This work was supported by German Research Foundation (DFG; Clinician Scientist Program In Vascular Medicine (PRIME), MA 2186/14-1 to P. Tomsits and D. Sch&#252;ttler; VO1568/3-1, IRTG1816, and SFB1002 project A13 to N. Voigt), German Research Foundation under Germany&#8217;s Excellence Strategy (EXC 2067/1- 390729940 to N. Voigt), German Centre for Cardiovascular Research (DZHK; 81X2600255 to S. Clauss and N. Voigt; 81Z0600206 to S. K&#228;&#228;b), the Corona Foundation (S199/10079/2019 to S. Clauss), the ERA-NET on Cardiovascular Diseases (ERA-CVD; 01KL1910 to S. Clauss), the Heinrich-and-Lotte-M&#252;hlfenzl Stiftung (to S. Clauss) and the Else-Kr&#246;ner-Fresenius Foundation (EKFS 2016_A20 to N. Voigt). The funders had no role in manuscript preparation.

All animal procedures were approved by the Lower Saxony Animal Review Board (LAVES, AZ-18/2900) and were conducted in accordance with all institutional, national, and international guidelines for animal welfare.
1. Prearrangements
<ol>
	<li>Prepare 1 L of 10x perfusion buffer (Table 1), 500 mL of 1x perfusion buffer (Table 2), 50 mL of digestion buffer (Table 3), 10 mL of stop buffer (Table 4), 1 L of Tyrode solution (Table 5), 10 mL of each calcium step solution (Tyrode solution with glucose and respective amount of calcium as indicated), 1 L of 4-AP solution (Table 5), 100 mL of pipette solution (Table 6) and 5 mL of pluronic acid&#160;according to the provided recipes. 
	NOTE: Bath solutions (Tyrode and 4-AP solution) can be prepared (without glucose) in advance and stored at +4 &#176;C, glucose is added on the experimental day. Pipette solution can be stored at -20 &#176;C, calcium indicator is added on the experimental day and solution then stored on ice until further use. 10x perfusion buffer can be stored at room temperature, 1x perfusion buffer, digestion buffer and stopping buffer should be freshly prepared on experimental day.</li>
	<li>Turn on the water bath and roller pump.</li>
	<li>Prefill Langendorff apparatus with perfusion buffer; make sure it is air-free.</li>
	<li>Prepare aortic cannula by fixing it under the dissection microscope, connect with a 1 mL syringe filled with perfusion buffer and clear air by rinsing the cannula. 
	NOTE: It is crucial to avoid any air inside the perfusion system, as this will directly affect coronary perfusion and thus digestion effectiveness. A bubble trap might be added to the setup if necessary, to safely avoid any air trapping.</li>
	<li>Prepare Petri dishes with enough perfusion buffer for organ collection and microscopy (buffer should securely cover the entire organ, a few millilitres &#8211; depending on the used Petri dish size &#8211; should be enough).</li>
	<li>Prepare 3 Petri dishes with digestion buffer for tissue dissociation and microscopy, buffer should cover the organ for dissection under the microscope within the respective Petri dish, amount depends on the used Petri dish size. For the dissociation use 3 mL for ventricular tissue and 1.5 mL for atrial tissue.</li>
</ol>
2. Organ harvest
<ol>
	<li>Inject mouse with 0.1 mL of heparin (1,000 U/mL) i.p. using a 1 mL syringe with a 27 G cannula and wait for 5-10 min.</li>
	<li>Place the mouse into an induction chamber along with a small tissue soaked in approximately 500 &#181;L of isoflurane. The animal should not be in contact with the tissue. To avoid that, one can use a plastic biopsy-embedding cassette to cover the tissue. Once the animal is fully anesthetized, check for toe pinch reflex and as soon as it is not present anymore, quickly euthanize the mouse by cervical dislocation.</li>
	<li>Place the mouse on a platform on its back (e.g., on Styrofoam covered with a paper towel) and fix the paws down with cannulas to hold it in place.</li>
	<li>Remove fur and skin covering the chest and part of the abdomen with a clear cut from jugulum towards symphysis and open the abdomen right under the xiphoid without injuring any organ structure using scissors. Lift the sternum with surgical forceps and cut the diaphragm with scissors along the edge of the ribs, then cut the ribs in medial axillary line and remove the rib cage to expose the heart.</li>
	<li>Carefully remove the pericardium using blunt forceps and quickly remove the heart by lifting it with blunt forceps from below and by cutting the large vessels with one single cut using scissors.</li>
	<li>Put the heart into room temperature perfusion buffer and cannulate the aorta with a blunt end needle under the microscope as quickly as possible. 
	NOTE: Remove any lung tissue and fatty tissue attached to the organ without losing too much time on it. While cannulating, make sure that the end of the needle does not extend through the aortic valve, as this will impair results by preventing buffers from entering the coronary arteries.</li>
	<li>Tie the heart with a piece of suturing silk firmly to the needle and disconnect from syringe. 
	NOTE: The entire procedure from obtaining the heart (the moment when the large vessels are cut) until suturing the aorta to the needle should take as little time as possible. It is recommended not taking longer than 90-180 s from removing the heart until start of perfusion.</li>
</ol>
3. Enzymatic digestion
<ol>
	<li>After aortic cannulation, immediately connect the cannulated heart to the Langendorff apparatus avoiding any air entering the system. 
	NOTE: It can help to have a hanging drop of perfusion buffer at the bottom of the Langendorff apparatus as well as a drop of perfusion buffer sitting on the top of the needle in order to avoid any air entering the system.</li>
	<li>Perfuse the heart with perfusion buffer for 1 min at a temperature of exactly 37 &#176;C and a perfusion rate of exactly 4 mL/min. 
	NOTE: In order to have a temperature of 37 &#176;C at the tip of the perfusion needle, water bath temperature has to be set slightly above at approximately 40 &#176;C. This should be tested regularly by measuring the temperature at the perfusion tip.</li>
	<li>Switch perfusion to digestion buffer and perfuse for exactly 9 min at a temperature of exactly 37 &#176;C and a perfusion rate of exactly 4 mL/min.</li>
	<li>Transfer the digested heart to a Petri dish with enough digestion buffer to keep it fully covered. Then carefully dissect the atria and ventricles under the microscope.</li>
	<li>Transfer the atria into a Petri dish with 1.5 mL of digestion buffer and the ventricles into another Petri-dish with 3 mL of digestion buffer.</li>
	<li>Atrial dissection
	<ol>
		<li>Carefully, but without loss of time, pull the atria apart into tiny pieces using blunt forceps.</li>
		<li>Dissolve the tissue by carefully pipetting up and down using a 1,000 &#181;L pipette tip, which has previously been cut to widen the tip opening.</li>
		<li>Transfer the solution to a 15 mL centrifuge tube and add an equivalent amount of stop buffer (1.5 mL) by carefully pipetting down at the side of the tube to end the reaction.</li>
		<li>Carefully pass all 3 mL of cell/tissue-solutions through a 200 &#181;m nylon mesh to remove remaining larger tissue pieces that have not been fully digested. 
		NOTE: A successful digestion will leave almost no solid chunks.</li>
	</ol>
	</li>
	<li>Ventricular dissection
	<ol>
		<li>Quickly chop the ventricular tissue into tiny pieces using dissection scissors and pipette up and down to dissolve. Use another 1,000 &#181;L pipette tip for pipetting up and down, it may be shortened to widen the opening.</li>
		<li>Transfer cell/tissue-solution into a 15 mL centrifuge tube and add an equivalent amount of stop buffer (3 mL) by carefully pipetting down at the side of the tube to end the reaction.</li>
		<li>Carefully pass all 6 mL of cell/tissue-solution through a 200 &#181;m nylon mesh to remove larger tissue pieces that have not been fully digested. 
		NOTE: A successful digestion will leave almost no solid chunks.</li>
	</ol>
	</li>
	<li>Leave both tubes (atrial and ventricular cell suspension) on the bench at room temperature for 6 min to settle.</li>
	<li>Centrifuge both 15 mL tubes at 5 x g for 2 min.</li>
</ol>
4. Calcium reintroduction
NOTE: The following steps are identical for both atrial and ventricular cells (unless otherwise mentioned) and are performed at room temperature.
<ol>
	<li>Discard the supernatant using a plastic Pasteur pipette and carefully resuspend pellet in 10 mL of calcium free Tyrode solution.</li>
	<li>Leave cells for 8 min for sedimentation.</li>
	<li>Centrifuge at 5 x g for 1 min (only atrial cells, sedimentation is enough for ventricular cells).</li>
	<li>Discard supernatant and carefully resuspend pellet in 10 mL of Tyrode solution with 100 &#181;M calcium concentration.</li>
	<li>Leave cells for 8 min for sedimentation.</li>
	<li>Centrifuge at 5 x g for 1 min (only atrial cells, sedimentation is enough for ventricular cells).</li>
	<li>Discard the supernatant and carefully resuspend pellet in 10 mL of Tyrode solution with 400 &#181;M calcium concentration.</li>
	<li>Leave cells for 8 min for sedimentation.</li>
	<li>Centrifuge at 5 x g for 1 min (only atrial cells, sedimentation is enough for ventricular cells).</li>
	<li>Discard the supernatant and carefully resuspend the pellet in 1 mL (atrial)/ 5 mL (ventricular) of Tyrode solution with 1 mM calcium concentration.</li>
</ol>
5. Loading of myocytes with fluorescent calcium-indicator Fluo-3 AM
NOTE: Due to the light sensitivity of the fluorescent calcium indicator, the following steps should be executed protected from light (e.g., by covering tubes with aluminium foil).
<ol>
	<li>Prepare Fluo-3 AM stock solution by adding 44 &#181;L of 20% pluronic F-127 in anhydrous DMSO to 50 &#181;g of Fluo-3AM (can be stored at -20 &#176;C protected from light).</li>
	<li>Add 10 &#181;L of Fluo-3 AM stock solution to 1 mL of cell suspension and incubate for 10 min at room temperature protected from light.</li>
	<li>Centrifuge at 5 x g for 1 min.</li>
	<li>Discard the supernatant using a plastic Pasteur pipette and resuspend the pellet in a reasonable amount of bath solution to obtain a good working concentration (1-5 mL of bath solution depending on the cell density).</li>
	<li>Leave for 30 min for de-esterification before starting with experiments.</li>
</ol>
6. Simultaneous patch-clamp and epifluorescent Ca2+-transient measurements as previously described16
NOTE: Patch clamp measurements are not the topic of this article, the interested reader may be referred to major publications providing in depth descriptions of this method17,18,19,20,21,22. Nevertheless, for a better overall understanding, a summary on a protocol to measure L-type calcium currents along with current induced calcium transients is provided.
<ol>
	<li>Transfer myocytes into a cell chamber and superfuse with bath solution at 37 &#176;C.</li>
	<li>Block potassium currents by adding 4-aminopyridine and barium chloride to the bath solution as indicated in Table 5.</li>
	<li>Ensure that borosilicate microelectrodes have a tip resistance of 2-5 M&#8486; filled with pipette solution (Table 6).</li>
	<li>Setup measurements to allow for simultaneous recording of both electrical signals and epifluorescence at the same time. Voltage clamp mode is used to measure L-type Ca2+-current with a protocol holding the cell at -80 mV and a 600 ms ramp-pulse to -40 mV to inactivate the fast Na+-current, followed by a 100 ms test-pulse to +10 mV at 0.5 Hz (Figure 2).</li>
	<li>Use excitation at 488 nm, emitted light at &#60;520 nm to detected and convert to [Ca2+]I assuming 
	<img alt="Equation 1" src="/files/ftp_upload/61964/61964equ01.jpg" /> 
	Where kd = dissociation constant of Fluo-3 (864 nM), F = Fluo-3 fluorescence; Fmax = Ca2+-saturated fluorescence obtained at the end of each experiment19.</li>
</ol>

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This article provides an easy and functional way to obtain high quality atrial and ventricular myocytes from the same mouse for patch-clamp studies with simultaneous calcium transient recordings. The quality of the obtained data highly depends on the quality of the cell isolation. As mentioned above, many methods to isolate murine cardiomyocytes have been described previously9,10,11,12. The iso...

Cardiac arrhythmias are common and one of the current major healthcare challenges since they affect millions of people worldwide. Arrhythmias are associated with high morbidity and mortality1,2 and represent the underlying cause for the majority of sudden cardiac deaths3. Up to date treatment options have improved patient survival but are still mainly symptomatic treatments rather than targeting the underlying mechanisms. Thus, these treatments have limited efficacy and may frequently cause severe side effects4,5,6. An improvement of current treatment options requires insight into the underlying pathophysiology, creating the need for suitable models to study. Small animal models - and specifically mouse models - play a crucial role in arrhythmia research as they allow to study key mechanisms of arrhythmogenesis, for example the genetic impact on cellular electrophysiology, ion channel function or calcium handling7,8.
For this purpose, isolated atrial and ventricular cardiomyocytes of sufficient quantity and viability are required. A broad spectrum of different isolation approaches to obtain atrial and ventricular myocytes has been previously described9,10,11,12,13 and some groups have presented data from simultaneous measurements of L-type current and calcium current induced calcium transients from either atrial14 or ventricular15 murine cardiomyocytes. However, to our best knowledge there is no data available of atrial as well as ventricular measurements from one animal. Researchers focus on a broad variety of topics ranging from electrophysiology to proteomics, functional studies as cell contractility or protein interactions, mitochondrial function, or genetics &#8211; all in need of isolated cardiomyocytes. Many of the published protocols thus have not been specifically developed for patch clamp studies, leading to limited yields and insufficient cell quality for patch clamp studies. Thus, simultaneous patch clamp and calcium transient measurements of atrial and ventricular cells isolated from one animal cannot be performed with established protocols.
Isolation of murine &#8211; especially atrial &#8211; myocytes for patch clamp experiments remains challenging. This article provides a simple and fast method for the isolation of high-quality murine atrial and ventricular myocytes via retrograde enzyme based Langendorff perfusion, which subsequently allows simultaneous measurements of both net membrane current and current induced calcium transients from one animal. This article elaborates a protocol for the isolation of atrial and ventricular myocytes derived from wild type mice and mice carrying genetic mutations. This protocol can be used for male and female mice alike. The myocyte isolation, images, and representative results described below were obtained from wild type C57Bl/6 mice at the age of 6 (&#177; 1) months. Nevertheless, this protocol has successfully been used for mice at various ages ranging from 2 to 24 months with different genotypes. Figure 1 shows the isolation setup and a close-up of a cannulated heart during enzyme perfusion.

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			<td>2,3-Butanedione monoxime</td>
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			<td>Anhydrous DMSO</td>
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			<td>D12345</td>
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			<td>Aortic cannula</td>
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			<td>130163-20</td>
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			<td>Sigma-Aldrich</td>
			<td>342920</td>
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			<td>INS650915-4</td>
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			<td>Bovine Calf Serum</td>
			<td>Sigma-Aldrich</td>
			<td>12133C</td>
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			<td>CaCl2</td>
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			<td>Circulating heated water bath</td>
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			<td>ME</td>
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			<td>Collagenase Type II</td>
			<td>Worthington</td>
			<td>LS994177</td>
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			<td>disscetion scissors</td>
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			<td>INS600124</td>
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			<td>DL-aspartat K+-salt</td>
			<td>Sigma-Aldrich</td>
			<td>A2025</td>
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			<td>EGTA</td>
			<td>Sigma-Aldrich</td>
			<td>E4378</td>
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			<td>Fluo-3</td>
			<td>Invitrogen</td>
			<td>F3715</td>
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			<td>Fluo-3 AM</td>
			<td>Invitrogen</td>
			<td>F1242</td>
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			<td>Glucose</td>
			<td>Sigma-Aldrich</td>
			<td>G8270</td>
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			<td>Guanosine 5&#8242;-triphosphate tris salt</td>
			<td>Sigma-Aldrich</td>
			<td>G9002</td>
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			<td>Heating coil</td>
			<td>Radnoti</td>
			<td>158821</td>
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			<td>Heparin</td>
			<td>Ratiopharm</td>
			<td>25.000 IE/5ml</td>
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			<td>HEPES</td>
			<td>Sigma-Aldrich</td>
			<td>H9136</td>
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			<td>induction chamber</td>
			<td>CWE incorporated</td>
			<td>13-40020</td>
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			<td>Isoflurane</td>
			<td>Cp-pharma</td>
			<td>1214</td>
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			<td>Jacketed heart chamber</td>
			<td>Radnoti</td>
			<td>130160</td>
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			<td>KCl</td>
			<td>Merck</td>
			<td>1049360250</td>
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			<td>KH2PO4</td>
			<td>Sigma-Aldrich</td>
			<td>P5655</td>
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			<td>MgCl x 6H2O</td>
			<td>Sigma-Aldrich</td>
			<td>M0250</td>
			<td></td>
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			<td>MgSO4 x 7H2O</td>
			<td>Sigma-Aldrich</td>
			<td>M9397</td>
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			<td>Na2ATP</td>
			<td>Sigma-Aldrich</td>
			<td>A2383</td>
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			<td>Na2HPO4 x 2H2O</td>
			<td>Sigma-Aldrich</td>
			<td>S5136</td>
			<td></td>
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			<td>NaCl</td>
			<td>Sigma-Aldrich</td>
			<td>S3014</td>
			<td></td>
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			<td>NaHCO3</td>
			<td>Sigma-Aldrich</td>
			<td>S5761</td>
			<td></td>
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			<td>Nylon mesh (200 &#181;m)</td>
			<td>VWR-Germany</td>
			<td>510-9527</td>
			<td></td>
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			<td>pasteur pipette</td>
			<td>Sigma Aldrich</td>
			<td>Z331759</td>
			<td></td>
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			<td>petri-dishes</td>
			<td>Thermo Fisher</td>
			<td>150318</td>
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			<td>Pluronic Acid F-127</td>
			<td>Sigma-Aldrich</td>
			<td>P2443</td>
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			<td>Sigma-Aldrich</td>
			<td>P8761</td>
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			<td>Ismatec</td>
			<td>ISM597D</td>
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			<td>INS650908-4</td>
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			<td>surgical scissors</td>
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			<td>INS700540</td>
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			<td>Merck</td>
			<td>Z683531-100EA</td>
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			<td>Taurin</td>
			<td>Sigma-Aldrich</td>
			<td>86330</td>
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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 yield is determined after calcium reintroduction by pipetting 10 &#181;L of cell suspension onto a microscope slide. More than 100 viable, rod-shaped, non-contracting cells/10 &#181;L for atrial cell isolation and more than 1,000 viable, rod-shaped, non-contracting cells/10 &#181;L for ventricular cell isolation are considered as sufficient yield and are commonly obtained using this protocol. Atrial cells obtained with this protocol showed a variety of different cell types containing cells of the cardiac conduc...

Watch this Scientific Journal Video about Isolation of High Quality Murine Atrial and Ventricular Myocytes for Simultaneous Measurements of Ca2+ Transients and L-Type Calcium Current at JoVE.com

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.

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 ...

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2.9K Views.  Ludwig-Maximilians University Munich (LMU). 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.

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

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 ...

Isolation and Kv Channel Recordings in Murine Atrial and Ventricular Cardiomyocytes

KCNE genes encode for a small family of Kv channel ancillary subunits that form heteromeric complexes with Kv channel alpha subunits to modify their functional properties. Mutations in KCNE genes have been found in patients with cardiac arrhythmias such as the long QT syndrome and/or atrial fibrillation. However, the precise molecular pathophysiology that leads to these diseases remains elusive. In previous studies the electrophysiological properties of the disease causing mutations in these genes have mostly been studied in heterologous expression systems and we cannot be sure if the reported effects can directly be translated into native cardiomyocytes. In our laboratory we therefore use a different approach. We directly study the effects of KCNE gene deletion in isolated cardiomyocytes from knockout mice by cellular electrophysiology - a unique technique that we describe in this issue of the Journal of Visualized Experiments. The hearts from genetically engineered KCNE mice are rapidly excised and mounted onto a Langendorff apparatus by aortic cannulation. Free Ca2+ in the myocardium is bound by EGTA, and dissociation of cardiac myocytes is then achieved by retrograde perfusion of the coronary arteries with a specialized low Ca2+ buffer containing collagenase. Atria, free right ventricular wall and the left ventricle can then be separated by microsurgical techniques. Calcium is then slowly added back to isolated cardiomyocytes in a multiple step comprising washing procedure. Atrial and ventricular cardiomyocytes of healthy appearance with no spontaneous contractions are then immediately subjected to electrophysiological analyses by patch clamp technique or other biochemical analyses within the first 6 hours following isolation.

Kv channel dysfunction is associated with cardiac arrhythmias. In order to study the molecular mechanisms that lead to such arrhythmias we utilize a systematic protocol for isolation of atrial and ventricular cardiomyocytes from Kv channel ancillary subunit knockout mice. Isolated cardiomyocytes can then immediately be used for cellular electrophysiological studies, biochemical or immunofluorescence (IF) assays.

KCNE genes encode for a small family of Kv channel ancillary subunits that form heteromeric complexes with Kv channel alpha subunits to modify their ...

Isolation of Human Atrial Myocytes for Simultaneous Measurements of Ca2+ Transients and Membrane Currents

The study of electrophysiological properties of cardiac ion channels with the patch-clamp technique and the exploration of cardiac cellular Ca2+ handling abnormalities requires isolated cardiomyocytes. In addition, the possibility to investigate myocytes from patients using these techniques is an invaluable requirement to elucidate the molecular basis of cardiac diseases such as atrial fibrillation (AF).1 Here we describe a method for isolation of human atrial myocytes which are suitable for both patch-clamp studies and simultaneous measurements of intracellular Ca2+ concentrations. First, right atrial appendages obtained from patients undergoing open heart surgery are chopped into small tissue chunks ("chunk method") and washed in Ca2+-free solution. Then the tissue chunks are digested in collagenase and protease containing solutions with 20 &mu;M Ca2+. Thereafter, the isolated myocytes are harvested by filtration and centrifugation of the tissue suspension. Finally, the Ca2+ concentration in the cell storage solution is adjusted stepwise to 0.2 mM. We briefly discuss the meaning of Ca2+ and Ca2+ buffering during the isolation process and also provide representative recordings of action potentials and membrane currents, both together with simultaneous Ca2+ transient measurements, performed in these isolated myocytes.

Isolation of Human Atrial Myocytes for Simultaneous Measurements of Ca2+ Transients and Membrane Currents

We describe the isolation of human atrial myocytes which can be used for intracellular Ca2+ measurements in combination with electrophysiological patch-clamp studies.

The study of  electrophysiological properties of cardiac ion channels with the patch-clamp  technique and the exploration of cardiac cellular Ca2+ ...

Cytosolic Calcium Measurements in Renal Epithelial Cells by Flow Cytometry

A variety of cellular processes, both physiological and pathophysiological, require or are governed by calcium, including exocytosis, mitochondrial function, cell death, cell metabolism and cell migration to name but a few. Cytosolic calcium is normally maintained at low nanomolar concentrations; rather it is found in high micromolar to millimolar concentrations in the endoplasmic reticulum, mitochondrial matrix and the extracellular compartment. Upon stimulation, a transient increase in cytosolic calcium serves to signal downstream events. Detecting changes in cytosolic calcium is normally performed using a live cell imaging set up with calcium binding dyes that exhibit either an increase in fluorescence intensity or a shift in the emission wavelength upon calcium binding. However, a live cell imaging set up is not freely accessible to all researchers. Alternative detection methods have been optimized for immunological cells with flow cytometry and for non-immunological adherent cells with a fluorescence microplate reader. Here, we describe an optimized, simple method for detecting changes in epithelial cells with flow cytometry using a single wavelength calcium binding dye. Adherent renal proximal tubule epithelial cells, which are normally difficult to load with dyes, were loaded with a fluorescent cell permeable calcium binding dye in the presence of probenecid, brought into suspension and calcium signals were monitored before and after addition of thapsigargin, tunicamycin and ionomycin.

Calcium is involved in numerous physiological and pathophysiological signaling pathways. Live cell imaging requires specialized equipment and can be time consuming. A quick, simple method using a flow cytometer to determine relative changes in cytosolic calcium in adherent epithelial cells brought into suspension was optimized.

A variety of cellular processes, both physiological and pathophysiological, require or are governed by calcium, including exocytosis, mitochondrial ...

Voltage and Calcium Dual Channel Optical Mapping of Cultured HL-1 Atrial Myocyte Monolayer

Optical mapping has proven to be a valuable technique to detect cardiac electrical activity on both intact ex vivo hearts and in cultured myocyte monolayers. HL-1 cells have been widely used as a 2-Dimensional cellular model for studying diverse aspects of cardiac physiology. However, it has been a great challenge to optically map calcium (Ca) transients and action potentials simultaneously from the same field of view in a cultured HL-1 atrial cell monolayer. This is because special handling and care is required to prepare healthy cells that can be electrically captured and optically mapped. Therefore, we have developed an optimal working protocol for dual channel optical mapping. In this manuscript, we have described in detail how to perform the dual channel optical mapping experiment. This protocol is a useful tool to enhance the understanding of action potential propagation and Ca kinetics in arrhythmia development.

This article describes the technique used to perform dual channel optical mapping in cultured HL-1 atrial cell monolayers. This unique protocol allows the simultaneous visualization of both calcium (Ca) and voltage (Vm) activity in the same area for the detailed detection and analysis of electrophysiological properties of culture monolayers.

Optical mapping has proven to be a valuable technique to detect cardiac electrical activity on both intact ex vivo hearts and in cultured myocyte ...

Isolation of Mitochondria from Minimal Quantities of Mouse Skeletal Muscle for High Throughput Microplate Respiratory Measurements

Dysfunctional skeletal muscle mitochondria play a role in altered metabolism observed with aging, obesity and Type II diabetes. Mitochondrial respirometric assays from isolated mitochondrial preparations allow for the assessment of mitochondrial function, as well as determination of the mechanism(s) of action of drugs and proteins that modulate metabolism. Current isolation procedures often require large quantities of tissue to yield high quality mitochondria necessary for respirometric assays. The methods presented herein describe how high quality purified mitochondria (~ 450 &#181;g) can be isolated from minimal quantities (~75-100 mg) of mouse skeletal muscle for use in high throughput respiratory measurements. We determined that our isolation method yields 92.5&#177; 2.0% intact mitochondria by measuring citrate synthase activity spectrophotometrically. In addition, Western blot analysis in isolated mitochondria resulted in the faint expression of the cytosolic protein, GAPDH, and the robust expression of the mitochondrial protein, COXIV. The absence of a prominent GAPDH band in the isolated mitochondria is indicative of little contamination from non-mitochondrial sources during the isolation procedure. Most importantly, the measurement of O2 consumption rate with micro-plate based technology and determining the respiratory control ratio (RCR) for coupled respirometric assays shows highly coupled (RCR; &#62;6 for all assays) and functional mitochondria. In conclusion, the addition of a separate mincing step and significantly reducing motor driven homogenization speed of a previously reported method has allowed the isolation of high quality and purified mitochondria from smaller quantities of mouse skeletal muscle that results in highly coupled mitochondria that respire with high function during microplate based respirometirc assays.

Here, we present a modification of a previously reported method that allows for the isolation of high quality and purified mitochondria from smaller quantities of mouse skeletal muscle. This procedure results in highly coupled mitochondria that respire with high function during microplate based respirometirc assays.

Dysfunctional skeletal muscle mitochondria play a role in altered metabolism observed with aging, obesity and Type II diabetes. Mitochondrial ...

Development and Characterization of In Vitro Microvessel Network and Quantitative Measurements of Endothelial [Ca2+]i and Nitric Oxide Production

Endothelial cells (ECs) lining the blood vessel walls in vivo are constantly exposed to flow, but cultured ECs are often grown under static conditions and exhibit a pro-inflammatory phenotype. Although the development of microfluidic devices has been embraced by engineers over two decades, their biological applications remain limited. A more physiologically relevant in vitro microvessel model validated by biological applications is important to advance the field and bridge the gaps between in vivo and in vitro studies. Here, we present detailed procedures for the development of cultured microvessel network using a microfluidic device with a long-term perfusion capability. We also demonstrate its applications for quantitative measurements of agonist-induced changes in EC [Ca2+]i and nitric oxide (NO) production in real time using confocal and conventional fluorescence microscopy. The formed microvessel network with continuous perfusion showed well-developed junctions between ECs. VE-cadherin distribution was closer to that observed in intact microvessels than statically cultured EC monolayers. ATP-induced transient increases in EC [Ca2+]i and NO production were quantitatively measured at individual cell levels, which validated the functionality of the cultured microvessels. This microfluidic device allows ECs to grow under a well-controlled, physiologically relevant flow, which makes the cell culture environment closer to in vivo than that in the conventional, static 2D cultures. The microchannel network design is highly versatile, and the fabrication process is simple and repeatable. The device can be easily integrated to the confocal or conventional microscopic system enabling high resolution imaging. Most importantly, because the cultured microvessel network can be formed by primary human ECs, this approach will serve as a useful tool to investigate how pathologically altered blood components from patient samples affect human ECs and provide insight into clinical issues. It also can be developed as a platform for drug screening.

Development and Characterization of In Vitro Microvessel Network and Quantitative Measurements of Endothelial [Ca2+]i and Nitric Oxide Production

Primary human umbilical vein endothelial cells (HUVECs) were grown to confluence within a microfluidic network device. The endothelial cell junction and F-actin distributions were illustrated and the changes in intracellular calcium concentration and nitric oxide production in response to adenosine triphosphate (ATP) were quantified in real-time at individual cell levels.

Endothelial cells (ECs) lining the blood vessel walls in vivo are constantly exposed to flow, but cultured ECs are often grown under static conditions and ...

Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ concentration. To do this, mitochondria utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester Ca2+. The influx of Ca2+ into the mitochondria stimulates three rate-limiting dehydrogenases of the citric acid cycle, increasing electron transfer through the oxidative phosphorylation (OXPHOS) complexes. This stimulation maintains &#916;&#936;m, which is temporarily dissipated as the positive calcium ions cross the mitochondrial inner membrane into the mitochondrial matrix.
We describe here a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using confocal microscopy. By permeabilizing the cells, mitochondrial Ca2+ can be measured using the fluorescent Ca2+ indicator Fluo-4, AM, with measurement of &#916;&#936;m using the fluorescent dye tetramethylrhodamine, methyl ester, perchlorate (TMRM). The benefit of this system is that there is very little spectral overlap between the fluorescent dyes, allowing accurate measurement of mitochondrial Ca2+ and &#916;&#936;m simultaneously. Using the sequential addition of Ca2+ aliquots, mitochondrial Ca2+ uptake can be monitored, and the concentration at which Ca2+ induces mitochondrial membrane permeability transition and the loss of &#916;&#936;m determined.

Mitochondria can utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester calcium (Ca2+), allowing them to shape cytosolic Ca2+ signaling within the cell. We describe a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using fluorescent dyes and confocal microscopy.

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ ...

Simultaneous Isolation of High Quality Cardiomyocytes, Endothelial Cells, and Fibroblasts from an Adult Rat Heart

The rat is an important animal model used in cardiovascular research, and rat cardiac cells are used routinely for in vitro analysis of the molecular mechanisms of cardiovascular disease progression such as cardiac hypertrophy, fibrosis, and atherosclerosis. Although several attempts with variable success have been made to develop immortalized cell lines from the cardiovascular system to understand these cellular mechanisms, primary cells offer a more natural and close to in vivo environment for such studies. Therefore, different laboratories working on a particular cell type have developed protocols to isolate individual types of rat cardiac cells of interest. A protocol that allows the isolation of more than one cell type, however, is missing. Here an optimized protocol is described that allows the isolation of high-quality major cardiac cell types (cardiomyocytes, endothelial cells, and fibroblasts) from a single preparation and enables their use for cellular analyses. This permits the most efficient use of available resources, which may save time and reduce research costs.

Several protocols have been developed and described for the isolation of different cardiac cell types from a rat heart. Here, an optimized protocol is described that allows the isolation of high-quality major cardiac cell types (cardiomyocytes, endothelial cells, and fibroblasts) from a single preparation, reducing the experimental costs.

The rat is an important animal model used in cardiovascular research, and rat cardiac cells are used routinely for in vitro analysis of the molecular ...

Monitoring ER/SR Calcium Release with the Targeted Ca2+ Sensor CatchER+

Intracellular calcium (Ca2+) transients evoked by extracellular stimuli initiate a multitude of biological processes in living organisms. At the center of intracellular calcium release are the major intracellular calcium storage organelles, the endoplasmic reticulum (ER) and the more specialized sarcoplasmic reticulum (SR) in muscle cells. The dynamic release of calcium from these organelles is mediated by the ryanodine receptor (RyR) and the inositol 1,4,5-triphosphate receptor (IP3R) with refilling occurring through the sarco/endoplasmic reticulum calcium ATPase (SERCA) pump. A genetically encoded calcium sensor (GECI) called CatchER was created to monitor the rapid calcium release from the ER/SR. Here, the detailed protocols for the transfection and expression of the improved, ER/SR-targeted GECI CatchER+ in HEK293 and C2C12 cells and its application in monitoring IP3R, RyR, and SERCA pump-mediated calcium transients in HEK293 cells using fluorescence microscopy is outlined. The receptor agonist or inhibitor of choice is dispersed in the chamber solution and the intensity changes are recorded in real time. With this method, a decrease in ER calcium is seen with RyR activation with 4-chloro-m-cresol (4-cmc), the indirect activation of IP3R with adenosine triphosphate (ATP), and inhibition of the SERCA pump with cyclopiazonic acid (CPA). We also discuss protocols for determining the in situ Kd and quantifying basal [Ca2+] in C2C12 cells. In summary, these protocols, used in conjunction with CatchER+, can elicit receptor mediated calcium release from the ER with future application in studying ER/SR calcium related pathologies.

Monitoring ER/SR Calcium Release with the Targeted Ca2+ Sensor CatchER+

We present protocols for the application of our targeted genetically-encoded calcium indicator (GECI) CatchER+ for monitoring rapid calcium transients in the endoplasmic/sarcoplasmic reticulum (ER/SR) of HEK293 and skeletal muscle C2C12 cells using real-time fluorescence microscopy. A protocol for the in situ Kd measurement and calibration is also discussed.

Intracellular calcium (Ca2+) transients evoked by extracellular stimuli initiate a multitude of biological processes in living organisms. At the center of ...