This work is supported by National Institutes of Health (NIH) grant R01 HL144777 (MVW).

Studies performed on rodents followed the Public Health Service Policy on Humane Care and Use of Laboratory Animals and were approved by the University of Michigan Institutional Animal Care and Use Committee. For this study, myocytes were isolated from 3-34-month-old Sprague-Dawley and F344BN rats weighing &#8805; 200 g5. Both male and female rates were used.
1. Myocyte pacing for contractile function studies
<ol>
	<li>Make fresh M199-based media for each set of isolated myocytes (Table of Materials). 1 day before pacing, plate 2 x 104 Ca2+-tolerant, rod-shaped myocytes on 22 mm2 laminin-coated glass coverslips (CSs), as described earlier8.</li>
	<li>To prepare the chambers, soak each chamber in 10% bleach for 1 h. Then, flush the chambers with running tap water for 30-45 min, followed by distilled water replaced every 5 min for 30 min, followed by deionized water replaced every 5 min for 30 min, and a final rinse in deionized water.</li>
	<li>Dry the chambers on a clean surface for 20-30 min (Supplemental Figure 1A-C). Next, UV treat the chambers in a biosafety cabinet for 5 min in each direction (total = 20 min). This step is not needed for pre-sterilized chambers. 
	CAUTION: Leave the area to minimize UV exposure.</li>
	<li>Remove the cover from the chamber, add 3 mL of M199-based media (see Table of Materials) to each well (Supplemental Figure 1B), replace the cover, and preheat the chamber in a 37 &#176;C incubator with 5% CO2 and 20% O2.</li>
	<li>Sterilize forceps in a hot bead sterilizer at 250 &#176;C for 20-25 s. Transfer the preheated stimulation chamber from the incubator to a biosafety cabinet.</li>
	<li>Transfer each coverslip (CS) containing cardiac myocytes to individual wells in the stimulation chamber using sterile forceps. Transfer CSs four at a time, followed by heating for 10 min prior to the transfer of the remaining four CSs to maintain the media temperature.</li>
	<li>Attach banana jacks to the stimulation chamber on one end (Supplemental Figure 1C) and to the stimulator on the other end (Supplemental Figure 1D).</li>
	<li>Set the stimulator frequency to 0.2 Hz and set the voltage (~12 V) to achieve contraction in ~10%-20% of all the rod-shaped cardiac myocytes within a well. To determine the number of contracting myocytes, place the chamber under an inverted microscope with 4x to 10x magnification.</li>
	<li>Place the stimulation chamber in an incubator at 37 &#176;C in 5% CO2 (Supplemental Figure 1E). Change the media every 12 h using media preheated in a 37 &#176;C, 5% CO2 incubator for 20-25 min. Continue electrical stimulation for up to 3 days with the media changed every 12 h.</li>
</ol>
2. Contractile function analysis of adult rat cardiac myocytes
<ol>
	<li>Preheat a gel pack and media in a 37 &#176;C incubator for 30 min prior to the experiment.</li>
	<li>Turn on the contractile function platform components listed in the Table of Materials and shown in Supplemental Figure 2, with special attention to the key components, which include an anti-vibration table (Supplemental Figure 2A-1) with an inverted microscope with a deep red (590 nm) filter (Supplemental Figure 2A-2,3) and CCD camera (Supplemental Figure 2A-4) connected to a CCD controller (Supplemental Figure 2A-5 and Supplemental Figure 2C-5) and integrated with a PC computer (Supplemental Figure 2B-14).</li>
	<li>Assemble tubing into the peristaltic pump (Supplemental Figure 2A-8; Supplemental Figure 2A-8), transfer the preheated gel pack into the tube holder (Supplemental Figure 2A-9), and turn on the vacuum (Supplemental Figure 2A-11) and power to the cell stimulator (Supplemental Figure 2B-13).</li>
	<li>Place a 50 mL tube with media in the insulated tube holder (Supplemental Figure 2A-9) and begin perfusion at ~0.5 mL/min through the peristaltic pump tubing (Supplemental Figure 2E-8).</li>
	<li>Add a small amount of preheated media to a small weigh boat (~2 mL). Transfer one CS containing myocytes from the stimulation chamber to the weigh boat. Return the stimulation chamber to 37 &#176;C in the 5% CO2 incubator.</li>
	<li>Remove the CS from the weigh boat and gently wipe the underside. Transfer the CS to a freshly greased CS chamber (Supplemental Figure 2A,F-10; black arrow) and gently add a drop of media (~200 &#956;L) onto the CS.</li>
	<li>Place a platinum electrode mount over the CS (Supplemental Figure 2F; gray arrow) and lay a new CS over the top using forceps. Place a top mount (Supplemental Figure 2F; white arrow) over the CS sandwich and then install two or four #0 pan-head screws to finish assembling the chamber.</li>
	<li>Once the media is present throughout the pump tubing, attach the tubing to the preheater, and then connect the preheater to the CS chamber assembled in step 2.7. Begin perfusion at 0.5 mL/min and place a tissue wipe below the chamber to make sure there are no leaks.</li>
	<li>Place the CS chamber in the stage adapter. The perfused media is collected at the opposite end of the chamber with tubing connected to a vacuum system. Turn on the heating system (Supplemental Figure 2A,D-7) and equilibrate the chamber, objective, and preheater for ~5-10 min to achieve a constant media temperature of 37 &#176;C. Visualize myocytes with the 40x water immersion objective on the microscope during this equilibration time.</li>
	<li>Activate the pacing stimulator (Supplemental Figure 2B-13) by setting the voltage to 1.5x the setting needed for 80% of rod-shaped cells to contract, which is typically 35-40 V. Set the stimulation frequency to 0.2 Hz to optimize contractile function and myocyte survival.</li>
	<li>Collect contractile shortening and re-lengthening traces from continuously perfused and paced myocytes. To collect shortening measurements, use a CCD camera (Supplemental Figure 2A-4) connected to a CCD controller (Supplemental Figure 2B-5,C) and a dedicated computer with an I/O controller board (Supplemental Figure 2B-14; see Table of Materials). The platform measures sarcomere shortening in rat myocytes, although similar results are obtained from myocyte measurements collected from the longitudinal edges of myocytes (see Ecklerle et al.14).</li>
	<li>To collect sarcomere shortening traces, open the software on the computer, select OK, and then File &#62; New. Prepare a screen template by selecting Traces &#62; Edit User Limits &#62; Sarcomere Length and then setting maximum and minimum values, which are typically between 2.0 &#956;m and 1.5 &#956;m. Calibrate the CCD camerawith a 0.01 mm graticule prior to making measurements in the myocytes (Figure 1D).</li>
	<li>To record sarcomere length traces, select File &#62; New. Identify a contracting myocyte and position the myocyte along the longitudinal axis of the camera so the striation pattern is vertical (Figure 1B).</li>
	<li>Use the computer mouse to place the region of interest (ROI) box (pink box) over the myocyte (Figure 1C). Select Collect and Start to record contractile function. Record shortening for 60 s from each myocyte at low stimulation frequencies (&#8804;0.5 Hz).</li>
	<li>Record sarcomere shortening from 5-10 cells per CS. Perform measurements at 1 cell/CS if studies include treatment with agonists/antagonists. Prepare separate pre-warmed media and a different, dedicated piece of perfusion tubing loaded in a multi-channel peristaltic pump and follow steps 2.9.-2.14. to measure the impact of drug or neurohormone delivery on myocyte contractile function.</li>
	<li>To measure shortening over a range of frequencies, pace myocytes at each frequency to obtain steady-state shortening prior to recording5. For these studies, double the perfusion rate and begin recording 15-20 s after stimulation at a new frequency to obtain consistent steady-state contractile responses for frequencies in the range of 0.2 Hz and 2 Hz. Typically, record no more than 2 myocytes/CS for shortening across the given range of stimulation frequencies.</li>
	<li>Record at least 7 contractions/cell to obtain reliable signal-averaged data when analyzing the recordings (see step 3.).</li>
</ol>
3. Data analysis of contractile function in isolated myocytes
<ol>
	<li>To begin, select File, choose a recorded trace, and then select Open. Select Tab 1 (left side) of the base trace and then set the yellow panel at the top of the trace to Sarc-Length. Select Traces and the screen template prepared in step 2.12.</li>
	<li>To prepare an analysis template, select Operations, Monotonic Transient Analysis Options, TTL Event Mark in the Definition of t0 box, as well as the following options from the menu: Baseline (resting sarcomere length), Departure velocity relative to t0 (dep v), Peak (peak height and %peak height/baseline or bl%peak h), Return velocity relative to tPeak (ret v), Time to Peak 50% (TTP50%) using t0, and Time to Baseline 50% (TTR50%) using tPeak. Save these analysis options by selecting Templates &#62; Save Analysis Template with an identifier. Load this analysis template prior to analyzing each trace.</li>
	<li>To begin data analysis, select Marks &#62; Gate &#62; Add Transient &#62; Convert from Event Mark &#62; Analysis Range. Now set the time range from -0.01 s to 1.20 s for myocytes paced at 0.2 Hz (Figure 2A, red marks on the lower portion of the raw trace). Select a shorter time range for myocytes stimulated at higher frequencies.</li>
	<li>Produce a signal-averaged trace by selecting Operations &#62; Average Events. A signal-averaged recording appears below the original base trace.</li>
	<li>Select Tab 1 of the signal-averaged trace (left side of the screen), and then select Marks on the upper menu, followed by Add Transient. Select Operations from the top menu and Monotonic Transient Analysis to display the signal-averaged values for baseline sarcomere length, peak height, bl%peak h, dep v, ret v, TTP50%, and TTR50% in the signal-averaged display panel.</li>
	<li>Transfer each sarcomere transient analysis to a spreadsheet for the composite analysis of multiple myocytes by selecting Export &#62; Monotonic Transient Analysis &#62; Clipboard Current. Paste this data into the spreadsheet for each trace.</li>
	<li>To copy the signal-averaged trace, select Export &#62; Current Trace &#62; Clipboard &#62; Options and set the decimal places to 5, then select Tabs Delimiter, and click OK, and OK. Paste the signal-averaged traces for each myocyte recording into a second spreadsheet.</li>
</ol>
4. Recording Ca2+ transients in rat adult cardiac myocytes
<ol>
	<li>Prior to measuring Ca2+ transients, turn on the platform components described in step 2.2., along with additional fluorescence imaging components (see additional components for Ca2+ imaging analysis in the Table of Materials; Supplemental Figure 2A-5,6; 2B-12).</li>
	<li>Prepare Fura-2AM stock solution containing 1 mM Fura-2AM plus 0.5 M probenecid in dimethylsulfoxide (DMSO). Probenecid minimizes Fura-2AM leakage15.</li>
	<li>Dilute the stock solution to 5 &#956;M Fura-2AM and 2.5 mM probenecid in 2.5 mL of media (see Table of Materials) in one well of a 6-well plate. Add 2.5 mL media alone (no Fura-2AM) to three additional wells in the plate, cover the plate with aluminum foil, and preheat the media to 37 &#176;C.</li>
	<li>Transfer a CS containing myocytes from the pacing chamber to the well containing Fura-2AM, cover, and return the plate to the 37 &#176;C incubator with 5% CO2 for 4 min. Mount tubing in the perfusion pump and perfuse with media during the Fura-2AM loading period, as described in step 2.4.</li>
	<li>Turn off or minimize room lights to reduce fluorescence photobleaching and to optimize the fluorescence signal. Remove the 6-well plate from the incubator, and then wash the CS for 10 s in each of the three wells containing media alone. Transfer the CS to a small weigh boat containing prewarmed media (no added Fura-2AM).</li>
	<li>Mount the CS to freshly greased CS chamber after gently wiping the underside of the CS. Gently add a drop of media to the CS, and then assemble the electrode mount over the CS, followed by a second CS and the top mount. Use #0 pan-head screws to finish assembling the cell chamber, as described in steps 2.6.-2.7.</li>
	<li>Connect the pump tubing filled with media to the preheater, connect the preheater to the CS chamber, and place in a stage adapter, as described in step 2.8. Collect the media with the vacuum tubing system and turn on the heating system (Supplemental Figure 2A,D-7) to equilibrate the chamber to 37&#176;C, as described in steps 2.8.-2.9.</li>
	<li>Activate the pacing stimulator (Supplemental Figure 2B-13) set to 35-40 V and 0.2 Hz, as described in step 2.10.</li>
	<li>Prior to recording a Ca2+ transient, turn on a small flashlight or booklight mounted near the computer keyboard to help see the keyboard. In addition, record the fluorescence background in an area without cardiac myocytes. To start, select File &#62; Start &#62; New, as described in step 2.12.</li>
	<li>Select a ROI, and set one tab (or trace bar, left side) for raw numerator and a second tab for raw denominator, defined in the upper-center of each tab. Record the background for 10-20 s. Right-click the mouse and drag over one trace bar panel to obtain the raw signal-averaged numerator and denominator values. Enter these values by selecting the Operations &#62; Constants, and enter the raw values.</li>
	<li>Prepare a new screen template to collect both shortening and Ca2+ transients. For sarcomere length, select Tab 1 and use the same process as described in step 2.13. For Ca2+ imaging, select Tab 2 &#62; Ratio (upper-center of tab) &#62; Traces &#62; Edit User Limits to set the minimum and maximum levels for the ratio, which are typically set between 1 and 5. Use the same process to define the numerator and denominator ranges for tabs 3 and 4 (left side).</li>
	<li>Position a ROI box over a myocyte image, as described in step 2.14. Select File &#62; New &#62; Collect. Now select Start to collect shortening and ratiometric Ca2+ data. Record &#8805;7 contractions/cell for signal-averaged analysis.</li>
	<li>Repeat the background trace recording described in step 4.10. after recording 2-4 myocytes. Typically, record 4-5 myocytes/CS or fewer myocytes if there are significant changes in the background reading.</li>
</ol>
5. Data analysis of Ca2+&#160;transients in isolated myocytes.
<ol>
	<li>Open the desired file for analysis and select Templates &#62; Screen Template for shortening plus Ca2+ transients. Next, select tabs 1 and 2 (left) to show sarcomere length and Ca2+ ratio, respectively. Select Operations &#62; Constants and enter the numerator and denominator values from the Ca2+ background trace.</li>
	<li>Prepare analysis templates for shortening plus Ca2+ transient data. Select tab 1 (left side) and use the same process as described in step 3.2. to set the sarcomere length analysis template.</li>
	<li>Select tab 2 (left side), and select Operations, Monotonic Transient Analysis Options, TTL event mark in the Definition of t0 box, and the following options from the menu: Baseline (basal ratio), Departure velocity relative to t0 (dep v), Peak (peak height and %peak height/baseline or bl%peak h), Decay velocity relative to tPeak (ret v), Time to Peak 50% (TTP50%) using t0, and Time to Baseline 50% (TTR50%) using tPeak. Save these analysis options by selecting Templates and Save Analysis Template using a new identifier name. Load this analysis template prior to analyzing each trace.</li>
	<li>Use the same process as described in steps 3.3.-3.6. to average the signal and then analyze sarcomere length, followed by the same steps for Ca2+ transients. Set separate marks for sarcomere length and for the Ca2+ transient ratio trace.</li>
</ol>

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The authors have no competing financial interests or other conflicts of interest.

The chronic pacing protocol outlined in step 1 extends the useful time for studying isolated myocytes and assessing the impact of longer treatments. In our lab, consistent results were obtained up to 4 days post isolation when measuring contractile function using sarcomere length on chronically paced myocytes. However, myocyte contractile function deteriorates quickly when using media older than 1 week to pace myocytes.
For contractile function studies, the data are collected at 37 &#176;C, wh...

The analysis of cardiac pump function often requires a range of approaches to gain adequate insight, especially for animal models of heart failure (HF). Echocardiography or hemodynamic measurements provide insight into in vivo cardiac dysfunction1, while in vitro approaches are often employed to identify whether dysfunction arises from changes in the myofilament and/or the Ca2+ transient responsible for coupling excitation, or the action potential, with contractile function (e.g., excitation-contraction [E-C] coupling). In vitro approaches also provide an opportunity to screen the functional response to neurohormones, vector-induced genetic alterations, as well as potential therapeutic agents2 prior to pursuing costly and/or laborious in vivo treatment strategies.
Several approaches are available to investigate in vitro contractile function, including force measurements in intact trabeculae3 or permeabilized myocytes4, as well as unloaded shortening and Ca2+ transients in intact myocytes in the presence and absence of HF5,6. Each of these approaches focuses on cardiac myocyte contractile function, which is directly responsible for cardiac pump function2,7. However, the analysis of both contraction and E-C coupling together is most often performed by measuring shortening of the muscle length and Ca2+ transients in isolated, Ca2+ tolerant adult myocytes. The laboratory utilizes a detailed published protocol to isolate myocytes from rat hearts for this step8.
Both the Ca2+ transient and myofilaments contribute to shortening and re-lengthening in intact myocytes and can contribute to contractile dysfunction2,7. Thus, this approach is recommended when in vitro functional analysis requires an intact myocyte containing the Ca2+ cycling machinery plus the myofilaments. For example, intact isolated myocytes are desirable for studying contractile function after modifying the myofilament or Ca2+ cycling function via gene transfer9. In addition, an intact myocyte approach is suggested for analyzing the functional impact of neurohormones when studying the impact of downstream second messenger signaling pathways and/or response to therapeutic agents2. An alternative measurement of load-dependent force in single myocytes is most often performed after membrane permeabilization (or skinning) at low temperatures (&#8804;15 &#176;C) to remove the Ca2+ transient contribution and focus on myofilament function10. The measurement of load-dependent force plus Ca2+ transients in intact myocytes is rare due largely to the complex and technical challenge of the approach11, especially when higher throughput is needed, such as for measuring responses to neurohormone signaling or as a screen for therapeutic agents. The analysis of cardiac trabeculae overcomes these technical challenges but also may be influenced by non-myocytes, fibrosis, and/or extracellular matrix remodeling2. Each of the approaches described above requires a preparation containing adult myocytes because neonatal myocytes and myocytes derived from inducible pluripotent stem cells (iPSCs) do not yet express the full complement of adult myofilament proteins and usually lack the level of myofilament organization present in the adult rod-shaped myocyte2. To date, evidence in iPSCs indicates that the full transition to adult isoforms exceeds more than 134 days in culture12.
Given the focus of this collection on HF, the protocols include approaches and analysis to differentiate contractile function in failing versus non-failing intact myocytes. Representative examples are provided from rat myocytes studied 18-20 weeks after a supra-renal coarctation, described earlier5,13. Comparisons are then made to myocytes from sham-treated rats.
The protocol and imaging platform described here are used to analyze and monitor changes in shortening and Ca2+ transients in rod-shaped cardiac myocytes during the development of HF. For this analysis, 2 x 104 Ca2+-tolerant, rod-shaped myocytes are plated on 22 mm2 laminin-coated glass coverslips (CSs) and cultured overnight, as described earlier8. The components assembled for this imaging platform, along with the media and buffers used for optimal imaging, are provided in the Table of Materials. A guide for data analysis using a software and the representative results are also provided here. The overall protocol is broken down into separate sub-sections, with the first three sections focusing on isolated rat myocytes and data analysis, followed by cellular Ca2+ transient experiments and data analysis in myocytes.

<table><tbody><tr><td>&lt;strong&gt;MEDIA&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Bovine serum albumin</td><td>Sigma (Roche)</td><td>3117057001</td><td>Final concentration = 0.2% (w/v)</td></tr><tr><td>Glutathione</td><td>Sigma</td><td>G-6529</td><td>Final concentration = 10 mM</td></tr><tr><td>HEPES</td><td>Sigma</td><td>H-7006</td><td>Final concentration = 15 mM</td></tr><tr><td>M199</td><td>Sigma</td><td>M-2520</td><td>1 bottle makes 1 L; pH 7.45</td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Sigma</td><td>S-8875</td><td>Final concentration = 4 mM</td></tr><tr><td>Penicillin/streptomycin</td><td>Fisher</td><td>15140122</td><td>Final concentration = 100 U/mL penicillin, 100 &amp;mu;g/mL streptomycin</td></tr><tr><td>&lt;strong&gt;REAGENTS SPECIFICALLY FOR Ca&lt;sup&gt;2+&lt;/sup&gt; IMAGING&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Dimethylsulfoxide (DMSO)</td><td>Sigma&amp;nbsp;</td><td>D2650</td><td></td></tr><tr><td>Fura-2AM</td><td>Invitrogen (Molecular Probes)</td><td>F1221</td><td>50 &amp;mu;g/vial;&amp;nbsp; Prepare stock solution of 1 mM Fura-2AM + 0.5 M probenicid in DMSO;&amp;nbsp; Final Fura2-AM concentration in media is&amp;nbsp; 5 &amp;mu;M</td></tr><tr><td>Probenicid</td><td>Invitrogen (Fisher)</td><td>P36400</td><td>Add 7.2 mg probenicid (0.5 M) to 1 mM Fura-2AM stock; Final concentration in media is 2.5 mM</td></tr><tr><td>&lt;strong&gt;MATERIALS FOR RAT MYOCYTE PACING&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>#1 22&amp;nbsp; mm&lt;sup&gt;2&lt;/sup&gt; glass coverslips</td><td>Corning</td><td>2845-22</td><td></td></tr><tr><td>3 x 36 inch cables with banana jacks</td><td>Pomona Electronics</td><td>B-36-2</td><td>Supplemental Figure 1, panel C</td></tr><tr><td>37&lt;sup&gt;o&lt;/sup&gt;C Incubator&amp;nbsp; with 95% O&lt;sub&gt;2&lt;/sub&gt;:5% CO&lt;sub&gt;2&lt;/sub&gt;&amp;nbsp;</td><td>Forma</td><td>3110</td><td>Supplemental Figure 1, panel E.&amp;nbsp; Multiple models are appropriate</td></tr><tr><td>Class II A/B3 Biosafety cabinet with UV lamp</td><td>Forma</td><td>1286</td><td>Multiple models are appropriate</td></tr><tr><td>Forceps - Dumont #5 5/45</td><td>Fine Science Tools</td><td>11251-35</td><td></td></tr><tr><td>Hot bead sterilizer</td><td>Fine Science Tools</td><td>1800-45</td><td></td></tr><tr><td>Low magnification inverted microscope</td><td>Leica</td><td>DM-IL&amp;nbsp;</td><td>Position this microscope adjacent to the incubator to monitor paced myocytes for contraction at the start of pacing and after media changes; &amp;nbsp;4X and 10X objectives recommended</td></tr><tr><td>Pacing chamber</td><td>Custom</td><td></td><td>Supplemental Figure 1, panel A. The Ionoptix C-pace system is a commercially available alternative or see &lt;sup&gt;22&lt;/sup&gt;</td></tr><tr><td>Stimulator</td><td>Ionoptix</td><td>Myopacer</td><td>Supplemental Figure 1, panel D.</td></tr><tr><td>&lt;strong&gt;MATERIALS FOR CONTRACTILE FUNCTION and/or Ca&lt;sup&gt;2+&lt;/sup&gt; IMAGING ANALYSIS&lt;/strong&gt;</td><td></td><td></td><td>ID in Supplemental Figure 2 &amp;amp; Alternatives/Recommended Options</td></tr><tr><td>Additional components for Ca&lt;sup&gt;2+&lt;/sup&gt; imaging analysis</td><td>Ionoptix</td><td>Essential system components: -- Photon counting system&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; --&amp;nbsp; Xenon power supply with dual excitation light source&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; --&amp;nbsp; Fluorescence interface</td><td>&amp;nbsp;- The photon counting system contains a photomultiplier (PMT) tube and dichroic mirrorand is installed adjacent to the CCD camera&amp;nbsp; (panel A #4).&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; - The power supply for the xenon bulb light source (see panel A #5 and panel C, left) is integrated with a dual excitation interface (340/380 nm&amp;nbsp; excitation and 510 nm emission) shown in panel A #6.&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; - The fluorescence interface between the computer and&amp;nbsp; light source is shown in panel B, #12.</td></tr><tr><td>CCD camera with image acquisition&amp;nbsp; hardware and software (240 frames/s)</td><td>Ionoptix&amp;nbsp;</td><td>Myocam with CCD controller&amp;nbsp;</td><td>Myocam and CCD controller are shown in Supplemental Fig. 2, panel A #4&amp;nbsp; and&amp;nbsp; panel A #5 &amp;amp; panel C #5 (right), respectively.&amp;nbsp; The controller is integrated with a PC computer system (panel B #14).&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; &amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;</td></tr><tr><td>&amp;nbsp;Chamber stimulator</td><td>Ionoptix&amp;nbsp;</td><td>Myopacer</td><td>Panel B, #13; Alternative:&amp;nbsp; Grass model S48</td></tr><tr><td>Coverslip mounted perfusion chamber</td><td>Custom chamber for 22 mm&lt;sup&gt;2&lt;/sup&gt; coverslip with silicone adapter and 2-4 Phillips pan-head #0 screws&amp;nbsp; (arrow, panel F)</td><td></td><td>Panel A #10 &amp;amp; panel F;&amp;nbsp;&amp;nbsp; Chamber temperature is calibrated to 37&lt;sup&gt;o&lt;/sup&gt;C using a TH-10Km probe and the TC&lt;sup&gt;2&lt;/sup&gt;BIP temperature controller (see temperature controller).&amp;nbsp; Commercial alternatives:&amp;nbsp; Ionoptix FHD or C-stim cell&amp;nbsp; chambers; Cell MicroControls culture stimulation system</td></tr><tr><td>Dedicated computer &amp;amp; software for data collection and analysis of function/Ca2+ transients</td><td>Ionoptix</td><td>PC with Ionwizard PC board and software</td><td>Panel B, #14; Contractile function is measured using either SarcLen (sarcomere length) or SoftEdge (myocyte length) acquisition modules of the IonWizard software. The Ionwizard software also includes PMT acquisition software for ratiometric&amp;nbsp; Ca&lt;sup&gt;2+&lt;/sup&gt; imaging in Fura-2AM-loaded myoyctes.&lt;br/&gt; - A 4 post electronic rack mount cabinet and shelves are recommended for housing the somputer and cell stimulator.&amp;nbsp; The fluorescence interface for Ca&lt;sup&gt;2+&lt;/sup&gt; imaging also is housed in this cabinet (see below).</td></tr><tr><td>Forceps - Dumont #5 TI</td><td>Fine Science Tools</td><td>11252-40</td><td>Panel F</td></tr><tr><td>Insulated tube holder for media</td><td>Custom</td><td></td><td>Panel A #9; This holder is easily assembled using styrofoam &amp;amp; a pre-heated gel pack to keep media warm</td></tr><tr><td>Inverted brightfield microscope</td><td>Nikon</td><td>TE-2000S</td><td>Install a rotating turret for epi-fluorescence (Panel A #2) for Ca2+ imaging.&amp;nbsp; &amp;nbsp;A deep red (590 nm) condenser filter also is recommended to minimize fluorescence bleaching during Ca&lt;sup&gt;2+&lt;/sup&gt; imaging.</td></tr><tr><td>Isolator Table</td><td>TMC Vibration Control</td><td>30 x 36 inches</td><td>Panel A, #1; Desirable:&amp;nbsp; elevated shelving, Faraday shielding&amp;nbsp;&amp;nbsp;</td></tr><tr><td>Microscope eyepieces &amp;amp; objective</td><td>Nikon</td><td>10X CFI eyepieces 40X water CFI Plan Fluor objective</td><td>Panel A #3; 40X objective:&amp;nbsp; n.a. 0.08; w.d. 2 mm.&amp;nbsp; A Cell MicroControls HLS-1 objective heater is mounted around the objective (see temperature controller below).&amp;nbsp; &amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;NOTE:&amp;nbsp; water immersion dispensers also are now available for water-based objectives.&amp;nbsp;&amp;nbsp;</td></tr><tr><td>Peristaltic pump</td><td>Gilson</td><td>Minipuls 3</td><td>Panel A #8 and panel E</td></tr><tr><td>small weigh boat</td><td>Fisher&amp;nbsp;</td><td>08-732-112</td><td></td></tr><tr><td>Temperature controller</td><td>Cell MicroControls</td><td>TC&lt;sup&gt;2&lt;/sup&gt;BIP</td><td>Panel A #7; Panel D. This temperature controller heats the coverslip chamber to 37&lt;sup&gt;o&lt;/sup&gt;C.&amp;nbsp;&amp;nbsp;&amp;nbsp; A preheater and objective heater are recommended for this platform.&amp;nbsp; A Cell MicroControls HPRE2 preheater and&amp;nbsp; HLS-1 objective heater are controlled&amp;nbsp; by the TC&lt;sup&gt;2&lt;/sup&gt;BIP temperature controller for our studies.</td></tr><tr><td>Under cabinet LED light with motion sensor&amp;nbsp;</td><td>Sylvania</td><td>#72423 LED light</td><td>Recommended for data collection during Ca&lt;sup&gt;2+&lt;/sup&gt; transient imaging under minimal room light..&amp;nbsp; Alternative:&amp;nbsp; A clip on flashlight/book light with flexible neck - multiple suppliers are available.</td></tr><tr><td>Vacuum line with in-line Ehrlenmeyer flask &amp;amp; protective filter</td><td>Fisher</td><td>Tygon tubing - E363;&amp;nbsp; polypropylene Ehrlenmeyer flask - 10-182-50B; Vacuum filter - 09-703-90</td><td>Panel A #11</td></tr></tbody></table>

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.

Contractile function studies are performed on rat myocytes starting the day after isolation (day 2) up to 4 days post isolation. Although myocytes can be recorded the day after isolation (i.e., day 2), longer culture times are often required after gene transfer or treatments to modify contractile function8. For myocytes cultured for more than 18 h after isolation, the pacing protocol described in section 1 helps maintain t-tubules and consistent shortening and re-lengthening results.
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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.

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 ...

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Research

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Medicine

1.3K Views.  University of Michigan, Ann Arbor. 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.

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

Analysis of Tubular Membrane Networks in Cardiac Myocytes from Atria and Ventricles

In cardiac myocytes a complex network of membrane tubules - the transverse-axial tubule system (TATS) - controls deep intracellular signaling functions. While the outer surface membrane and associated TATS membrane components appear to be continuous, there are substantial differences in lipid and protein content. In ventricular myocytes (VMs), certain TATS components are highly abundant contributing to rectilinear tubule networks and regular branching 3D architectures. It is thought that peripheral TATS components propagate action potentials from the cell surface to thousands of remote intracellular sarcoendoplasmic reticulum (SER) membrane contact domains, thereby activating intracellular Ca2+ release units (CRUs). In contrast to VMs, the organization and functional role of TATS membranes in atrial myocytes (AMs) is significantly different and much less understood. Taken together, quantitative structural characterization of TATS membrane networks in healthy and diseased myocytes is an essential prerequisite towards better understanding of functional plasticity and pathophysiological reorganization. Here, we present a strategic combination of protocols for direct quantitative analysis of TATS membrane networks in living VMs and AMs. For this, we accompany primary cell isolations of mouse VMs and/or AMs with critical quality control steps and direct membrane staining protocols for fluorescence imaging of TATS membranes. Using an optimized workflow for confocal or superresolution TATS image processing, binarized and skeletonized data are generated for quantitative analysis of the TATS network and its components. Unlike previously published indirect regional aggregate image analysis strategies, our protocols enable direct characterization of specific components and derive complex physiological properties of TATS membrane networks in living myocytes with high throughput and open access software tools. In summary, the combined protocol strategy can be readily applied for quantitative TATS network studies during physiological myocyte adaptation or disease changes, comparison of different cardiac or skeletal muscle cell types, phenotyping of transgenic models, and pharmacological or therapeutic interventions.

In cardiac myocytes, tubular membrane structures form intracellular networks. We describe optimized protocols for i) isolation of myocytes from mouse heart including quality control, ii) live cell staining for state-of-the-art fluorescence microscopy, and iii) direct image analysis to quantify the component complexity and the plasticity of intracellular membrane networks.

In cardiac myocytes a complex network of membrane tubules - the transverse-axial tubule system (TATS) - controls deep intracellular signaling functions. ...

Bioengineering

Optical Imaging of Isolated Murine Ventricular Myocytes

The ability to isolate adult cardiac myocytes has permitted researchers to study a variety of cardiac pathologies at the single cell level. While advances in calcium sensitive dyes have permitted the robust optical recording of single cell calcium dynamics, recording of robust transmembrane optical voltage signals has remained difficult. Arguably, this is because of the low single to noise ratio, phototoxicity, and photobleaching of traditional potentiometric dyes. Therefore, single cell voltage measurements have long been confined to the patch clamp technique which while the gold standard, is technically demanding and low throughput. However, with the development of novel potentiometric dyes, large, fast optical responses to changes in voltage can be obtained with little to no phototoxicity and photobleaching. This protocol describes in detail how to isolate adult murine myocytes which can be used for cellular shortening, calcium, and optical voltage measurements. Specifically, the protocol describes how to use a ratiometric calcium dye, a single-excitation calcium dye, and a single excitation voltage dye. This approach can be used to assess the cardiotoxicity and arrhythmogenicity of various chemical agents. While phototoxicity is still an issue at the single cell level, methodology is discussed on how to reduce it.

We present the methodology for the isolation of murine myocytes and how to obtain voltage or calcium traces simultaneously with sarcomere shortening traces using fluorescence photometry with simultaneous digital cell geometry measurements.

The ability to isolate adult cardiac myocytes has permitted researchers to study a variety of cardiac pathologies at the single cell level. While advances ...

Isolation, Culture, and Functional Characterization of Adult Mouse Cardiomyoctyes

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

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

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

Biology

Isolation and Physiological Analysis of Mouse Cardiomyocytes

Cardiomyocytes, the workhorse cell of the heart, contain exquisitely organized cytoskeletal and contractile elements that generate the contractile force used to pump blood. Individual cardiomyocytes were first isolated over 40 years ago in order to better study the physiology and structure of heart muscle. Techniques have rapidly improved to include enzymatic digestion via coronary perfusion. More recently, analyzing the contractility and calcium flux of isolated myocytes has provided a vital tool in the cellular and sub-cellular analysis of heart failure. Echocardiography and EKGs provide information about the heart at an organ level only. Cardiomyocyte cell culture systems exist, but cells lack physiologically essential structures such as organized sarcomeres and t-tubules required for myocyte function within the heart. In the protocol presented here, cardiomyocytes are isolated via Langendorff perfusion. The heart is removed from the mouse, mounted via the aorta to a cannula, perfused with digestion enzymes, and cells are introduced to increasing calcium concentrations. Edge and sarcomere detection software is used to analyze contractility, and a calcium binding fluorescent dye is used to visualize calcium transients of electrically paced cardiomyocytes; increasing understanding of the role cellular changes play in heart dysfunction. Traditionally used to test drug effects on cardiomyocytes, we employ this system to compare myocytes from WT mice and mice with a mutation that causes dilated cardiomyopathy. This protocol is unique in its comparison of live cells from mice with known heart function and known genetics. Many experimental conditions are reliably compared, including genetic or environmental manipulation, infection, drug treatment, and more. Beyond physiologic data, isolated cardiomyocytes are easily fixed and stained for cytoskeletal elements. Isolating cardiomyocytes via perfusion is an extremely versatile method, useful in studying cellular changes that accompany or lead to heart failure in a variety of experimental conditions.

Individual cardiomyocytes from wild type and mutant mice can be isolated from the heart in order to study their contractility and calcium transients. This allows characterization of the contribution of cellular dysfunction to heart dysfunction from any cause.

Cardiomyocytes, the workhorse cell of the heart, contain exquisitely organized cytoskeletal and contractile elements that generate the contractile force ...

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 ...

Electromechanical Assessment of Optogenetically Modulated Cardiomyocyte Activity

Over the past two decades, optogenetic tools have been established as potent means to modulate cell-type specific activity in excitable tissues, including the heart. While Channelrhodopsin-2 (ChR2) is a common tool to depolarize the membrane potential in cardiomyocytes (CM), potentially eliciting action potentials (AP), an effective tool for reliable silencing of CM activity has been missing. It has been suggested to use anion channelrhodopsins (ACR) for optogenetic inhibition. Here, we describe a protocol to assess the effects of activating the natural ACR GtACR1 from Guillardia theta in cultured rabbit CM. Primary readouts are electrophysiological patch-clamp recordings and optical tracking of CM contractions, both performed while applying different patterns of light stimulation. The protocol includes CM isolation from rabbit heart, seeding and culturing of the cells for up to 4 days, transduction via adenovirus coding for the light-gated chloride channel, preparation of patch-clamp and carbon fiber setups, data collection and analysis. Using the patch-clamp technique in whole-cell configuration allows one to record light-activated currents (in voltage-clamp mode, V-clamp) and AP (current-clamp mode, I-clamp) in real time. In addition to patch-clamp experiments, we conduct contractility measurements for functional assessment of CM activity without disturbing the intracellular milieu. To do so, cells are mechanically preloaded using carbon fibers and contractions are recorded by tracking changes in sarcomere length and carbon fiber distance. Data analysis includes assessment of AP duration from I-clamp recordings, peak currents from V-clamp recordings and force calculation from carbon fiber measurements. The described protocol can be applied to the testing of biophysical effects of different optogenetic actuators on CM activity, a prerequisite for the development of a mechanistic understanding of optogenetic experiments in cardiac tissue and whole hearts.

We present a protocol for evaluating the electromechanical effects of GtACR1 activation in rabbit cardiomyocytes. We provide detailed information on cell isolation, culturing and adenoviral transduction, and on functional experiments with the patch-clamp and carbon-fiber techniques.

Over the past two decades, optogenetic tools have been established as potent means to modulate cell-type specific activity in excitable tissues, including ...

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

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

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

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

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

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

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

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

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

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 ...

Isolation of High Quality Murine Atrial and Ventricular Myocytes for Simultaneous Measurements of Ca2+ Transients and L-Type Calcium Current

Mouse models play a crucial role in arrhythmia research and allow studying key mechanisms of arrhythmogenesis including altered ion channel function and calcium handling. For this purpose, atrial or ventricular cardiomyocytes of high quality are necessary to perform patch-clamp measurements or to explore calcium handling abnormalities. However, the limited yield of high-quality cardiomyocytes obtained by current isolation protocols does not allow both measurements in the same mouse. This article describes a method to isolate high-quality murine atrial and ventricular myocytes via retrograde enzyme-based Langendorff perfusion, for subsequent simultaneous measurements of calcium transients and L-type calcium current from one animal. Mouse hearts are obtained, and the aorta is rapidly cannulated to remove blood. Hearts are then initially perfused with a calcium-free solution (37 &#176;C) to dissociate the tissue at the level of intercalated discs and afterwards with an enzyme solution containing little calcium to disrupt extracellular matrix (37 &#176;C). The digested heart is subsequently dissected into atria and ventricles. Tissue samples are chopped into small pieces and dissolved by carefully pipetting up and down. The enzymatic digestion is stopped, and cells are stepwise reintroduced to physiologic calcium concentrations. After loading with a fluorescent Ca2+-indicator, isolated cardiomyocytes are prepared for simultaneous measurement of calcium currents and transients. Additionally, isolation pitfalls are discussed and patch-clamp protocols and representative traces of L-type calcium currents with simultaneous calcium transient measurements in atrial and ventricular murine myocytes isolated as described above are provided.

Isolation of High Quality Murine Atrial and Ventricular Myocytes for Simultaneous Measurements of Ca2+ Transients and L-Type Calcium Current

Mouse models allow studying key mechanisms of arrhythmogenesis. For this purpose, high quality cardiomyocytes are necessary to perform patch-clamp measurements. Here, a method to isolate murine atrial and ventricular myocytes via retrograde enzyme-based Langendorff perfusion, which allows simultaneous measurements of calcium-transients and L-type calcium current, is described.

Mouse models play a crucial role in arrhythmia research and allow studying key mechanisms of arrhythmogenesis including altered ion channel function and ...

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

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