This work was supported by National Institutes of Health Grants R01 HL114940 (Biesiadecki), R01 AG060542 (Ziolo), and T32 HL134616 (Sturgill and Salyer).

All procedures performed in this study were approved by the Institutional Animal Care and Use Committee at the Ohio State University in accordance with NIH guidelines.
1. Solution preparation
NOTE: Please see Table 2 for buffer concentrations.
<ol>
	<li>Pre-isolation (up to 2 weeks in advance of myocyte isolation)
	<ol>
		<li>Perfusion buffer base: Prepare 1 L of perfusion buffer base (NaCl, KCl, NaH2PO4, HEPES, Glucose, and BDM) in 1 L of ultrapure 18.2 M&#937;&#183;cm H2O. Adjust to pH 7.4 prior to 0.22 &#181;m sterile filtration. Store at 4 &#176;C until the day of isolation.</li>
		<li>Clearing buffer: Prepare 1 L of clearing buffer (NaCl, KCl, NaH2PO4, HEPES, Glucose, EDTA, and BDM) in 1 L of ultrapure 18.2 M&#937;&#183;cm H2O. Adjust to pH 7.4 prior to 0.22 &#181;m sterile filtration. Store at 4 &#176;C until the day of isolation.</li>
		<li>100 mM CaCl2 stock solution: Weigh out 1.11 g of CaCl2 and dissolve into 100 mL of ultrapure 18.2 M&#937;&#183;cm H2O. Store at room temperature until the day of isolation.</li>
		<li>Liberase aliquots: Dissolve 5 mg of digestive enzymes in 5 mL of sterile, RNase-free water. Prepare 0.8 mL aliquots and store at -20 &#176;C until the day of isolation.</li>
		<li>Laminate the dishes: Apply 100 &#181;L of 40 &#181;g/mL mouse laminin to sterile, glass-bottom 30 mL Petri dishes. Allow to sit for 30 min and remove excess laminin. Set the laminin on a Petri dish with UV incubation at room temperature for 30 min. Store at 4 &#176;C until the day of isolation (laminated Petri dishes can be used for up to 2 weeks).</li>
		<li>DMEM media: In a biosafety cabinet, add 2.5 mL of FBS, 0.5 mL of Penicillin-Streptomycin (50 U/mL-50 &#181;g), and 1 mL of 500 mM BDM to 46 mL of DMEM. Store at 4 &#176;C until the day of isolation (media can be used for up to 2 weeks).</li>
		<li>M199 media: Prepare 1 L of M199 media (NaHCO3, HEPES, L-glutathione, M199, BDM, and BSA) in 1 L of ultrapure 18.2 M&#937;&#183;cm H2O. Adjust to pH 7.4 prior to sterile filtration (0.22 &#181;m filter). Store at 4 &#176;C until the day of isolation.</li>
		<li>500 mM BDM stock: Add 0.5 g of BDM to 10 mL of ultrapure 18.2 M&#937;&#183;cm H2O. Store at 4 &#176;C until the day of isolation.</li>
		<li>Stabilizing platform creation: Mold playdough around the base of the needle screwed into the Luer stopcock. Mold to the Petri dish that will contain the heart and ensure that the needle is at proper height for ventricle injection (~5 mm above bottom of dish).</li>
	</ol>
	</li>
	<li>The day of isolation
	<ol>
		<li>Perfusion buffer: On the day of the isolation, add 9.5 mg of MgCl2 to 100 mL of perfusion buffer base. Allow to mix completely before removing 30 mL of completed perfusion buffer and place in a clean, wide-mouth, 100 mL glass bottle with a screw cap lid (digestion buffer). 
		&#8203;NOTE: These additions can be made on the day of isolation without additional pH adjustments as their addition negligibly alters the pH.
		<ol>
			<li>Remove 12 mL of perfusion buffer and set to the side (stop buffer). Pour the remaining 58 mL of perfusion buffer into another clean, wide-mouth, 100 mL glass bottle with a screw cap lid. Store the remaining perfusion buffer at 37 &#176;C throughout isolation.</li>
		</ol>
		</li>
		<li>Digestion buffer: Add 7.5 &#181;L of 100 mM CaCl2 to 30 mL of perfusion buffer for a final concentration of 25 &#181;M. Immediately prior to the isolation, add 770 &#181;L of Liberase to the digestion buffer for a final concentration of 26 &#181;g/mL.</li>
		<li>Stop buffer: Dissolve 240 mg of BSA in 12 mL of perfusion buffer. Store at 37 &#176;C throughout isolation.</li>
		<li>Clearing buffer: Prepare a 3 mL syringe of clearing buffer and clear the needle of bubbles. Store on ice until isolation.</li>
	</ol>
	</li>
</ol>
2. Manifold preparation
<ol>
	<li>Clear the temperature-controlled manifold with freshly prepared perfusion buffer, being careful to ensure no bubbles remain. Using a Luer lock connector, screw in a 27 G needle, and clear of bubbles. A cleared manifold is essential for a successful isolation.</li>
</ol>
3. Animal preparation
<ol>
	<li>Inject the mouse intraperitoneally with 100 mg/kg ketamine and 20 mg/kg xylazine by body weight immediately prior to the isolation. This procedure used a male, 4-month-old C57Bl/6 mouse.</li>
	<li>If needed, place the mouse on a heated surgical pad to encourage sedation. Tent the arms of the mouse, and tape the limbs and base of the tail to a blue lab diaper. Ensure that the animal is fully anesthetized&#160;by the withdrawal of&#160;toe-pinch reflex. Apply a sterile drape around the surgical area.</li>
</ol>
4. Cardiomyocyte isolation procedure
<ol>
	<li>Expose the sternum of the mouse and, lateral to the midline, cut proximally through the ribs and to the axilla. Gently cut fully through the diaphragm, ensuring shallow cuts to avoid the heart. Clamp the sternum with a hemostat and fold the ribs backwards to expose the thoracic cavity.</li>
	<li>Gently remove the pericardium from the heart and cut fully through the inferior vena cava, immediately distal to the heart. Use a 3 mL syringe with a 27 G needle to quickly inject 3 mL of ice-cold clearing buffer into the right ventricle of the heart over 1 min.</li>
	<li>Gently hold the heart using tweezers and pull away from the body, exposing as much of the aorta as possible. Using a hemostat, clamp the ascending aorta, being careful not to clamp the atria, and excise the heart from the chest.</li>
	<li>Quickly transfer the clamped heart to the lid of a polypropylene Petri dish with approximately 10 mL of warm perfusion buffer. Place the clamped heart into the supporting platform and use a stabilized 27 G needle attached to the manifold to inject 10 mL of temperature-controlled 37 &#176;C perfusion buffer into the left ventricle over 5 min.</li>
	<li>Transfer the clamped heart and supporting platform to a Petri dish with approximately 5 mL of digestion buffer. Exchange input syringes for a 50 mL syringe with 25 mL of digestion buffer.</li>
	<li>Clear the manifold of any bubbles and remaining perfusion buffer before injection. Carefully replace the needle into the same needle position in the left ventricular apex.</li>
	<li>Use a perfusion pump to inject temperature-controlled 37 &#176;C digestion buffer into the left ventricle for 15 min using a 50 mL syringe. Control the temperature of the solution with a water jacket previously calibrated to eject 37 &#176;C solution at the end of the needle.</li>
	<li>Remove the ventricles from the atria and transfer to a 10 mL beaker. Add 3 mL of digestion buffer to the beaker and, using sharp scissors, cut the ventricles into large chunks. Cover the beaker with aluminum foil and place in a shaking water bath preheated to 37 &#176;C for 5 min.</li>
	<li>Discard the supernatant, being careful to avoid removing any chunks of tissue. Resuspend the tissue chunks in 3 mL of digestion buffer and triturate for approximately 4 min or until a homogenous mixture is achieved.</li>
	<li>Filter the cells through a 70 &#181;m nylon cell strainer into a 50 mL polypropylene tube.</li>
	<li>Return the filtrate to a 14 mL round-bottom polypropylene tube and centrifuge for 1 min at 100 rpm. Discard the supernatant and resuspend the pellet in 3 mL of stop buffer.</li>
	<li>Add 54 &#181;L of 100 mM CaCl2 stock to the resuspended cells for a final CaCl2 concentration of 1.8 mM. This step can also be performed stepwise to increase cell yield.</li>
	<li>Allow live cells to settle by gravity for 10 min before removing the supernatant containing dead cells. Resuspend the pellet in storage solution (perfusion solution with 200 &#181;M CaCl2). These cells can now be used for functional experiments (i.e., calcium imaging/contraction, patch clamp, etc.), culture, etc. 
	&#8203;NOTE: The cells may also be resuspended in lab- or experiment-dependent solutions.</li>
</ol>
5. Cell culture
<ol>
	<li>Count the cells using a hemocytometer or by a field view count using a gridded cover slip. As myocytes are very large, counting using a hemocytometer is not always accurate. This is used to get an idea of approximately how many cells were obtained from the isolation.</li>
	<li>Dilute the cells to ~25,000 cells/mL with DMEM media. Add 1 mL of cell solution to each well.</li>
	<li>Incubate at 37 &#176;C, 95% O2, 5% CO2 for 2 h.</li>
	<li>Gently aspirate DMEM media from each well and add 2 mL of M199 media.</li>
	<li>Incubate at 37 &#176;C, 95% O2, 5% CO2 for up to 24 h.</li>
</ol>

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</ol>

No conflicts of interest to disclose.

The principal advantage of our Langendorff-free cardiomyocyte isolation technique is that it limits hypoxia and ischemic time by not requiring cannulation to a Langendorff apparatus. Alternatively to classical Langendorff techniques that take several minutes to remove, clean, and hang the heart, often resulting in ischemic damage to the myocyte, our method includes an in vivo clearing of blood via an ice-cold clearing solution. The ice-cold clearing buffer contains ethylenediaminetetraacetic acid (EDTA)...

For decades, an essential idea in cardiac biology literature is the molecular mechanism of action. The mechanism of action must be established in order to publish reliable studies. A well-established strategy to determine molecular mechanism is isolated cardiomyocyte studies, which require high-quality cardiomyocytes for attaining trustable data. Cellular and molecular experiments performed on cardiomyocytes to determine mechanism of action are the gold standard for investigating contraction1, electrophysiology2, calcium (Ca2+) cycling3, myofilament Ca2+ sensitivity4, cytoskeleton5, metabolism6, effects of hormones7, signaling molecules8, drug studies9, etc. The mouse has become the species of choice for most cardiac biology experiments due to the ease of genetic manipulation, its small size, its relatively short lifespan, low cost, etc10. However, the reliable isolation of high-quality mouse cardiomyocytes is not trivial with current techniques.
Labs have been isolating cardiomyocytes for almost 70 years11. Virtually all techniques to isolate cardiomyocytes rely on digestion of the heart via various enzymes (collagenase, protease, trypsin, etc.). In the early periods (1950s-1960s), the chunk method was employed, which involved removing the heart, cutting into much smaller pieces and incubating in solution with collagenase/protease/trypsin12. In the 1970s labs implemented the ameliorated &#8220;Langendorff&#8221; method13, which isolated cardiomyocytes using a coronary artery perfusion-based isolation technique (retrograde perfusion with enzyme via the Langendorff apparatus); this technique remains the dominant method of myocyte isolation in the field today, ~50 years later14,15,16. Recent work has shifted to cannulating the heart in vivo to limit hypoxia time and ischemic damage resulting in superior cardiomyocyte isolations (better yields and higher quality)17. Recently, this has evolved into performing in vivo, Langendorff-free heart perfusions18,19,20,21,22. We have evolved the Langendorff-free cardiomyocyte isolation technique based on the Ackers-Johnson et al.18 technique and adapted various components from the many previous isolation techniques. These key adaptations include the injection of an ice-cold clearing buffer and the incorporation of a supporting platform to stabilize the needle, allowing for decreased manipulation of the heart. Also detailed in this technique is&#160; temperature control of injected buffers (37 &#176;C), which decreased the time between in vivo injection and digestion due to less EDTA perfusion as previously published18. By decreasing manipulation of the heart and therefore minimizing puncture site size, thorough and constant perfusion of the coronary arteries is obtained. We also refined the technique with a secondary chunk method digestion,&#160;the amount of EDTA in the injected clearing buffer, and changed the pH. Our described technique is more reliable, more efficient, and does not require the extensive training/practice compared to the using the Langendorff apparatus (Table 1).

<table><tbody><tr><td>10 cc Bd Luer-Lok Syringe</td><td>Fisher Sci</td><td>14-827-52</td><td></td></tr><tr><td>10 mL Pyrex Low-Form Beaker</td><td>Cole-Palmer</td><td>UX-34502-01</td><td></td></tr><tr><td>100 mL polypropylene cap glass media storage bottle</td><td>DWK Life Sciences</td><td>UX-34523-00</td><td></td></tr><tr><td>14 mL Round-Bottom Polypropylene Test Tubes With Cap</td><td>Fisher Sci</td><td>14-959-11B</td><td></td></tr><tr><td>2,3-Butanedione Monoxime</td><td>Sigma</td><td>B0753</td><td>&gt;98%</td></tr><tr><td>3 cc BD Luer-Lok Syringe</td><td>Fisher Sci</td><td>14-823-435</td><td></td></tr><tr><td>35 mm glass bottom dishes</td><td>MatTek Corporation</td><td>P35G-1.0-20-C</td><td></td></tr><tr><td>50 mL BD Syringe without Needle</td><td>Fisher Sci</td><td>13-689-8</td><td></td></tr><tr><td>50 mL Conical Centrifuge Tubes</td><td>Cole-Palmer</td><td>EW-22999-84</td><td></td></tr><tr><td>95% O&lt;sub&gt;2&lt;/sub&gt; 5% CO&lt;sub&gt;2&lt;/sub&gt;</td><td></td><td></td><td></td></tr><tr><td>AIMS Space Gel Heating Pad</td><td>Fisher Sci</td><td>14-370-223</td><td></td></tr><tr><td>BD PrecisionGlide 27 G X 1/2&quot; Hypodermic Needles</td><td>Becton Dickinson</td><td>305109</td><td></td></tr><tr><td>Bovine Serum Albumin</td><td>Sigma</td><td>A3803</td><td>Heat shock fraction, lyophilized powder, essentially fatty acid free, &gt;98%</td></tr><tr><td>Calcium Chloride dihydrate</td><td>Sigma</td><td>C7902</td><td>&gt;99%</td></tr><tr><td>D-(+)-Glucose</td><td>Sigma</td><td>G7021</td><td>Suitable for cell culture, &gt;99.5%</td></tr><tr><td>DMEM</td><td>Fisher Sci</td><td>11965092</td><td></td></tr><tr><td>EDTA</td><td>Fisher Sci</td><td>AAA1071336</td><td></td></tr><tr><td>Falcon 100 mm TC-treated Cell Culture Dish</td><td>Corning</td><td>353003</td><td></td></tr><tr><td>FBS</td><td>R&amp;amp;D Systems (Bio-techne)</td><td>S11195</td><td></td></tr><tr><td>Fisherbrand Isotemp Heated Immersion Circulators</td><td>Fisher Sci</td><td>13-874-432</td><td></td></tr><tr><td>Hartman Mosquito Hemostatic Forceps</td><td>World Precision Instruments</td><td>15921</td><td></td></tr><tr><td>Hausser Scientific Hy-Lite Counting Chamber Set</td><td>Fisher Sci</td><td>02-671-11</td><td></td></tr><tr><td>HEPES</td><td>Sigma</td><td>H4034</td><td>&gt;99.5%</td></tr><tr><td>Labeling Tape</td><td>Fisher Sci</td><td>15-901-10R</td><td></td></tr><tr><td>Legato 100 Syringe Pump</td><td>kdScientific</td><td>788100</td><td></td></tr><tr><td>L-glutathione</td><td>Fisher Sci</td><td>ICN19467980</td><td></td></tr><tr><td>Liberase TH Research Grade</td><td>Sigma</td><td>5401135001</td><td>High thermolysin concentration</td></tr><tr><td>M199</td><td>Fisher Sci</td><td>MT10060CV</td><td></td></tr><tr><td>Magnesium Chloride</td><td>Invitrogen</td><td>AM9530G</td><td></td></tr><tr><td>Mouse Laminin</td><td>Corning</td><td>354232</td><td></td></tr><tr><td>Pen/Strep</td><td>Fisher Sci</td><td></td><td></td></tr><tr><td>Potassium Chloride</td><td>Sigma</td><td>P5405</td><td>&gt;99%</td></tr><tr><td>Precision Digital Reciprocating Water Bath</td><td>ThermoFisher Scientific</td><td>TSCIR19</td><td></td></tr><tr><td>Sodium Bicarbonate</td><td>Sigma</td><td>S5761</td><td>Suitable for cell culture</td></tr><tr><td>Sodium Chloride</td><td>Sigma</td><td>S5886</td><td>&gt;99%</td></tr><tr><td>Sodium phosphate monobasic</td><td>Sigma</td><td>S5011</td><td>&gt;99%</td></tr><tr><td>Sterile Cell Strainer 70 &amp;micro;m</td><td>Fisher Sci</td><td>22-363-548</td><td></td></tr><tr><td>Student Fine Scissors</td><td>Fine Science Tools</td><td>91460-11</td><td></td></tr><tr><td>VWR Absorbent Underpads</td><td>Fisher Sci</td><td>NC9481815</td><td></td></tr></tbody></table>

a simple and effective method to consistently isolate mouse cardiomyocytes

The need for reproducible yet technically simple methods yielding high-quality cardiomyocytes is essential for research in cardiac biology. Cellular and molecular functional experiments (e.g., contraction, electrophysiology, calcium cycling, etc.) on cardiomyocytes are the gold standard for establishing mechanism(s) of disease. The mouse is the species of choice for functional experiments and the described technique is specifically for the isolation of mouse cardiomyocytes. Previous methods requiring a Langendorff apparatus require high levels of training and precision for aortic cannulation, often resulting in ischemia. The field is shifting toward Langendorff-free isolation methods that are simple, are reproducible, and yield viable myocytes for physiological data acquisition and culture. These methods greatly diminish ischemia time compared to aortic cannulation and result in reliably obtained cardiomyocytes. Our adaptation to the Langendorff-free method includes an initial perfusion with ice-cold clearing solution, use of a stabilizing platform that ensures a steady needle during perfusion, and additional digestion steps to ensure reliably obtained cardiomyocytes for use in functional measurements and culture. This method is simple and quick to perform and requires little technical skill.

There are a few elements to examine when determining the success of an isolation. First, the cardiomyocytes must be rod-shaped with no membrane blebs, such as the cells isolated in Figure 1. A typical isolation will yield ~80% of the myocytes being rod-shaped. If the isolation yields anything less than 50% rod-shaped cells, then it is considered an unsuccessful isolation and cardiomyocytes are not used. Lastly, the cardiomyocytes should be quiescent. Spontaneously contraction myocytes demons...

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A Simple and Effective Method to Consistently Isolate Mouse Cardiomyocytes

The gold standard in cardiology for cellular and molecular functional experiments are cardiomyocytes. This article describes adaptations to the non-Langendorff technique to isolate mouse cardiomyocytes.

The need for reproducible yet technically simple methods yielding high-quality cardiomyocytes is essential for research in cardiac biology. Cellular and ...

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2.8K Views.  The Ohio State University. Deze techniek is betrouwbaarder en efficiënter en vereist geen uitgebreide oefening of training in vergelijking met de meest gebruikte Langendorff-apparaattechnieken. Het grote voordeel van deze techniek is dat het de ischemische tijd beperkt, omdat de initiële perfusie in vivo plaatsvindt. Tijdens het uitvoeren van deze techniek is het essentieel om bubbels in het spruitstuk te vermijden en overmatige beweging van het hart tijdens de spijsvertering te voorkomen, deze kunnen de opbrengst ve...

Video: A Simple and Effective Method to Consistently Isolate Mouse Cardiomyocytes

Isolation and Culture of Neonatal Mouse Cardiomyocytes

Cultured neonatal cardiomyocytes have long been used to study myofibrillogenesis and myofibrillar functions. Cultured cardiomyocytes allow for easy investigation and manipulation of biochemical pathways, and their effect on the biomechanical properties of spontaneously beating cardiomyocytes.	
The following 2-day protocol describes the isolation and culture of neonatal mouse cardiomyocytes. We show how to easily dissect hearts from neonates, dissociate the cardiac tissue and enrich cardiomyocytes from the cardiac cell-population. We discuss the usage of different enzyme mixes for cell-dissociation, and their effects on cell-viability. The isolated cardiomyocytes can be subsequently used for a variety of morphological, electrophysiological, biochemical, cell-biological or biomechanical assays. We optimized the protocol for robustness and reproducibility, by using only commercially available solutions and enzyme mixes that show little lot-to-lot variability. We also address common problems associated with the isolation and culture of cardiomyocytes, and offer a variety of options for the optimization of isolation and culture conditions.

Primary mouse cardiomyocyte cultures are one of the pivotal tools for the investigation of myofibrillar organization and function. The following protocol describes the isolation and culture of primary cardiomyocytes from neonatal mouse hearts. The resulting cardiomyocyte cultures may be subsequently used for a variety of biomechanical, biochemical and cell-biological assays.

Cultured neonatal cardiomyocytes have long been used to study  myofibrillogenesis and myofibrillar functions. Cultured cardiomyocytes allow  for easy ...

Biology

Isolation, Culture, and Functional Characterization of Adult Mouse Cardiomyoctyes

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

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

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

Isolation and Physiological Analysis of Mouse Cardiomyocytes

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

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

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

Isolation of Atrial Myocytes from Adult Mice

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

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

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

Isolation and Culture of Adult Mouse Cardiomyocytes for Cell Signaling and in vitro Cardiac Hypertrophy

Technological advances have made genetically modified mice, including transgenic and gene knockout mice, an essential tool in many research fields. Adult cardiomyocytes are widely accepted as a good model for cardiac cellular physiology and pathophysiology, as well as for pharmaceutical intervention. Genetically modified mice preclude the need for complicated cardiomyocyte infection processes to generate the desired genotype, which are inefficient due to cardiomyocytes&#8217; terminal differentiation. Isolation and culture of high quantity and quality functional cardiomyocytes will dramatically benefit cardiovascular research and provide an important tool for cell signaling transduction research and drug development. Here, we describe a well-established method for isolation of adult mouse cardiomyocytes that can be implemented with little training. The mouse heart is excised and cannulated to an isolated heart system, then perfused with a calcium-free and high potassium buffer followed by type II collagenase digestion in Langendorff retrograde perfusion mode. This protocol yields a consistent result for the collection of functional adult mouse cardiomyocytes from a variety of genetically modified mice.

Isolation and Culture of Adult Mouse Cardiomyocytes for Cell Signaling and in vitro Cardiac Hypertrophy

We describe a reliable method for isolation of adult mouse cardiomyocytes. This protocol yields a consistent result for the culture of functional adult cardiomyocytes from a variety of genetically modified mice.

Technological advances have made genetically modified mice, including transgenic and gene knockout mice, an essential tool in many research fields. Adult ...

Isolation, Culture and Transduction of Adult Mouse Cardiomyocytes

Cultured cardiomyocytes can be used to study cardiomyocyte biology using techniques that are complementary to in vivo systems. For example, the purity and accessibility of in vitro culture enables fine control over biochemical analyses, live imaging, and electrophysiology. Long-term culture of cardiomyocytes offers access to additional experimental approaches that cannot be completed in short term cultures. For example, the in vitro investigation of dedifferentiation, cell cycle re-entry, and cell division has thus far largely been restricted to rat cardiomyocytes, which appear to be more robust in long-term culture. However, the availability of a rich toolset of transgenic mouse lines and well-developed disease models make mouse systems attractive for cardiac research. Although several reports exist of adult mouse cardiomyocyte isolation, few studies demonstrate their long-term culture. Presented here, is a step-by-step method for the isolation and long-term culture of adult mouse cardiomyocytes. First, retrograde Langendorff perfusion is used to efficiently digest the heart with proteases, followed by gravity sedimentation purification. After a period of dedifferentiation following isolation, the cells gradually attach to the culture and can be cultured for weeks. Adenovirus cell lysate is used to efficiently transduce the isolated cardiomyocytes. These methods provide a simple, yet powerful model system to study cardiac biology.

This protocol describes a step-by-step method for the reproducible isolation and long-term culture of adult mouse cardiomyocytes with high yield, purity, and viability.

Cultured cardiomyocytes can be used to study cardiomyocyte biology using techniques that are complementary to in vivo systems. For example, the purity and ...

Isolation, Transfection, and Long-Term Culture of Adult Mouse and Rat Cardiomyocytes

Ex vivo culture of the adult mammalian cardiomyocytes (CMs) presents the most relevant experimental system for the in vitro study of cardiac biology. Adult mammalian CMs are terminally differentiated cells with minimal proliferative capacity. The post-mitotic state of adult CMs not only restricts cardiomyocyte cell cycle progression but also limits the efficient culture of CMs. Moreover, the long-term culture of adult CMs is necessary for many studies, such as CM proliferation and analysis of gene expression.
The mouse and the rat are the two most preferred laboratory animals to be used for cardiomyocyte isolation. While the long-term culture of rat CMs is possible, adult mouse CMs are susceptible to death and cannot be cultured more than five days under normal conditions. Therefore, there is a critical need to optimize the cell isolation and long-term culture protocol for adult murine CMs. With this modified protocol, it is possible to successfully isolate and culture both adult mouse and rat CMs for more than 20 days. Moreover, the siRNA transfection efficiency of isolated CM is significantly increased compared to previous reports. For adult mouse CM isolation, the Langendorff perfusion method is utilized with an optimal enzyme solution and sufficient time for complete extracellular matrix dissociation. In order to obtain pure ventricular CMs, both atria were dissected and discarded before proceeding with the disassociation and plating. Cells were dispersed on a laminin coated plate, which allowed for efficient and rapid attachment. CMs were allowed to settle for 4-6 h before siRNA transfection. Culture media was refreshed every 24 h for 20 days, and subsequently, CMs were fixed and stained for cardiac-specific markers such as Troponin and markers of cell cycle such as KI67.

Here, we present a protocol for the isolation, transfection, and long-term culture of adult mouse and rat cardiomyocytes.

Ex vivo culture of the adult mammalian cardiomyocytes (CMs) presents the most relevant experimental system for the in vitro study of cardiac biology. ...

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 ...

Simultaneous Isolation and Culture of Atrial Myocytes, Ventricular Myocytes, and Non-Myocytes from an Adult Mouse Heart

The isolation and culturing of cardiac myocytes from mice has been essential for furthering the understanding of cardiac physiology and pathophysiology. While isolating myocytes from neonatal mouse hearts is relatively straightforward, myocytes from the adult murine heart are preferred. This is because compared to neonatal cells, adult myocytes more accurately recapitulate cell function as it occurs in the adult heart in vivo. However, it is technically difficult to isolate adult mouse cardiac myocytes in the necessary quantities and viability, which contributes to an experimental impasse. Furthermore, published procedures are specific for the isolation of either atrial or ventricular myocytes at the expense of atrial and ventricular non-myocyte cells. Described here is a detailed method for isolating both atrial and ventricular cardiac myocytes, along with atrial and ventricular non-myocytes, simultaneously from a single mouse heart. Also provided are the details for optimal cell-specific culturing methods, which enhance cell viability and function. This protocol aims not only to expedite the process of adult murine cardiac cell isolation, but also to increase the yield and viability of cells for investigations of atrial and ventricular cardiac cells.

A method is described for the simultaneous isolation of myocytes and non-myocytes from both the atria and ventricles of a single adult mouse heart. This protocol results in consistent yields of highly viable cardiac myocytes and non-myocytes and details optimal cell-specific culture conditions for phenotyping and in vitro analysis.

The isolation and culturing of cardiac myocytes from mice has been essential for furthering the understanding of cardiac physiology and pathophysiology. ...

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.

A Simple and Effective Method to Consistently Isolate Mouse Cardiomyocytes

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