We thank Dana Morgenstern for careful proofreading of the manuscript.

All methods and procedures described in this protocol have been approved by the Institutional Animal Care and Use Committee (IACUC) of Case Western Reserve University.
1. Preparation of Solutions, Instruments, and Coverslips
NOTE: 1x solutions can be used for up to a month.
<ol>
	<li>Make 10x Krebs-Henseleit buffer HEPES buffer without calcium (KHB-HB) by adding 68.96 g of NaCl, 3.57 g of KCl, 59.58 g of HEPES, 2.18 g of K2HPO4, 3.08 g of MgSO4 and 19.82 g of glucose to 800 mL of double distilled water in a 1,000 mL flask. After contents are fully dissolved, bring up to volume in a 1,000 mL volumetric flask. 
	NOTE: Traditional Krebs Henseleit solution uses sodium bicarbonate as a buffer and the solution in this protocol uses Krebs Henseleit solution with HEPES buffer. Solution is stable for 6 months if sterile filtered.</li>
	<li>Make 10x Tyrode&#39;s solution by adding 86.51 g of NaCl, 0.552 g of NaH2PO4, 2.03 g of MgCl2, 9.91 g of glucose, 4.03 g of KCl, 2.65 g of CaCl2, and 35.76 g of HEPES to 800 mL of double distilled water in a 1000 mL flask. After contents are fully dissolved, bring up to volume in a 1000 mL volumetric flask. 
	NOTE: Solution is stable for 6 months if sterile filtered.</li>
	<li>Make 1x KHB-HB by measuring out 100 mL of the 10x stock and adding to 875 mL of double distilled water in a 1,000 mL flask. Place flask in 37 &#176;C water bath. Once the solution has reached 37 &#176;C, use NaOH to increase the pH to 7.39. After adjusting the pH, bring solution to volume in a 1000 mL volumetric flask. Sterile filter the solution using a vacuum filtration system.</li>
	<li>Make 1x Tyrode&#39;s solution by measuring out 100 mL of the 10x stock and adding to 875 mL of double distilled water in a 1,000 mL flask. Place flask in 37 &#176;C water bath. Once the solution has reached 37 &#176;C, use NaOH to increase the pH to 7.39. After adjusting the pH, bring solution to volume in a 1,000 mL volumetric flask. Sterile filter using a vacuum filtration system.</li>
	<li>Make 1x modified Tyrode&#39;s solution by measuring out 100 mL of the 10x stock and adding to 875 mL of double distilled water in a 1,000 mL flask. Dissolve 3.07 g of L-glutathione reduced in flask. Place flask in 37 &#176;C water bath. Once the solution has reached 37 &#176;C, use NaOH to increase the pH to 7.39. After adjusting the pH, bring solution to volume in a 1,000 mL volumetric flask. Sterile filter the solution using a vacuum filtration system.</li>
	<li>Make 100 mM blebbistatin stock solution by adding 855 &#181;L of dimethyl sulfoxide (DMSO) to 25 mg of powder. Aliquot out in 20 &#181;L increments and store in a -80 &#176;C freezer for up to six months.</li>
	<li>Make stopping buffer by adding 2 g of bovine serum albumin (BSA) and 1 vial of aliquoted blebbistatin stock to 100 mL of 1x KHB-HB and sterile filter the solution using a vacuum filtration system.</li>
	<li>Make plating buffer by adding 5 mL of fetal bovine serum and 1 vial of the aliquoted blebbistatin stock to 95 mL of M199 HEPES. Sterile filter the solution using a vacuum filtration system.</li>
	<li>Make myocyte culture buffer by adding one 1 vial of the aliquoted blebbistatin stock and 4 mls penicillin-streptomycin to 396 mL of M199 (25 mM HEPES). Sterile filter the solution using a vacuum filtration system.</li>
	<li>Autoclave 2 pairs of Dumont tweezers, 2 pairs of Iris curved scissors, 2 hemostats, one pair of plastic surgery forceps, 6 black braided silk 4-0 sutures arranged to be used as a surgical double-throw knot, and four 100 mL beakers.</li>
	<li>Sterilize 22 x 22 mm2 glass coverslips. First, place a single coverslip in each well of a six well plate. Afterwards, with the lid removed, turn on the UV lamp of the biosafety cabinet and expose the coverslips to UV light for 1 h.</li>
	<li>Make working laminin stock solution by first thawing the bottle on ice. Add contents of one bottle to enough cold sterile phosphate-buffered saline (PBS) to reach a final concentration of 0.04 mg/mL. Aliquot out 1.3 &#181;L into autoclaved 1.5 mL centrifuge tubes. Store at -80 &#176;C. 
	NOTE: Each tube has enough laminin for a single six well plate. Avoid multiple freeze thaw cycles.</li>
	<li>Coat sterilized coverslip by first thawing the working laminin solution on ice. Using a P1000 pipette, aspirate 200 &#181;L of laminin. Gently drag the pipette tip along one edge of the coverslip to allow capillary action to pull out a minuscule amount of laminin to facilitate coverslip attachment to the six well plate.</li>
	<li>Then, expel the remaining laminin in the center of the coverslip. In a circular motion, spread the laminin droplet across the coverslip. Place in a 37 &#176;C incubator at least 1 h and up to 24 h before the isolation.</li>
</ol>
2. Preparation of the Langendorff Apparatus
NOTE: The individual components of the Langendorff apparatus used in this protocol are listed in Table of Materials.
<ol>
	<li>Turn on the circulating water bath. Set temperature so that perfusate has a temperature of 37 &#176;C. 
	NOTE: With the solution reservoirs set to a height of 60 cm, the circulating waterbirth needs to be set to 41 &#176;C to have the perfusate be 37 &#176;C. Unlike previously reported protocols, the height of the reservoir does not need to be changed.</li>
	<li>Rinse the Langendorff apparatus with 70% ethanol followed by two rinses with autoclaved double distilled water. After rinsing, fill reservoir with KHB-HB and oxygenate with 100% oxygen.</li>
	<li>Prime the system by allowing oxygenated KHB-HB to first flow into a 100 mL beaker. Once 50 mL of solution has flowed into the beaker, switch the 3-way stop-cock position to stop flow from the KHB-HB reservoir. Pour 50 mL of oxygenated KHB-HB from the beaker into the collagenase reservoir.</li>
	<li>Let the KHB-HB drain from the digestion reservoir until 5 mL remains in collagenase reservoir. While priming collagenase reservoir, switch the 3-way stop cock repeatedly between reservoirs to allow the lines to degas. After the system is primed, remember to use the degassing trap located on top of the heating coil to allow any remaining air to exit the system.</li>
	<li>Make the collagenase solution. For rats combine 100 mg of type II collagenase, 100 mL of oxygenated KHB-HB, and 2 vials of the blebbistain stock. For mice, combine 100 mg of type II collagenase, 40 mL of oxygenated KHB-HB, and 2 vials of the blebbistain stock. Once mixed, the solution should be stable for 1 h. 
	NOTE: Myocyte viability can vary between type II collagenase lots. Take advantage of a collagenase sampling program to test a lot before bulk ordering.</li>
</ol>
3. Myocyte Isolation
<ol>
	<li>Inject the animal with 1,000 U of heparin. Wait 5 min. 
	NOTE: Mice and rats of any age can be used. However, in general the older or more diseased the animal, the lower the myocyte yield.</li>
	<li>Sacrifice the animal by first anesthetizing it with isoflurane using the open-drop method (1 cc of Isoflurane per 500 cc volume) before euthanizing the animal with a pentobarbital mixture (150 mg/kg intraperitoneal).</li>
	<li>Rapidly excise the heart by first grabbing the fur above the xiphoid process. With the iris scissors, make a small incision immediately below the xiphoid process and pull the fur upwards toward the head exposing the skin.</li>
	<li>Grab the xiphoid process and cut the diaphragm exposing the thoracic cavity. Make a trap door incision, pull the sternum back using a hemostat, and use the curved forceps to excise the heart above the ascending aorta and place in cold KHB-HB.</li>
	<li>Cannulate the heart using a stereo microscope and number 5 forceps. Make sure the heart is submerged and the cannula was primed before heart excision to prevent emboli. Confirm proper positioning of the cannula by visualizing the tip of the cannula approximately 1 mm above aortic insertion into the ventricle. 
	NOTE: The faster the cannulation time, the better the myocyte yield.</li>
	<li>Start the flow of KHB-HB by rotating the stopcock on the Langendorff. Connect the cannula to the Langendorff. Perfuse the heart for 5 min. 
	NOTE: Since perfusion is supplied by a gravity-based system, flow through the heart will be a function of coronary artery compliance.</li>
	<li>Switch perfusion from the KHB-HB reservoir to the digestion buffer reservoir. Once the digestion buffer reaches the heart, set a timer (5 min for mouse or 15 min for rat). Make sure to collect the perfusate in a sterile 100 mL beaker. Refill the digestion buffer reservoir as needed with the perfusate until the digestion time has expired.</li>
	<li>After digestion, separate the chambers of the heart with forceps and the iris scissors in a sterile 100 mL beaker. Place each chamber into a separate well of a six well plate. Pour 5 mL of collagenase solution into each well.</li>
	<li>Immediately start mincing the heart tissue using scissors. Tissue chunks should be approximately 1 mm3. Using sterile transfer pipettes, gently triturate the minced heart tissue. The solution should turn cloudy.</li>
	<li>Once the tissue chunks become white and feathery, examine the cells using an inverted microscope. If the number of viable cells is greater than 80%, proceed to strain the cells into a 50 mL conical tube using a 100 &#181;m cell strainer. Use a different tube and strainer for each chamber of the heart.
	<ol>
		<li>If the number of viable cells is less than 80%, check the time it took to cannulate. If the cannulation time is over 5 min, try another heart. If not, assay new collagenase lots through the collagenase sampling program.</li>
	</ol>
	</li>
	<li>Pellet the cells by centrifuging at 215 x g for 2 min. The pellet should be compact and not loose. If the pellet is loose, the preparation contains many dead cells. In a tissue culture hood, resuspend the pellet in 10 mL of stopping buffer.</li>
	<li>Pellet the cells by centrifuging at 215 x g for 2 min. The pellet should be compact and not loose. If the pellet is loose, the preparation contains many dead cells.</li>
	<li>Resuspend the cells in 5 mL of plating buffer. Perform a cell count. Adjust the milliliters of plating buffer to reach a final myocyte concentration of 2 x 104 cells per mL.</li>
	<li>Remove the laminin-coated coverslips from the incubator. Aspirate the laminin droplet.</li>
	<li>Plate 200 &#181;L of myocyte suspension on each coverslip. Place in a 37 &#176;C incubator (21% O2, 5% CO2) for 2 h to allow attachment. After 2 h, aspirate the unattached cells, add 2 mL of culture media, and culture for up to 4 days.</li>
</ol>
4. Fura-2 Dye Loading
<ol>
	<li>Make a 2 mM fura-2 acetoxymethyl ester (fura-2 AM) stock solution by adding 25 &#181;L of DMSO to 50 &#181;g of fura-2 AM powder (1 vial). Aliquot out into 6 &#181;L aliquots. Take 1 aliquot of fura-2 AM and add to 6 mL of plating medium. Vortex to mix.</li>
	<li>Remove 1 six well plate of myocytes from the incubator. Aspirate media. Add 1 mL of fura-2 media mixture to each well. Cover plate with foil, leave the plate at room temperature, and wait 15 min.</li>
	<li>Aspirate fura-2 media mixture and add 1 mL of Tyrode&#39;s solution to each well. Cover with foil. Wait 20 min at room temperature to allow for dye washout before imaging.</li>
</ol>
5. Fluo-4 Dye Loading
<ol>
	<li>Make a 1.82 mM fluo-4 acetoxymethyl ester (fluo-4 AM) stock solution by adding 25 &#181;L of DMSO to 50 &#181;g of fluo-4 AM powder (1 vial). Aliquot out into 8.333 &#181;L aliquots. Take 1 aliquot of fluo-4 AM stock and add to 6 mL of plating medium. Vortex to mix.</li>
	<li>Remove 1 six well plate of myocytes from the incubator. Aspirate media. Add 1 mL of fluo-4 AM media mixture to each well. Cover plate with foil, leave the plate at room temperature, and wait 15 min.</li>
	<li>Aspirate fluo-4 AM media mixture and add 1 mL of Tyrode&#39;s solution to each well. Cover with foil. Wait 20 min at room temperature to allow for dye washout before imaging.</li>
</ol>
6. Membrane Potential Dye Loading
<ol>
	<li>Remove component A and component B from the membrane potential kit. In a 15 mL conical tube, combine 50 &#181;L of component B and 5 &#181;L of component A. Vortex to mix. Add 10 mL of plating media to the 15 mL conical tube containing the voltage dye mixture. Vortex to mix.</li>
	<li>Remove 1 six well plate of myocytes from the incubator. Aspirate the media. Add 800 &#181;L of the membrane potential dye mixture to each well. Cover plate with foil, leave the plate at room temperature, and wait 15 min.</li>
	<li>Aspirate dye media mixture and add 1 mL of modified-Tyrode&#39;s solution to each well. Cover with foil.</li>
</ol>
7. Photometry and Charge Coupled Device Recordings
<ol>
	<li>Turn on the equipment in the following order: microscope, arc lamp, hyperswitch, fluorescence interface system, Myocam power supply, field stimulator, and computer.</li>
	<li>Make sure the excitation/emission filter sets are appropriate for the imaging dye. 
	NOTE: Fura-2 is excited at 340 nm and 380 nm of light. It emits at 510 nm of light. Fluo-4 and the voltage membrane dye are excited at 485 nm of light and emit at 520 nm of light.</li>
	<li>Prime the system by turning on the vacuum, fully opening the hose clamp, and gently plunging each 60 mL syringe being used in the manifold. For calcium recordings use Tyrode&#39;s solution. For voltage recordings use modified Tyrode&#39;s solution.</li>
	<li>Turn heater on and set flow by adjusting the roller clamp on the perfusion tubing. Make recordings at 36 &#177; 1 &#176;C.</li>
	<li>Open the acquisition software. Make sure the parameters are set for the correct imaging dye.</li>
	<li>In the dark, remove the foil from the six well plate and place a coverslip in the pacing chamber. Make sure the stimulator is off during this step. Focus on the myocytes using the 10x objective.</li>
	<li>Once in focus, start pacing by field stimulating at 1 Hz, 0.2 V. Gradually increase the voltage until 1:1 pacing is obtained. Then increase the voltage until 1.5x the threshold is reached. 
	NOTE: Because excitation-contraction coupling is temperature dependent, make sure the cells have been perfused for 15 min before recording. This allows for myocytes to recover from the shock of going from room temperature back to 37 &#176;C as well as loosely attached cells to float away.</li>
	<li>Switch from the 10x objective to the 40x objective. Focus in on a cell that is following a 1:1 pacing. Adjust the plastic shades so only one cell is in the field of view.</li>
	<li>Using the software, place the area of interest box on well-defined sarcomeres. Start the acquisition software to initiate the excitation light. Using the neutral density filters, adjust the intensity setting accordingly to obtain an adequate SNR.</li>
</ol>

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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Can+optical+recordings+of+membrane+potential+be+used+to+screen+for+drug-induced+action+potential+prolongation+in+single+cardiac+myocytes.">Can optical recordings of membrane potential be used to screen for drug-induced action potential prolongation in single cardiac myocytes.</a> Journal of Pharmacological and Toxicological Methods. 54 (2), 173-182 (2006).</li><li>Tian, Q., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical+action+potential+screening+on+adult+ventricular+myocytes+as+an+alternative+QT-screen.">Optical action potential screening on adult ventricular myocytes as an alternative QT-screen.</a> Cellular Physiology and Biochemistry. 27 (3-4), 281-290 (2011).</li><li>Warren, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-precision+recording+of+the+action+potential+in+isolated+cardiomyocytes+using+the+near-infrared+fluorescent+dye+di-4-ANBDQBS.">High-precision recording of the action potential in isolated cardiomyocytes using the near-infrared fluorescent dye di-4-ANBDQBS.</a> The American Journal of Physiology-Heart and Circulatory Physiology. 299 (4), 1271-1281 (2010).</li><li>FluoVolt&#8482; Membrane Potential Kit. ThermoFisher Scientific Available from: <a target="_blank" href="https://assets.thermofisher.com/TFS-Assets/LSG/manuals/MAN0009668_FluoVolt_Membrane_Potential_Kit_UG.pdf">https://assets.thermofisher.com/TFS-Assets/LSG/manuals/MAN0009668_FluoVolt_Membrane_Potential_Kit_UG.pdf</a> (2018)</li><li>Fluorescence SpectraViewer. ThermoFisher Scientific Available from: <a target="_blank" href="https://www.thermofisher.com/us/en/home/life-science/cell-analysis/labeling-chemistry/fluorescence-spectraviewer.html">https://www.thermofisher.com/us/en/home/life-science/cell-analysis/labeling-chemistry/fluorescence-spectraviewer.html</a> (2019)</li><li>Paredes, R. M., Etzler, J. C., Watts, L. T., Zheng, W., Lechleiter, J. 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The authors have nothing to disclose.

Being able to isolate cardiac myocytes is a powerful method that can be used to understand cardiac physiology, pathology, and toxicology. In the above protocol, we described a method that utilizes a constant gravity pressure Langendorff apparatus to obtain single cardiac myocytes. Afterwards, using the fluorescence photometry system, we describe how to simultaneously acquire either calcium and shortening or voltage and shortening traces.
Because of the different kinetics between calcium dyes, care must be taken on which dye to select. For this protocol, both the fura-2 and fluo-4 used were engineered with AM esters necessitating a wash step to allow for intracellular esterases time to cleave the AM group and trap the dye in the cell. While both fura-2 and fluo-4 are considered high affinity calcium dyes, the Kd for fura-2 is 145 nM compared to the 345 nM for fluo-49. Further, fura-2 is ratiometric. Because of this, it can be used to quantify intracellular calcium levels9,12. Fluo-4 on the other hand is a single wave calcium probe. The advantage of using fluo-4 is it produces a brighter fluorescence signal. Regardless of which calcium dye is used, compared to the calcium dye, membrane voltage probes have a lower SNR.
As shown in Figure 4 and Figure 5, voltage traces compared to calcium traces are smaller in amplitude. Using the software&#39;s digital trace filtering, it is possible to increase the SNR and quantify the data (Figure 4 and Figure 7). Once quantified, both calcium transients and optical APDs demonstrate restitution, shortening their duration at faster pacing frequencies (Figure 2, Figure 3, Figure 6, and Figure 7). Shorter APDs during faster pacing cycles are necessary to allow enough time for ventricular filling during diastole. Alterations in this phenomenon is thought to be indicative of an increase in the risk of arrythmias13,14,15,16. While alterations in APD can be caused by disease, they can also be caused by chemicals. As shown in Figure 7, when the predominant murine repolarizing potassium current, Ito, is blocked, the optical APD becomes longer.
Still, as reported previously with voltage sensitive dyes, light intensity and duration can alter the APD2,5,17. This is believed to be the result of the generation of reactive oxidative species (ROS)5. Previously, it has been shown that the addition of antioxidants to the recording solution can prevent voltage sensitive dye cytotoxicity5. As a result, we added the antioxidant L-glutathione (10 mM), to Tyrode&#39;s solution. Shown in Figure 8 is the last 11 s of a 20 s recording obtained at 1 Hz pacing. As indicated by the red arrows, alterations in the APD did not occur until 15 s into the recording; therefore, while the modified Tyrode&#39;s solution did not prevent phototoxicity it delayed it significantly. Using modified Tyrode&#39;s solution, using a low light intensity setting and keeping the duration of the recording to under 5 s, it is possible to avoid any dye induced alterations in APD. This is important because without taking care to avoid phototoxicity, the data could be misinterpreted as causing early or delayed after depolarizations. In addition to limiting the exposure to blue light, there are additional precautions that can be taken to prevent misinterpretation of the data.
The first is to only record from cells that follow one to one pacing and have a resting sarcomere length greater than or equal to 1.75 &#181;m. The 1.75 &#181;m cutoff is taken from the observation by Gordon et al.18 that tension rapidly declines once the sarcomere length is below this amount. Nonetheless, certain pathologies may result in significant alterations in resting sarcomere length. To be sure that the phenotype is real and not an artifact of the isolation, the following trouble shooting approaches should be taken.
If myocytes are consistently not following 1:1 pacing, have sarcomere lengths below 1.75 &#181;m, heavy membrane blebbing, or do not survive the isolation, the first thing to check is the time it took to cannulate the heart. The longer the cannulation time, the lower the yield will be. If a long cannulation time is required, viability can be improved by placing the heart in a cardioplegic solution19. Nonetheless, because the collagenase is an enzyme, the activity and specificity of a specific lot change over time. If the overall yields progressively become worse despite good cannulation times, new lots should be assayed. While our protocol was optimized for 5 s recordings, if longer voltage traces are needed, additional neutral density filters will need to be purchased. The system described in the protocol comes with neutral density filters that reduce the transmitted light by 37%, 50%, 75%, 90%, and 95%.
In summary, we described a methodology that allowed for the isolation of adult murine ventricular myocytes that were used for calcium, voltage, and sarcomere shortening measurements.

In order to study the heart during healthy and pathological states, it is often useful to examine the phenotype at the single cell level. While scientific advances have permitted the robust measurement of single cell calcium dynamics, single cell optical voltage measurements have remained scarce1. Arguably, this is because of the low signal to noise ratio (SNR), phototoxicity, and photobleaching of traditional potentiometric dyes2,3. Nonetheless, isolated myocyte optical action potentials have been obtained2,3,4. Further, with advances in the chemistry and the physics of voltage sensitive dyes, the SNR has improved5. Newer membrane potential probes (Table of Materials) respond to changes in membrane potential in sub-milliseconds and have a fluorogenic response range of approximately 25% per 100 mV. Further, the excitation/emission of the membrane potential kit (e.g., FluoVolt; Table of Materials) used in this protocol works with standard fluorescein isothiocyanate (FITC) or green fluorescent protein (GFP) settings6.
The FITC and GFP excitation/emission spectra overlap with the fluo-4 calcium bound spectra7. Simultaneous acquisition of fluorescence photometry with digital cell geometry measurements traditionally has been used for the simultaneous acquisition of calcium and cellular shortening measurements8. This protocol describes in detail how to isolate murine myocytes and how to record calcium or voltage signals using standard FITC settings. Additionally, it describes how a simple switch in excitation/emission filters on the imaging workstation can be used to obtain calcium and shortening measurements using the ratio metric calcium dye fura-2. Compared to fluo-4, fura-2 has a higher affinity for calcium and is relatively resistant to photobleaching9. Consequently, using a single workstation this protocol allows for a thorough examination of singly myocyte excitation-contraction coupling.

<table><tbody><tr><td>0.25 Liter Water Jacketed Reservoir</td><td>Radnoti, LLC</td><td>120142-025</td><td></td></tr><tr><td>1 liter volumetric flask</td><td>Fisher Scientific</td><td>10-205F</td><td></td></tr><tr><td>100 ml beaker</td><td>Fisher Scientific</td><td>FB-100-100</td><td></td></tr><tr><td>100 ml graduated cylinder</td><td>Fisher Scientific</td><td>08 562 5C</td><td></td></tr><tr><td>1000 ml flask</td><td>Fisher Scientific</td><td>FB-500-1000</td><td></td></tr><tr><td>2-Bar Lab Stand with Stabilizer Bar and 24&quot; Stainless Steel Rods</td><td>Radnoti, LLC</td><td>159951-2</td><td></td></tr><tr><td>4-Aminopyridine</td><td>Sigma-Aldrich</td><td>275875</td><td></td></tr><tr><td>40X Oly UApo/340 Non-Immersion Objective (NA 0.9, WD 0.2mm)</td><td>IonOptix</td><td>MSCP1-40 (b)</td><td></td></tr><tr><td>60-mL syringe, BD Luer-Lok tip</td><td>BD</td><td>309650</td><td></td></tr><tr><td>Aortic Metal Cannulae</td><td>Harvard Apparatus</td><td>73-0112</td><td></td></tr><tr><td>Bovine Serum Albumin</td><td>Fisher Scientific</td><td>BP9703-100</td><td></td></tr><tr><td>C-6 Standard Heating Circulator</td><td>Chemyx</td><td>A30006</td><td></td></tr><tr><td>CaCl2</td><td>Fisher Scientific</td><td>BP510500</td><td></td></tr><tr><td>Cell framing adapter</td><td>IonOptix</td><td>CFA300</td><td></td></tr><tr><td>CellPr0 Vacuum Filtration System, 1 liter, 0.22&amp;micro;m,Cs/12</td><td>Labratory Product Sales, Inc</td><td>V100022</td><td></td></tr><tr><td>CellPro Vacuum Filtration System, 250mL, 0.22&amp;micro;m,Cs/12</td><td>Labratory Product Sales, Inc</td><td>V25022</td><td></td></tr><tr><td>CellPro Vacuum Filtration System, 500mL, 0.22&amp;micro;m,12/CS</td><td>Labratory Product Sales, Inc</td><td>V50022</td><td></td></tr><tr><td>CMC (mTCII) Temp Control w/ inline flow heater</td><td>IonOptix</td><td>TEMPC2</td><td></td></tr><tr><td>Cole-Parmer Large-bore 3-way, male-lock, stopcocks</td><td>Cole-Parmer</td><td>EW-30600-23</td><td></td></tr><tr><td>Cole-Parmer Luer fittings, Large-bore stopcocks, male lock, 4-way</td><td>Cole-Parmer</td><td>EW-30600-12</td><td></td></tr><tr><td>Cole-Parmer Stopcocks with Luer Connections; 1-way; male slip</td><td>Cole-Parmer</td><td>EW-30600-01</td><td></td></tr><tr><td>Collagenase Type II</td><td>Worthington</td><td>LS004177</td><td></td></tr><tr><td>Corning Sterile Cell Strainers</td><td>Fisher Scientific</td><td>07-201-432</td><td></td></tr><tr><td>Dell Optiplex 790 mini-tower, 4G RAM, 250G HD, Windows 7 Pro</td><td>IonOptix</td><td>CPUD7M</td><td></td></tr><tr><td>DMSO</td><td>Fisher Scientific</td><td>50980367</td><td></td></tr><tr><td>Dumont Tweezers Style 5</td><td>Amazon</td><td>B00F70ZDEQ</td><td></td></tr><tr><td>FHD Rapid Change Stimulation Chamber</td><td>IonOptix</td><td>FHDRCC1</td><td></td></tr><tr><td>Fluo-3/4 Optics Package</td><td>IonOptix</td><td>IonOP-Fluo</td><td></td></tr><tr><td>Fluorescence system interface &amp;ndash; (w PCI-I/O card)</td><td>IonOptix</td><td>FSI700</td><td></td></tr><tr><td>Gibco Penicillin-Streptomycin (10,000 U/mL)</td><td>Fisher Scientific</td><td>15-140-122</td><td></td></tr><tr><td>Glucose</td><td>Fisher Scientific</td><td>D16-1</td><td></td></tr><tr><td>Hemostat, Curved 5-1/2&quot;</td><td>Amazon</td><td>B00GGAAPD0</td><td></td></tr><tr><td>HEPES</td><td>Fisher Scientific</td><td>BP310500</td><td></td></tr><tr><td>HyperSwitch dual excitation light source</td><td>IonOptix</td><td>HSW400</td><td></td></tr><tr><td>Inverted Motic Fluorescence Microscope</td><td>IonOptix</td><td>MSCP1-40 (a)</td><td></td></tr><tr><td>IonWizard Core + Analysis</td><td>IonOptix</td><td>IONWIZ</td><td></td></tr><tr><td>Iris Scissors, curved</td><td>Amazon</td><td>B018KRRMY6</td><td></td></tr><tr><td>K2HPO4</td><td>Fisher Scientific</td><td>P288-100</td><td></td></tr><tr><td>KCl</td><td>Fisher Scientific</td><td>BP3661</td><td></td></tr><tr><td>L-Glutathione reduced</td><td>Sigma-Aldrich</td><td>G4251</td><td></td></tr><tr><td>LOOK Silk Spool, Black Braided, 4-0, 100yds</td><td>SouthernAnesthesiaSurgical Inc.</td><td>SP116-EA</td><td></td></tr><tr><td>M199 Media</td><td>Fisher Scientific</td><td>12 340 030</td><td></td></tr><tr><td>MgCl2</td><td>Fisher Scientific</td><td>MP021914215</td><td></td></tr><tr><td>MgSO4</td><td>Fisher Scientific</td><td>BP2131</td><td></td></tr><tr><td>MyoCam-S Digital CCD video system</td><td>IonOptix</td><td>MCS100</td><td></td></tr><tr><td>MyoPacer Field Stimulator</td><td>IonOptix</td><td>MYP100</td><td></td></tr><tr><td>NaCl</td><td>Fisher Scientific</td><td>BP358212</td><td></td></tr><tr><td>NaH2PO4</td><td>Fisher Scientific</td><td>56-754-9250GM</td><td></td></tr><tr><td>Oxygenator Bubbler with Fluid Inlet for 0.25 Liter</td><td>Radnoti, LLC</td><td>140143-025</td><td></td></tr><tr><td>Photomultiplier sub-system</td><td>IonOptix</td><td>PMT400</td><td></td></tr><tr><td>PMT Acquisition add-on</td><td>IonOptix</td><td>PMTACQ</td><td></td></tr><tr><td>Radnoti Heating Coil 5 mL with Degasing Trap</td><td>Radnoti, LLC</td><td>158830</td><td></td></tr><tr><td>Ring Clamp 60 &amp;ndash; 80mm Dia. for 250ml Reservoir</td><td>Radnoti, LLC</td><td>120141-025</td><td></td></tr><tr><td>Ring Clamp for Bubble Trap Compliance Chamber</td><td>Radnoti, LLC</td><td>120149RC</td><td></td></tr><tr><td>Saint-Gobain ACF000010 5/32 in.9/32 in.</td><td>Fisher Scientific</td><td>14-171-214</td><td></td></tr><tr><td>Saint-Gobain ACF000013 3/16 in.3/8 in.</td><td>Fisher Scientific</td><td>14-171-217</td><td></td></tr><tr><td>Saint-Gobain ACF000016 1/4 in.5/16 in.</td><td>Fisher Scientific</td><td>14-171-219</td><td></td></tr><tr><td>Saint-Gobain ACF000025 5/16 in.5/8 in.</td><td>Fisher Scientific</td><td>14-171-226</td><td></td></tr><tr><td>Saint-Gobain ACF00003 1/16 in.3/16 in.</td><td>Fisher Scientific</td><td>14-171-209</td><td></td></tr><tr><td>Saint-Gobain ACF00005 1/16 in.3/32 in.</td><td>Fisher Scientific</td><td>14-171-210</td><td></td></tr><tr><td>Saint-Gobain ACF00009 5/32 in.7/32 in.</td><td>Fisher Scientific</td><td>14-171-213</td><td></td></tr><tr><td>Sarcomere Length Recording add-on</td><td>IonOptix</td><td>SARACQ</td><td></td></tr><tr><td>T/C Adson Tissue Platic Surgery Forceps 4.75&quot;</td><td>Amazon</td><td>B00JDWRBGC</td><td></td></tr><tr><td>VETUS Anti-Static Curved Tip Tweezers</td><td>Amazon</td><td>B07QMZC94J</td><td></td></tr><tr><td>Vistek 3200 Motic Vibration Isolation Platform</td><td>IonOptix</td><td>ISO100</td><td></td></tr></tbody></table>

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.

Figure 1A shows the Langendorff apparatus. The oxygenator is in the KHB-HB reservoir. The collagenase solution is in the middle 60 mL syringe reservoir. The degassing line is connected to the empty 60 mL syringe reservoir. After a successful isolation, most of the cells should be rod shaped and striated. Under a 40x objective, most myocytes should have clear striations visible. Figure 1B,C shows examples of healthy rat myocytes. Once isolated, cells can be cultured up to 4 days while maintaining their morphology and electrical properties.
To measure excitation-contraction coupling, the cells are then placed in a heated pacing chamber. Because myocytes are sensitive to changes in temperature, it is important to allow the coverslip to equilibrate for 15 min in the chamber before recording. For fluorescence recordings, the excitation wavelength is generated by a 75 W xenon-arc bulb. Xenon-arc bulbs produce a light spectrum that mimics natural sunlight. The intensity of the light and the wavelength are controlled by neutral density/emission filers. The excitation light then passes through the objective to the myocyte. The emission wavelength is then collected by a photomultiplier tube. Using the system described here, both the excitation and emission filters need to be changed manually.
Shortening on the other hand is obtained by a charge coupled device sensor. Measuring in real time up to 1,000 times per second, the acquisition software performs an average of the lines within an area of interest to create a well resolved striation pattern. A fast Fourier transform (FFT) is then calculated. The peak within the power spectrum represents the average sarcomere spacing. Changes in the sarcomere spacing during pacing are then plotted and subsequently quantified.
Figure 2 shows calcium and shortening traces recorded from a C57/B6 mouse myocyte loaded with the calcium dye fura-2. The pacing protocol is a modification of pacing protocols described previously10,11. Healthy mouse myocytes should be able to be paced at their resting heart rate 10 Hz. Figure 3 is quantification of ensembled averaged data obtained from a C57/B6 mice and their transgenic (TG) littermates who had a point mutation introduced into a potassium channel. Notice there is no difference between the groups except for the relaxation time at 10 Hz pacing.
Unlike fura-2 which is a dual excitation dye, the voltage dye and fluo-4 are single wavelength excitation dyes whose excitation/emission work with standard FITC excitation and emission spectrum (494/506 nm). Therefore, recordings of calcium and sarcomere shortening or voltage and sarcomere shortening can be obtained using this filter set.
Figure 4A shows a voltage tracing recorded from a C57/B6 mouse myocyte paced at 10 Hz. Compared to calcium signals, single cell voltage tracings are smaller in amplitude and need post-processing to obtain a useable signal. Figure 4B shows an ensembled averaged action potential (AP) made from the APs in Figure 4A. Figure 4C,D shows an ensembled average AP after a low pass Butterworth or a Savitzky-Golay digital filter was applied. Care must be taken when filtering the signal as not to distort the real data. Notice the subtle differences in the shape of the APs in Figure 4B-D.
Figure 5 shows traces recorded from rat myocytes paced at 1 Hz. In addition to the voltage signal being lower than the calcium signal, the contraction kinetics are different as well. This is because calcium dyes buffer calcium while voltage dyes do not.
As with the calcium transient (Figure 3), myocytes demonstrated pacing dependent changes in their optical action potential duration (APD) as well (Figure 6). While the fura-2 traces were ensembled averaged before being quantified, the voltage traces were filtered with a Savitzky-Golay polynomial smoothing filter (width 5, order 2) before being ensembled averaged and quantified.
As quantified in Figure 6 and Figure 7, in addition to demonstrating pacing induced changes in APD, they also demonstrated drug induced prolongation of the AP. At 4 Hz pacing, concentration dependent blockade of the transient outward current (Ito) with 4-aminopyridine resulted in prolongation of the APD.
Finally, care must be taken to avoid cytotoxicity. Figure 8 is the last 11 s of a 20 s recording. Indicated by the red arrows in Figure 8, prolonged exposure of myocytes to blue light leads to triggered activity.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60196/60196fig1a.jpg" /> 
Figure 1: Constant pressure Langendorff apparatus. (A) The Langendorff Apparatus with each component labeled in white lettering. (B) Isolated Sprague-Dawley rat myocytes viewed through a 10x objective. (C) Isolated rat myocytes viewed through a 40x objective. <a href="https://www.jove.com/files/ftp_upload/60196/60196fig1large.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/60196/60196fig2a.jpg" /> 
Figure 2: Representative calcium and sarcomere shortening traces recorded from C57/B6 myoyctes using fura-2. Calcium and sarcomere shortening traces recorded at 1, 2, 4, 10, 0.5 and 0.75 Hz. <a href="https://www.jove.com/files/ftp_upload/60196/60196fig2large.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/60196/60196fig3a.jpg" /> 
Figure 3: Quantification of sarcomere shortening, peak calcium, relaxation time, and reuptake time recorded from a C57/B6 wild type (WT) and transgenic (TG) mice. (A) Sarcomere shortening. (B) Peak calcium. (C) Relaxation time defined as 90% return to baseline of the shortening trace. (D) Reuptake time defined as 90% return to baseline of the calcium trace. <a href="https://www.jove.com/files/ftp_upload/60196/60196fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60196/60196fig4a.jpg" /> 
Figure 4: Optical action potential recorded from a C57/B6 mouse myocyte paced at 10 Hz. (A) 1 second unfiltered trace. (B) Ensembled averaged optical action potential. (C) Ensembled averaged optical action potential after a lowpass Butterworth filter was applied. (D) Ensembled averaged optical action potential after a Savitzky-Golay polynomial smoothing filter was applied. <a href="https://www.jove.com/files/ftp_upload/60196/60196fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60196/60196fig5a.jpg" /> 
Figure 5: Representative calcium, voltage, and sarcomere shortening traces recorded from Sprague-Dawley rat myocytes paced at 1 Hz. (A) Calcium and sarcomere shortening traces recorded at 1 Hz pacing using fluo-4. (B) Voltage and sarcomere shortening traces recorded at 1 Hz pacing using the voltage dye. <a href="https://www.jove.com/files/ftp_upload/60196/60196fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/60196/60196fig6a.jpg" /> 
Figure 6: Optical action potentials recorded from Sprague-Dawley rat myocytes paced at 1, 2, and 4 Hz pacing. (A) Filtered trace recorded at 1 Hz pacing. (B) Filtered trace recorded at 2 Hz pacing. (C) Filtered trace recorded at 4 Hz pacing. (D) Action potential duration 10, measured as 10% return to baseline. (E) Action potential duration 50, measured as 50% return to baseline. (F) Action potential duration 90, measured as 90% return to baseline. <a href="https://www.jove.com/files/ftp_upload/60196/60196fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/60196/60196fig7a.jpg" /> 
Figure 7: The Effects of 4-aminopyridine on Sprague-Dawley rat optical action potentials recorded at 4 Hz pacing. (A) Ensembled averaged trace recorded at 4 Hz pacing with no 4-Aminopyridine in the solution. (B) Ensembled averaged trace recorded at 4 Hz pacing with 1 &#181;M 4-Aminopyridine in the solution. (C) Ensembled averaged trace recorded at 4 Hz pacing with 10 &#181;M 4-Aminopyridine in the solution. (D) Action potential duration 10, measured as 10% return to baseline. (E) Action potential duration 50, measured as 50% return to baseline. (F) Action potential duration 90, measured as 90% return to baseline. <a href="https://www.jove.com/files/ftp_upload/60196/60196fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/60196/60196fig8a.jpg" /> 
Figure 8: Voltage dye induced phototoxicity in Sprague-Dawley rat myocytes after 20 s of continuous light exposure. Red arrows indicate cytotoxic events. <a href="https://www.jove.com/files/ftp_upload/60196/60196fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Optical Imaging of Isolated Murine Ventricular Myocytes at JoVE.com

Optical Imaging of Isolated Murine Ventricular Myocytes

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 ...

optical-imaging-of-isolated-murine-ventricular-myocytes

Research

JoVE Journal

Bioengineering

6.1K Views.  University of Science and Technology of China, Hefei, Anhui, China. 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.

Video: Optical Imaging of Isolated Murine Ventricular Myocytes

Optical Mapping of Action Potentials and Calcium Transients in the Mouse Heart

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. Cardiovascular physiological phenotyping of the mouse heart can be easily done using fluorescence imaging employing various probes for transmembrane potential (Vm), calcium transients (CaT), and other parameters. Excitation-contraction coupling is characterized by action potential and intracellular calcium dynamics; therefore, it is critically important to map both Vm and CaT simultaneously from the same location on the heart1-4. Simultaneous optical mapping from Langendorff perfused mouse hearts has the potential to elucidate mechanisms underlying heart failure, arrhythmias, metabolic disease, and other heart diseases. Visualization of activation, conduction velocity, action potential duration, and other parameters at a myriad of sites cannot be achieved from cellular level investigation but is well solved by optical mapping1,5,6. In this paper we present the instrumentation setup and experimental conditions for simultaneous optical mapping of Vm and CaT in mouse hearts with high spatio-temporal resolution using state-of-the-art CMOS imaging technology. Consistent optical recordings obtained with this method illustrate that simultaneous optical mapping of Langendorff perfused mouse hearts is both feasible and reliable.

This paper details the dissection procedure, instrumental setup, and experimental conditions during optical mapping of transmembrane potential (Vm) and intracellular calcium transient (CaT) in intact isolated Langendorff perfused mouse hearts.

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. ...

Optical Mapping of Langendorff-perfused Rat Hearts

Optical mapping of the cardiac surface with voltage-sensitive fluorescent dyes has become an important tool to investigate electrical excitation in experimental models that range in scale from cell cultures to whole-organs[1, 2]. Using state-of-the-art optical imaging systems, generation and propagation of action potentials during normal cardiac rhythm or throughout initiation and maintenance of arrhythmias can be visualized almost instantly[1]. The latest commercially-available systems can provide information at exceedingly high spatiotemporal resolutions and were based on custom-built equipment initially developed to overcome the obstacles imposed by more conventional electrophysiological methods[1]. Advancements in high-resolution and high-speed complementary metal-oxide-semiconductor (CMOS) cameras and intensely-bright, light-emitting diodes (LEDs) as well as voltage-sensitive dyes, optics, and filters have begun to make electrical signal acquisition practical for cardiovascular cell biologists who are more accustomed to working with microscopes. Although the newest generation of CMOS cameras can acquire 10,000 frames per second on a 16,384 pixel array, depending on the type of sample preparation, long-established fluorescence acquisition technologies such as photodiode arrays, laser scanning systems, and cooled charged-coupled device (CCD) cameras still have some distinct advantages with respect to dynamic range, signal-to-noise ratio, and quantum efficiency[1, 3]. In the present study, Lewis rat hearts were perfused ex vivo with a crystalloid perfusate (Krebs-Henseleit solution) at 37&deg;C on a modified Langendorff apparatus. After a 20 minute stabilization period, the hearts were intermittently perfused with 11 mMol/L 2,3-butanedione monoxime to eliminate contraction-associated motion during image acquisition. For optical mapping, we loaded hearts with the fast-response potentiometric probe di-8-ANEPPS[4] (5 &mu;Mol/L) and briefly illuminated the preparation with 475&plusmn;15 nm excitation light. During a typical 2 second period of illumination, &gt;605 nm light emitted from the cardiac preparation was imaged with a high-speed CMOS camera connected to a horizontal macroscope. For this demonstration, hearts were paced at 300 beats per minute with a coaxial electrode connected to an isolated electrical stimulation unit. Simultaneous bipolar electrographic recordings were acquired and analyzed along with the voltage signals using readily-available software. In this manner, action potentials on the surface of Langendorff-perfused rat hearts can be visualized and registered with electrographic signals.

This article describes a high temporal and spatial resolution technique to optically image action potential movement on the surface of Langendorff-perfused rat hearts using a potentiometric dye (di-8-ANEPPS).

Optical mapping of the cardiac surface with voltage-sensitive fluorescent dyes has become an important tool to investigate electrical excitation in ...

Biology

Isolation and Genetic Manipulation of Adult Cardiac Myocytes for Confocal Imaging

Cardiac myocytes isolated from adult hearts are widely accepted as a model somewhere half way between embryonic and neonatal muscle cells on one side and a working heart on the other. Thus, cardiomyocytes serve as good models for cardiac cellular physiology and pathophysiology, for pharmaceutical investigations as well as for the exploration of transgenic animal models. Here we describe a method of isolating the cells from the heart. Furthermore we show how a genetic manipulation on cardiac myocytes can be performed without breeding a transgenic animal: This is the combination of long term culture (1 week) and adenoviral gene transfer. The latter one is described from the construction of the virus to the transduction of the cells. It can be used for the expression of genetically encoded biosensors (GEBs), fluorescent fusion proteins, but also for protein over expression and down regulation, e.g. using RNAi. Here we provide an example for the expression of a fusion protein staining a subcellular structure (Golgi). Such protein expression can be visualized by confocal imaging of z-stacks for a 3D-reconstruction of subcellular structures. The protocol comprises state-of-the-art in cell culture, molecular biology and biophysics and thus provides an approach for exploring new horizons in cellular cardiology.

Adult cardiac myocytes are primary cells that can be isolated from animal hearts and cultured for several days. Within this culture period adenoviral gene transfer can be used to express genetically encoded biosensors (GEBs) or fluorescent fusion proteins. Both approaches allow cellular investigations by means of confocal microscopy.

Cardiac myocytes isolated from adult hearts are widely accepted as a model somewhere half way between embryonic and neonatal muscle cells on one side and ...

Murine Short Axis Ventricular Heart Slices for Electrophysiological Studies

Murine cardiomyocytes have been extensively used for in vitro studies of cardiac physiology and new therapeutic strategies. However, multicellular preparations of dissociated cardiomyocytes are not representative of the complex in vivo structure of cardiomyocytes, non-myocytes and extracellular matrix, which influences both mechanical and electrophysiological properties of the heart. Here we describe a technique to prepare viable ventricular slices of adult mouse hearts with a preserved in vivo like tissue structure, and demonstrate their suitability for electrophysiological recordings. After excision of the heart, ventricles are separated from the atria, perfused with Ca2+-free solution containing 2,3-butanedione monoxime and embedded in a 4% low-melt agarose block. The block is placed on a microtome with a vibrating blade, and tissue slices with a thickness of 150-400 &#181;m are prepared keeping the vibration frequency of the blade at 60-70 Hz and moving the blade forward as slowly as possible. Thickness of the slices depends on the further application. Slices are stored in ice cold Tyrode&#39;s solution with 0.9 mM Ca2+ and 2,3-butanedione monoxime (BDM) for 30 min. Afterwards, slices are transferred to 37 &#176;C DMEM for 30 min to wash out the BDM. Slices can be used for electrophysiological studies with sharp electrodes or micro electrode arrays, for force measurements to analyze contractile function or to investigate the interaction of transplanted stem cell-derived cardiomyocytes and host tissue. For sharp electrode recordings, a slice is placed into a 3 cm cell culture dish on the heating plate of an inverted microscope. The slice is stimulated with a unipolar electrode, and intracellular action potentials of cardiomyocytes within the slice are recorded with a sharp glass electrode.

Here, we describe the preparation of viable ventricular slices from adult mice and their use for sharp electrode action potential recordings. These multicellular preparations provide a preserved in vivo like tissue structure, which makes them a valuable model for electrophysiological and pharmacological studies in vitro.

Murine cardiomyocytes have been extensively used for in vitro studies of cardiac physiology and new therapeutic strategies. However, multicellular ...

Medicine

Electrophysiological Assessment of Murine Atria with High-Resolution Optical Mapping

Recent genome-wide association studies targeting atrial fibrillation (AF) have indicated a strong association between the genotype and electrophysiological phenotype in the atria. That encourages us to utilize a genetically-engineered mouse model to elucidate the mechanism of AF. However, it is difficult to evaluate the electrophysiological properties in murine atria due to their small size. This protocol describes the electrophysiological evaluation of atria using an optical mapping system with a high temporal and spatial resolution in Langendorff perfused murine hearts. The optical mapping system is assembled with dual high-speed complementary metal oxide semiconductor cameras and high magnification objective lenses, to detect the fluorescence of a voltage-sensitive dye and Ca2+ indicator. To focus on the assessment of murine atria, optical mapping is performed with an area of 2 mm &#215; 2 mm or 10 mm x 10 mm, with a 100 &#215; 100 resolution (20 &#181;m/pixel or 100 &#181;m/pixel) and sampling rate of up to 10 kHz (0.1 ms) at maximum. A 1-French size quadripolar electrode pacing catheter is placed into the right atrium through the superior vena cava avoiding any mechanical damage to the atrium, and pacing stimulation is delivered through the catheter. An electrophysiological study is performed with programmed stimulation including constant pacing, burst pacing, and up to triple extrastimuli pacing. Under a spontaneous or pacing rhythm, the optical mapping recorded the action potential duration, activation map, conduction velocity, and Ca2+ transient individually in the right and left atria. In addition, the programmed stimulation also determines the inducibility of atrial tachyarrhythmias. Precise activation mapping is performed to identify the propagation of the excitation in the atrium during an induced atrial tachyarrhythmia. Optical mapping with a specialized setting enables a thorough electrophysiological evaluation of the atrium in murine pathological models.

This protocol describes the electrophysiological evaluation of murine atria utilizing an optical mapping system with a high temporal and spatial resolution, including dual recordings of the membrane voltage and Ca2+ transient under programmed stimulation through a specialized electrode catheter.

Recent genome-wide association studies targeting atrial fibrillation (AF) have indicated a strong association between the genotype and ...

Modifications of the Langendorff Method for Simultaneous Isolation of Atrial and Ventricular Myocytes from Adult Mice

A single cardiomyocyte is a vital tool in the cellular and subcellular level studies of cardiac biology and diseases as a fundamental unit of contraction and electrical activity. Hence, isolating viable, high-quality cardiomyocytes from the heart is the initial and most crucial experimental step. Comparing the various protocols for isolating the cardiomyocytes of adult mice, the Langendorff retrograde perfusion is the most successful and reproducible method reported in the literature, especially for isolating ventricular myocytes. However, isolating quality atrial myocytes from the perfused heart remains challenging, and few successful isolation reports are available. Solving this complicated problem is extremely important because apart from ventricular disease, atrial disease accounts for a large part of heart diseases. Therefore, further investigations on the cellular level to reveal the mechanisms are warranted. In this paper, a protocol based on the Langendorff retrograde perfusion method is introduced and some modifications in the depth of aorta cannulation and the steps that may affect the digestion process to isolate atrial and ventricular myocytes were simultaneously made. Moreover, the isolated cardiomyocytes are confirmed to be amenable to patch clamp investigation.

This protocol describes modifications of the Langendorff method including the depth of aorta cannulation for simultaneous Isolation of atrial and ventricular myocytes from adult mice.

A single cardiomyocyte is a vital tool in the cellular and subcellular level studies of cardiac biology and diseases as a fundamental unit of contraction ...

Revealing Murine Pulmonary Veins Myocardial Architecture

Microdissection and Immunofluorescence Staining of Myocardial Sleeves in Murine Pulmonary Veins

Pulmonary veins (PVs) are the major source of ectopic beats in atrial arrhythmias and play a crucial role in the development and progression of atrial fibrillation (AF). PVs contain myocardial sleeves (MS) composed of cardiomyocytes. MS are implicated in the initiation and maintenance of AF, as they preserve similarities to the cardiac working myocardium, including the ability to generate ectopic electrical impulses. Rodents are widely used and may represent excellent animal models to study the pulmonary vein myocardium since cardiomyocytes are widely present all over the vessel wall. However, precise microdissection and preparation of murine PVs is challenging due to the small organ size and intricate anatomy.
We demonstrate a microscopy-guided microdissection protocol for isolating the murine left atrium (LA) together with the PVs. Immunofluorescence staining using cardiac Troponin-T (cTNT) and connexin 43 (Cx43) antibodies is performed to visualize the LA and PVs in full length. Imaging at 10x and 40x magnification provides a comprehensive view of the PV structure as well as detailed insights into the myocardial architecture, particularly highlighting the presence of connexin 43 within the MS.

Author Spotlight: Developing a Translational Model for Atrial Fibrillation Research Across Species 

This protocol demonstrates microscopy-guided isolation and immunofluorescence staining of murine pulmonary veins. We prepare tissue samples containing the left atrium, pulmonary veins, and the corresponding lungs and stain them for cardiac Troponin T and Connexin 43.

Pulmonary veins (PVs) are the major source of ectopic beats in atrial arrhythmias and play a crucial role in the development and progression of atrial ...

Isolation of Human Ventricular Cardiomyocytes from Vibratome-Cut Myocardial Slices

The isolation of ventricular cardiac myocytes from animal and human hearts is a fundamental method in cardiac research. Animal cardiomyocytes are commonly isolated by coronary perfusion with digestive enzymes. However, isolating human cardiomyocytes is challenging because human myocardial specimens usually do not allow for coronary perfusion, and alternative isolation protocols result in poor yields of viable cells. In addition, human myocardial specimens are rare and only regularly available at institutions with on-site cardiac surgery. This hampers the translation of findings from animal to human cardiomyocytes. Described here is a reliable protocol that enables efficient isolation of ventricular myocytes from human and animal myocardium. To increase the surface-to-volume ratio while minimizing cell damage, myocardial tissue slices 300 &#181;m thick are generated from myocardial specimens with a vibratome. Tissue slices are then digested with protease and collagenase. Rat myocardium was used to establish the protocol and quantify yields of viable, calcium-tolerant myocytes by flow-cytometric cell counting. Comparison with the commonly used tissue-chunk method showed significantly higher yields of rod-shaped cardiomyocytes (41.5 &#177; 11.9 vs. 7.89 &#177; 3.6%, p &#60; 0.05). The protocol was translated to failing and non-failing human myocardium, where yields were similar as in rat myocardium and, again, markedly higher than with the tissue-chunk method (45.0 &#177; 15.0 vs. 6.87 &#177; 5.23 cells/mg, p &#60; 0.05). Notably, with the protocol presented it is possible to isolate reasonable numbers of viable human cardiomyocytes (9&#8211;200 cells/mg) from minimal amounts of tissue (&#60;50 mg). Thus, the method is applicable to healthy and failing myocardium from both human and animal hearts. Furthermore, it is possible to isolate excitable and contractile myocytes from human tissue specimens stored for up to 36 h in cold cardioplegic solution, rendering the method particularly useful for laboratories at institutions without on-site cardiac surgery.

Presented is a protocol for the isolation of human and animal ventricular cardiomyocytes from vibratome-cut myocardial slices. High yields of calcium-tolerant cells (up to 200 cells/mg) can be obtained from small amounts of tissue (&#60;50 mg). The protocol is applicable to myocardium exposed to cold ischemia for up to 36 h.

The isolation of ventricular cardiac myocytes from animal and human hearts is a fundamental method in cardiac research. Animal cardiomyocytes are commonly ...

An Antegrade Perfusion Method for Cardiomyocyte Isolation from Mice

In basic research using mouse heart, isolating viable individual cardiomyocytes is a crucial technical step to overcome. Traditionally, isolating cardiomyocytes from rabbits, guinea pigs or rats has been performed via retrograde perfusion of the heart with enzymes using a Langendorff apparatus. However, a high degree of skill is required when this method is used with a small mouse heart. An antegrade perfusion method that does not use a Langendorff apparatus was recently reported for the isolation of mouse cardiomyocytes. We herein report a complete protocol for the improved antegrade perfusion of the excised heart to isolate individual heart cells from adult mice (8 - 108 weeks old). Antegrade perfusion is performed by injecting perfusate near the apex of the left ventricle of the excised heart, the aorta of which was clamped, using an infusion pump. All procedures are carried out on a pre-warmed heater mat under a microscope, which allows for the injection and perfusion processes to be monitored. The results suggest that ventricular and atrial myocytes, and fibroblasts can be well isolated from a single adult mouse simultaneously.

We developed a simple method for isolating high quality individual mouse heart cells by the antegrade perfusion technique. This method is Langendorff-free and useful for isolating ventricular and atrial myocytes or interstitial cells, such as cardiac fibroblasts or progenitors.

In basic research using mouse heart, isolating viable individual cardiomyocytes is a crucial technical step to overcome. Traditionally, isolating ...

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 ...