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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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% CO2</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.&#160; 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.&#160; 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.&#160; 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 (O2)</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&#160;</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&#34; 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/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/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/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/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/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

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1.1K Views.  Wayne State University. 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. 

Video: Mechanical Control of Relaxation Using Intact Cardiac Trabeculae

Quantitative Visualization and Detection of Skin Cancer Using Dynamic Thermal Imaging

In 2010 approximately 68,720 melanomas will be diagnosed in the US alone, with around 8,650 resulting in death 1. To date, the only effective treatment for melanoma remains surgical excision, therefore, the key to extended survival is early detection 2,3. Considering the large numbers of patients diagnosed every year and the limitations in accessing specialized care quickly, the development of objective in vivo diagnostic instruments to aid the diagnosis is essential. New techniques to detect skin cancer, especially non-invasive diagnostic tools, are being explored in numerous laboratories. Along with the surgical methods, techniques such as digital photography, dermoscopy, multispectral imaging systems (MelaFind), laser-based systems (confocal scanning laser microscopy, laser doppler perfusion imaging, optical coherence tomography), ultrasound, magnetic resonance imaging, are being tested. Each technique offers unique advantages and disadvantages, many of which pose a compromise between effectiveness and accuracy versus ease of use and cost considerations. Details about these techniques and comparisons are available in the literature 4.
Infrared (IR) imaging was shown to be a useful method to diagnose the signs of certain diseases by measuring the local skin temperature. There is a large body of evidence showing that disease or deviation from normal functioning are accompanied by changes of the temperature of the body, which again affect the temperature of the skin 5,6. Accurate data about the temperature of the human body and skin can provide a wealth of information on the processes responsible for heat generation and thermoregulation, in particular the deviation from normal conditions, often caused by disease. However, IR imaging has not been widely recognized in medicine due to the premature use of the technology 7,8 several decades ago, when temperature measurement accuracy and the spatial resolution were inadequate and sophisticated image processing tools were unavailable. This situation changed dramatically in the late 1990s-2000s. Advances in IR instrumentation, implementation of digital image processing algorithms and dynamic IR imaging, which enables scientists to analyze not only the spatial, but also the temporal thermal behavior of the skin 9, allowed breakthroughs in the field.
In our research, we explore the feasibility of IR imaging, combined with theoretical and experimental studies, as a cost effective, non-invasive, in vivo optical measurement technique for tumor detection, with emphasis on the screening and early detection of melanoma 10-13. In this study, we show data obtained in a patient study in which patients that possess a pigmented lesion with a clinical indication for biopsy are selected for imaging. We compared the difference in thermal responses between healthy and malignant tissue and compared our data with biopsy results. We concluded that the increased metabolic activity of the melanoma lesion can be detected by dynamic infrared imaging.

We demonstrated that malignant pigmented lesions with increased metabolic activity generate quantifiable amounts of heat and the measurement of the transient thermal response of the skin to a cooling excitation allows quantitative identification of melanoma and other skin cancers (vs. non-proliferative nevi) at an early stage of the disease.

In 2010 approximately 68,720 melanomas will be diagnosed in the US alone, with around 8,650 resulting in death 1. To date, the only effective treatment ...

Mesenteric Artery Contraction and Relaxation Studies Using Automated Wire Myography

Proximal resistance vessels, such as the mesenteric arteries, contribute substantially to the peripheral resistance. These small vessels of between 100-400 &mu;m in diameter function primarily in directing blood flow to various organs according to the overall requirements of the body. The rat mesenteric artery has a diameter greater than 100 &mu;m. The myography technique, first described by Mulvay and Halpern1, was based on the method proposed by Bevan and Osher2. The technique provides information about small vessels under isometric conditions, where substantial shortening of the muscle preparation is prevented. Since force production and sensitivity of vessels to different agonists is dependent on the extent of stretch, according to active tension-length relation, it is essential to conduct contraction studies under isometric conditions to prevent compliance of the mounting wires. Stainless steel wires are preferred to tungsten wires because of oxidation of the latter, which affects recorded responses3.The technique allows for the comparison of agonist-induced contractions of mounted vessels to obtain evidence for normal function of vascular smooth muscle cell receptors.
We have shown in several studies that isolated mesenteric arteries that are contracted with phenylyephrine relax upon addition of cumulative concentrations of extracellular calcium (Ca2+e). The findings led us to conclude that perivascular sensory nerves, which express the G protein-coupled Ca2+-sensing receptor (CaR), mediate this vasorelaxation response. Using an automated wire myography method, we show here that mesenteric arteries from Wistar, Dahl salt-sensitive(DS) and Dahl salt-resistant (DR) rats respond differently to Ca2+e. Tissues from Wistar rats showed higher Ca2+-sensitivity compared to those from DR and DS. Reduced CaR expression in mesenteric arteries from DS rats correlates with reduced Ca2+e-induced relaxation of isolated, pre-contracted arteries. The data suggest that the CaR is required for relaxation of mesenteric arteries under increased adrenergic tone, as occurs in hypertension, and indicate an inherent defect in the CaR signaling pathway in Dahl animals, which is much more severe in DS.
The method is useful in determining vascular reactivity ex vivo in mesenteric resistance arteries and similar small blood vessels and comparisons between different agonists and/or antagonists can be easily and consistently assessed side-by-side6,7,8.

An automated myography method for force measurements in isolated mesenteric arteries is described. It employs a Mulvany-Halpern Auto Dual Wire Myograph 510A to determine responses to phenylephrine and extracellular calcium. The method allows consistent determination of isometric responses to agonists in small vessels of diameters of 60 - 300 &mu;m, independently.

Proximal resistance vessels, such as the mesenteric arteries, contribute substantially to the peripheral resistance. These small vessels of between ...

Direct Pressure Monitoring Accurately Predicts Pulmonary Vein Occlusion During Cryoballoon Ablation

Cryoballoon ablation (CBA) is an established therapy for atrial fibrillation (AF). Pulmonary vein (PV) occlusion is essential for achieving antral contact and PV isolation and is typically assessed by contrast injection. We present a novel method of direct pressure monitoring for assessment of PV occlusion.
Transcatheter pressure is monitored during balloon advancement to the PV antrum. Pressure is recorded via a single pressure transducer connected to the inner lumen of the cryoballoon. Pressure curve characteristics are used to assess occlusion in conjunction with fluoroscopic or intracardiac echocardiography (ICE) guidance. PV occlusion is confirmed when loss of typical left atrial (LA) pressure waveform is observed with recordings of PA pressure characteristics (no A wave and rapid V wave upstroke). Complete pulmonary vein occlusion as assessed with this technique has been confirmed with concurrent contrast utilization during the initial testing of the technique and has been shown to be highly accurate and readily reproducible.
We evaluated the efficacy of this novel technique in 35 patients. A total of 128 veins were assessed for occlusion with the cryoballoon utilizing the pressure monitoring technique; occlusive pressure was demonstrated in 113 veins with resultant successful pulmonary vein isolation in 111 veins (98.2%). Occlusion was confirmed with subsequent contrast injection during the initial ten procedures, after which contrast utilization was rapidly reduced or eliminated given the highly accurate identification of occlusive pressure waveform with limited initial training.
Verification of PV occlusive pressure during CBA is a novel approach to assessing effective PV occlusion and it accurately predicts electrical isolation. Utilization of this method results in significant decrease in fluoroscopy time and volume of contrast.

Effective pulmonary vein isolation utilizing a cryoballoon depends on complete pulmonary vein occlusion. The point of occlusion can be effectively predicted by direct analysis of pulmonary vein pressure waveform analysis during balloon inflation using a simple and reproducible technique.

Cryoballoon  ablation (CBA) is an established therapy for atrial fibrillation (AF).  Pulmonary vein (PV) occlusion is essential for achieving antral ...

Early Detection of Drug-Induced Renal Hemodynamic Dysfunction Using Sonographic Technology in Rats

The kidney normally functions to maintain hemodynamic homeostasis and is a major site of damage caused by drug toxicity. Drug-induced nephrotoxicity is estimated to contribute to 19- 25% of all clinical cases of acute kidney injury (AKI) in critically ill patients. AKI detection has historically relied on metrics such as serum creatinine (sCr) or blood urea nitrogen (BUN) which are demonstrably inadequate in full assessment of nephrotoxicity in the early phase of renal dysfunction. Currently, there is no robust diagnostic method to accurately detect hemodynamic alteration in the early phase of AKI while such alterations might actually precede the rise in serum biomarker levels. Such early detection can help clinicians make an accurate diagnosis and help in in decision making for therapeutic strategy. Rats were treated with Cisplatin to induce AKI. Nephrotoxicity was assessed for six days using high-frequency sonography, sCr measurement and upon histopathology of kidney. Hemodynamic evaluation using 2D and Color-Doppler images were used to serially study nephrotoxicity in rats, using the sonography. Our data showed successful drug-induced kidney injury in adult rats by histological examination. Color-Doppler based sonographic assessment of AKI indicated that resistive-index (RI) and pulsatile-index (PI) were increased in the treatment group; and peak-systolic velocity (mm/s), end-diastolic velocity (mm/s) and velocity-time integral (VTI, mm) were decreased in renal arteries in the same group. Importantly, these hemodynamic changes evaluated by sonography preceded the rise of sCr levels. Sonography-based indices such as RI or PI can thus be useful predictive markers of declining renal function in rodents. From our sonography-based observations in the kidneys of rats that underwent AKI, we showed that these noninvasive hemodynamic measurements may consider as an accurate, sensitive and robust method in detecting early stage kidney dysfunction. This study also underscores the importance of ethical issues associated with animal use in research.

Early stage hemodynamic dysfunction is critical to the development of kidney disease. Yet, detection methodologies are limited. Recent advances in sonography provide a noninvasive, accurate option for early detection of kidney injury. This study outlines a step-by-step, sonographic methodology for detecting kidney dysfunction using a drug-induced nephrotoxicity rat model.

The kidney normally functions to maintain hemodynamic homeostasis and is a major site of damage caused by drug toxicity. Drug-induced nephrotoxicity is ...

Visualization of Streptococcus pneumoniae within Cardiac Microlesions and Subsequent Cardiac Remodeling

During bacteremia Streptococcus pneumoniae can translocate across the vascular endothelium into the myocardium and form discrete bacteria-filled microscopic lesions (microlesions) that are remarkable due to the absence of infiltrating immune cells. Due to their release of cardiotoxic products, S. pneumoniae within microlesions are thought to contribute to the heart failure that is frequently observed during fulminate invasive pneumococcal disease in adults. Herein is demonstrated a protocol for experimental mouse infection that leads to reproducible cardiac microlesion formation within 30 hr. Instruction is provided on microlesion identification in hematoxylin &#38; eosin stained heart sections and the morphological distinctions between early and late microlesions are highlighted. Instruction is provided on a protocol for verification of S. pneumoniae within microlesions using antibodies against pneumococcal capsular polysaccharide and immunofluorescent microscopy. Last, a protocol for antibiotic intervention that rescues infected mice and for the detection and assessment of scar formation in the hearts of convalescent mice is provided. Together, these protocols will facilitate the investigation of the molecular mechanisms underlying pneumococcal cardiac invasion, cardiomyocyte death, cardiac remodeling as a result of exposure to S. pneumoniae, and the immune response to the pneumococci in the heart.

Visualization of Streptococcus pneumoniae within Cardiac Microlesions and Subsequent Cardiac Remodeling

Streptococcus pneumoniae forms discrete non-purulent microscopic lesions in the heart. Outlined is the protocol for a murine model of cardiac microlesion formation. Instruction is provided on microlesion visualization using microscopy, discrimination between early and late microlesions, and methods to detect cardiac remodeling in hearts of convalescent animals.

During bacteremia Streptococcus pneumoniae can translocate across the vascular endothelium into the myocardium and form discrete bacteria-filled ...

Noninvasive Determination of Vortex Formation Time Using Transesophageal Echocardiography During Cardiac Surgery

Trans-mitral blood flow produces a three-dimensional rotational body of fluid, known as a vortex ring, that enhances the efficiency of left ventricular (LV) filling compared with a continuous linear jet. Vortex ring development is most often quantified with vortex formation time (VFT), a dimensionless parameter based on fluid ejection from a rigid tube. Our group is interested in factors that affect LV filling efficiency during cardiac surgery. In this report, we describe how to use standard two-dimensional (2D) and Doppler transesophageal echocardiography (TEE) to noninvasively derive the variables needed to calculate VFT. We calculate atrial filling fraction (&#946;) from velocity-time integrals of trans-mitral early LV filling and atrial systole blood flow velocity waveforms measured in the mid-esophageal four-chamber TEE view. Stroke volume (SV) is calculated as the product of the diameter of the LV outflow track measured in the mid-esophageal long axis TEE view and the velocity-time integral of blood flow through the outflow track determined in the deep transgastric view using pulse-wave Doppler. Finally, mitral valve diameter (D) is determined as the average of major and minor axis lengths measured in orthogonal mid-esophageal bicommissural and long axis imaging planes, respectively. VFT is then calculated as 4 &#215; (1-&#946;) &#215; SV/(&#960;D3). We have used this technique to analyze VFT in several groups of patients with differing cardiac abnormalities. We discuss our application of this technique and its potential limitations and also review our results to date. Noninvasive measurement of VFT using TEE is straightforward in anesthetized patients undergoing cardiac surgery. The technique may allow cardiac anesthesiologists and surgeons to assess the impact of pathological conditions and surgical interventions on LV filling efficiency in real time.

We describe a protocol to measure vortex formation time, an index of left ventricular filling efficiency, using standard transesophageal echocardiography techniques in patients undergoing cardiac surgery. We apply this technique to analyze vortex formation time in several groups of patients with differing cardiac pathologies.

Trans-mitral blood flow produces a three-dimensional rotational body of fluid, known as a vortex ring, that enhances the efficiency of left ventricular ...

Normothermic Ex Situ Heart Perfusion in Working Mode: Assessment of Cardiac Function and Metabolism

The current standard method for organ preservation (cold storage, CS), exposes the heart to a period of cold ischemia that limits the safe preservation time and increases the risk of adverse post-transplantation outcomes. Moreover, the static nature of CS does not allow for organ evaluation or intervention during the preservation interval. Normothermic ex situ heart perfusion (ESHP) is a novel method for preservation of the donated heart that minimizes cold ischemia by providing oxygenated, nutrient-rich perfusate to the heart. ESHP has been shown to be non-inferior to CS in the preservation of standard-criteria donor hearts and has also facilitated the clinical transplantation of the hearts donated after the circulatory determination of death. Currently, the only available clinical ESHP device perfuses the heart in an unloaded, non-working state, limiting assessments of myocardial performance. Conversely, ESHP in working mode provides the opportunity for comprehensive evaluation of cardiac performance by assessment of functional and metabolic parameters under physiologic conditions. Moreover, earlier experimental studies have suggested that ESHP in working mode may result in improved functional preservation. Here, we describe the protocol for ex situ perfusion of the heart in a large mammal (porcine) model, which is reproducible for different animal models and heart sizes. The software program in this ESHP apparatus allows for real-time and automated control of the pump speed to maintain desired aortic and left atrial pressure and evaluates a variety of functional and electrophysiological parameters with minimal need for supervision/manipulation.

Normothermic ex situ heart perfusion (ESHP), preserves the heart in a beating, semi-physiologic state. When performed in a working mode, ESHP provides the opportunity to perform sophisticated assessments of donor heart function and organ viability. Here, we describe our method for myocardial performance evaluation during ESHP.

The current standard method for organ preservation (cold storage, CS), exposes the heart to a period of cold ischemia that limits the safe preservation ...

In Vitro Assessment of Cardiac Function Using Skinned Cardiomyocytes

In this article, we describe the steps required to isolate a single permeabilized (&#34;skinned&#34;) cardiomyocyte and attach it to a force-measuring apparatus and a motor to perform functional studies. These studies will allow measurement of cardiomyocyte stiffness (passive force) and its activation with different calcium (Ca2+)-containing solutions to determine, amongst others: maximum force development, myofilament Ca2+-sensitivity (pCa50), cooperativity (nHill) and the rate of force redevelopment (ktr). This method also enables determination of the effects of drugs acting directly on myofilaments and of the expression of exogenous recombinant proteins on both active and passive properties of cardiomyocytes. Clinically, skinned cardiomyocyte studies highlight the pathophysiology of many myocardial diseases and allow in vitro assessment of the impact of therapeutic interventions targeting the myofilaments. Altogether, this technique enables the clarification of cardiac pathophysiology by investigating correlations between in vitro and in vivo parameters in animal models and human tissue obtained during open heart or transplant surgery.

This protocol aims to describe step-by-step the technique of extraction and assessment of cardiac function using skinned cardiomyocytes. This methodology allows measurement and acutemodulation of myofilament function using small frozen biopsies that can be collected from different cardiac locations, from mice to men.

In this article, we describe the steps required to isolate a single permeabilized (&#34;skinned&#34;) cardiomyocyte and attach it to a force-measuring ...

An Intact Pericardium Ischemic Rodent Model

This protocol has shown that the pericardium and its contents play an essential anti-fibrotic role in the ischemic rodent model (coronary ligation to induce myocardial injury). The majority of pre-clinical myocardial infarction models require the disruption of pericardial integrity with loss of the homeostatic cellular milieu. However, recently a methodology has been developed by us to induce myocardial infarction, which minimizes pericardial damage and retains the heart&#39;s resident immune cell population. An improved cardiac functional recovery in mice with an intact pericardial space following coronary ligation has been observed. This method provides an opportunity to study inflammatory responses in the pericardial space following myocardial infarction. Further development of the labeling techniques can be combined with this model to understand the fate and function of pericardial immune cells in regulating the inflammatory mechanisms that drive remodeling in the heart, including fibrosis.

This protocol outlines the steps for inducing myocardial infarction in mice while preserving the pericardium and its contents.

This protocol has shown that the pericardium and its contents play an essential anti-fibrotic role in the ischemic rodent model (coronary ligation to ...

Analysis of Cardiac Contractile Dysfunction and Ca2+ Transients in Rodent Myocytes

Contractile dysfunction and Ca2+ transients are often analyzed at the cellular level as part of a comprehensive assessment of cardiac-induced injury and/or remodeling. One approach for assessing these functional alterations utilizes unloaded shortening and Ca2+ transient analyses in primary adult cardiac myocytes. For this approach, adult myocytes are isolated by collagenase digestion, made Ca2+ tolerant, and then adhered to laminin-coated coverslips, followed by electrical pacing in serum-free media. The general protocol utilizes adult rat cardiac myocytes but can be readily adjusted for primary myocytes from other species. Functional alterations in myocytes from injured hearts can be compared to sham myocytes and/or to in vitro therapeutic treatments. The methodology includes the essential elements needed for myocyte pacing, along with the cell chamber and platform components. The detailed protocol for this approach incorporates the steps for measuring unloaded shortening by sarcomere length detection and cellular Ca2+ transients measured with the ratiometric indicator Fura-2 AM, as well as for raw data analysis.

Analysis of Cardiac Contractile Dysfunction and Ca2+ Transients in Rodent Myocytes

A set of protocols are presented that describe the measurement of contractile function via sarcomere length detection along with calcium (Ca2+) transient measurement in isolated rat myocytes. The application of this approach for studies in animal models of heart failure is also included.

Contractile dysfunction and Ca2+ transients are often analyzed at the cellular level as part of a comprehensive assessment of cardiac-induced injury ...