results in failure at the growth plate rather than in the tendon or at the tendon enthesis. Similarly, biomechanical testing of the Achilles tendon is challenging. Although the Achilles tendon is larger than other murine tendons, the calcaneus is small, making gripping of this bone difficult. The bone can be removed, followed by gripping the two tendon ends; however, this precludes the testing of the tendon-to-bone attachment. Other groups report gripping the calcaneus bone using custom-made fixtures19,20, anchoring by clamps21, fixing in self curing plastic cement22 or using a conical shape slot22, yet these prior methods remain limited by low reproducibility, high gripping failure rates, and tedious preparation requirements.
The objective of the current study was to develop an accurate and efficient method for tensile biomechanical testing of murine tendons, focusing on the supraspinatus and Achilles tendons as examples. Using a combination of 3D reconstructions from native bone anatomy, solid modeling, and additive manufacturing, a novel method was developed to grip the bones. These fixtures effectively secured the bones, prevented growth plate failure, decreased specimen preparation time, and increased testing reproducibility. The new method is readily adaptable to test other murine tendons as well as tendons in rats and other animals.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr >
			<td >Agarose</td>
			<td >Fisher Scientific</td>
			<td >BP160-100</td>
			<td >Dissovle 1g in 100 ml ultrapure water to make 1% agarose&#160;</td>
		</tr>
		<tr >
			<td >Bruker microCT&#160;</td>
			<td >Bruker BioSpin Corp</td>
			<td >Skyscan 1272&#160;</td>
			<td>Used by authors</td>
		</tr>
		<tr >
			<td >ElectroForce&#160;</td>
			<td >TA Instruments</td>
			<td >3200</td>
			<td>Testing platform</td>
		</tr>
		<tr >
			<td >Ethanol 200 Proof</td>
			<td >Fisher Scientific</td>
			<td >A4094</td>
			<td>Dilute to 70% and use as suggested in protocol</td>
		</tr>
		<tr >
			<td >Fixture to attach grips</td>
			<td >Custom made</td>
			<td ></td>
			<td>Used by authors</td>
		</tr>
		<tr >
			<td >Kimwipes</td>
			<td >Kimberly-Clark</td>
			<td >&#160;S-8115</td>
			<td>As suggested in protocol</td>
		</tr>
		<tr >
			<td >MicroCT CT-Analyser (Ctan)</td>
			<td >Bruker BioSpin Corp</td>
			<td ></td>
			<td>Used by authors for visualizing and analyzing micro-CT scans&#160;</td>
		</tr>
		<tr >
			<td >MilliQ water (Ultrapure water)</td>
			<td>Millipore Sigma</td>
			<td>QGARD00R1 (or related purifier)</td>
			<td>100 ml&#160;</td>
		</tr>
		<tr >
			<td >Meshmixer</td>
			<td >Autodesk</td>
			<td >http://www.meshmixer.com/</td>
			<td >Free engineering software used by authors to refine mesh</td>
		</tr>
		<tr >
			<td >Objet EDEN 260VS&#160;</td>
			<td >Stratasys LTD</td>
			<td ></td>
			<td>Precision Prototyping</td>
		</tr>
		<tr >
			<td >Objet Studio</td>
			<td >Stratasys LTD</td>
			<td ></td>
			<td>Used by authors with 3D printer</td>
		</tr>
		<tr >
			<td >PBS - Phosphate-Buffered Saline</td>
			<td >ThermoFisher Scientific</td>
			<td >10010031</td>
			<td >2.5 L of 10% PBS&#160;</td>
		</tr>
		<tr >
			<td >S&#38;T Forceps</td>
			<td >Fine Science Tools</td>
			<td >00108-11</td>
			<td>Used by authors</td>
		</tr>
		<tr >
			<td >Scalpel Blade - #11</td>
			<td >Fine Science Tools</td>
			<td >10011-00</td>
			<td>Used by authors</td>
		</tr>
		<tr >
			<td >Scalpel Handle - #3</td>
			<td >Fine Science Tools</td>
			<td >10003-12</td>
			<td>Used by authors</td>
		</tr>
		<tr >
			<td >SkyScan 1272</td>
			<td >Bruker BioSpin Corp</td>
			<td ></td>
			<td>Used by authors for visualizing and analyzing micro-CT scans&#160;</td>
		</tr>
		<tr >
			<td >Skyscan CT-Vox</td>
			<td >Bruker BioSpin Corp</td>
			<td ></td>
			<td>Used by authors for visualizing and analyzing micro-CT scans&#160;</td>
		</tr>
		<tr >
			<td >SkyScan NRecon</td>
			<td >Bruker BioSpin Corp</td>
			<td ></td>
			<td>Used by authors for visualizing and analyzing micro-CT scans&#160;</td>
		</tr>
		<tr >
			<td >SolidWorks CAD</td>
			<td >Dassault Syst&#232;mes</td>
			<td >SolidWorks Research Subsription</td>
			<td>Solid modeling computer-aided design used by authors</td>
		</tr>
		<tr >
			<td >SuperGlue</td>
			<td >Loctite</td>
			<td >234790</td>
			<td>As suggested in protocol</td>
		</tr>
		<tr >
			<td >Testing bath</td>
			<td >Custom made</td>
			<td ></td>
			<td>Used by authors</td>
		</tr>
		<tr >
			<td >Thin film grips&#160;</td>
			<td >Custom made</td>
			<td ></td>
			<td>Used by authors</td>
		</tr>
		<tr >
			<td >VeroWhitePlus</td>
			<td >Stratasys LTD</td>
			<td >NA</td>
			<td>3D printing material used by authors</td>
		</tr>
		<tr >
			<td >WinTest&#160;</td>
			<td >WinTest Software</td>
			<td ></td>
			<td>Used by authors to collect data</td>
		</tr>
	</tbody>
</table>

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.

3D-printed fixtures were used to test 8-week old murine supraspinatus and Achilles tendons. All mechanically tested samples failed at the enthesis, as characterized by microCT scans, visual inspection, and video analysis after tensile tests. A one-to-one comparison of the previous and current methods for supraspinatus tendon testing in our laboratory is shown in Figure 3. In the previous method28,29,30</s...

Watch this Scientific Journal Video about Biomechanical Testing of Murine Tendons at JoVE.com

Biomechanical Testing of Murine Tendons

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, ...

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Bioengineering

13.0K Views.  Columbia University. The protocol describes efficient and reproducible tensile biomechanical testing methods for murine tendons through the use of custom-fit 3D printed fixtures.

Video: Biomechanical Testing of Murine Tendons

Directed Cellular Self-Assembly to Fabricate Cell-Derived Tissue Rings for Biomechanical Analysis and Tissue Engineering

Each year, hundreds of thousands of patients undergo coronary artery bypass surgery in the United States.1 Approximately one third of these patients do not have suitable autologous donor vessels due to disease progression or previous harvest. The aim of vascular tissue engineering is to develop a suitable alternative source for these bypass grafts. In addition, engineered vascular tissue may prove valuable as living vascular models to study cardiovascular diseases. Several promising approaches to engineering blood vessels have been explored, with many recent studies focusing on development and analysis of cell-based methods.2-5 Herein, we present a method to rapidly self-assemble cells into 3D tissue rings that can be used in vitro to model vascular tissues.
To do this, suspensions of smooth muscle cells are seeded into round-bottomed annular agarose wells. The non-adhesive properties of the agarose allow the cells to settle, aggregate and contract around a post at the center of the well to form a cohesive tissue ring.6,7 These rings can be cultured for several days prior to harvesting for mechanical, physiological, biochemical, or histological analysis. We have shown that these cell-derived tissue rings yield at 100-500 kPa ultimate tensile strength8 which exceeds the value reported for other tissue engineered vascular constructs cultured for similar durations (&lt;30 kPa).9,10 Our results demonstrate that robust cell-derived vascular tissue ring generation can be achieved within a short time period, and offers the opportunity for direct and quantitative assessment of the contributions of cells and cell-derived matrix (CDM) to vascular tissue structure and function.

This article outlines a versatile method to create cell-derived tissue rings by cellular self-assembly. Smooth muscle cells seeded into ring-shaped agarose wells aggregate and contract to form robust three-dimensional (3D) tissues within 7 days. Millimeter-scale tissue rings are conducive to mechanical testing and serve as building blocks for tissue assembly.

Each year, hundreds of thousands of patients undergo coronary artery bypass surgery in the United States.1 Approximately one third of these patients do ...

Determination of the Transport Rate of Xenobiotics and Nanomaterials Across the Placenta using the ex vivo Human Placental Perfusion Model

Decades ago the human placenta was thought to be an impenetrable barrier between mother and unborn child. However, the discovery of thalidomide-induced birth defects and many later studies afterwards proved the opposite. Today several harmful xenobiotics like nicotine, heroin, methadone or drugs as well as environmental pollutants were described to overcome this barrier. With the growing use of nanotechnology, the placenta is likely to come into contact with novel nanoparticles either accidentally through exposure or intentionally in the case of potential nanomedical applications. Data from animal experiments cannot be extrapolated to humans because the placenta is the most species-specific mammalian organ 1. Therefore, the ex vivo dual recirculating human placental perfusion, developed by Panigel et al. in 1967 2 and continuously modified by Schneider et al. in 1972 3, can serve as an excellent model to study the transfer of xenobiotics or particles.
Here, we focus on the ex vivo dual recirculating human placental perfusion protocol and its further development to acquire reproducible results.
The placentae were obtained after informed consent of the mothers from uncomplicated term pregnancies undergoing caesarean delivery. The fetal and maternal vessels of an intact cotyledon were cannulated and perfused at least for five hours. As a model particle fluorescently labelled polystyrene particles with sizes of 80 and 500 nm in diameter were added to the maternal circuit. The 80 nm particles were able to cross the placental barrier and provide a perfect example for a substance which is transferred across the placenta to the fetus while the 500 nm particles were retained in the placental tissue or maternal circuit. The ex vivo human placental perfusion model is one of few models providing reliable information about the transport behavior of xenobiotics at an important tissue barrier which delivers predictive and clinical relevant data.

Determination of the Transport Rate of Xenobiotics and Nanomaterials Across the Placenta using the ex vivo Human Placental Perfusion Model

The ex vivo dual recirculating human placental perfusion model can be used to investigate the transfer of xenobiotics and nanoparticles across the human placenta. In this video protocol we describe the equipment and techniques required for a successful execution of a placenta perfusion.

Decades ago the human placenta was thought to be an impenetrable barrier between mother and unborn child. However, the discovery of thalidomide-induced ...

Biomechanical Characterization of Human Soft Tissues Using Indentation and Tensile Testing

Regenerative medicine aims to engineer materials to replace or restore damaged or diseased organs. The mechanical properties of such materials should mimic the human tissues they are aiming to replace; to provide the required anatomical shape, the materials must be able to sustain the mechanical forces they will experience when implanted at the defect site. Although the mechanical properties of tissue-engineered scaffolds are of great importance, many human tissues that undergo restoration with engineered materials have not been fully biomechanically characterized. Several compressive and tensile protocols are reported for evaluating materials, but with large variability it is difficult to compare results between studies. Further complicating the studies is the often destructive nature of mechanical testing. Whilst an understanding of tissue failure is important, it is also important to have knowledge of the elastic and viscoelastic properties under more physiological loading conditions.
This report aims to provide a minimally destructive protocol to evaluate the compressive and tensile properties of human soft tissues. As examples of this technique, the tensile testing of skin and the compressive testing of cartilage are described. These protocols can also be directly applied to synthetic materials to ensure that the mechanical properties are similar to the native tissue. Protocols to assess the mechanical properties of human native tissue will allow a benchmark by which to create suitable tissue-engineered substitutes.

Tissue biomechanics is important for maintaining cell shape and function and for determining phenotype. This report demonstrates non-destructive mechanical protocols for characterizing elastic and viscoelastic properties of human soft tissues, which can be directly applied to tissue-engineered substrates to allow a close matching of engineered materials to native tissue.

Regenerative medicine aims to engineer materials to replace or restore damaged or diseased organs. The mechanical properties of such materials should ...

An Experimental and Finite Element Protocol to Investigate the Transport of Neutral and Charged Solutes across Articular Cartilage

Osteoarthritis (OA) is a debilitating disease that is associated with degeneration of articular cartilage and subchondral bone. Degeneration of articular cartilage impairs its load-bearing function substantially as it experiences tremendous chemical degradation, i.e. proteoglycan loss and collagen fibril disruption. One promising way to investigate chemical damage mechanisms during OA is to expose the cartilage specimens to an external solute and monitor the diffusion of the molecules. The degree of cartilage damage (i.e. concentration and configuration of essential macromolecules) is associated with collisional energy loss of external solutes while moving across articular cartilage creates different diffusion characteristics compared to healthy cartilage. In this study, we introduce a protocol, which consists of several steps and is based on previously developed experimental micro-Computed Tomography (micro-CT) and finite element modeling. The transport of charged and uncharged iodinated molecules is first recorded using micro-CT, which is followed by applying biphasic-solute and multiphasic finite element models to obtain diffusion coefficients and fixed charge densities across cartilage zones.

We propose a protocol to investigate the transport of charged and uncharged molecules across articular cartilage with the aid of recently developed experimental and numerical methods.

Osteoarthritis (OA) is a debilitating disease that is associated with degeneration of articular cartilage and subchondral bone. Degeneration of articular ...

A Test Bed to Examine Helmet Fit and Retention and Biomechanical Measures of Head and Neck Injury in Simulated Impact

Conventional wisdom and the language in international helmet testing and certification standards suggest that appropriate helmet fit and retention during an impact are important factors in protecting the helmet wearer from impact-induced injury. This manuscript aims to investigate impact-induced injury mechanisms in different helmet fit scenarios through analysis of simulated helmeted impacts with an anthropometric test device (ATD), an array of headform acceleration transducers and neck force/moment transducers, a dual high speed camera system, and helmet-fit force sensors developed in our research group based on Bragg gratings in optical fiber. To simulate impacts, an instrumented headform and flexible neck fall along a linear guide rail onto an anvil. The test bed allows simulation of head impact at speeds up to 8.3 m/s, onto impact surfaces that are both flat and angled. The headform is fit with a crash helmet and several fit scenarios can be simulated by making context specific adjustments to the helmet position index and/or helmet size. To quantify helmet retention, the movement of the helmet on the head is quantified using post-hoc image analysis. To quantify head and neck injury potential, biomechanical measures based on headform acceleration and neck force/moment are measured. These biomechanical measures, through comparison with established human tolerance curves, can estimate the risk of severe life threatening and/or mild diffuse brain injury and osteoligamentous neck injury. To our knowledge, the presented test-bed is the first developed specifically to assess biomechanical effects on head and neck injury relative to helmet fit and retention.

Using an anthropometric head and neck, optical fiber-based fit force transducers, an array of head acceleration and neck force/moment transducers, and a dual high speed camera system, we present a test bed to study helmet retention and effects on biomechanical measures of head and neck injury secondary to head impact.

Conventional wisdom and the language in international helmet testing and certification standards suggest that appropriate helmet fit and retention during ...

Methods for In Vivo Biomechanical Testing on Brachial Plexus in Neonatal Piglets

Neonatal brachial plexus palsy (NBPP) is a stretch injury that occurs during the birthing process in nerve complexes located in the neck and shoulder regions, collectively referred to as the brachial plexus (BP). Despite recent advances in obstetrical care, the problem of NBPP continues to be a global health burden with an incidence of 1.5 cases per 1,000 live births. More severe types of this injury can cause permanent paralysis of the arm from the shoulder down. Prevention and treatment of NBPP warrants an understanding of the biomechanical and physiological responses of newborn BP nerves when subjected to stretch. Current knowledge of the newborn BP is extrapolated from adult animal or cadaveric BP tissue instead of in vivo neonatal BP tissue. This study describes an in vivo mechanical testing device and procedure to conduct in vivo biomechanical testing in neonatal piglets. The device consists of a clamp, actuator, load cell, and camera system that apply and monitor in vivo strains and loads until failure. The camera system also allows monitoring of the failure location during rupture. Overall, the presented method allows for a detailed biomechanical characterization of neonatal BP when subjected to stretch.

Methods for In Vivo Biomechanical Testing on Brachial Plexus in Neonatal Piglets

Presented here are methods to perform in vivo biomechanical testing on brachial plexus in a neonatal piglet model.

Neonatal brachial plexus palsy (NBPP) is a stretch injury that occurs during the birthing process in nerve complexes located in the neck and shoulder ...

Biaxial Basal Tone and Passive Testing of the Murine Reproductive System Using a Pressure Myograph

The female reproductive organs, specifically the vagina and cervix, are composed of various cellular components and a unique extracellular matrix (ECM). Smooth muscle cells exhibit a contractile function within the vaginal and cervical walls. Depending on the biochemical environment and the mechanical distension of the organ walls, the smooth muscle cells alter the contractile conditions. The contribution of the smooth muscle cells under baseline physiological conditions is classified as a basal tone. More specifically, a basal tone is the baseline partial constriction of smooth muscle cells in the absence of hormonal and neural stimulation. Furthermore, the ECM provides structural support for the organ walls and functions as a reservoir for biochemical cues. These biochemical cues are vital to various organ functions, such as inciting growth and maintaining homeostasis. The ECM of each organ is composed primarily of collagen fibers (mostly collagen types I, III, and V), elastic fibers, and glycosaminoglycans/proteoglycans. The composition and organization of the ECM dictate the mechanical properties of each organ. A change in ECM composition may lead to the development of reproductive pathologies, such as pelvic organ prolapse or premature cervical remodeling. Furthermore, changes in ECM microstructure and stiffness may alter smooth muscle cell activity and phenotype, thus resulting in the loss of the contractile force.
In this work, the reported protocols are used to assess the basal tone and passive mechanical properties of the nonpregnant murine vagina and cervix at 4-6 months of age in estrus. The organs were mounted in a commercially available pressure myograph and both pressure-diameter and force-length tests were performed. Sample data and data analysis techniques for the mechanical characterization of the reproductive organs are included. Such information may be useful for constructing mathematical models and rationally designing therapeutic interventions for women&#8217;s health pathologies.

This protocol utilized a commercially available pressure myograph system to perform pressure myograph testing on the murine vagina and cervix. Utilizing media with and without calcium, the contributions of the smooth muscle cells (SMC) basal tone and passive extracellular matrix (ECM) were isolated for the organs under estimated physiological conditions.

The female reproductive organs, specifically the vagina and cervix, are composed of various cellular components and a unique extracellular matrix (ECM). ...

The Quantification of Injectability by Mechanical Testing

Injectable biomaterials are becoming increasingly popular for the minimally invasive delivery of drugs and cells. These materials are typically more viscous than traditional aqueous injections and may be semi-solid, therefore, their injectability cannot be assumed. This protocol describes a method to objectively assess the injectability of these materials using a standard mechanical tester. The syringe plunger is compressed by the crosshead at a set rate, and the force is measured. The maximum or plateau force value can then be used for comparison between samples, or to an absolute force limit. This protocol can be used with any material, and any syringe and needle size or geometry. The results obtained may be used to make decisions about formulations, syringe and needle sizes early in the translational process. Further, the effects of altering formulations on injectability may be quantified, and the optimum time to inject temporally changing materials determined. This method is also suitable as a reproducible way to examine the effects of injection on a material, to study phenomena such as self-healing and filter pressing or study the effects of injection on cells. This protocol is faster and more directly applicable to injectability than rotational rheology, and requires minimal post processing to obtain key values for direct comparisons.

Presented here is a protocol for quantitatively evaluating the injectability of a material through a syringe-needle system using a standard mechanical testing rig.

Injectable biomaterials are becoming increasingly popular for the minimally invasive delivery of drugs and cells. These materials are typically more ...

Optical Coherence Tomography Based Biomechanical Fluid-Structure Interaction Analysis of Coronary Atherosclerosis Progression

In this paper, we present a complete workflow for the biomechanical analysis of atherosclerotic plaque in the coronary vasculature. With atherosclerosis as one of the leading causes of global death, morbidity and economic burden, novel ways of analyzing and predicting its progression are needed. One such computational method is the use of fluid-structure interaction (FSI) to analyze the interaction between the blood flow and artery/plaque domains. Coupled with in vivo imaging, this approach could be tailored to each patient, assisting in differentiating between stable and unstable plaques. We outline the three-dimensional reconstruction process, making use of intravascular Optical Coherence Tomography (OCT) and invasive coronary angiography (ICA). The extraction of boundary conditions for the simulation, including replicating the three-dimensional motion of the artery, is discussed before the setup and analysis is conducted in a commercial finite element solver. The procedure for describing the highly nonlinear hyperelastic properties of the artery wall and the pulsatile blood velocity/pressure is outlined along with setting up the system coupling between the two domains. We demonstrate the procedure by analyzing a non-culprit, mildly stenotic, lipid-rich plaque in a patient following myocardial infarction. Established and emerging markers related to atherosclerotic plaque progression, such as wall shear stress and local normalized helicity, respectively, are discussed and related to the structural response in the artery wall and plaque. Finally, we translate the results to potential clinical relevance, discuss limitations, and outline areas for further development. The method described in this paper shows promise for aiding in the determination of sites at risk of atherosclerotic progression and, hence, could assist in managing the significant death, morbidity, and economic burden of atherosclerosis.

There is a need to determine which atherosclerotic lesions will progress in the coronary vasculature to guide intervention before myocardial infarction occurs. This article outlines the biomechanical modeling of arteries from Optical Coherence Tomography using fluid-structure interaction techniques in a commercial finite element solver to help predict this progression.

In this paper, we present a complete workflow for the biomechanical analysis of atherosclerotic plaque in the coronary vasculature. With atherosclerosis ...

Construction of a Human Aorta Smooth Muscle Cell Organ-On-A-Chip Model for Recapitulating Biomechanical Strain in the Aortic Wall

Conventional two-dimensional cell culture techniques and animal models have been used in the study of human thoracic aortic aneurysm and dissection (TAAD). However, human TAAD sometimes cannot be characterized by animal models. There is an apparent species gap between clinical human studies and animal experiments that may hinder the discovery of therapeutic drugs. In contrast, the conventional cell culture model is unable to simulate in vivo biomechanical stimuli. To this end, microfabrication and microfluidic techniques have developed greatly in recent years, providing novel techniques for establishing organoids-on-a-chip models that replicate the biomechanical microenvironment. In this study, a human aorta smooth muscle cell organ-on-a-chip (HASMC-OOC) model was developed to simulate the pathophysiological parameters of aortic biomechanics, including the amplitude and frequency of cyclic strain experienced by human aortic smooth muscle cells (HASMCs) that play a vital role in TAAD. In this model, the morphology of HASMCs became elongated in shape, aligned perpendicularly to the strain direction, and presented a more contractile phenotype under strain conditions than under static conventional conditions. This was consistent with the cell orientation and phenotype in native human aortic walls. Additionally, using bicuspid aortic valve-related TAAD (BAV-TAAD) and tricuspid aortic valve-related TAAD (TAV-TAAD) patient-derived primary HASMCs, we established BAV-TAAD and TAV-TAAD disease models, which replicate HASMC characteristics in TAAD. The HASMC-OOC model provides a novel in vitro platform that is complementary to animal models for further exploring the pathogenesis of TAAD and discovering therapeutic targets.

Here, we developed a human aorta smooth muscle cell organ-on-a-chip model to replicate the in vivo biomechanical strain of smooth muscle cells in the human aortic wall.

Conventional two-dimensional cell culture techniques and animal models have been used in the study of human thoracic aortic aneurysm and dissection ...