This research was supported by Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Education, Science and Technology (2013068067); by the Brain Korea 21 Graduate Programme of the Korean Ministry of Education, Science and Technology, Seoul National University Hospital, the Korean Society of Hypertension (2013), SK Telecom Research Fund (no. 3420130290) and from the National Natural Science Foundation of China (NSFC&#160;31460265; NSFC 81260035).

The protocol is in accordance with the Guide for the Care and Use of Laboratory Animals published by the UN National Institutes of Health (NIH Publication No. 85-23, revised 1996). It was approved by the Institutional Animal Care and Use Committee (IACUC) of Seoul National University (IACUC approval no.: SNU-101213-1).
1. Buffer Preparation (Table Materials and Equipment)
<ol>
	<li>Prepare 300 ml of fresh isolation solution on the day of the experiment (in mM: NaCl, 135; KCl, 5.4; MgCl2, 3.5; glucose, 5; HEPES, 5; Na2HPO4, 0.4; taurine, 20; pH 7.4 with NaOH).</li>
	<li>Prepare 100 ml of fresh storage solution on the day of the experiment (in mM: NaCl 120; KCl 5.4; MgSO4 5; CaCl2 0.2; sodium-pyruvate 5; glucose 5.5; taurine 20; HEPES 10; mannitol 29; pH 7.4 with NaOH).</li>
	<li>Prepare 1L of perfusion solution (in mM: NaCl, 141.4; KCl, 4; NaH2PO4, 0.33; MgCl2, 1; HEPES, 10; glucose, 5.5; CaCl2, 1.8; mannitol, 14.5; pH 7.4 with NaOH).</li>
	<li>Prepare 30 ml of collagenase solution 1 by adding collagenase (1 mg/ml), protease (0.1 mg/ml), bovine serum albumin (BSA, 1.67mg/ml), and Ca2+ (0.05 mM) to 25 ml of isolation solution.</li>
	<li>Prepare 20 ml of collagenase solution 2 by adding collagenase (1 mg/ml), BSA (1.67 mg/ml), and Ca2+ (0.05 mM) to 16.7 ml of isolation solution.</li>
	<li>Prepare 40 ml of BSA solution by adding 0.4 g of BSA to 40 ml of isolation solution. Separate 10 ml and 30 ml into two beakers. Add Ca2+ to 30 ml of BSA solution so that the final Ca2+ concentration in the BSA solution is 1 mM.</li>
</ol>
2. Preparation for the Isolation of Left Ventricular (LV) Myocytes
<ol>
	<li>Transfer 8-12 week-old, male Sprague-Dawley (SD) rats in clean transport cages from the animal facility to the preparation and isolation room.</li>
	<li>Heat two water baths to 37 o C. 
	Note: Use one water bath for the water - jacketed reservoir and the perfusion tubes of the Langendorff perfusion system. Use the other water bath to agitate myocardial tissue trunks in order to separate single myocytes.</li>
	<li>Add 5 ml and 3.3 ml of BSA solution (without Ca2+) to 25 ml of collagenase solution 1 and 16.7 ml collagenase solution 2 to make up the volume to 30 ml and 20 ml, respectively.</li>
	<li>Add isolation solution (100 ml) and collagenase solution 1 (30 ml) to column 1 (Col. 1) and column 2 (Col. 2) in the Langendorff perfusion system, respectively (Figure 1A).</li>
	<li>Oxygenate the isolation solution and collagenase solution 1 in Col. 1 and Col.2 via the oxygen connection tube in the Langendorff perfusion system (Figure 1A). Similarly, oxygenate collagenase solution 2 and the BSA solution in the shaking water bath with 100% O2&#160;via the oxygen connection tube.</li>
</ol>
3. Isolation of LV Myocytes
<ol>
	<li>Anesthetize a SD rat via intraperitoneal injection of pentobarbital sodium (30 mg/kg) and confirm the anesthetic status by toe pinching and lack of withdrawal reflection.</li>
	<li>Move the rat to a dissection tray. In the supine position, secure the four legs to the sides of the body with tape.</li>
	<li>Apply thoracic mid-axial incisions with surgical scissors to open the chest, ensuring not to damage the heart. Use another pair of clean scissors to dissect the heart from the connecting vessels (e.g., superior and inferior vena cava, pulmonary vessels, and aorta) and pericardial membrane13.</li>
	<li>Leave a portion of the aorta of a sufficient length (5 - 8 mm), clamp the aorta with fine forceps, and rapidly mount the cannula of the Langendorff perfusion system within 1 min (Figure 1A). Tie suture thread 4/0 tightly over the aorta.</li>
	<li>Turn on the valve on Col. 1 and perfuse the isolated heart with pre-warmed and oxygenated isolation solution for 10 min (perfusion rate: 12-14 ml/min). Turn off the valve on Col. 1, turn on the valve on Col. 2 and perfuse the heart with collagenase solution 1 for 8 - 10 min.</li>
	<li>Dismount the digested heart by cutting the aorta and transfer it by holding the aorta with forceps into the flask containing fresh isolation solution (Figure 1Bi). Use fine scissors and forceps to cut most of the LV free wall (including the septum) into smaller pieces (~22 mm, Figure 1Bii).</li>
	<li>Transfer the pieces into a flask containing pre-warmed and oxygenated fresh collagenase solution 2 (Figure 1Biii). Shake for 10 min. Keep delivering oxygen to the myocyte-containing collagenase solution 2.</li>
	<li>Move the myocyte - suspension to a 10 ml centrifuge tube using droppers with a suction bulb and add Ca2+ - containing BSA solution (1:1 in volume). Centrifuge at 30 g for 2 min and discard the supernatant. Re-suspend the myocyte pellet in 5 ml of BSA solution. Centrifuge and discard the supernatant.</li>
	<li>Disperse the myocyte pellets and keep myocytes in 10 ml of pre-oxygenated storage solution at RT (Figure 1Biv-vi).</li>
	<li>Repeat the procedures (steps 3.7 - 3.9) with the remaining LV tissue in the flask. 
	Note: Keep repeating the procedures (steps 3.7 - 3.9) again until most of the LV tissue disappears to obtain a good yield of myocytes.</li>
	<li>Keep the myocytes in storage solution for 6-8 hr at RT until the end of experiments.</li>
</ol>
4. Simultaneous Measurements of Intracellular Ca2+ Transients and Myocyte Contraction
<ol>
	<li>Load LV myocytes with the acetoxymethyl ester Fura-2 AM (2 &#181;M). 
	Note: Perform all loading procedures and experiments with loaded myocytes in a dark tube (Figure 1Bvii).
	<ol>
		<li>Centrifuge the myocyte suspension (1 ml) at 2,000 x g for 10 sec. Discard the supernatant and re-suspend the myocyte pellet in 1 ml of Tyrode solution with a low Ca2+ concentration (250 &#181;M, Table Materials and Equipment).</li>
		<li>Add Fura-2AM and poloxamer 407 (2 &#181;l), gently disperse the myocyte suspension, and keep the mixture at RT (20 - 24o C) for 15 min&#160;(Figure 1Bvii).</li>
		<li>Centrifuge the mixture for 5 sec, discard the supernatant and disperse the myocyte pellet in 1 ml of perfusion solution containing 500 &#181;M Ca2+. After 10 min, centrifuge the mixture for 5 sec and discard the supernatant.</li>
		<li>Add fresh perfusion solution (500 &#181;M Ca2+, 1 ml) and keep the Fura-2AM - loaded myocyte pellet in this solution for recordings.</li>
	</ol>
	</li>
	<li>Measurement of LV myocyte contraction and intracellular Ca2+
	<ol>
		<li>Before recording, fill the perfusion tube that runs through a water jacket with the Tyrode solution, which is pre-warmed to 36 o C.</li>
		<li>Place a few drops of the Fura 2 AM - loaded LV myocyte suspension on the chamber of an inverted fluorescence microscope for 5 - 8 min. Slowly perfuse the Tyrode solution (2.2 ml/min).</li>
		<li>Press the &#34;start&#34; button on the front panel of the digital stimulator to start field stimulation (2 Hz). 
		Note: The output voltage (10 V, 5 msec duration) is applied to the myocytes in the chamber through platinum wires positioned on either side of the chamber.
		<ol>
			<li>Select myocytes that contract stably (not those displaying hyper- or hypo-contraction) for recording.</li>
		</ol>
		</li>
		<li>Adjust the myocyte of choice in the horizontal position of the video-based sarcomere detection system and adjust the focus of the microscope to obtain optimal images of sarcomeres. Position the purple rectangular box on the area in which clear sarcomeres are clearly observed until the average of sarcomere lengths (red peak) displays a singular sharp peak (Figure 2A, lower image).</li>
		<li>Record the changes of sarcomere length in response to field stimulation (Figure 2A and B). 
		Note: Use the loaded myocytes within 1 - 2 hr.</li>
		<li>Adjust the aperture of the camera so that the field in the video-based sarcomere detection system is the size of the myocyte (Figure 2A). Stimulate the myocyte with excitation at 360 nm/380 nm and emission at 510 nm (acquisition frequency 1,000 Hz). Record sarcomere shortening and the Fura2 AM ratio with field stimulation (Figure 2B).</li>
	</ol>
	</li>
</ol>
5. Assessment of Myofilament Ca2+ Sensitivity
<ol>
	<li>Average the sarcomere lengths and Ca2+ transients at steady-state (10 - 20 traces) and plot the phase-plane loop of the Fura-2 ratio vs. sarcomere length of the same myocyte (both with measured values and delta changes; Figure 2C).</li>
	<li>In each plot, define Fura -2 ratio at 50% relaxation (EC50, Figure 2C). Compare both the loop and EC50 of each intervention.</li>
</ol>

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The authors declare that they have no competing financial interests.

Here, we describe the protocols to assess changes of myofilament Ca2+ sensitivity in single isolated cardiac myocyte and emphasize the importance of measuring this parameter alongside electrophysiological properties, intracellular Ca2+ transients, and myofilament dynamics. This is because the recordings of one or two of the parameters may not explicate the mechanisms underlying cardiac contraction and relaxation. Unlike conventional methods that measure myocyte contraction and the intracellular Ca<s...

Cardiac excitation-contraction (E-C) coupling is the fundamental scheme for analyzing mechanical properties of the myocardium, i.e., the contractile function of the heart1,2. E-C coupling is initiated by membrane depolarization secondary to the activities of sarcolemmal ion channels (e.g., the voltage-gated Na+ channel, which can be measured via patch-clamp techniques). Subsequent activation of voltage-gated L-type Ca2+ channels (LTCCs) and Ca2+ influx via LTCCs trigger the bulk of Ca2+ release through ryanodine receptors (RyRs), increasing the cytosolic Ca2+ concentration from the nanomolar (nM) to micromolar (&#181;M) level. Such an increase in cytosolic Ca2+ promotes Ca2+ binding to troponin C (TnC) in thin filaments and elicits conformational changes of the filament complex to facilitate the actin-myosin interaction and attains myocardial contraction3. Conversely, the cytosolic Ca2+ is re-uptaken&#160;back into the sarcoplasmic reticulum (SR) through the Ca2+-ATPase in SR (SERCA2a) or is extruded out of the myocyte via the Na+/Ca2+ exchanger and the plasmalemmal Ca2+ ATPase1,2. Consequently, the decline in cytosolic Ca2+ instigates conformational changes of thin filaments back to the original state, resulting in the dissociation of actin-myosin and myocyte relaxation1-3. In this scheme, the activity of SERCA2a is generally considered to determine the speed of myocardial relaxation because it accounts for 70 - 90% of cytosolic Ca2+ removal in most mammalian heart cells1. As such, abnormal Ca2+ handling by LTCC, RyR and SERCA2a, etc. has been considered the primary mechanisms for impaired contractility and relaxation in the diseased heart1-4.
In reality, free cytosolic Ca2+ that functions as the messenger in E-C coupling accounts for around 1% of total intracellular Ca2+ and the majority of Ca2+ is bound to intracellular Ca2+ buffers5,6. This is due to the fact that various Ca2+ buffers are abundant in cardiac myocytes, e.g., membrane phospholipids, ATP, phosphocreatine, calmodulin, parvalbumin, myofibril TnC, myosin, SERCA2a, and calsequestrin in the SR.5,6,7. Among them, SERCA2a and TnC are the predominant Ca2+ buffers5,6,7. Furthermore, Ca2+ binding to its buffers is a dynamic process during twitch (e.g., 30-50% of Ca2+ binds to TnC and dissociate from it during Ca2+ transients7) and the change in Ca2+ binding cause additional &#34;release&#34; of free Ca2+ to the cytosol, results in the alterations of the intracellular Ca2+ concentration. Consequently, perturbation of the intracellular Ca2+ level induces abnormal myofilament movements, which are the precursors of contractile dysfunction and arrhythmias8,9. Many factors (both physiological and pathological) can be the sources of post-transcriptional modifications of myofilament proteins, which influence myofilament Ca2+ buffering and myofilament Ca2+ sensitivity8-10. Recently, it was reported that mutations in myofilament proteins increase the Ca2+ binding affinity and intracellular Ca2+ handling, triggering pause-dependent potentiation of Ca2+ transients, abnormal Ca2+ release, and arrhythmias8. In line with this concept, we have also shown that myofilament Ca2+ desensitization in hypertensive rat hearts secondary to the up-regulation of neuronal nitric oxide synthase is associated with elevated diastolic and systolic Ca2+ levels11, which in turn, increases the vulnerability of the LTCC to Ca2+-dependent inactivation12. Hence, myofilament Ca2+ sensitivity is an &#34;active&#34; regulator of intracellular Ca2+ homeostasis and myocyte contractile function. It has become necessary to analyze interactions between myofilament and Ca2+ handling proteins for thorough investigation of myocyte E-C coupling and cardiac function.
Here, we describe a protocol that assesses the changes of myofilament Ca2+ sensitivity in isolated cardiac myocytes. Comprehensive analysis of&#160;intracellular Ca2+ profile, myofilament Ca2+ sensitivity and contraction will unearth novel mechanisms underlying myocardial mechanics.

<table><tbody><tr><td>Sprague Dawley rat</td><td>Koatech</td><td></td><td>8-12 weeks</td></tr><tr><td>Pentobarbital Sodium</td><td>Hanlim Pharmaceutical (Korea)</td><td>AHN901</td><td>Insurance code:645301220</td></tr><tr><td>NaCl</td><td>Sigma</td><td>S9625</td><td></td></tr><tr><td>KCl</td><td>Sigma</td><td>P4504</td><td></td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>S8282</td><td></td></tr><tr><td>HEPES</td><td>Sigma</td><td>H3375</td><td></td></tr><tr><td>Glucose</td><td>Sigma</td><td>G8270</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Biosesang</td><td>C2002</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Biosesang</td><td>M2001</td><td></td></tr><tr><td>Mannitol</td><td>Sigma</td><td>M4125</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>M5921</td><td></td></tr><tr><td>Sodium Pyruvate</td><td>Sigma</td><td>P2256</td><td></td></tr><tr><td>Taurine</td><td>Merck</td><td>8.08616.1000</td><td></td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&amp;nbsp;&lt;/sub&gt;</td><td>Sigma</td><td>71649</td><td></td></tr><tr><td>Bovine Fetal Albumin</td><td>Sigma</td><td>A7906</td><td></td></tr><tr><td>Collagenase Type 2</td><td>Worthington</td><td>LS004177</td><td></td></tr><tr><td>Protease</td><td>Sigma</td><td>P6911</td><td></td></tr><tr><td>Fura-2 (AM)</td><td>Molecular Probes</td><td>F1221</td><td></td></tr><tr><td>Pluronic F127 20% solution in DMSO</td><td>Invitrogen</td><td>P3000MP</td><td></td></tr><tr><td>Shaking Water Bath</td><td>Chang Shin Scientific</td><td>Model: C-108</td><td></td></tr><tr><td>IonWizard Softwae Suite</td><td>IonOptix Ltd</td><td>Experimental Builder</td><td>Acquisition and Analysis of EC Coupling Data in Myocytes</td></tr><tr><td>Myocyte Calcium &amp;amp; Contractility Recording System</td><td>IonOptix Ltd</td><td></td><td></td></tr><tr><td>Circulating Water Bath</td><td>BS-Tech</td><td>BW2-8</td><td></td></tr><tr><td>Myocyte Fluorescence Microscope</td><td>Nikon</td><td>DIATPHOTO 200</td><td></td></tr><tr><td>MyoCam-S Power</td><td>IonOptix</td><td></td><td></td></tr><tr><td>Fluorescence &amp;amp; Video Detection</td><td>IonOptix</td><td>MyoCam-S</td><td></td></tr><tr><td>CFA300</td><td></td><td></td><td></td></tr><tr><td>PMT400</td><td></td><td></td><td></td></tr><tr><td>Fluorescence &amp;amp; System Interface</td><td>IonOptix</td><td>FSI700</td><td></td></tr><tr><td>Excitation Light Source</td><td>IonOptix</td><td>mSTEP</td><td></td></tr><tr><td>High intensity ARC Lamp Power supply</td><td>Cairn Reseach</td><td></td><td></td></tr><tr><td>Filter wheel controller</td><td>IonOptix</td><td>GB/MUS200</td><td></td></tr><tr><td>Digital Stimulator</td><td>Medical Systems Corportion</td><td>S-98 Mutimode</td><td></td></tr><tr><td>&lt;strong&gt;Compositions of Experimental Solutions&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>Isolation Solution (pH: 7.4, NaOH)</td><td></td><td></td><td></td></tr><tr><td>NaCl</td><td>Sigma</td><td>S9625</td><td>Concentration (mmol) 135</td></tr><tr><td>KCl</td><td>Sigma</td><td>P4504</td><td>Concentration (mmol) 5.4</td></tr><tr><td>HEPES</td><td>Sigma</td><td>H3375</td><td>Concentration (mmol) 5</td></tr><tr><td>Glucose</td><td>Sigma</td><td>G8270</td><td>Concentration (mmol) 5</td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Biosesang</td><td>M2001</td><td>Concentration (mmol) 3.5</td></tr><tr><td>Taurine</td><td>Sigma</td><td>CB2742654</td><td>Concentration (mmol) 20</td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&amp;nbsp;&lt;/sub&gt;</td><td>Sigma</td><td>71649</td><td>Concentration (mmol) 0.4</td></tr><tr><td>Storage Solution (pH: 7.4, NaOH)</td><td></td><td></td><td></td></tr><tr><td>NaCl</td><td>Sigma</td><td>S9625</td><td>Concentration (mmol) 120</td></tr><tr><td>KCl</td><td>Sigma</td><td>P4504</td><td>Concentration (mmol) 5.4</td></tr><tr><td>HEPES</td><td>Sigma</td><td>H3375</td><td>Concentration (mmol) 10</td></tr><tr><td>Glucose</td><td>Sigma</td><td>G8270</td><td>Concentration (mmol) 5.5</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Biosesang</td><td>C2002</td><td>Concentration (mmol) 0.2</td></tr><tr><td>Mannitol</td><td>Sigma</td><td>M4125</td><td>Concentration (mmol) 29</td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>M5921</td><td>Concentration (mmol) 5</td></tr><tr><td>Sodium Pyruvate</td><td>Sigma</td><td>P2256</td><td>Concentration (mmol) 5</td></tr><tr><td>Taurine</td><td>Sigma</td><td>CB2742654</td><td>Concentration (mmol) 20</td></tr><tr><td>Perfusion Solution (Tyrode solution, pH: 7.4, NaOH)</td><td></td><td></td><td></td></tr><tr><td>NaCl</td><td>Sigma</td><td>S9625</td><td>Concentration (mmol) 141.4</td></tr><tr><td>KCl</td><td>Sigma</td><td>P4504</td><td>Concentration (mmol) 4</td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>S8282</td><td>Concentration (mmol) 0.33</td></tr><tr><td>HEPES</td><td>Sigma</td><td>H3375</td><td>Concentration (mmol) 10</td></tr><tr><td>Glucose</td><td>Sigma</td><td>G8270</td><td>Concentration (mmol) 5.5</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Biosesang</td><td>C2002</td><td>Concentration (mmol) 1.8&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; For Fura 2AM loading, CaCl2 concentrations are 0.25 mM and 0.5 mM</td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Biosesang</td><td>M2001</td><td>Concentration (mmol) 1</td></tr><tr><td>Mannitol</td><td>Sigma</td><td>M4125</td><td>Concentration (mmol) 14.5</td></tr><tr><td>Collangenase Solution 1</td><td></td><td></td><td></td></tr><tr><td>Isolation Solution (30mL)</td><td></td><td></td><td></td></tr><tr><td>Bovine Fetal Albumin (BSA solution 5 ml)</td><td></td><td></td><td>Concentration (mmol) 1.67 mg/mL</td></tr><tr><td>Collagenase Type 2</td><td>Worthington</td><td>LS004177</td><td>Concentration (mmol) 1 mg/mL</td></tr><tr><td>Protease</td><td>Sigma</td><td>P6911</td><td>Concentration (mmol) 0.1 mg/mL</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Biosesang</td><td>C2002</td><td>Concentration (mmol) 0.05 mM</td></tr><tr><td>Collangenase Solution 2</td><td></td><td></td><td></td></tr><tr><td>Isolation Solution (20mL)</td><td></td><td></td><td></td></tr><tr><td>Bovine Fetal Albumin (BSA solution 3.3 mL)</td><td></td><td></td><td>Concentration (mmol) 1.67 mg/mL</td></tr><tr><td>Collagenase Type 2</td><td>Worthington</td><td>LS004177</td><td>Concentration (mmol) 1 mg/mL</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Biosesang</td><td>C2002</td><td>Concentration (mmol) 0.05 mM</td></tr><tr><td>BSA solution</td><td></td><td></td><td></td></tr><tr><td>Isolation Solution (40mL)</td><td></td><td></td><td></td></tr><tr><td>Bovine Fetal Albumin</td><td>Sigma</td><td>A7906</td><td>Concentration (mmol) 400 mg</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Biosesang</td><td>C2002</td><td>Concentration (mmol) 1mM</td></tr></tbody></table>

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.

LV myocytes are isolated from normal and hypertensive rat hearts. Rod-shaped myocytes with clear striations (representing sarcomeres) and stable contractions in response to field stimulation are considered to be the optimal myocytes and are selected for recordings (Figure 2A). In the example shown in Figure 2A, a Fura 2 AM -loaded LV myocyte is positioned horizontally and the aperture of the camera is adjusted so that the myocyte occupies most of the reco...

Watch this Scientific Journal Video about Assessment of Myofilament Ca2+ Sensitivity Underlying Cardiac Excitation-contraction Coupling at JoVE.com

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.

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 ...

assessment-myofilament-ca2-sensitivity-underlying-cardiac-excitation

Research

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Biology

9.4K Views.  Seoul National University College of Medicine. 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. 

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

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 ...

Bioengineering

Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles

Described here is a method to measure contractility of isolated skeletal muscles. Parameters such as muscle force, muscle power, contractile kinetics, fatigability, and recovery after fatigue can be obtained to assess specific aspects of the excitation-contraction coupling (ECC) process such as excitability, contractile machinery and Ca2+ handling ability. This method removes the nerve and blood supply and focuses on the isolated skeletal muscle itself. We routinely use this method to identify genetic components that alter the contractile property of skeletal muscle though modulating Ca2+ signaling pathways. Here, we describe a newly identified skeletal muscle phenotype, i.e., mechanic alternans, as an example of the various and rich information that can be obtained using the in vitro muscle contractility assay. Combination of this assay with single cell assays, genetic approaches and biochemistry assays can provide important insights into the mechanisms of ECC in skeletal muscle.

Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles

We describe a method to directly measure muscle force, muscle power, contractile kinetics and fatigability of isolated skeletal muscles in an in vitro system using field stimulation. Valuable information on Ca2+ handling properties and contractile machinery of the muscle can be obtained using different stimulating protocols.

 
Described here is a method to measure contractility of isolated skeletal muscles. Parameters such as muscle force, muscle power, contractile kinetics, ...

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 ...

Medicine

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 ...

Optical Mapping of Intra-Sarcoplasmic Reticulum Ca2+ and Transmembrane Potential in the Langendorff-perfused Rabbit Heart

Sarcoplasmic reticulum (SR) Ca2+ handling plays a key role in normal excitation-contraction coupling and aberrant SR Ca2+ handling is known to play a significant role in certain types of arrhythmia. Because arrhythmias are spatially distinct, emergent phenomena, they must be investigated at the tissue level. However, methods for directly probing SR Ca2+ in the intact heart remain limited. This article describes the protocol for dual optical mapping of transmembrane potential (Vm) and free intra-SR [Ca2+] ([Ca2+]SR) in the Langendorff-perfused rabbit heart. This approach takes advantage of the low-affinity Ca2+ indicator Fluo-5N, which has minimal fluorescence in the cytosol where intracellular [Ca2+] ([Ca2+]i) is relatively low but exhibits significant fluorescence in the SR lumen where [Ca2+]SR is in the millimolar range. In addition to revealing SR Ca2+ characteristics spatially across the epicardial surface of the heart, this approach has the distinct advantage of simultaneous monitoring of Vm, allowing for investigations into the bidirectional relationship between Vm and SR Ca2+ and the role of SR Ca2+ in arrhythmogenic phenomena.

Optical Mapping of Intra-Sarcoplasmic Reticulum Ca2+ and Transmembrane Potential in the Langendorff-perfused Rabbit Heart

This article describes the detailed protocol and equipment necessary for dual optical mapping of transmembrane potential (Vm) and free intra-sarcoplasmic reticulum (SR) Ca2+ in the Langendorff-perfused rabbit heart. This method allows for direct observation and quantification of Vm and SR Ca2+ dynamics in the intact heart.

Sarcoplasmic reticulum (SR) Ca2+ handling plays a key role in normal excitation-contraction coupling and aberrant SR Ca2+ handling is known to play a ...

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 ...

Isolating Myofibrils from Skeletal Muscle Biopsies and Determining Contractile Function with a Nano-Newton Resolution Force Transducer

Striated muscle cells are indispensable for the activity of humans and animals. Single muscle fibers are comprised of myofibrils, which consist of serially linked sarcomeres, the smallest contractile units in muscle. Sarcomeric dysfunction contributes to muscle weakness in patients with mutations in genes encoding for sarcomeric proteins. The study of myofibril mechanics allows for the assessment of actin-myosin interactions without potential confounding effects of damaged, adjacent myofibrils when measuring the contractility of single muscle fibers. Ultrastructural damage and misalignment of myofibrils might contribute to impaired contractility. If structural damage is present in the myofibrils, they likely break during the isolation procedure or during the experiment. Furthermore, studies in myofibrils provide the assessment of actin-myosin interactions in the presence of the geometrical constraints of the sarcomeres. For instance, measurements in myofibrils can elucidate whether myofibrillar dysfunction is the primary effect of a mutation in a sarcomeric protein. In addition, perfusion with calcium solutions or compounds is almost instant due to the small diameter of the myofibril. This makes myofibrils eminently suitable to measure the rates of activation and relaxation during force production. The protocol described in this paper employs an optical force probe based on the principle of a Fabry-P&#233;rot interferometer capable of measuring forces in the nano-Newton range, coupled to a piezo length motor and a fast-step perfusion system. This setup enables the study of myofibril mechanics with high resolution force measurements.

Presented here is a protocol to assess the contractile properties of striated muscle myofibrils with nano-Newton resolution. The protocol employs a setup with an interferometry-based, optical force probe. This setup generates data with a high signal-to-noise ratio and enables the assessment of the contractile kinetics of myofibrils.

Striated muscle cells are indispensable for the activity of humans and animals. Single muscle fibers are comprised of myofibrils, which consist of ...

Functional Site-Directed Fluorometry in Native Skeletal Muscle Cells

Functional Site-Directed Fluorometry in Native Cells to Study Skeletal Muscle Excitability

Functional site-directed fluorometry has been the technique of choice to investigate the structure-function relationship of numerous membrane proteins, including voltage-gated ion channels. This approach has been used primarily in heterologous expression systems to simultaneously measure membrane currents, the electrical manifestation of the channels&#39; activity, and fluorescence measurements, reporting local domain rearrangements. Functional site-directed fluorometry combines electrophysiology, molecular biology, chemistry, and fluorescence into a single wide-ranging technique that permits the study of real-time structural rearrangements and function through fluorescence and electrophysiology, respectively. Typically, this approach requires an engineered voltage-gated membrane channel that contains a cysteine that can be tested by a thiol-reactive fluorescent dye. Until recently, the thiol-reactive chemistry used for the site-directed fluorescent labeling of proteins was carried out exclusively in Xenopus oocytes and cell lines, restricting the scope of the approach to primary non-excitable cells. This report describes the applicability of functional site-directed fluorometry in adult skeletal muscle cells to study the early steps of excitation-contraction coupling, the process by which muscle fiber electrical depolarization is linked to the activation of muscle contraction. The present protocol describes the methodologies to design and transfect cysteine-engineered voltage-gated Ca2+ channels (CaV1.1) into muscle fibers of the flexor digitorum brevis of adult mice using in vivo electroporation and the subsequent steps required for functional site-directed fluorometry measurements. This approach can be adapted to study other ion channels and proteins. The use of functional site-directed fluorometry of mammalian muscle is particularly relevant to studying basic mechanisms of excitability.

Author Spotlight: Functional Site-Directed Fluorometry in Native Cells to Study Skeletal Muscle Excitability  

Functional site-directed fluorometry is a method to study protein domain motions in real time. Modification of this technique for its application in native cells now allows the detection and tracking of single voltage-sensor motions from voltage-gated Ca2+ channels in murine isolated skeletal muscle fibers.

Functional site-directed fluorometry has been the technique of choice to investigate the structure-function relationship of numerous membrane proteins, ...

Excitation-Contraction Coupling in Skeletal Muscles

Excitation-contraction coupling is a series of events that occur between generating an action potential and initiating a muscle contraction. It occurs at the triad, a structure found in skeletal muscle fibers that comprise a T-tubule and terminal cisternae of the sarcoplasmic reticulum on each side. These triads are visible in longitudinally sectioned muscle fibers. They are typically located at the A-I junction &#8212; the junction between the A and I bands of the sarcomere.
When an action potential reaches a triad, it triggers calcium release from the sarcoplasmic reticulum. This temporary change in calcium permeability lasts for about 0.03 seconds, significantly increasing calcium concentration in and around the sarcomere. Troponin acts as a lock, preventing interaction between thick and thin filaments within a sarcomere, the smallest functional unit of muscle cells. Upon calcium binding, troponin undergoes a conformational change, allowing tropomyosin to move away from the active sites on actin filaments. This change marks the beginning of the contraction cycle.
Once the active sites are exposed, the myosin heads attach to them, creating cross-bridges between thick and thin filaments. The connection between the head and tail of the myosin molecule acts as a hinge, allowing the heads to pivot. This pivoting motion, fueled by the energy released from ATP hydrolysis, is called the power stroke and is a crucial step in muscle contraction. The myosin heads slide the thin filaments from both ends of a sarcomere toward the M line. This inward movement causes the filaments to move towards the center of the sarcomere. The filaments may often overlap, causing the I band and H zone to narrow and disappear when the muscle fully contracts. However, the A band and lengths of the thick and thin filaments do not change. Since the thin filaments at each side of the sarcomere attach to Z discs, their inward movement results in the shortening of the sarcomere and, as a result, the entire muscle fiber.

Excitation-contraction Coupling in Skeletal Muscles

Excitation-contraction coupling is a series of events that occur between generating an action potential and initiating a muscle contraction. It occurs at ...

Education

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Anatomy and Physiology

Unit 1

Muscle Tissue

KEY TERMS AND CONCEPTS

Relaxation of Skeletal Muscles

The period of muscle contraction primarily influences the duration of stimulation at the neuromuscular junction (NMJ), the presence of free calcium ions in the sarcoplasm, and the availability of energy or ATP to support contractions.
When an action potential reaches the axon terminal, it depolarizes the membrane and opens voltage-gated sodium channels. Sodium ions enter the cell, further depolarizing the presynaptic membrane. This depolarization causes voltage-gated calcium channels to open. Calcium ions entering the cell initiate a signaling cascade that causes small membrane-bound vesicles, called synaptic vesicles, containing neurotransmitter molecules to fuse with the presynaptic membrane. The fusion of a vesicle with the presynaptic membrane causes neurotransmitters to be released into the synaptic cleft, the extracellular space between the presynaptic and postsynaptic membranes.
A single action potential reaching the presynaptic end of the neuromuscular junction triggers the release of acetylcholine in the synaptic cleft. Once the receptors present on the postsynaptic membrane recognize acetylcholine, the neurotransmitter quickly dissociates from the receptor and is degraded by acetylcholinesterase present in the synaptic cleft. The swift removal and inactivation of acetylcholine from the receptor prevent overstimulation of the postsynaptic membrane from excessive firing of action potentials. As a result, the muscle membrane repolarizes, and the calcium concentration in the muscle fiber returns to a normal resting level. There are two mechanisms involved in restoring normal intracellular calcium levels: the active transport of calcium into the sarcoplasmic reticulum (SR) and of calcium across the sarcolemma into the extracellular fluid. Of the two mechanisms, the transport into the SR is more important. The SR promptly resumes normal permeability and actively reabsorbs calcium ions from the surrounding cytosol. When calcium concentration in the cytosol decreases, the contraction process reverses. Troponin releases calcium ions and reverts to its original conformation. As a result, the tropomyosin moves back, covering the myosin-binding sites on actin. Therefore, a single action potential only has a brief impact on the muscle fiber.
When there is a rapid succession of action potentials at the presynaptic end of the neuromuscular junction, there is a continuous release of acetylcholine. This, in turn, triggers a series of action potentials in the sarcolemma at the post-synaptic end and results in continuous high levels of calcium in the cytosol. Under these circumstances, the contraction cycle repeats itself without muscle fibers undergoing relaxation.

The period of muscle contraction primarily influences the duration of stimulation at the neuromuscular junction (NMJ), the presence of free calcium ions ...

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

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