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, which is near the body temperature for rats. To optimize myocyte viability and obtain consistent traces for each myocyte, these experiments are performed at a pacing frequency lower than the physiological heart rate of ~5 Hz in rats. These settings produce consistent contractile function data, but chamber temperature and/or pacing frequency can be adjusted if the shortening traces remain consistent across multiple CSs containing the same treatment group of myocytes. In addition, optimal results are obtained by choosing myocytes that lack blebs, are fully adhered to a CS, and contract at both ends of a rod-shaped cell. Cells attached at only one end, those with multiple blebs, and/or myocytes contracting at only one end are not desirable and are often more difficult to record due to background movement. For these studies, &#8805;20 sarcomeres are included in the ROI to achieve a sharp power spectrum peak, reproducible resting sarcomere lengths, and an optimal signal/noise ratio for the shortening trace. A ROI with as few as seven sarcomeres can be used if the power spectrum peak remains sharp. The resting sarcomere length in isolated rat myocytes usually ranges from 2.00-1.80 &#956;m. If a resting sarcomere length is less than 1.5 &#956;m, an active contraction is not realistic based on our understanding of sarcomere architecture18 and usually is the result of sub-optimal myocyte orientation.
The measurement of contractile function with or without Ca2+ transient measurements in intact myocytes is a higher throughput approach requiring less investment in developing the technical expertise needed for force measurements in intact single cells11. Intact myocyte studies also focus exclusively on myocyte function, without the contribution of other cell types in multi-cellular preparations2. An emerging area in this field is the development of higher throughput analysis by simultaneously measuring shortening and/or Ca2+ transients in multiple intact myocytes to accelerate the screening of therapeutic agents at the cellular level prior to more extensive in vivo analysis.
Contractile function and Ca2+ transient measurements also are possible in intact myocytes isolated from other rodent models such as mice, although there are some important considerations and differences compared to rat myocytes. Mouse myocytes are viable in culture for 24-36 h, but viability progressively decreases in myocytes cultured for more than 48 h19. This truncated culture interval should be considered when using vector-based gene transfer, especially for myofilament proteins that have a half-life in days rather than hours20. Unlike rat myocytes, the successful isolation and culture of mouse myocytes also depend on the addition of myosin ATPase inhibitors, such as 2,3-butanedione monoxime or blebbistatin, to culture media19. As a result, chronic pacing is not a viable option in mouse myocyte cultures. A longer equilibration time of 15-20 min is also needed to remove the functional impact of these myosin inhibitors on isolated mouse myocytes. Finally, mouse myocytes are optimally paced at 0.5 Hz to obtain reproducible shortening traces compared to the 0.2 Hz utilized for rat myocytes.
In studies on PO-treated rats, contractile dysfunction is consistently detected with low-frequency pacing in rat myocytes. A greater frequency-dependent reduction in shortening also is detected in myocytes after PO compared to sham rats when there is in vivo contractile dysfunction5,13. During a study using a range of frequencies, doubling the pump speed to provide adequate media produced steady-state shortening within 15-20 s after changing the stimulation frequency between 0.2 and 2 Hz. Rat myocytes also respond to short-term stimulation at higher frequencies up to 8 Hz, although cell viability quickly deteriorates at these higher frequencies. Overall, myocyte contractile function has proven to be a reproducible approach to confirm or validate in vivo cardiac dysfunction in rat models, such as PO- compared to sham-treated rats (Table 1; see Kim et al.13). In addition, this platform can test whether a treatment targeting myocytes restores or impairs contractile function prior to pursuing more expensive and/or laborious in vivo approaches. Both acute delivery and longer exposure to therapeutic agents and/or after gene delivery during primary culture are possible using this approach. For example, gene transfer experiments to replace endogenous cTnI with cTnIT144D indicate no significant change in shortening amplitude in myocytes from sham rats. In contrast, cTnIT144D restores peak shortening amplitude toward sham values in myocytes from PO-treated rats (Table 1). An inherent weakness of this approach is the absence of the typical load present in myocytes under in vivo conditions. As a result, responses may differ from load-dependent functional responses, and the example experiment indicates that in vivo approaches are needed to further investigate whether cTnIT144D improves cardiac performance during PO.
The measurement of both shortening and Ca2+ transients in the same myocyte is an advantage of this platform. For example, the direction of change may differ for Ca2+ transients compared to contractile function. As shown in Table 2, peak shortening increases significantly after gene transfer of cTnIT144D into myocytes from 2-3-month-old rats. This outcome differs from the data obtained in older, sham-treated myocytes (Table 1) and may be related to rat age5. However, further testing is needed to prove this interpretation. The data in Table 2 also show that cTnIT144D does not change the peak Ca2+ amplitude and, instead, tends to slow the Ca2+ decay rate and elevate resting Ca2+. These findings suggest that cTnIT144D expression enhances contractile function, while the Ca2+ cycling machinery compensates to dampen this effect. These results highlight the ability of this approach to distinguish between changes in contractile function and Ca2+ cycling. While the specific results presented here are not yet definitive, they provide a solid rationale for developing a genetic animal model to express myocardial cTnIT144D and evaluate whether its expression blunts dysfunction caused by PO in the future. A final observation from the Ca2+ transient data is that fluorescent dyes such as Fura-2AM change the contractile function kinetics in myocytes21. Although the relative response produced by a given treatment or animal model is comparable within Fura-2-loaded myocytes, these results should not be combined with shortening measurements made in the absence of Fura-2AM. To optimize the use of Fura-2AM for Ca2+ transient analysis, the laboratory most often measures shortening in the absence of Fura-2AM and performs a subset of experiments to measure Ca2+ transients.

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>MEDIA</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>NaHCO3</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 &#956;g/mL streptomycin</td>
		</tr>
		<tr>
			<td>REAGENTS SPECIFICALLY FOR Ca2+ IMAGING</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dimethylsulfoxide (DMSO)</td>
			<td>Sigma&#160;</td>
			<td>D2650</td>
			<td></td>
		</tr>
		<tr>
			<td>Fura-2AM</td>
			<td>Invitrogen (Molecular Probes)</td>
			<td>F1221</td>
			<td>50 &#956;g/vial;&#160; Prepare stock solution of 1 mM Fura-2AM + 0.5 M probenicid in DMSO;&#160; Final Fura2-AM concentration in media is&#160; 5 &#956;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>MATERIALS FOR RAT MYOCYTE PACING</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>#1 22&#160; mm2 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>37oC Incubator&#160; with 95% O2:5% CO2&#160;</td>
			<td>Forma</td>
			<td>3110</td>
			<td>Supplemental Figure 1, panel E.&#160; 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&#160;</td>
			<td>Position this microscope adjacent to the incubator to monitor paced myocytes for contraction at the start of pacing and after media changes; &#160;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 22</td>
		</tr>
		<tr>
			<td>Stimulator</td>
			<td>Ionoptix</td>
			<td>Myopacer</td>
			<td>Supplemental Figure 1, panel D.</td>
		</tr>
		<tr>
			<td>MATERIALS FOR CONTRACTILE FUNCTION and/or Ca2+ IMAGING ANALYSIS</td>
			<td></td>
			<td></td>
			<td>ID in Supplemental Figure 2 &#38; Alternatives/Recommended Options</td>
		</tr>
		<tr>
			<td>Additional components for Ca2+ imaging analysis</td>
			<td>Ionoptix</td>
			<td>Essential system components: -- Photon counting system&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; --&#160; Xenon power supply with dual excitation light source&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; --&#160; Fluorescence interface</td>
			<td>&#160;- The photon counting system contains a photomultiplier (PMT) tube and dichroic mirrorand is installed adjacent to the CCD camera&#160; (panel A #4).&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; - 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&#160; excitation and 510 nm emission) shown in panel A #6.&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; - The fluorescence interface between the computer and&#160; light source is shown in panel B, #12.</td>
		</tr>
		<tr>
			<td>CCD camera with image acquisition&#160; hardware and software (240 frames/s)</td>
			<td>Ionoptix&#160;</td>
			<td>Myocam with CCD controller&#160;</td>
			<td>Myocam and CCD controller are shown in Supplemental Fig. 2, panel A #4&#160; and&#160; panel A #5 &#38; panel C #5 (right), respectively.&#160; The controller is integrated with a PC computer system (panel B #14).&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; &#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;</td>
		</tr>
		<tr>
			<td>&#160;Chamber stimulator</td>
			<td>Ionoptix&#160;</td>
			<td>Myopacer</td>
			<td>Panel B, #13; Alternative:&#160; Grass model S48</td>
		</tr>
		<tr>
			<td>Coverslip mounted perfusion chamber</td>
			<td>Custom chamber for 22 mm2 coverslip with silicone adapter and 2-4 Phillips pan-head #0 screws&#160; (arrow, panel F)</td>
			<td></td>
			<td>Panel A #10 &#38; panel F;&#160;&#160; Chamber temperature is calibrated to 37oC using a TH-10Km probe and the TC2BIP temperature controller (see temperature controller).&#160; Commercial alternatives:&#160; Ionoptix FHD or C-stim cell&#160; chambers; Cell MicroControls culture stimulation system</td>
		</tr>
		<tr>
			<td>Dedicated computer &#38; 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&#160; Ca2+ imaging in Fura-2AM-loaded myoyctes. 
			- A 4 post electronic rack mount cabinet and shelves are recommended for housing the somputer and cell stimulator.&#160; The fluorescence interface for Ca2+ 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 &#38; 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.&#160; &#160;A deep red (590 nm) condenser filter also is recommended to minimize fluorescence bleaching during Ca2+ imaging.</td>
		</tr>
		<tr>
			<td>Isolator Table</td>
			<td>TMC Vibration Control</td>
			<td>30 x 36 inches</td>
			<td>Panel A, #1; Desirable:&#160; elevated shelving, Faraday shielding&#160;&#160;</td>
		</tr>
		<tr>
			<td>Microscope eyepieces &#38; objective</td>
			<td>Nikon</td>
			<td>10X CFI eyepieces 40X water CFI Plan Fluor objective</td>
			<td>Panel A #3; 40X objective:&#160; n.a. 0.08; w.d. 2 mm.&#160; A Cell MicroControls HLS-1 objective heater is mounted around the objective (see temperature controller below).&#160; &#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;NOTE:&#160; water immersion dispensers also are now available for water-based objectives.&#160;&#160;</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&#160;</td>
			<td>08-732-112</td>
			<td></td>
		</tr>
		<tr>
			<td>Temperature controller</td>
			<td>Cell MicroControls</td>
			<td>TC2BIP</td>
			<td>Panel A #7; Panel D. This temperature controller heats the coverslip chamber to 37oC.&#160;&#160;&#160; A preheater and objective heater are recommended for this platform.&#160; A Cell MicroControls HPRE2 preheater and&#160; HLS-1 objective heater are controlled&#160; by the TC2BIP temperature controller for our studies.</td>
		</tr>
		<tr>
			<td>Under cabinet LED light with motion sensor&#160;</td>
			<td>Sylvania</td>
			<td>#72423 LED light</td>
			<td>Recommended for data collection during Ca2+ transient imaging under minimal room light..&#160; Alternative:&#160; A clip on flashlight/book light with flexible neck - multiple suppliers are available.</td>
		</tr>
		<tr>
			<td>Vacuum line with in-line Ehrlenmeyer flask &#38; protective filter</td>
			<td>Fisher</td>
			<td>Tygon tubing - E363;&#160; 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.
A representative portion of a CS containing myocytes for shortening studies is shown in Figure 1A, along with an appropriately positioned myocyte prior to setting the ROI (Figure 1B). Once a&#160;ROI is identified (Figure 1C; pink box), the algorithm information shown below the myocyte also helps to optimize myocyte positioning prior to recording. Specifically, the linear optical density (LOD; black line) is an indicator for the number and spacing of sarcomeres, and a sharp power spectrum in the fast Fourier transform trace (FFT, red line) helps to achieve optimal alignment for the recording of shortening and re-lengthening. The graticule pattern used for the calibration of sarcomere length (and edge detection) is shown in Figure 1D. A typical aligned recording of sarcomere length shortening is shown in Figure 2A (top panel), along with the signal-averaged analysis described in section 3 (lower panel).
Dysfunction is often detected in myocytes when there is in vivo dysfunction in animal models. For example, systolic dysfunction observed by echocardiography in response to pressure overload (PO)13 is also detected in myocyte shortening studies5. To illustrate data traces, representative raw (Figure 2; upper panel) and signal-averaged (Figure 2;&#160;lower panel) traces obtained at 0.2 Hz are shown for myocytes from sham- (Figure 2A) and PO-treated (Figure 2B) rats. To test whether myocyte function could be rescued after PO, viral-mediated gene transfer at the time of myocyte isolation also was used to replace endogenous cardiac troponin I (cTnI) with a phospho-mimetic cTnI T144D substitution (T144D) in the sarcomere16. The initial analysis shows the PO-induced reduction in contractile function (Table 1, upper panel) returned toward sham levels in PO myocytes 4 days after gene transfer of cTnIT144D (Table 1; lower panel).
This platform also can be used to measure Ca2+ transients, along with shortening in isolated myocytes. Shortening and Ca2+ were not recorded after PO because PO reduces adherence of rat myocytes to laminin13, and a comparable model previously showed altered Ca2+ handling develops at a similar time point17. Instead, representative experiments were performed in Fura-2AM-loaded myocytes isolated from 2-3-month-old rats. A representative recording and signal-averaged traces are shown in Figure 3, along with the data analysis in Table 2. For this set of experiments, myocytes isolated from adult rats were studied 4 days after adenoviral-mediated gene transfer (multiplicity of infection = 100) of cTnIT144D or wild-type cTnI. Both sarcomere shortening and Ca2+ transients were measured after loading myocytes with Fura-2AM. Gene transfer of cTnIT144D enhanced peak shortening and elevated diastolic Ca2+ levels compared to cTnI in these initial studies (Table 2). While more extensive analysis is needed, the initial results suggest in vivo replacement with cTnIT144D could produce a complex cardiac phenotype due to changes in both contractile function and Ca2+ handling over time.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64055/64055fig01.jpg" /> 
Figure 1: Adult rat cardiac myocytes used for functional studies. (A)&#160;Representative isolated cardiac myocytes from an adult rat (scale bar = 50 &#956;m). Arrow points to a representative myocyte imaged for contractile function analysis. (B)&#160;Representative myocyte with a ROI (pink) located on the side (scale bar = 20 &#956;m). (C)&#160;Representative myocyte with a ROI (upper panel), the sarcomere pattern for this myocyte (lower panel, blue; scale bar = 20 &#956;m), and the sharp peak of the power spectrum (lower panel, red). (D)&#160;Screen capture of 0.01 mm graticule to calibrate the shortening measurements. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64055/64055fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64055/64055fig02.jpg" /> 
Figure 2: Representative recordings from rat myocytes. A raw recording (upper panel) and signal-averaged (lower panel) trace recorded at 0.2 Hz in myocytes from (A) sham- and (B) pressure overload (PO)-treated rats. Myocytes were isolated 18-20 weeks after surgery, with PO produced by supra-renal coarctation13. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64055/64055fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64055/64055fig03.jpg" /> 
Figure 3: Representative recording and analysis of sarcomere length (SL) and Ca2+ transients from Fura-2AM-loaded adult cardiac myocytes. (A)&#160;Raw traces for SL, Ca2+ transient ratio (ratio), and the numerator and denominator traces used to produce the Ca2+ transient ratio. (B)&#160;An example of signal-averaged traces for SL (upper trace) and the Ca2+ transient ratio (lower trace). (C)&#160;Traces are analyzed for sarcomere length (SL) and the Ca2+ transient ratio using the monotonic trace algorithm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64055/64055fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Rat group</td>
			<td>&#160;Sham (n=30)</td>
			<td>PO (n=32)</td>
		</tr>
		<tr>
			<td>Resting sarcomere length (mm; SL)</td>
			<td>&#160; 1.761 + 0.006</td>
			<td>&#160; 1.748 + 0.004</td>
		</tr>
		<tr>
			<td>&#160;peak height (% of baseline)</td>
			<td>&#160; 9.003 + 0.409</td>
			<td>&#160; 6.680 + 0.552*</td>
		</tr>
		<tr>
			<td>Peak amplitude (mm)</td>
			<td>0.159 + 0.007</td>
			<td>0.117 + 0.010*</td>
		</tr>
		<tr>
			<td>Shortening rate (mm/s)</td>
			<td>&#160; -4.674 + 0.285</td>
			<td>&#160; -4.143 + 0.335</td>
		</tr>
		<tr>
			<td>Re-lengthening rate (mm/s)</td>
			<td>3.251 + 0.223</td>
			<td>&#160; 2.706 + 0.273</td>
		</tr>
		<tr>
			<td>Time to peak (ms; TTP)</td>
			<td>60 + 2</td>
			<td>59 + 3</td>
		</tr>
		<tr>
			<td>Time to 50% re-lengthening (ms; TTR50%)</td>
			<td>35 + 2</td>
			<td>37 + 3</td>
		</tr>
		<tr>
			<td>Rat group</td>
			<td>Sham + cTnIT144D (n=14)</td>
			<td>PO + cTnIT144D (n=17)</td>
		</tr>
		<tr>
			<td>Resting sarcomere length (mm; SL)</td>
			<td>1.768 + 0.009</td>
			<td>&#160; 1.776 + 0.004</td>
		</tr>
		<tr>
			<td>&#160;peak height (% of baseline)</td>
			<td>9.038 + 1.339</td>
			<td>&#160; 8.414 + 0.960</td>
		</tr>
		<tr>
			<td>Peak amplitude (mm)</td>
			<td>0.160 + 0.023</td>
			<td>0.149 + 0.016</td>
		</tr>
		<tr>
			<td>Shortening rate (mm/s)</td>
			<td>&#160; -5.972 + 0.711</td>
			<td>&#160; -4.173 + 0.726</td>
		</tr>
		<tr>
			<td>Re-lengthening rate (mm/s)</td>
			<td>3.925 + 0.577</td>
			<td>3.055 + 0.403</td>
		</tr>
		<tr>
			<td>Time to peak (ms; TTP)</td>
			<td>&#160;51 + 5</td>
			<td>&#160;59 + 2</td>
		</tr>
		<tr>
			<td>Time to 50% re-lengthening (ms; TTR50%)</td>
			<td>&#160;31 + 3</td>
			<td>32 + 2</td>
		</tr>
	</tbody>
</table>
Table 1: Comparison of contractile function in cardiac myocytes in response to pressure overload (PO) and gene transfer. Myocyte shortening results are from sham and PO rat hearts 18-20 weeks post surgery (upper panel)5,13 and myocytes from sham and PO rats 4 days after cTnIT144D gene transfer (lower panel). Contractile function is measured 4 days after myocyte isolation/gene transfer in all groups. Results are expressed as mean &#177; SEM (n = number of myocytes). Each set of sham and PO data is compared by a Student&#39;s t-test, with *p &#60; 0.05 considered statistically significant. Peak amplitude results for sham and PO myocytes alone were reported earlier in&#160;Ravichandran et al.5.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td colspan="2">Sarcomere length analysis</td>
		</tr>
		<tr>
			<td>Gene transfer group</td>
			<td>cTnI (n=21)</td>
			<td>cTnIT144D (n=16)</td>
		</tr>
		<tr>
			<td>Resting sarcomere length (mm; SL)</td>
			<td>&#160;&#160; 1.81 + 0.01</td>
			<td>&#160;&#160; 1.81 + 0.01</td>
		</tr>
		<tr>
			<td>Peak amplitude (mm)</td>
			<td>&#160;&#160; 0.11 + 0.01</td>
			<td>&#160;&#160; 0.14 + 0.02*</td>
		</tr>
		<tr>
			<td>Shortening rate (mm/s)</td>
			<td>&#160; -4.29 + 0.42</td>
			<td>&#160;-4.67 + 0.77</td>
		</tr>
		<tr>
			<td>Re-lengthening rate (mm/s)</td>
			<td>&#160; 2.92 + 0.43</td>
			<td>&#160;3.71 + 0.64&#160;&#160;&#160;&#160;&#160;</td>
		</tr>
		<tr>
			<td>Time to peak (ms; TTP)</td>
			<td>50 + 4</td>
			<td>84 + 28</td>
		</tr>
		<tr>
			<td>Time to 50% re-lengthening (ms; TTR50%)</td>
			<td>37 + 4</td>
			<td>35 + 6</td>
		</tr>
		<tr>
			<td></td>
			<td colspan="2">Ca2+ transient analysis</td>
		</tr>
		<tr>
			<td>Gene transfer group</td>
			<td>cTnI (n=21)</td>
			<td>cTnIT144D (n=19)</td>
		</tr>
		<tr>
			<td>Resting Ca2+ Ratio</td>
			<td>0.89 + 0.02</td>
			<td>&#160;1.04 + 0.04*</td>
		</tr>
		<tr>
			<td>Peak Ca2+ Ratio</td>
			<td>0.50 + 0.05</td>
			<td>&#160;0.42 + 0.07</td>
		</tr>
		<tr>
			<td>Ca2+ Transient Rate (D/sec)&#160;</td>
			<td>45.24 + 5.85</td>
			<td>&#160;40.53 + 10.95</td>
		</tr>
		<tr>
			<td>Ca2+ Decay Rate (D/s)&#160;</td>
			<td>&#160;-4.91 + 0.99</td>
			<td>&#160;-2.87 + 0.57</td>
		</tr>
		<tr>
			<td>Time to peak Ca2+ (ms; TTP)</td>
			<td>34 + 2</td>
			<td>41 + 3</td>
		</tr>
		<tr>
			<td>Time to 50% Ca2+ Decay(ms; TTD50%)</td>
			<td>101 + 8</td>
			<td>117 + 6</td>
		</tr>
	</tbody>
</table>
Table 2: Contractile function and Ca2+ transients in adult rat myocytes 4 days after cTnIT144D gene transfer. An analysis of contractile function (upper panel) and Ca2+ transients (lower panel) is shown in myocytes after gene transfer of cTnIT144D compared to wild-type cTnI. Myocytes are isolated from 2-3-month-old adult rats, and data are presented as mean &#177; SEM (n = number of myocytes). Statistical comparisons of contractile function (left) and Ca2+ transients (right) are performed using an unpaired Student&#39;s t-test with significance set to *p &#60; 0.05.
Supplemental Figure 1: Components of the pacing system for myocytes plated on laminin-coated CSs. (A)&#160;Custom pacing chamber containing platinum electrodes in each chamber. (B)&#160;Pacing chamber with the first four chambers filled with media. (C)&#160;Pacing chamber attached to banana-jack cables, which are connected to the chamber and (D) the stimulator. (E)&#160;The connection between the stimulator (right) and the 37&#160;&#176;C incubator (left). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64055/SupplementalFigure1rev.zip" target="_blank">Please click here to download this File.</a>
Supplemental Figure 2: Components needed for myocyte contractile function and/or Ca2+ transient measurements. (A) The contractile function platform showing each component, with the numbered items explained in more detail in the Table of Materials. Components in the platform include an anti-vibration table (#1), inverted brightfield microscope (#2,3), CCD camera (#4), CCD controller and xenon power supply (#5), dual emission light source (#6), temperature controller (#7), peristaltic pump (#8), insulated tube holder (#9), coverslip-mounted perfusion chamber (#10), and vacuum system (#11). (B) Additional components include the fluorescence interface (#12), chamber stimulator (#13), and PC computer (#14), which are explained in more detail in the Table of Materials. Close-up views of numbered items in panel A are shown in C-F. (C)&#160;View of the xenon power supply (left) and CCD controller (right). (D) Temperature controller. (E)&#160;Peristaltic pump. (F)&#160;View of the base for the CS perfusion chamber (black arrow), the platinum electrode mount (gray arrow), and the top mount (white arrow) with #0 pan-head screws. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64055/SupplementalFigure2rev.zip" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Analysis of Cardiac Contractile Dysfunction and Ca2+ Transients in Rodent Myocytes at JoVE.com

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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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

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

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

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

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

Intrauterine Telemetry to Measure Mouse Contractile Pressure In Vivo

A complex integration of molecular and electrical signals is needed to transform a quiescent uterus into a contractile organ at the end of pregnancy. Despite the discovery of key regulators of uterine contractility, this process is still not fully understood. Transgenic mice provide an ideal model in which to study parturition. Previously, the only method to study uterine contractility in the mouse was ex vivo isometric tension recordings, which are suboptimal for several reasons. The uterus must be removed from its physiological environment, a limited time course of investigation is possible, and the mice must be sacrificed. The recent development of radiometric telemetry has allowed for longitudinal, real-time measurements of in vivo intrauterine pressure in mice. Here, the implantation of an intrauterine telemeter to measure pressure changes in the mouse uterus from mid-pregnancy until delivery is described. By comparing differences in pressures between wild type and transgenic mice, the physiological impact of a gene of interest can be elucidated. This technique should expedite the development of therapeutics used to treat myometrial disorders during pregnancy, including preterm labor.

Intrauterine Telemetry to Measure Mouse Contractile Pressure In Vivo

This manuscript provides a protocol for implanting telemeters in the mouse for the purpose of measuring intrauterine pressures during pregnancy.

A complex integration of molecular and electrical signals is needed to transform a quiescent uterus into a contractile organ at the end of pregnancy. ...

A Flow Cytometry-based Assay for Measuring Mitochondrial Membrane Potential in Cardiac Myocytes After Hypoxia/Reoxygenation

Timely and efficient reperfusion of the occluded coronary artery is the best strategy for decreasing myocardial infarct size in patients with a ST-segment elevated myocardial infarction. However, reperfusion per se can result in further cardiomyocyte death, a phenomenon known as reperfusion injury. The opening of the mitochondrial permeability transition pore (mPTP), with the decrease of the mitochondrial membrane potential (MMP), or mitochondrial depolarization, is universally recognized as the final step of reperfusion injury and is responsible for mitochondrial and cardiomyocyte death. JC-1 is a lipophilic cationic dye that accumulates in mitochondria depending on the value of MMP. The higher the MMP is, the more JC-1 accumulates in the mitochondria. The increasing amounts of JC-1 in mitochondria can be reflected by a fluorescence emission shift from green (~530 nm) to red (~590 nm). Therefore, the reduction of the red/green fluorescence intensity ratio can indicate the depolarization of mitochondria. Here, we take advantage of JC-1 to measure MMP, or the opening of mPTP in human cardiac myocytes after hypoxia/reoxygenation, detected by flow cytometry.

Here, we present a protocol that uses JC-1 dye to assess the mitochondrial membrane potential of cells after being exposed to hypoxia/reoxygenation with or without a protective agent.

Timely and efficient reperfusion of the occluded coronary artery is the best strategy for decreasing myocardial infarct size in patients with a ST-segment ...

Isolation of Atrial Myocytes from Adult Mice

The electrophysiological properties of atrial myocytes importantly affect overall cardiac function. Alterations in the underlying ionic currents responsible for the action potential can cause pro-arrhythmic substrates that underlie arrhythmias, such as atrial fibrillation, which are highly prevalent in many conditions and disease states. Isolating adult mouse atrial cardiomyocytes for use in patch-clamp experiments has greatly advanced our knowledge and understanding of the cellular electrophysiology in the healthy atrial myocardium and in the setting of atrial pathophysiology. In addition, studies using genetic mouse models have elucidated the role of a vast array of proteins in regulating atrial electrophysiology. Here we provide a detailed protocol for the isolation of cardiomyocytes from the atrial appendages of adult mice using a combination of enzymatic digestion and mechanical dissociation of these tissues. This approach consistently and reliably yields isolated atrial cardiomyocytes that can then be used to characterize cellular electrophysiology by measuring action potentials and ionic currents in patch-clamp experiments under a number of experimental conditions.

This protocol is used to isolate single atrial cardiomyocytes from the adult mouse heart using a chunk digestion approach. This approach is used to isolate right or left atrial myocytes that can be used to characterize atrial myocyte electrophysiology in patch-clamp studies.

The electrophysiological properties of atrial myocytes importantly affect overall cardiac function. Alterations in the underlying ionic currents ...

On-Site Sampling and Extraction of Brain Tumors for Metabolomics and Lipidomics Analysis

Despite the variety of tools available for cancer diagnosis and classification, methods that enable fast and simple characterization of tumors are still in need. In recent years, mass spectrometry has become a method of choice for untargeted profiling of discriminatory compound as potential biomarkers of a disease. Biofluids are generally considered as preferable matrices given their accessibility and easier sample processing while direct tissue profiling provides more selective information about a given cancer. Preparation of tissues for the analysis via traditional methods is much more complex and time-consuming, and, therefore, not suitable for fast on-site analysis. The current work presents a protocol combining sample preparation and extraction of small molecules on-site, immediately after tumor resection. The sampling device, which is of the size of an acupuncture needle, can be inserted directly into the tissue and then transported to the nearby laboratory for instrumental analysis. The results of metabolomics and lipidomics analyses demonstrate the capability of the approach for the establishment of phenotypes of tumors related to the histological origin of the tumor, malignancy, and genetic mutations, as well as for the selection of discriminating compounds or potential biomarkers. The non-destructive nature of the technique permits subsequent performance of routinely used tests e.g., histological tests, on the same samples used for SPME analysis, thus enabling attainment of more comprehensive information to support personalized diagnostics.

The manuscript presents on-site sampling of human brain tumors with solid phase microextraction followed by their biochemical profiling towards the discovery of biomarkers.

Despite the variety of tools available for cancer diagnosis and classification, methods that enable fast and simple characterization of tumors are still ...

In vitro Assessment of Myocardial Protection following Hypothermia-Preconditioning in a Human Cardiac Myocytes Model

Ischemia/reperfusion-derived myocardial dysfunction is a common clinical scenario in patients after cardiac surgery. In particular, the sensitivity of cardiomyocytes to ischemic injury is higher than that of other cell populations. At present, hypothermia affords considerable protection against an expected ischemic insult. However, investigations into complex hypothermia-induced molecular changes remain limited. Therefore, it is essential to identify a culture condition similar to in vivo conditions that can induce damage similar to that observed in the clinical condition in a reproducible manner. To mimic ischemia-like conditions in vitro, the cells in these models were treated by oxygen/glucose deprivation (OGD). In addition, we applied a standard time-temperature protocol used during cardiac surgery. Furthermore, we propose an approach to use a simple but comprehensive method for the quantitative analysis of myocardial injury. Apoptosis and expression levels of apoptosis-associated proteins were assessed by flow cytometry and using an ELISA kit. In this model, we tested a hypothesis regarding the effects of different temperature conditions on cardiomyocyte apoptosis in vitro. The reliability of this model depends on strict temperature control, controllable experimental procedures, and stable experimental results. Additionally, this model can be used to study the molecular mechanism of hypothermic cardioprotection, which may have important implications for the development of complementary therapies for use with hypothermia.

The distinct effects of different degrees of hypothermia on myocardial protection have not been thoroughly evaluated. The goal of the present study was to quantify the levels of cell death following different hypothermia treatments in a human cardiomyocyte-based model, laying the foundation for future in-depth molecular research.

Ischemia/reperfusion-derived myocardial dysfunction is a common clinical scenario in patients after cardiac surgery. In particular, the sensitivity of ...

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Ovarian function progressively declines during aging and in some pathophysiological conditions including karyotype abnormality, autoimmune diseases, chemo- and radiation-therapies, as well as ovarian surgeries. In unmarried women with severe ovarian dysfunction, fertility preservation is important for future pregnancies. Although oocyte cryopreservation is an established method for fertility preservation, these patients could only preserve a limited number of oocytes even after ovarian hyperstimulation, leading to repeated stimulations to ensure sufficient oocytes to guarantee future pregnancy. To solve this issue, we have recently developed a drug-free in vitro activation (IVA) procedure, which enable us to stimulate early stages of ovarian follicles to develop to the preantral follicle stage. These preantral follicles can respond to the unique protocol of gonadotropin stimulation, resulting in increased number of retrieved oocytes per ovarian stimulation for cryopreservation. The drug-free IVA comprised from the surgical approach and ovarian stimulation. We removed a part of cortex from one or both ovaries from patients under laparoscopic surgery. The ovarian cortical tissues were cut into small cubes to disrupt the Hippo signaling pathway and stimulate the development of early stage follicles. These cubes were grafted orthotropically into remaining ovaries as well as beneath the serosa of both Fallopian tubes. We have already published the surgical procedure of the drug-free IVA and the protocol of subsequent ovarian stimulation, but herein we present the details of laboratory methods required for drug-free IVA.

We present details of laboratory procedures for drug-free in vitro activation (IVA) of ovarian follicles for patients with severe ovarian dysfunction. This method could increase the number of retrievable oocytes per ovarian hyperstimulation and benefit fertility preservation for those patients.

Ovarian function progressively declines during aging and in some pathophysiological conditions including karyotype abnormality, autoimmune diseases, ...

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

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

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A Rodent Model of The Ross Operation: Syngeneic Pulmonary Artery Graft Implantation in A Systemic Position

The Ross operation for aortic valve disease has regained new interest due to its outstanding long-term results. Nonetheless, when employed as freestanding root replacement, the possible dilation of the pulmonary autograft and subsequent aortic regurgitation is described. Several animal models have been proposed. However, these are usually limited to ex-vivo models or in-vivo experiments with relatively expensive large animal models. In this study, we sought to establish a rodent model of pulmonary artery graft (PAG) implantation in a systemic position. A total of 39 adult Lewis rats were included. Immediately after euthanasia, the pulmonary root was harvested from a donor animal (n=17). Syngeneic recipient (n=17) and sham-operated (n=5) rats were sedated and ventilated. In the recipient group, the PAG was implanted with an end-to-end anastomosis in infra-renal abdominal aortic position. Sham-operated rats underwent only transection and re-anastomosis of the aorta. Animals were followed with serial ultrasound studies for two months and post-mortem histological analysis. The median PAG diameter in the native position was 3.20 mm (IQR=3.18-3.23). At follow-up, the median diameter of the PAG was 4.03 mm (IQR=3.74-4.13) at 1 week, 4.07 mm (IQR=3.80-4.28) at 1 month, and 4.27 mm (IQR=3.90-4.35) at 2 months (p&lt;0.01). Peak systolic velocity was 220.07 mm/s (IQR=210.43-246.41) at 1 week, 430.88 mm/s (IQR=375.28-495.56) at 1 month, and 373.68 mm/s (IQR=305.78-429.81) at 2 months (p=0.02) and did not differ from the sham-operated group at the end of the experiment (p=0.5). Histological analysis did not show any sign of endothelial thrombosis. This study showed that rodent models may allow for the evaluation of the long-term adaptation of the pulmonary root to a high-pressure system. A systemically placed syngeneic PAG implantation represents a simple and feasible platform for the development and evaluation of novel surgical techniques and drug therapies to further improve the outcomes of the Ross operation.

We demonstrate how to establish a murine model of pulmonary root implantation into the descending aorta to simulate the Ross procedure. This model enables the medium/long-term evaluation of pulmonary autograft remodeling in a systemic position, representing the basis of developing therapeutic strategies to promote its adaptation.

The Ross operation for aortic valve disease has regained new interest due to its outstanding long-term results. Nonetheless, when employed as freestanding ...

Improved Rodent Model of Myocardial Ischemia and Reperfusion Injury

Myocardial ischemia and reperfusion injury (MIRI), induced by coronary heart disease (CHD), causes damage to the cardiomyocytes. Furthermore, evidence suggests that thrombolytic therapy or primary percutaneous coronary intervention (PPCI) does not prevent reperfusion injury. There is still no ideal animal model for MIRI. This study aims to improve the MIRI model in rats to make surgery easier and more feasible. A unique method for establishing MIRI is developed by using a soft tube during a key step of the ischemic period. To explore this method, thirty rats were randomly divided into three groups: sham group (n = 10); experimental model group (n = 10); and existing model group (n = 10). Findings of triphenyltetrazolium chloride staining, electrocardiography, and percent survival are compared to determine the accuracies and survival rates of the operations. Based on the study results, it has been concluded that the improved surgery method is associated with a higher survival rate, elevated ST-T segment, and larger infarct size, which is expected to mimic the pathology of MIRI better.

Myocardial ischemia-reperfusion model of rat heart is improved by using a self-made retractor, polyvinyl chloride tube, and a unique knotting method. Electrocardiogram, triphenyltetrazolium chloride and histological staining, and percent survival analysis results showed that the improved model group has higher success and survival rates than the already existing model group.

Myocardial ischemia and reperfusion injury (MIRI), induced by coronary heart disease (CHD), causes damage to the cardiomyocytes. Furthermore, evidence ...

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

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