This work is supported by the National Institutes of Health (1R01HL151738) and the American Heart Association (18TPA34170169).

The following protocol has been approved by the Institutional Animal Care and Use Committee of Wayne State University. The protocol here describes steps for utilizing rodent experimental subjects, but can be adapted for use in other model organisms.
1. Preparation
<ol>
	<li>Obtain the experimental subject and allow it to acclimate to the lab.</li>
	<li>Prepare 250 mL of perfusion solution and 250 mL of modified Tyrode&#39;s solution (Table 1), by oxygenating each to 10-20 ppm of O2, with pH 7.3.</li>
	<li>Prepare a 5 mL syringe with a cannula attached. Fill the syringe with perfusion solution. Use cannulas that are 23 G for a mouse, and 18 or 16 G for a small or large rat, respectively. Slide a 1 mm length of polyethylene (PE) tubing on the end of a blunted needle (Table of Materials), to help prevent the aorta from slipping off the cannula after being tied.</li>
	<li>Complete preparation of the dissection and cannulation area (Figure 1)
	<ol>
		<li>Fill two clean weigh boats at least halfway with perfusion solution. Submerge the tip of the cannula in perfusion solution with at least one double-looped suture placed around the cannula. Hold the syringe in place with a bar clamp.</li>
		<li>Place surgical scissors, a hemostat, iris forceps, a pair of small curved scissors, and two #3 forceps for easy access.</li>
	</ol>
	</li>
	<li>Prepare the experimental system (Figure 2) by priming and circulating the modified Tyrode&#39;s solution throughout the experimental system. Ensure that the chamber is completely full and stable.</li>
	<li>Turn on all data acquisition boxes, including the force transducer, the length motor, the pacing system, the temperature control system, and the data acquisition computer.</li>
	<li>Copy and rename a template folder containing the required *.dap and pointer files to reference the current experiment. Open the data acquisition software.</li>
	<li>Prepare the trabecula isolation tools (Vannas scissors, #5 or #55 forceps, glass probe) by submerging their tips in 10% (w/v) bovine serum albumin (BSA) in ultrapure water or perfusion solution, to coat the metal with protein (Figure 2B). 
	&#8203;NOTE: The entire experimental system must be prepared prior to dissection.</li>
</ol>
2. Gross dissection and cannulation
<ol>
	<li>Record the identification, body weight, and other relevant information of the animal subject.</li>
	<li>Optionally, inject 1,000 U/kg of heparin in sterile saline to the rodent via intraperitoneal injection at least 10 min prior to euthanasia, to minimize risks of clotting before coronary perfusion.</li>
	<li>Place the animal in an induction chamber and induce general anesthesia using 3%-5% isoflurane vaporized in 100% oxygen, according to standard procedure. 
	NOTE: Turn on any external scavengers, down draft tables, or fume hoods used to minimize exposure to isoflurane.</li>
	<li>Once the rodent loses its righting reflex and the breathing rate has slowed:
	<ol>
		<li>(For a rat) remove the animal from the induction chamber and place it supine on a dissection pad, with continued anesthesia through a nose cone.</li>
		<li>(For a mouse) perform cervical dislocation immediately after removing the animal from the induction chamber. The nose cone is not necessary. Turn the isoflurane vaporizer to 0%.</li>
	</ol>
	</li>
	<li>If needed, affix the upper limbs of the rodent to the dissection pad away from the chest wall using tape supported by pins, taking care not to pierce into the animal. Check for proper anesthesia depth (lack of toe-pinch response) prior to proceeding to the dissection.</li>
	<li>Using surgical scissors (Mayo for rats, Metzenbaum for mice), just below the xyphoid process, cut the skin transverse across the full width of the rodent&#39;s abdomen into the peritoneal space.</li>
	<li>Using surgical scissors, make two vertical cuts parasagittal (up the sides of the chest wall) on both the left and right sides of the chest wall from the transverse cut. Then, cut across the diaphragm, connecting the parasagittal cuts, and freeing the thoracic space.</li>
	<li>Clamp and lift the xyphoid process using a hemostat toward the rodent&#39;s head, to move the sternum and chest wall up toward the rodent&#39;s head, exposing the thoracic space and the heart. If needed, quickly extend the parasagittal cuts to near the second vertebral space, to fully expose the heart and/or break the pericardial membrane.</li>
	<li>Using curved iris forceps, carefully lift the heart to visualize the great vessels. Place the forceps between the myocardium and the spine of the rodent, clamp down on the greater vessels, and lift the heart, taking care not to clamp the atria or ventricular segments.
	<ol>
		<li>While slightly lifting the heart, quickly place the curved iris scissors (concave up) between the curved iris forceps and the spine of the rodent, and cut the great vessels and lungs away from the heart.&#160;Quickly move the heart into a weigh boat or beaker containing fresh perfusion solution, and shake to help clear blood off of the heart.&#160;</li>
	</ol>
	</li>
	<li>Turn the isoflurane vaporizer to 0%, if not already done. If large segments of lung, pericardial, or other tissues remain attached to the heart, carefully cut away and discard them at this time to minimize interference with the cannulation.</li>
	<li>Using the curved iris forceps, move the heart into a clean weigh boat or beaker, with the prepared cannula submerged. Cannulate the aorta, secure the aorta by tightening the looped silk suture, and flush with up to 5 mL of perfusion solution.</li>
	<li>Remove the heart from the cannula and place it in a silicone elastomer-coated weigh dish to prepare for trabecula isolation.</li>
</ol>
3. Isolation and equilibration of trabecula
<ol>
	<li>Place the heart in the silicone elastomer-coated dish under a stereomicroscope and illuminate it.</li>
	<li>Locate the right ventricular outflow tract. Pin the left atrium and the ventricular apex to the silicone elastomer in the dish.</li>
	<li>Using long Vannas scissors, cut from the right ventricular outflow tract to the apex along the septum. Cut from the right ventricular outflow tract to the right atrium near the aorta, then cut through the right atrium (Figure 3B).</li>
	<li>Using forceps, carefully pull open the right ventricular (RV) free wall from the outflow tract, being careful not to stretch the tissue. 
	NOTE: Experimenters may find thin, white connective tissue strands, which can be cut without concern. Larger, pink (tissue) colored strands should be evaluated with care, as they may be trabeculae that can be isolated.</li>
	<li>Pin the free wall of the right ventricle triangle to the dish, to expose the right ventricle (Figure 3C).</li>
	<li>Using a glass pipette that has been melted and formed with a thin (&#60;500 &#181;m diameter) but not sharp end, search the exposed endocardium for free-standing trabeculae (Figure 3C). 
	NOTE: A free-standing trabecula is a strip of muscle that one can fully probe underneath. Use trabeculae that have parallel sides (constant width), avoiding triangular papillary muscles. Be careful during this process and subsequent dissection, as applying strain on the trabecula can cause damage, reducing the developed force.</li>
	<li>Dissect the trabecula using small Vannas scissors. Leave a &#8805;1 mm cubed piece of tissue at each end of the trabecula to allow for attachment. Do not stretch the trabecula where possible, and minimize contact of metal tools, as both can also cause damage to the muscle. Relocate the pins holding the heart as needed to minimize strain on the muscle near the trabeculae identified.</li>
	<li>Cut ~2 inches off the end of a 7 mL transfer pipette, slowly draw the trabecula into the pipette, and transfer it into a new weigh dish containing 50% perfusion solution and 50% modified Tyrode&#39;s solution. Allow the muscle to equilibrate to the increase in extracellular calcium within the mixed solution for several minutes.
	<ol>
		<li>Repeat steps 3.7-3.8 to dissect additional trabecula at this time, to obtain additional trabecula as backup.</li>
	</ol>
	</li>
	<li>Turn off the pump supplying superfusion and/or suction to the experimental chamber. Using the large bore transfer pipette, move the trabecula into the experimental chamber filled with Tyrode&#39;s solution.</li>
	<li>Pin one &#62;1 mm cube piece of tissue at the end of the trabecula to a hook on the force transducer, then pin the second cube to the motor. 
	NOTE: Separate the motor and force transducer when mounting the first side for better access to the transducer, then move the hooks as close together as possible when mounting the second cube of tissue while the trabecula remains slack.</li>
	<li>Restart superfusion and begin pacing the muscle to determine the threshold voltage. Pace at 20% above the threshold voltage for approximately 1 h to allow the trabecula to equilibrate.</li>
	<li>At the end of this equilibrium period, slowly stretch the muscle using the micrometer connected to the motor until optimal developed stress generation is achieved, by observing the developed (minimum to peak) tension. Stop increasing the muscle length when the passive diastolic tension rises faster than the peak tension, indicating the optimal length has been passed.</li>
	<li>Turn off the transmitted microscope illumination, and illuminate the trabecula using a gooseneck illuminator at a steep angle. Using a camera connected through the microscope optics that has previously been calibrated, capture an image of the trabecula during diastole into the experimental folder. If the trabecula is wider than the camera&#39;s field of view, take multiple images across the muscle.</li>
	<li>Open the image(s) in an imaging software that reports pixel distances.
	<ol>
		<li>Measure the pixels distance of the muscle diameter four times along its length. Measure the muscle (trabecula) length in pixels, excluding the large cube of tissue.</li>
		<li>If the muscle length is longer than the field of view, use reference points along the muscle to measure the entire length.</li>
	</ol>
	</li>
	<li>Using the template in the experimental folder, average the diameter measures, and convert the diameter and lengths from pixels to mm using a previously obtained calibration. Calculate the cross-sectional area as &#960;*diameter2/4 in mm2, and the muscle length in &#181;m.</li>
</ol>
4. Data acquisition
<ol>
	<li>Once the muscle is equilibrated, open the data acquisition software, select Experiments | Length Control, and enter the calibrated trabecula length (FL) and cross-sectional area (area [m2]) in the Calibration box.</li>
	<li>From the template folder (Step 1.7), ensure that the freeform_file.txt points to the correct folder, and open the freeform.dap file in a text editor. Set the isotonic level (isoton) to 32,000 in the *.dap file.</li>
	<li>In the Length Control box, select the Freeform tab and browse to the appropriate Freeform list file. Ensure the Data Filing path is also the correct folder to save data. Begin acquiring data from fully isometric twitches by pressing Run Experiment, before acquiring load-clamp data using a feedback control with proportional and integration gain parameters.</li>
	<li>Acquire load-clamped data by defining the afterload (isoton) in the *.dap file, and iterate values of proportional gain (propgain) and integration (Ki) parameters by saving the file in the text editor. Press Run Experiment in the data acquisition software interface.
	<ol>
		<li>Ensure that the clamp includes relengthening back to its original length (sometimes referred to as relaxation loading5) during this iteration to attain the maximal range of strain rates, by setting mode (flswitch) from one and increasing the threshold for ending the load-clamp (flthreshold) from zero.</li>
	</ol>
	</li>
	<li>Control the end of the load-clamp by changing the mode (flswitch) from one to zero. Repeat the acquisition while changing the end of the load-clamp from zero relengthening to complete relengthening back to the starting length.
	<ol>
		<li>To increase the length, incrementally increase the threshold for ending the load-clamp (flthreshold) from zero until the muscle nearly relengthens back to its original length.</li>
	</ol>
	</li>
	<li>Reset the mode (flswitch = 1) and threshold (flthreshold = 0), and run one final data acquisition.</li>
	<li>If desired, modify the afterload in the study by repeating steps 4.4-4.6. This acquisition may be done immediately.</li>
	<li>If desired, modify the preload by stretching or shortening the muscle, or treat the muscle by adding compounds to the Tyrode&#39;s solution. If altering the preload or adding compounds, wait a minimum of 20 min to ensure that the slow-force response9,15 has stabilized and/or for the compound to fully penetrate the muscle.</li>
	<li>Once the data acquisition is completed, remove the trabecula. If needed, freeze or fix the trabecula for biochemical or histological analysis.</li>
	<li>Clean the work areas and experimental system, flush all tubing with water, and turn off all components.</li>
</ol>
5. Data analysis
<ol>
	<li>Open the data files using a quantitative analysis program by directing the program to the appropriate file path.</li>
	<li>Quantify relaxation by ensuring that the data analysis program analyzes the clamped beat, and that the program correctly acquires the start of the load-clamp.
	<ol>
		<li>Ensure that the end of the load-clamp is identified, so that the relaxation rate (1/&#964;) is quantified from the peak negative derivative of stress during an isometric relaxation.</li>
		<li>Use the Glantz method16 to determine this exponential time constant, or other appropriate quantification of relaxation (minimum derivative of stress, logistic time constant17, or kinematic model18).</li>
	</ol>
	</li>
	<li>Ensure that the data analysis program calculates the strain rate by taking the time derivative of the strain, where strain is calculated as the length as a function of time divided by length at optimal contraction.</li>
	<li>Repeat the above steps for all traces of a given condition.</li>
	<li>Plot the relationship between the relaxation rate and strain rate, limiting the maximum data to a physiological strain rate of less than 1 s-1. Exclude data at low strain rates (Figure 4C), as the relaxation phase may not reflect the exponential decay17,18.</li>
	<li>Obtain the slope of the line between the relaxation rate and the strain rate, and record the slope as the index of MCR.</li>
	<li>Repeat the above analysis for each condition interrogated.</li>
</ol>

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Mechanical control of relaxation (MCR) quantifies the dependence of the myocardial relaxation rate on the strain rate of the muscle proceeding relaxation8,9. Strain rate, rather than afterload, is both necessary and sufficient to modify the relaxation rate8. As interventions to modify the calcium rate have not been proven to substantially improve cardiac relaxation, mechanical intervention may provide new insights into the mechanism and provide a new treatment for diastolic dysfunction.
The protocol to modify the myocardial strain rate described here utilizes an isotonic load-clamp8,9. A strength of the isotonic load-clamp is the quantitative control of the afterload stress. Windkessel-like protocols could be used to further probe changes in afterload, preload, and cardiac work2,6,7. A ramp not controlled by the load-clamp could also be used to better isolate the change in strain from strain rate. Regardless, the afterload itself does not appear to be a strong modifier of the relaxation rate8.
The protocol may also be adapted to approach more physiological conditions for temperature and pacing rate. The current protocol details were used to show the presence of the MCR. Conducting experiments in physiological conditions is generally recommended, depending on the experimental question. However, experiments performed at 37 &#176;C, or at high pacing rates, can more rapidly induce rundown (damage) to the muscle. A solution with improved oxygen carrying capacity may be needed. Further, the data acquisition must be able to sample the length and force fast enough to resolve the rapid twitches and provide feedback control.
The current protocol does not describe the measurement of calcium or the measurement and control of sarcomere lengths. Calcium measurements have been addressed in other protocols11, while sarcomere length measurement can be added with appropriate equipment. Sarcomere length control is not utilized in current MCR studies, because muscle length is the most correlative parameter to the clinical condition19. Further sarcomere length control (vs. muscle length control) would provide specific answers to kinetic questions, but is unlikely to add to translational knowledge due to inter-sarcomere variation and the minimum understanding of sarcomere length changes in vivo.
Three experimental considerations are highlighted here to increase the reproducibility of the data.
First, free-standing cardiac trabeculae may be difficult to find in some animals (unpublished results and communications). While twitching muscles can be found in most rats, a reasonable success rate for obtaining data from trabecula in rats is one in three. Trabecula success may be higher with Brown Norway x Lewis F1 rats, which have also been used historically20 and reported to have more trabeculae (unpublished communications). For mice, success rates are likely to be lower, with less than one in 10 expected for mice from a BL/6 background; however, a higher rate is expected for mice from a FVBN background (unpublished communications and observations).
Second, damage to muscles can reduce output. If the developed forces are less than 10 mN mm-2 at 25 &#176;C and 0.5 Hz pacing, investigators may need to perform troubleshooting to assess if inadvertent stretching or contact between metal forceps and muscle is occurring, if solutions are not properly prepared, or if pacing or experimental equipment is functioning properly. Other protocols utilizing intact trabecula have suggested using Luer-lock syringes as transfer vessels11. While this is possible, especially if the user controls a very slow flow rate or smaller muscle segment, the current protocol utilizes a much larger bore transfer pipette to minimize possible damage. Another step where ischemic damage may occur is during dissection. The aorta should be cannulated and flushed with perfusion solution within 3 min of the first abdominal cut (rat) or cervical dislocation (mouse), similar to limits listed in cardiomyocyte isolation protocols21,22. This minimizes the time when the cardiac tissue is not exposed to the cardioplegia-like perfusion solution. Further, dissections lasting more than 30 min generally do not produce twitching trabecula. Thus, operators should practice rapid but careful dissection to minimize damage. A cross-sectional area above 0.2 mm2 (2 x 10-7 m2) may suffer from core ischemia20.
Third, the way in which muscles are attached to the motor and force transducer should be considered. This protocol currently focuses on hooks and free-standing trabeculae. The sometimes rapid strain rate of the stretch prior to relaxation can cause a muscle to slide if not affixed properly, which is why the current protocol does not use &#34;baskets&#34; to hold the trabecula23,24. Alternate mounting methods (adhesives, clips, etc25,26) can also be considered and validated. The protocol described here utilizes trabeculae and not papillary muscles. The chordae of the papillary muscle induce a series elasticity that can inhibit changes to the MCR9. However, the exact placement of the attachments in the muscle is unlikely to impact the measures, because the trabecula length (and diameter) vary substantially.
A limitation of piercing the muscle ends with hooks is that the mounting point itself can also be damaged. Possible tearing of the affixed muscle tissue with frequent contractions (depending on their strength) may change the length or series elasticity. This rate of tearing is difficult to control. Similarly, damage to the tissue and the hook can be exacerbated during stretching, also potentially causing problems. Visual inspection, and the developed force values remaining &#62;80% of the equilibrated isometric force, should be used to assess whether the preparation is damaged and should be excluded.
Another limitation or consideration affects what experimental questions can be answered by the method. For example, consider the use of 2,3-butanedione monoxime (BDM) in the perfusion solution. BDM is a phosphatase, which may alter the function of the muscle. In addition, the long period of unloading and lack of pacing mean that the latent phosphorylation state has likely changed. Thus, caution should be used if trying to directly assess the muscle contractility of an animal (vs. differences between genotypes or treatments), as the contractile state has likely changed. However, the impact of phosphorylation may be assessed pharmacologically by adding an agonist or antagonist of the pathway.
In summary, MCR provides insight into how relaxation is regulated by muscle movement (strain rate). MCR may help to provide better insight into the diagnosis and monitoring of diastolic disease, along with targets for pharmacological intervention, such as modifying myosin kinetics. The protocol and advice outlined here lay out knowledge developed over several years of trials, and should be applicable to other systems and models of cardiac disease.

Cardiac relaxation is impaired in nearly all forms of heart failure (including heart failure with reduced ejection fraction) and in many cardiovascular diseases. In addition to numerous methods to evaluate cardiac function in permeabilized muscles, the evaluation of intact cardiac muscles is gaining interest. Such tissues are assessed unloaded (ends free to contract) or loaded (length or force controlled). Historically, intact isolated myocytes have been evaluated in an unloaded condition, where the cell body is free to shorten during contraction. Intact cardiac trabeculae are often evaluated in isometric conditions, where the length is not allowed to change, but stress (force per cross-sectional area) is generated. Both intact myocyte and trabeculae methods are beginning to converge with modifications of load1,2.
Protocols for load-clamping a muscle (i.e., controlling a muscle&#39;s developed stress at a specified value that simulates physiologic afterloads) have been developed over several decades3,4,5. In intact cardiac tissues, load-clamps allow researchers to more closely mimic the in vivo cardiac cycle using isotonic or Windkessel-like afterloads6,7,8,9. The goal of this protocol is to obtain data used to quantify the MCR (i.e., the strain rate dependence of the relaxation rate)8,9.
While the MCR protocol has been adapted from prior work, the focus of this protocol (compared to similar protocols utilizing intact cardiac tissues) is on biomechanical mechanisms that modify relaxation. There are several protocols utilizing load-clamping3,4,5,7,10&#160;and protocols focusing on Windkessel models1,2,11, but this protocol describes specifically how stretch prior to relaxation modifies the relaxation rate. We have shown that this control occurs during a proto-diastolic period8, a phase originally described by Wiggers12. In normal healthy hearts, the myocardium undergoes a lengthening strain during ejection prior to aortic valve closure (i.e., prior to isovolumic relaxation)13. This is mimicked by prolonging the duration of afterload control until the muscle begins to stretch. Clinical evidence suggests that this lengthening may be attenuated or lost in disease states14, and the implications and mechanisms of altered end-systolic strain rates have not been fully elucidated. Given the sparse treatment options for diastolic diseases and heart failure with a preserved ejection fraction, we posit that MCR may provide insight into novel mechanisms underlying impaired relaxation.
While the gross dissection described here focuses on rodents, trabecula isolation can be performed from any intact heart, and has previously been used with a human cardiac trabecula8. Similarly, the data acquisition and analysis can also be applied to cardiomyocytes or other isolated muscle types1,10. The discussion includes commentary on possible alterations and adaptations to the method, along with limitations, such as caution against utilizing papillary muscles because of the mechanical properties of the chordae9.

<table><tbody><tr><td>18 or 16 gauge blunted needle/canula</td><td></td><td></td><td>for cannulation of rat aorta, use 1mm of PE160 or PE205 tubing as stop</td></tr><tr><td>2,3-Butanedione Monoxime</td><td>Sigma-Aldrich</td><td>B0753-25G</td><td></td></tr><tr><td>23 gauge blunted needle/canula</td><td></td><td></td><td>for cannulation of mouse aorta, use 1mm of PE50 tubing as stop</td></tr><tr><td>5 mL syringe</td><td>BD Luer-Lock</td><td>309646</td><td></td></tr><tr><td>95% Oxygen/5% CO&lt;sub&gt;2&lt;/sub&gt;</td><td>AirGas</td><td>Z02OX9522000043</td><td></td></tr><tr><td>Anethesia system</td><td>EZ Systems</td><td>EZ-SA800</td><td>Can use any appropriate anethesia method/system</td></tr><tr><td>Bovine Serum Albumin</td><td>Fisher BioReagents</td><td>BP-1600</td><td>to coat tips of fine forcepts, scissors</td></tr><tr><td>Calcium Chloride Dihydrate</td><td>Fisher Chemical</td><td>C79-500</td><td></td></tr><tr><td>Containers/dissection dishes</td><td>FisherBrand</td><td>08-732-113</td><td>Weigh dishes for creating dissection plates</td></tr><tr><td>Crile Hemostat</td><td>Fine Science Tools</td><td>13005-14</td><td>for mouse gross dissection</td></tr><tr><td>D-(+)-Glucose</td><td>Sigma-Aldrich</td><td>G8270-1KG</td><td></td></tr><tr><td>Data acquisition software</td><td>SLControl</td><td></td><td></td></tr><tr><td>Data acquisition system</td><td>MicrostarLabs</td><td>DAP5216a</td><td>Can use any DAQ.&amp;nbsp; This is a PCI based data acqusition for use with SLControl; must have a PC with a PCI slot</td></tr><tr><td>Data analysis software</td><td>Mathworks</td><td>Matlab</td><td>Custom Script</td></tr><tr><td>Dumont #3 Forceps</td><td>Fine Science Tools</td><td>11231-30</td><td>2x for cannulation of aorta</td></tr><tr><td>Dumont #5 Forceps</td><td>Fine Science Tools</td><td>11254-20</td><td>2x for trabecula isolation</td></tr><tr><td>Experimental system</td><td>Aurora Scientific</td><td>801C</td><td>Can use any appropriate experimental chamber with force and length control</td></tr><tr><td>Fine Scissors, curved</td><td>Fine Science Tools</td><td>14061-09</td><td>for removal of heart</td></tr><tr><td>Gooseneck Piggyback Illuminator</td><td>AmScope</td><td>LED-6WA</td><td></td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H3375-250G</td><td></td></tr><tr><td>Imaging software</td><td>IrfanView</td><td></td><td></td></tr><tr><td>Iris Forceps</td><td>World Precision Instruments</td><td>15915</td><td>for removal of heart</td></tr><tr><td>Isoflurane</td><td>VetOne</td><td>502017</td><td></td></tr><tr><td>Magnesium Chloride Hexahydrate</td><td>Sigma-Aldrich</td><td>M2670-100G</td><td></td></tr><tr><td>Magnesium Sulfate</td><td>Sigma-Aldrich</td><td>M7506-500G</td><td></td></tr><tr><td>Mayo Scissors</td><td>Fine Science Tools</td><td>14110-15</td><td>for rat gross dissection</td></tr><tr><td>Metzenbaum Scissors</td><td>Fine Science Tools</td><td>14116-14</td><td>for mouse gross dissection</td></tr><tr><td>Microscope connected camera</td><td>Flir</td><td>BFS-U3-27S5M-C</td><td>Includes acquisition software</td></tr><tr><td>Microscope/digital imaging system</td><td>Olympus</td><td>IX-73</td><td>Can use any appropriate microscope.&amp;nbsp; Needed to measure muscle length, cross sectional area</td></tr><tr><td>Mounting Pin/Needle</td><td>BD PrecisionGlide</td><td>305136</td><td>For holding heart to dish.&amp;nbsp; 27 G x 1-1/4</td></tr><tr><td>Mounting Pin/Needle</td><td>Fine Science Tools</td><td>26000-40</td><td>For holding heart to dish. 0.4mm diameter insect pin (Alt to 27G needle)</td></tr><tr><td>Oxygen (O&lt;sub&gt;2&lt;/sub&gt;)</td><td>AirGas</td><td>OX USP300</td><td></td></tr><tr><td>Peristaltic Pump</td><td>Rainin</td><td>Rabbit</td><td>Can be any means to create flow in experimental chamber</td></tr><tr><td>pH and Oxygen sensor</td><td>Mettler Toledo</td><td>SevenGo pH and DO</td><td></td></tr><tr><td>Potassium Bicarbonate</td><td>Sigma-Aldrich</td><td>237205-100G</td><td></td></tr><tr><td>Potassium Chloride</td><td>Fisher Chemical</td><td>P217-500</td><td></td></tr><tr><td>Potassium Phosphate Monobasic</td><td>Sigma-Aldrich</td><td>795488-500G</td><td></td></tr><tr><td>Rochester-pean Hemostat</td><td>World Precision Instruments</td><td>501708</td><td>for rat gross dissection</td></tr><tr><td>Silk Suture, Size: 4/0</td><td>Fine Science Tools</td><td>18020-40</td><td>cut to ~1.5 inch pieces, soaked in water</td></tr><tr><td>Sodium Bicarbonate</td><td>Sigma-Aldrich</td><td>S6297-250G</td><td></td></tr><tr><td>Sodium Chloride</td><td>Sigma-Aldrich</td><td>S9888-1KG</td><td></td></tr><tr><td>Sodium Hydroxide</td><td>Sigma-Aldrich</td><td>S8045-500G</td><td></td></tr><tr><td>Sodium Phosphate Dibasic</td><td>Sigma-Aldrich</td><td>S7907-100G</td><td></td></tr><tr><td>Stereomicroscope</td><td>AmScope</td><td>SM-1TX</td><td></td></tr><tr><td>Student Vannas Spring Scissors&amp;nbsp;</td><td>Fine Science Tools</td><td>91500-09</td><td>for opening of the RV</td></tr><tr><td>SYLGARD 184 Silicone Elastomer Base</td><td>Dow Corning</td><td>3097358-1004</td><td>For creating dissection plates</td></tr><tr><td>Syringe Holder</td><td>Harbor Frieght</td><td>Helping Hands 60501</td><td>Can be used as alternate for ring stand</td></tr><tr><td>Taurine</td><td>Sigma-Aldrich</td><td>T0625-1KG</td><td></td></tr><tr><td>Transfer Pipette</td><td>FisherBrand</td><td>13-711-7M</td><td>cut ~1&quot; from tip to widen bore</td></tr><tr><td>Vannas Spring Scissors</td><td>Fine Science Tools</td><td>15000-00</td><td>for trabecula isolation</td></tr></tbody></table>

mechanical control of relaxation using intact cardiac trabeculae

Diastolic dysfunction is a common phenotype across cardiovascular disease presentations. In addition to elevated cardiac stiffness (elevated left ventricular end-diastolic pressure), impaired cardiac relaxation is a key diagnostic indicator of diastolic dysfunction. While relaxation requires the removal of cytosolic calcium and deactivation of sarcomeric thin filaments, targeting such mechanisms has yet to provide effective treatments. Mechanical mechanisms, such as blood pressure (i.e., afterload), have been theorized to modify relaxation. Recently, we showed that modifying the strain rate of a stretch, not afterload, was both necessary and sufficient to modify the subsequent relaxation rate of myocardial tissue. The strain rate dependence of relaxation, called the mechanical control of relaxation (MCR), can be assessed using intact cardiac trabeculae. This protocol describes the preparation of a small animal model, experimental system and chamber, isolation of the heart and subsequent isolation of a trabecula, preparation of the experimental chamber, and experimental and analysis protocols. Evidence for lengthening strains in the intact heart suggests that MCR might provide new arenas for better characterization of pharmacological treatments, along with a method to assess myofilament kinetics in intact muscles. Therefore, studying the MCR may elucidate a path to novel approaches and new frontiers in the treatment of heart failure.

A representative data set is shown in Figure 4, and additional results can be found in prior publications8,9. Briefly, the strain rate is calculated from the derivative of the strain, just prior to isometric relaxation. Strain is the length as a function of time divided by the length of the muscle at optimal length. Relaxation rate is calculated as 1/&#964;, where &#964; is the exponential time constant16. Multiple strain rates and their resultant relaxation rates are required to determine the mechanical control of relaxation (MCR). These data are plotted on a relaxation rate versus strain rate graph. The slope of the line provides the MCR index.
Note that end systolic and diastolic strain rates are not likely to exceed 1 s-1. Therefore, the slope should only include strain rates &#60; 1 s-1. The relaxation rate at low strain rates can be confounded by changes in the minimum time derivative of stress (dStress/dtmin), and in these cases, data from stretches less than approximately 0.15 s-1 may be ignored.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64879/64879fig01.jpg" /> 
Figure 1: Setup of gross dissection and heart cannulation area. From right to left: a. The anesthesia induction chamber for the animal is located near the dissection area. b. A snorkel for optional volatile gas scavenger. c. A dissection pad, where the animal will be placed supine, is flanked by (clockwise)&#160;d. a nosecone, to provide continued anesthesia to the rodent, e. hemostats, f. surgical scissors, g. curved fine scissors placed for easy access with the dominant (cutting) hand, h. curved iris forceps placed for easy access with the non-dominant hand. Upper limbs of the rodent may be affixed to the dissection pad using tape, and i. a&#160;dissection dish (or small beaker) with perfusion solution should be placed nearby to rinse the heart. j. An area staged for cannulation must be placed nearby. A syringe with an appropriate cannula is mounted on a ring stand. k. (Inset) A closer image of a 16 G cannula, with 1 mm of PE205 tubing attached and a suture loosely knotted. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64879/64879fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64879/64879fig02.jpg" /> 
Figure 2: Experimental area and tools. (A) Experimental area. The experimental chamber and data acquisition system are prepared on a nearby inverted microscope (left). Trabecula isolation and mounting occurs under a stereoscope (right). (B) Tips of two forceps, large and small Vannas scissors, and a custom glass probe are prepared by soaking in 10% BSA solution. (C) Additional dissection tools. A silicone elastomer-plated dish allows for mounting the heart during fine dissection. A 7 mL transfer pipette with the end cut is shown below the dish and glass probe. The cut tip of the transfer pipette is discarded, and an enlarged bore is used to transfer the muscle, with minimal risk of stretching or dehydration. (D) Magnified image of the end of a&#160;glass pipette, which is approximately 2 mm long and 0.25-0.5 mm in diameter. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64879/64879fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64879/64879fig03.jpg" /> 
Figure 3: Guide to dissection. (A) Gross dissection should begin with a transverse cut (1. green) through the skin into the peritoneal cavity below the xyphoid process (lower ribs and xyphoid are indicated by a thin gray line). Parasagittal cuts from the peritoneal cavity up the ribcage should follow (2. teal and 3. blue lines), after which the diaphragm should be cut. A hemostat can then be used to clamp the xyphoid process and elevate the chest wall toward the head. (B) An intact rat heart pinned to a silicon-elastomer dish, oriented with a view of the right ventricular outflow tract (RVOT), right atrium (RA), aorta (Ao), and left atrium (LA). The first set of cuts should be from the RVOT to the apex along the septum (1. Yellow line). A second set of cuts should be from the RVOT along the base of the heart then across the right atrium (2. orange line). (C) The RVOT can be carefully pulled away from the aorta to open the heart and pin it back. Yellow and orange lines correspond to cuts described in B. Free standing trabeculae are often found near the base of the RV free wall and near the septum, but can occur anywhere (red lines indicate common locations). (D) Magnified view of the glass probe under a trabecula (yellow arrow indicates intersection of trabecula and probe). (E) A trabecula (red arrow) is mounted to the experimental system between a force transducer (left) and motor (center-right), and is flanked by two pacing leads (horizontal above and below the trabecula). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64879/64879fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64879/64879fig04.jpg" /> 
Figure 4: Representative traces and results. (A) A single cardiac trabecula illuminated using a gooseneck LED light at a steep (~75&#176;) angle from the axis of the microscope lens, which enhances contrast of the trabecula. In this example, two images are tiled to show the full length of the trabecula. (B) Stress-time (top) and strain-time (bottom) curves for the same trabecula. An isometric twitch, along with three load-clamp twitches at increasing end systolic strain rates are shown. (C) Representative MCR calculation. MCR is defined as the slope of the line between the relaxation rate and strain rate. As noted in the discussion, strain rates may be restricted to provide a quantitative, reproducible MCR. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64879/64879fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Table 1: Solutions. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64879/64879_Table1.xlsx" target="_blank">Please click here to download this Table.</a>

Watch this Scientific Journal Video about Mechanical Control of Relaxation Using Intact Cardiac Trabeculae at JoVE.com

Mechanical Control of Relaxation Using Intact Cardiac Trabeculae

Rapid myocardial and cardiac relaxation is essential for normal physiology. Mechanical relaxation mechanisms are now known to be dependent on strain rate. This protocol provides an overview of the acquisition and analysis of experiments to further study the mechanical control of relaxation.

Diastolic dysfunction is a common phenotype across cardiovascular disease presentations. In addition to elevated cardiac stiffness (elevated left ...

mechanical-control-of-relaxation-using-intact-cardiac-trabeculae

Nanyang Technological University

Research

JoVE Journal

Medicine

1.1K Views.  Wayne State University. Dit protocol beschrijft een methode om mechanische controle van ontspanning of de spanningsafhankelijkheid van spierontspanning in cardiale trabecula te meten. Het grote voordeel van deze techniek is dat deze zich richt op hartontspanning. Gezien het gebrek aan behandelingsopties voor diastolische ziekten, kan deze techniek worden gebruikt om nieuwe indexen en medicijndoelen met betrekking tot cardiale ontspanning te identificeren.Deze methode wordt getoond met een knaagdierhart, maar ...

Video: Mechanical Control of Relaxation Using Intact Cardiac Trabeculae

In vitro Measurements of Tracheal Constriction Using Mice

Transgenic and knockout mice have been powerful tools for the investigation of the physiology and pathophysiology of airways1,2. In vitro tensometry of isolated tracheal preparations has proven to be a useful assay of airway smooth muscle (ASM) contractile response in genetically modified mice. These in vitro tracheal preparations are relatively simple, provide a robust response, and retain both functional cholinergic nerve endings and muscle responses, even after long incubations.
Tracheal tensometry also provides a functional assay to study a variety of second messenger signaling pathways that affect contraction of smooth muscle. Contraction in trachea is primarily mediated by parasympathetic, cholinergic nerves that release acetylcholine onto ASM (Figure 1). The major ASM acetylcholine receptors are muscarinic M2 and M3 which are Gi/o and Gq coupled receptors, respectively3,4,5. M3 receptors evoke contraction by coupling to Gq to activate phospholipase C, increase IP3 production and IP3-mediated calcium release from the sarcoplasmic reticulum3,6,7. M2/Gi/o signaling is believed to enhance contractions by inhibition of adenylate cyclase leading to a decrease in cAMP levels5,8,9,10. These pathways constitute the so called "pharmaco-contraction coupling" of airway smooth muscle11. In addition, cholinergic signaling through M2 receptors (and modulated by M3 signaling) involves pathways that depolarize the ASM which in turn activate L-type, voltage-dependent calcium channels (Figure 1) and calcium influx (so called "excitation-contraction coupling")4,7. More detailed reviews on signaling pathways controlling airway constriction can be found4,12. The above pathways appear to be conserved between mice and other species. However, mouse tracheas differ from other species in some signaling pathways. Most prominent is their lack of contractile response to histamine and adenosine13,14, both well-known ASM modulators in humans and other species5,15.
Here we present protocols for the isolation of murine tracheal rings and the in vitro measurement of their contractile output. Included are descriptions of the equipment configuration, trachea ring isolation and contractile measurements. Examples are given for evoking contractions indirectly using high potassium stimulation of nerves and directly by depolarization of ASM muscle to activate voltage-dependent calcium influx (1. high K+, Figure 1). In addition, methods are presented for stimulations of nerves alone using electric field stimulation (2. EFS, Figure 1), or for direct stimulation of ASM muscle using exogenous neurotransmitter applied to the bath (3. exogenous ACH, Figure 1). This flexibility and ease of preparation renders the isolated trachea ring model a robust and functional assay for a number of signaling cascades involved in airway smooth muscle contraction.

In vitro Measurements of Tracheal Constriction Using Mice

Transgenic mice have been extremely useful in ascribing physiological function to genes. As such, research in general, and functional studies of airway, in particular, have undergone a remarkable shift toward murine models. Here we provide protocols for in vitro trachea constriction studies to evaluate smooth muscle function in murine airway.

Transgenic and knockout mice have been powerful tools for the investigation of the physiology and pathophysiology of airways1,2. In vitro tensometry of ...

Trabecular Meshwork Response to Pressure Elevation in the Living Human Eye

The mechanical characteristics of the trabecular meshwork (TM) are linked to outflow resistance and intraocular pressure (IOP) regulation. The rationale behind this technique is the direct observation of the mechanical response of the TM to acute IOP elevation. Prior to scanning, IOP is measured at baseline and during IOP elevation. The limbus is scanned by spectral-domain optical coherence tomography at baseline and during IOP elevation (ophthalmodynamometer (ODM) applied at 30 g force). Scans are processed to enhance visualization of the aqueous humor outflow pathway using ImageJ. Vascular landmarks are used to identify corresponding locations in baseline and IOP elevation scan volumes. Schlemm canal (SC) cross-sectional area (SC-CSA) and SC length from anterior to posterior along its long axis are measured manually at 10 locations within a 1 mm segment of SC. Mean inner to outer wall distance (short axis length) is calculated as the area of SC divided by its long axis length. To examine the contribution of adjacent tissues to the effect IOP elevations, measurements are repeated without and with smooth muscle relaxation with instillation of tropicamide. TM migration into SC is resisted by TM stiffness, but is enhanced by the support of its attachment to adjacent smooth muscle within the ciliary body. This technique is the first to measure the living human TM response to pressure elevation in situ under physiological conditions within the human eye.

Trabecular meshwork (TM) migration into Schlemm&#8217;s canal space can be induced by acute pressure elevation by ophthalmodynamometer, and observed by spectral domain optical coherence tomography. The goal of this method is to quantify the morphometric response of the living outflow tract to acute pressure elevation in living tissues in situ.

The mechanical characteristics of the trabecular meshwork (TM) are linked to outflow resistance and intraocular pressure (IOP) regulation. The rationale ...

Light-sheet Fluorescence Microscopy to Capture 4-Dimensional Images of the Effects of Modulating Shear Stress on the Developing Zebrafish Heart

The hemodynamic forces experienced by the heart influence cardiac development, especially trabeculation, which forms a network of branching outgrowths from the myocardium. Genetic program defects in the Notch signaling cascade are involved in ventricular defects such as Left Ventricular Non-Compaction Cardiomyopathy or Hypoplastic Left Heart Syndrome. Using this protocol, it can be determined that shear stress driven trabeculation and Notch signaling are related to one another. Using Light-sheet Fluorescence Microscopy, visualization of the developing zebrafish heart was possible. In this manuscript, it was assessed whether hemodynamic forces modulate the initiation of trabeculation via Notch signaling and thus, influence contractile function occurs. For qualitative and quantitative shear stress analysis, 4-D (3-D+time) images were acquired during zebrafish cardiac morphogenesis, and integrated light-sheet fluorescence microscopy with 4-D synchronization captured the ventricular motion. Blood viscosity was reduced via gata1a-morpholino oligonucleotides (MO) micro-injection to decrease shear stress, thereby, down-regulating Notch signaling and attenuating trabeculation. Co-injection of Nrg1 mRNA with gata1a MO rescued Notch-related genes to restore trabeculation. To confirm shear stress driven Notch signaling influences trabeculation, cardiomyocyte contraction was further arrested via tnnt2a-MO to reduce hemodynamic forces, thereby, down-regulating Notch target genes to develop a non-trabeculated myocardium. Finally, corroboration of the expression patterns of shear stress-responsive Notch genes was conducted by subjecting endothelial cells to pulsatile flow. Thus, the 4-D light-sheet microscopy uncovered hemodynamic forces underlying Notch signaling and trabeculation with clinical relevance to non-compaction cardiomyopathy.

Here, we present a protocol to visualize developing hearts in zebrafish in 4-Dimensions (4-D). 4-D imaging, via light-sheet fluorescence microscopy (LSFM), takes 3-Dimensional (3-D) images over time, to reconstruct developing hearts. We show qualitatively and quantitatively that shear stress activates endocardial Notch signaling during chamber development, which promotes cardiac trabeculation.

The hemodynamic forces experienced by the heart influence cardiac development, especially trabeculation, which forms a network of branching outgrowths ...

Bioengineering

Biaxial Mechanical Characterizations of Atrioventricular Heart Valves

Extensive biaxial mechanical testing of the atrioventricular heart valve leaflets can be utilized to derive&#160;optimal parameters used in constitutive models, which provide a mathematical representation of the mechanical function of those structures. This presented biaxial mechanical testing protocol involves (i) tissue acquisition, (ii) the preparation of tissue specimens, (iii) biaxial mechanical testing, and (iv) postprocessing of the acquired data. First, tissue acquisition requires obtaining porcine or ovine hearts from a local Food and Drug Administration-approved abattoir for later dissection to retrieve the valve leaflets. Second, tissue preparation requires using tissue specimen cutters on the leaflet tissue to extract a clear zone for testing. Third, biaxial mechanical testing of the leaflet specimen requires the use of a commercial biaxial mechanical tester, which consists of force-controlled, displacement-controlled, and stress-relaxation testing protocols to characterize the leaflet tissue&#39;s mechanical properties. Finally, post-processing requires the use of data image correlation techniques and force and displacement readings to summarize the tissue&#39;s mechanical behaviors in response to external loading. In general, results from biaxial testing demonstrate that the leaflet tissues yield a nonlinear, anisotropic mechanical response. The presented biaxial testing procedure is advantageous to other methods since the method presented here allows for a more comprehensive characterization of the valve leaflet tissue under one unified testing scheme, as opposed to separate testing protocols on different tissue specimens. The proposed testing method has its limitations in that shear stress is potentially present in the tissue sample. However, any potential shear is presumed negligible.

This protocol involves characterizations of atrioventricular valve leaflets with force-controlled, displacement-controlled, and stress-relaxation biaxial mechanical testing procedures. Results acquired with this protocol can be used for constitutive model development to simulate the mechanical behavior of functioning valves under a finite element simulation framework.

Extensive biaxial mechanical testing of the atrioventricular heart valve leaflets can be utilized to derive&#160;optimal parameters used in constitutive ...

Assessment of Sarcoplasmic Reticulum Calcium Reserve and Intracellular Diastolic Calcium Removal in Isolated Ventricular Cardiomyocytes

Intracellular calcium recycling plays a critical role in regulation of systolic and diastolic function in cardiomyocytes. Cardiac sarcoplasmic reticulum (SR) serves as a Ca2+ reservoir for contraction, which reuptakes intracellular Ca2+ during relaxation. The SR Ca2+ reserve available for beats is determinate for cardiac contractibility, and the removal of intracellular Ca2+ is critical for cardiac diastolic function. Under some pathophysiological conditions, such as diabetes and heart failure, impaired calcium clearance and SR Ca2+ store in cardiomyocytes may be involved in the progress of cardiac dysfunction. Here, we describe a protocol to evaluate SRCa2+ reserve and diastolic Ca2+ removal. Briefly, a single cardiomyocyte was enzymatically isolated, and the intracellular Ca2+ fluorescence indicated by Fura-2 was recorded by a calcium imaging system. To employ caffeine for inducing total SR Ca2+ release, we preset an automatic perfusion switch program by interlinking the stimulation system and the perfusion system. Then, the mono-exponential curve fitting was used for analyzing decay time constants of calcium transients and caffeine-induced calcium pulses. Accordingly, the contribution of the SR Ca2+-ATPase (SERCA) and Na+-Ca2+ exchanger (NCX) to diastolic calcium removal was evaluated.

Intracellular calcium recycling plays a critical role in regulation of systolic and diastolic function in cardiomyocytes. Here, we describe a protocol to evaluate sarcoplasmic reticulum Ca2+ reserve and diastolic calcium removal function in cardiomyocytes by a calcium imaging system.

Intracellular calcium recycling plays a critical role in regulation of systolic and diastolic function in cardiomyocytes. Cardiac sarcoplasmic reticulum ...

Simultaneous Brightfield, Fluorescence, and Optical Coherence Tomographic Imaging of Contracting Cardiac Trabeculae Ex Vivo

In cardiac muscle, intracellular Ca2+ transients activate contractile myofilaments, causing contraction, macroscopic shortening, and geometric&#160;deformation. Our understanding of the internal relationships between these events has been limited because we can neither &#39;see&#39; inside the muscle nor precisely track the spatio-temporal nature of excitation-contraction dynamics. To resolve these problems, we have constructed a device that combines a suite of imaging modalities. Specifically, it integrates a brightfield microscope to measure local changes of sarcomere length and tissue strain, a fluorescence microscope to visualize the Ca2+ transient, and an optical coherence tomograph to capture the tissue&#39;s geometric&#160;changes throughout the time-course of a cardiac cycle. We present here the imaging infrastructure and associated data collection framework. Data are collected from isolated rod-like tissue structures known as trabeculae carneae. In our instrument, a pair of position-controlled platinum hooks hold each end of an ex vivo muscle sample while it is continuously superfused with nutrient-rich saline solution. The hooks are under independent control, permitting real-time control of muscle length and force. Lengthwise translation enables the piecewise scanning of the sample, overcoming limitations associated with the relative size of the microscope imaging window (540 &#181;m by 540 &#181;m) and the length of a typical trabecula (&#62;2000 &#181;m). Platinum electrodes at either end of the muscle chamber stimulate the trabecula at a user-defined rate. We exploit the stimulation signal as a trigger for synchronizing the data from each imaging window to reconstruct the entire sample twitching under steady-state conditions. Applying image-processing techniques to these brightfield imaging data provides tissue displacement and sarcomere length maps. Such a collection of data, when incorporated into an experiment-modeling pipeline, will provide a deeper understanding of muscle contractile homogeneity and heterogeneity in physiology and pathophysiology.

Simultaneous Brightfield, Fluorescence, and Optical Coherence Tomographic Imaging of Contracting Cardiac Trabeculae Ex Vivo

This protocol presents a collection of sarcomere, calcium, and macroscopic geometric&#160;data from an actively contracting cardiac trabecula ex vivo. These simultaneous measurements are made possible by the integration of three imaging modalities.

In cardiac muscle, intracellular Ca2+ transients activate contractile myofilaments, causing contraction, macroscopic shortening, and ...

Isolation, Culture, and Functional Characterization of Adult Mouse Cardiomyoctyes

The use of primary cardiomyocytes (CMs) in culture has provided a powerful complement to murine models of heart disease in advancing our understanding of heart disease. In particular, the ability to study ion homeostasis, ion channel function, cellular excitability and excitation-contraction coupling and their alterations in diseased conditions and by disease-causing mutations have led to significant insights into cardiac diseases. Furthermore, the lack of an adequate immortalized cell line to mimic adult CMs, and the limitations of neonatal CMs (which lack many of the structural and functional biomechanics characteristic of adult CMs) in culture have hampered our understanding of the complex interplay between signaling pathways, ion channels and contractile properties in the adult heart strengthening the importance of studying adult isolated cardiomyocytes. Here, we present methods for the isolation, culture, manipulation of gene expression by adenoviral-expressed proteins, and subsequent functional analysis of cardiomyocytes from the adult mouse. The use of these techniques will help to develop mechanistic insight into signaling pathways that regulate cellular excitability, Ca2+ dynamics and contractility and provide a much more physiologically relevant characterization of cardiovascular disease.

Here we describe the isolation of adult mouse cardiomyoctyes using a Langendorff perfusion system. The resulting cells are Ca2+-tolerant, electrically quiescent and can be cultured and transfected with adeno- or lentiviruses to manipulate gene expression. Their functionality can also be analyzed using the MMSYS system and patch clamp techniques.

The use of primary cardiomyocytes (CMs) in culture has provided a powerful complement to murine models of heart disease in advancing our understanding of ...

Biology

Contractility Measurements on Isolated Papillary Muscles for the Investigation of Cardiac Inotropy in Mice

Papillary muscle isolated from adult mouse hearts can be used to study cardiac contractility during different physiological/pathological conditions. The contractile characteristics can be evaluated independently of external influences such as vascular tonus or neurohumoral status. It depicts a scientific approach between single cell measurements with isolated cardiac myocytes and in vivo studies like echocardiography. Thus, papillary muscle preparations serve as an excellent model to study cardiac physiology/pathophysiology and can be used for investigations like the modulation by pharmacological agents or the exploration of transgenic animal models. Here, we describe a method of isolating the murine left anterior papillary muscle to investigate cardiac contractility in an organ bath setup. In contrast to a muscle strip preparation isolated from the ventricular wall, the papillary muscle can be prepared in toto without damaging the muscle tissue severely. The organ bath setup consists of several temperature-controlled, gassed and electrode-equipped organ bath chambers. The isolated papillary muscle is fixed in the organ bath chamber and electrically stimulated. The evoked twitch force is recorded using a pressure transducer and parameters such as twitch force amplitude and twitch kinetics are analyzed. Different experimental protocols can be performed to investigate the calcium- and frequency-dependent contractility as well as dose-response curves of contractile agents such as catecholamines or other pharmaceuticals. Additionally, pathologic conditions like acute ischemia can be simulated.

Murine left ventricular papillary muscle can be used to investigate cardiac contractility in vitro. This article describes in detail the isolation and experimental protocols to study cardiac contractile characteristics.

Papillary muscle isolated from adult mouse hearts can be used to study cardiac contractility during different physiological/pathological conditions. The ...

Assessment of Myofilament Ca2+ Sensitivity Underlying Cardiac Excitation-contraction Coupling

Heart failure and cardiac arrhythmias are the leading causes of mortality and morbidity worldwide. However, the mechanism of pathogenesis and myocardial malfunction in the diseased heart remains to be fully clarified. Recent compelling evidence demonstrates that changes in the myofilament Ca2+ sensitivity affect intracellular Ca2+ homeostasis and ion channel activities in cardiac myocytes, the essential mechanisms responsible for the cardiac action potential and contraction in healthy and diseased hearts. Indeed, activities of ion channels and transporters underlying cardiac action potentials (e.g., Na+, Ca2+ and K+ channels and the Na+-Ca2+ exchanger) and intracellular Ca2+ handling proteins (e.g., ryanodine receptors and Ca2+-ATPase in sarcoplasmic reticulum (SERCA2a) or phospholamban and its phosphorylation) are conventionally measured to evaluate the fundamental mechanisms of cardiac excitation-contraction (E-C) coupling. Both electrical activities in the membrane and intracellular Ca2+ changes are the trigger signals of E-C coupling, whereas myofilament is the functional unit of contraction and relaxation, and myofilament Ca2+ sensitivity is imperative in the implementation of myofibril performance. Nevertheless, few studies incorporate myofilament Ca2+ sensitivity into the functional analysis of the myocardium unless it is the focus of the study. Here, we describe a protocol that measures sarcomere shortening/re-lengthening and the intracellular Ca2+ level using Fura-2 AM (ratiometric detection) and evaluate the changes of myofilament Ca2+ sensitivity in cardiac myocytes from rat hearts. The main aim is to emphasize that myofilament Ca2+ sensitivity should be taken into consideration in E-C coupling for mechanistic analysis. Comprehensive investigation of ion channels, ion transporters, intracellular Ca2+ handling, and myofilament Ca2+ sensitivity that underlie myocyte contractility in healthy and diseased hearts will provide valuable information for designing more effective strategies of translational and therapeutic value.

Assessment of Myofilament Ca2+ Sensitivity Underlying Cardiac Excitation-contraction Coupling

This paper describes a protocol that assesses the changes of myofilament Ca2+ sensitivity during contraction in isolated cardiac myocytes from rat heart. Together with cardiac electrophysiology, systolic/diastolic cytosol Ca2+ levels and contraction/relaxation, this measurement is imperative in underpinning the mechanisms mediating cardiac excitation-contraction coupling in healthy and diseased hearts.

Heart failure and cardiac arrhythmias are the leading causes of mortality and morbidity worldwide. However, the mechanism of pathogenesis and myocardial ...

Preparation of Human Myocardial Tissue for Long-Term Cultivation

Cardiomyocyte cultivation has seen a vast number of developments, ranging from two-dimensional (2D) cell cultivation to iPSC derived organoids. In 2019, an ex vivo way to cultivate myocardial slices obtained from human heart samples was demonstrated, while approaching in vivo condition of myocardial contraction. These samples originate mostly from heart transplantations or left-ventricular assist device placements. Using a vibratome and a specially developed cultivation system, 300 &#181;m thick slices are placed between a fixed and a spring wire, allowing for stable and reproducible cultivation for several weeks. During cultivation, the slices are continuously stimulated according to individual settings. Contractions can be displayed and recorded in real-time, and pharmacological agents can be readily applied. User-defined stimulation protocols can be scheduled and performed to assess vital contraction parameters like post-pause-potentiation, stimulation threshold, force-frequency relation, and refractory period. Furthermore, the system enables a variable pre- and afterload setting for a more physiological cultivation.
Here, we present a step-by-step guide on how to generate a successful long-term cultivation of human left ventricular myocardial slices, using a commercial biomimetic cultivation solution.

We present a protocol for ex vivo cultivation of human ventricular myocardial tissue. It allows for detailed analysis of contraction force and kinetics, as well as the application of pre- and afterload to mimic the in vivo physiological environment more closely.

Cardiomyocyte cultivation has seen a vast number of developments, ranging from two-dimensional (2D) cell cultivation to iPSC derived organoids. In 2019, ...

Mechanical Control of Relaxation Using Intact Cardiac Trabeculae

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