This work is supported by operating grants from the Canadian Institutes of Health Research (MOP 93718, 142486) and the Heart and Stroke Foundation of Canada to R.A. Rose.&#160; H.J. Jansen is the recipient of a Killam Postdoctoral Fellowship.

All animal procedures were approved by The University of Calgary Animal Care and Use Committee and were conducted in accordance with the guidelines of the Canadian Council on Animal Care. The atrial myocyte isolation, images, and representative results described below were obtained from a 15 week-old male wildtype C57Bl/6 mouse. We routinely use this protocol to isolate atrial myocytes from wildtype mice17,18, mice carrying genetic mutations19,20 and mouse models of disease such as chronic hypertension6,14. The protocol can be used similarly for male or female mice. We also utilized a similar version of this isolation procedure to isolate sinoatrial node myocytes from the mouse heart17,21,22,23. A flowchart of this experimental protocol is located in Figure 1.
1. Preparation of Stock Solutions and Equipment
<ol>
	<li>Prepare 1 dissecting dish by adding silicone elastomer according to the manufacturer&#8217;s instructions. Add enough silicone elastomer compound to cover the bottom of a 10 cm Petri dish to a depth of 1 cm. Allow to cure and then insert 6 insect pins into the dish. 
	NOTE: This silicone dissecting dish can be reused for months and stored at room temperature.</li>
	<li>Prepare 3 fire-polished Pasteur pipettes with an opening of 1 mm (small bore), 3 mm (medium bore), or 5 mm (large bore) in diameter as shown in Figure 2A. To make these pipettes, score a Pasteur pipette and then snap along the score mark to produce an opening that is slightly larger than the desired bore size. Use a metal file to smooth the surface and then fire-polish this opening using an open flame. 
	NOTE: This will produce a smooth, fire-polished edge with an opening of the desired diameter. It is important that the opening is free of cracks and rough surfaces. These fire-polished pipettes can be stored at room temperature and reused for months.</li>
	<li>Prepare stock solutions for Tyrode&#8217;s pH 6.9 solution and Tyrode&#8217;s pH 7.4 solution as listed in Table 1. Also prepare 10 mL each of 1 M MgCl2, 1 M CaCl2, and 100 mM CaCl2. Use ultrapure water for all solutions and store at 4 &#176;C for up to 2 months.</li>
	<li>Prepare 1 L of modified Kraft-Bru&#776;he (KB) solution as indicated in Table 2. Use ultrapure water. Divide the solution into 20 mL aliquots and store at -20 &#176;C for up to 2 months.</li>
</ol>
2. Preparation of Solutions and Isolation Setup for Atrial Myocyte Isolation
<ol>
	<li>Prepare 50 mL of a modified Tyrode&#8217;s pH 7.4 solution as described in Table 3 in a 125 mL Erlenmeyer flask, using 1 M CaCl2 and 1 M MgCl2 stock solutions. Place the Erlenmeyer flask in a 35 &#176;C water bath until use, as shown in Figure 2B.</li>
	<li>Prepare the modified Tyrode&#8217;s pH 6.9 solution as described in Table 4 in a 50 mL tube, using the 100 mM CaCl2 stock solution. Aliquot 2.5 mL of this solution into each of three 5 mL round bottom tubes. Place these tubes in a wire rack placed in a 35 &#176;C water bath until use, as shown in Figure 2B.</li>
	<li>Prepare the enzyme solution as described in Table 5 in a 14 mL round bottom tube. To make the protease solution, add 1 mg of protease per 100 &#956;L of ultrapure water. Place the tube containing this enzyme solution in the wire rack and incubate in a 35 &#176;C water bath until use.</li>
	<li>Thaw one aliquot of modified KB solution in a 35 &#176;C water bath. Aliquot 2.5 mL of KB solution into each of three 5 mL round bottom tubes and 2.5 mL into a 14 mL round bottom tube. Place these tubes in the wire rack and incubate in a 35 &#176;C water bath until use, as shown in Figure 2B.</li>
	<li>Lay out the dissecting plate, dissecting tools, a Pasteur pipette, and the fire-polished pipettes as shown in Figure 2B.</li>
</ol>
3. Dissection of Mouse Atrial Appendage(s)
<ol>
	<li>Inject the mouse with 0.2 mL of heparin (10 000 USP U/10 mL) via intraperitoneal injection and wait 5 min for absorption.</li>
	<li>Place the mouse in an induction chamber and anesthetize by isoflurane inhalation (3-4%). Isoflurane and oxygen are delivered using an anesthetic machine and waste anesthetic gas is scavenged. Once the mouse is anesthetized, and does not exhibit a toe pinch reflex, euthanize the mouse by rapid cervical dislocation. Place the mouse on a paper towel or a cork board and tape the paws down to hold the mouse in place.</li>
	<li>Wet the chest of the mouse with 70% ethanol. Remove the fur and skin covering the chest using curved scissors. Next, use rat tooth forceps to lift the sternum and then cut the diaphragm along the edge of the ribs. Remove the entire rib cage using curved scissors to expose the heart.</li>
	<li>To remove the atrial appendage (right or left), gently lift the appendage using fine dissecting forceps and cut it out with spring scissors. Immediately transfer the atrial appendage to a silicone coated dissecting dish containing 20 mL of the warmed modified Tyrode&#8217;s pH 7.4 solution described in step 2.1.</li>
	<li>Place one dissecting pin at the top and one pin at the bottom of the opening of the atrial appendage. Using a pasture pipette, flush the atria with the warmed modified Tyrode&#8217;s pH 7.4 solution to remove blood. Open the atrial appendage by cutting along the top and bottom edge of the atrial appendage. Next, pin the corners of the atrial appendage down to create a flat, rectangular piece of tissue, as shown in Figure 3A.</li>
</ol>
4. Isolation of Atrial Myocytes
NOTE: Steps in this section are all performed at 35 &#176;C, with tubes submerged in a 35 &#176;C water bath. Be careful when transferring tissue strips between round bottom tubes to ensure that only the tissue (and not the solution) is transferred between tubes.
<ol>
	<li>Cut the atrial appendage into approximately 8-10 equal sized strips (approximately 0.7 mm in width) using spring scissors and fine forceps. An example of the strips of atrial tissue is shown in Figure 3B. Note that the strips contract once they are cut free from the main piece of tissue. Using the small bore fire-polished pipette, transfer the tissue strips into the first tube containing warmed modified Tyrode&#8217;s pH 6.9 solution described in step 2.2. Wait 5 min.</li>
	<li>Wash the tissue strips by transferring them to the second and then the third round bottom tube containing modified Tyrode&#8217;s pH 6.9 solution prepared in step 2.2 using the medium bore fire-polished pipette.
	<ol>
		<li>To wash the tissue strips, cap the 5 mL round bottom tube and gently invert the tube 3 times. Let the tissue strips settle to the bottom of the tube before transferring the tissue strips to the next tube using the medium bore fire-polished pipette.</li>
	</ol>
	</li>
	<li>Transfer the tissue strips into the enzyme solution described in step 2.3 using a medium bore fire-polished pipette and incubate for 30 min. Swirl the tube every 3-5 min to prevent the tissue strips from adhering together. 
	NOTE: At the beginning of the enzymatic digestion, tissue strips settle quickly following swirling. At approximately 20 min of digestion, the tissue strips begin to float in the enzymatic solution following swirling. During this time, the atrial tissue strips also change in appearance from pale pink to white as they are digested.</li>
	<li>After enzymatic digestion, perform three washes using 2.5 mL of KB solution in the 5 mL round bottom tubes prepared in step 2.4. For each wash, gently invert the tube 3 times before moving the tissue to the next tube using the medium bore fire-polished pipette. Following the final wash, transfer the strips into the 14 mL round bottom tube containing 2.5 mL of KB solution. Wait 5 min.</li>
	<li>Gently triturate the tissue for 7.5 min using the wide bore fire-polished pipette. This will mechanically dissociate the tissue strips and yield a cloudy solution filled with individual atrial myocytes. 
	NOTE: During trituration, the tissue becomes white and the solution becomes cloudy. The force of trituration, achieved by altering both the frequency and velocity of expelling the tissue strips from the wide bore fire-polished pipette, should be tailored to the individual isolation. If trituration is too gentle, the cell yield will be low, while trituration that is too harsh will yield many dead cells. Avoid bubbles while triturating.</li>
	<li>Fill the 14 mL round bottom tube containing the triturated tissue strips with KB solution to a final volume of 7-10 mL depending on the desired density of cells for experimental use. Place this tube at room temperature for 1 h. Following this incubation period, cells can be used for a variety of experiments for up to 7 h. Cells can also be stored at 4 &#176;C for up to 7 h.</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+patterns+of+constitutive+phosphodiesterase+activity+in+mouse+sinoatrial+node+and+atrial+myocardium.">Distinct patterns of constitutive phosphodiesterase activity in mouse sinoatrial node and atrial myocardium.</a> PLoS One. 7 (10), e47652 (2012).</li><li>Egom, E. E., 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=Impaired+sinoatrial+node+function+and+increased+susceptibility+to+atrial+fibrillation+in+mice+lacking+natriuretic+peptide+receptor+C.">Impaired sinoatrial node function and increased susceptibility to atrial fibrillation in mice lacking natriuretic peptide receptor C.</a> Journal of Physiology. 593 (5), 1127-1146 (2015).</li><li>Hua, R., 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=Effects+of+Wild-Type+and+Mutant+Forms+of+Atrial+Natriuretic+Peptide+on+Atrial+Electrophysiology+and+Arrhythmogenesis.">Effects of Wild-Type and Mutant Forms of Atrial Natriuretic Peptide on Atrial Electrophysiology and Arrhythmogenesis.</a> Circulation: Arrhythmia and Electrophysiology. 8 (5), 1240-1254 (2015).</li><li>Krishnaswamy, P. 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The authors have to nothing to disclose.

Our lab routinely uses this protocol to isolate mouse atrial myocytes for use in patch-clamp experiments in order to investigate the effects of different forms of cardiovascular disease, genetic mutations, or pharmacological compounds on atrial myocyte electrophysiology. Although highly reproducible, the quality of the data obtained from the isolated atrial myocytes depends on the quality of the isolation. In addition, reintroduction of calcium following the atrial myocyte isolation will result in cell death for a population of isolated myocytes due to the calcium paradox16. Accordingly, isolating viable, high-quality atrial myocytes using this approach requires practice and optimization at multiple points throughout the isolation. Once optimized, it is estimated that between 70-90% of the total atrial myocytes isolated using this approach will be both calcium tolerant and rod shaped. The steps requiring the most practice and optimization are discussed below.
The speed and efficiency of the dissection will have downstream effects on the quality of the isolated cells. It is important to take time to ensure all blood is removed from the atrial tissue and that tissue strips are cut to a similar size. It should take approximately 5 min to remove the atrial appendage, cut the tissue into strips, and transfer the tissue strips into the first tube of modified Tyrode&#8217;s pH 6.9 solution. However, if this step takes too long, the quality of the tissue may be compromised.
It is also important that tissue strips are cut to a uniform size within an isolation and between hearts. If tissue strips are too large or too small, or if they are not uniform within an isolation, this can cause problems during both enzymatic digestion and trituration. This is because small strips will be more thoroughly digested and large strips will be under digested. It is equally important to consider the genotype and disease setting being studied as the size of the atrial appendage can vary between animals. For example, hypertrophic hearts have larger atrial appendages compared to healthy hearts, and therefore the experimenter can cut more strips in hypertrophic hearts compared to normal sized hearts. Accordingly, optimizing the size of the cut tissue strips and applying these dimensions to each individual atrial appendage will greatly improve the reproducibility of myocyte isolations between experimental conditions.
The delicate balance between the enzymatic digestion and mechanical dissociation is key to a successful atrial myocyte isolation using this protocol. If tissue is not adequately swirled during the enzymatic&#160;digestion the individual tissue strips will tend to clump and stick together, which will limit the effectiveness of the enzymatic digestion. If agitated too frequently or vigorously, this can damage the atrial tissue, which will result in the isolation of nonviable cells. Mechanical dissociation of isolated atrial myocytes from tissue strips during trituration is the most critical step to practice and optimize using this approach to isolate atrial myocytes. If trituration is too gentle, cell yield will be low. On the other hand, if trituration is too harsh, then an abundance of non-viable myocytes will be isolated, and the quality of data obtained during patch-clamp experiments will be jeopardized. In addition, the composition of the atria can affect the isolation. For example, if tissue is fibrotic, the enzymatic digestion and the trituration steps may need to be modified. It is therefore important to take the time to develop the skills required to obtain high quality cells during trituration that can be used for patch-clamp experiments.
As with all experimental techniques there are limitations. This technique requires practice in order to reproducibly isolate viable, high quality myocytes, which in turn will impact the feasibility of any experiments to be performed using these myocytes. This approach is also terminal and atrial myocytes isolated using this approach can be used on the day of the isolation only. Our lab uses the cells within 6-7 h of isolation.
This approach of isolating atrial cardiomyocytes has several applications. For example, this approach can be modified to isolate atrial myocytes (as well as cardiac fibroblasts) from other species including human atrial tissue biopsies. In addition, a benefit to using this chunk method for atrial myocyte isolation (in contrast to retrograde perfusion of the heart) is that it can be modified to isolate cardiomyocytes from other regions of the heart, such as the sinoatrial node or other specific regions of the atrial myocardium, or encompass the entire supraventricular region of the heart. Our lab uses atrial cardiomyocytes for patch-clamp experiments to measure action potentials and ionic currents, although this approach need not be limited to this technique. For example, isolated myocytes can be used to investigate alterations in calcium transients and contractility in a variety of experimental settings. Atrial cardiomyocytes can also be used in immunofluorescence studies to study the location of proteins or structures of interest. Accordingly, this approach is highly versatile with many possible applications.

The atria, which are the thin walled, low pressure chambers of the heart that receive blood from the superior and inferior vena cavae as well as the pulmonary veins, are integral in normal cardiac physiology. Like other regions of the heart, the atria contain a number of cell types, including cardiomyocytes, fibroblasts, endothelial cells, vascular smooth muscle cells, and others. Atrial myocytes are electrically excitable cells that play an essential role in the conduction of electrical signals through the heart, thereby ensuring proper atrial contraction during each heart beat1. Electrical dysfunction in the atria can lead to a number of atrial specific arrhythmias such as atrial flutter and atrial fibrillation2,3. These are highly prevalent, yet poorly understood, atrial arrhythmias that lead to significant morbidity and mortality. Atrial fibrillation can occur in association with genetic mutations, in association with aging or in the setting of acquired forms of heart disease, including hypertension, heart failure and diabetes2,4,5,6. These conditions can alter the electrical properties of atrial myocytes which can create a substrate that increases the prevalence of arrhythmogenesis1,2.
Normal electrical function in the atria, as well as atrial arrhythmogenesis, are importantly affected by the morphology of the action potential (AP) produced in atrial myocytes. The atrial AP is generated from the activity of a number of ionic currents, including the sodium current (INa, carried by NaV&#173;1.5 channels), the L-type calcium current (ICa,L, carried by Cav1.2 and CaV1.3 channels), and several potassium currents including the ultra-rapid delayed rectifier potassium current (IKur, carried by KV1.5 channels), the transient outward potassium current (Ito, carried by KV&#173;4.2 and KV4.3 channels), a steady state potassium current (IKss, carried by KV2.1 channels), and the inward rectifier potassium current (IK1, carried by Kir2.1 channels)1,7,8. Although they do not play a major role in the mouse atria, the rapid and slow components of the delayed rectifier K+ current (IKr and IKs) also contribute to AP repolarization in some species7. Alterations in one or more of these ionic currents can significantly alter the electrical properties of atrial myocytes, which can lead to atrial arrhythmias. For example, a reduction in INa can slow conduction velocity across the atria by reducing the AP upstroke velocity. On the other hand, a reduction in repolarizing potassium currents or an increase in either ICa,L or the late INa can result in the development of afterdepolarizations that can trigger spontaneous activity in the atria1,2,9.
It is important to recognize that there are differences in AP morphology in different parts of the atrial myocardium that are likely due to differences in the expression or regulation of these underlying ion channels. For example, differences in AP duration between the right and left atria in association with differences in Ito current density have been well described10,11,12,13. Also, we have recently demonstrated that there are distinct patterns of electrical remodeling in the right and left atria of mice with chronic hypertension6,14. The right atrial posterior wall also contains the sinoatrial node, which has its own distinct patterns of AP morphology and firing patterns15. The distinct properties of myocytes in each of these different parts of the atria can be investigated in detail using isolated myocytes from each of these regions.
There are different approaches that can be used to isolate atrial myocytes for patch-clamp electrophysiology studies16. One possibility is to use a retrograde perfusion approach where the heart is cannulated via the aorta for the delivery of enzymes. While this is a viable approach, it can produce variability in atrial myocyte quality due to inconsistencies in perfusion of the atria. We have adopted a &#8216;chunk&#8217; digestion approach for the isolation of atrial myocytes that eliminates the need for retrograde perfusion of the heart. Our approach uses a combination of enzymatic digestion and mechanical dissociation of atrial tissue that consistently and reliably yields large numbers of isolated atrial myocytes that are suitable for patch-clamp studies. While we describe our approach here using atrial appendage tissue, the approach can be used on any region of the atrial myocardium (i.e., right or left atrial appendages, free walls, posterior walls) that the investigator chooses. This approach is ideal for studies of atrial myocyte electrophysiology in genetically modified mice, in mouse models of cardiovascular disease, or for studying the effects of pharmacological compounds5,6,17,18,19.

<table>
	<tbody>
		<tr>
			<td>1, 2-Bis(2-Aminophenoxy)ethane-N, N, N&#39;, N&#39;-tetraacetic acid 98%</td>
			<td>Sigma</td>
			<td>A4926-1G</td>
			<td></td>
		</tr>
		<tr>
			<td>Adenosine 5&#39;-triphosphate disodium salt hydrate BioXtra, &#62; 99%, from microbial</td>
			<td>Sigma</td>
			<td>A7699-1G</td>
			<td></td>
		</tr>
		<tr>
			<td>Adenosine 5&#39;-triphosphate magnesium salt &#62; 95%, bacterial</td>
			<td>Sigma</td>
			<td>A9187-1G</td>
			<td></td>
		</tr>
		<tr>
			<td>Amphocetericin B from Streptomyces sp. ~80% (HPLC), powder</td>
			<td>Sigma</td>
			<td>A4888-500 MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine serum albumin</td>
			<td>Sigma</td>
			<td>A3059-50G</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate</td>
			<td>Sigma</td>
			<td>223506-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Cesium chloride ReagentPlus, 99.9%</td>
			<td>Sigma</td>
			<td>289329-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>Cesium hydroxide monohydrate &#62; 99.5% trace metals basis</td>
			<td>Sigma</td>
			<td>562505-1KG</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase Type 2</td>
			<td>Worthington Biochemical Corporation</td>
			<td>LS004176</td>
			<td></td>
		</tr>
		<tr>
			<td>Creatine anhydrous</td>
			<td>Sigma</td>
			<td>C0780</td>
			<td></td>
		</tr>
		<tr>
			<td>D-(+)-Glucose</td>
			<td>Sigma</td>
			<td>G7021-1KG</td>
			<td></td>
		</tr>
		<tr>
			<td>DL-Aspartic acid potassium salt</td>
			<td>Sigma</td>
			<td>A2025-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>Elastase suspension</td>
			<td>Worthington Biochemical Corporation</td>
			<td>LS002279</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethylene glycol-bis(2-amino-ethylether)-N,N,N&#39;,N&#39;-tetraacetic acid &#62;97.0%</td>
			<td>Sigma</td>
			<td>E4378-25G</td>
			<td></td>
		</tr>
		<tr>
			<td>Guanosine 5&#39;-triphosphate sodium salt hydrate &#62; 95% (HPLC), powder</td>
			<td>Sigma</td>
			<td>G8877-250MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin 10 000 USP units/10mL</td>
			<td>SANDOZ</td>
			<td>10750</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES &#62; 99.5% (titration)</td>
			<td>Sigma</td>
			<td>H3375-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>L-Glutamic acid potassium salt monohydrate &#62; 99% (HPLC), powder</td>
			<td>Sigma</td>
			<td>G1501-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium sulfate</td>
			<td>Sigma</td>
			<td>M2643-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Nifedipine &#62; 98% (HPLC), powder</td>
			<td>Sigma</td>
			<td>N7634-1G</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphocreatine disodium salt hydrate enzymatic, approx 98%</td>
			<td>Sigma</td>
			<td>P7936-5G</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium chloride ACS reagent, 99.0-100.5%</td>
			<td>Sigma</td>
			<td>P3911-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium hydroxide</td>
			<td>EM Science</td>
			<td>PX1480-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium phosphate monobasic</td>
			<td>EMD</td>
			<td>PX1565-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Protease from Streptomyces griseus, type XIV, &#62;3.5 units/mg solid, powder</td>
			<td>Sigma</td>
			<td>P5147-1G</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride ACS reagent, &#62; 99.0%</td>
			<td>Sigma</td>
			<td>S9888-2.5KG</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium hydroxide, pellets, 97+%, A.C.S. reagent</td>
			<td>Sigma</td>
			<td>221465-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Sylgard 184 silicone elastomer kit</td>
			<td>World Precision Instruments Inc</td>
			<td>SYLG184</td>
			<td></td>
		</tr>
		<tr>
			<td>Taurine</td>
			<td>Sigma</td>
			<td>T0625-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>Tetraethylammonium chloride &#62; 98% (titration)</td>
			<td>Sigma</td>
			<td>T2265-100G</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

The atrial myocytes isolated using this protocol can be used to characterize the electrophysiological properties of these cells using the patch-clamp technique. Aliquots of atrial myocytes in KB solution can be added to the recording chamber of a standard patch-clamp apparatus and superfused with solutions appropriate for the kind of recording the experimentalist wishes to perform. Atrial myocytes isolated using this protocol are best used for electrophysiological studies within 6-7 h of isolation. Representative patch-clamp data from our laboratory is presented below.
Figure 4 illustrates examples of isolated atrial myocytes from normal mice prepared using the protocol above. Isolated atrial myocytes are typically on the order of 100 &#181;m in length and 10 &#181;m in width with clear striations. The capacitance of isolated atrial myocytes is typically 40-70 pF.
Figure 5A illustrates an example of an atrial myocyte AP recorded using the perforated patch-clamp technique in current clamp mode, as we have described previously6,19,20. Summary data illustrating typical atrial myocyte AP parameters are provided in Table 6. Specifically, we present summary data for measurements of resting membrane potential (RMP), maximum upstroke velocity (Vmax), overshoot (OS) and AP duration at 50% (APD50), 70% (APD70) and 90% (APD90) repolarization time (Table 6). APs can also be recorded in the whole cell configuration14. The superfusion and pipette solutions for recording APs are available in Table 7 and Table 8.
Figure 5B illustrates a representative family of Na+ currents (INa) recorded in the whole cell configuration of the patch-clamp technique. These currents were recorded using 50 ms voltage clamp steps between -100 and +10 mV from a holding potential of -120 mV. We have described approaches and protocols for recording INa previously6,14,20. A summary INa IV relationship is also presented in Figure 5B. Solutions used to record INa are presented in Table 7 and Table 8.
Figure 5C illustrates a representative family of Ca2+ currents (ICa,L) recorded in the whole cell configuration of the patch-clamp technique. These currents were recorded using 250 ms voltage clamp steps between -60 and +80 mV from a holding potential of -70 mV. The experimental conditions that can be used to measure ICa,L have been previously described17,18,20. A summary ICa,L IV relationship is also presented in Figure 5C. The solutions used to record ICa,L are available in Table 7 and Table 8.
Figure 5D illustrates a representative family of K+ currents (IK) recorded in the whole cell configuration of the patch-clamp technique. These currents were recorded from a holding potential of -80 mV using 500 ms voltage clamp steps between -120 mV and +80 mV, as we have described previously6,14. The summary IV relationship for total IK is also presented in Figure 5D. The solutions used to record IK are available in Table 7 and Table 8.
Using these approaches to record APs and major families of ionic currents, including Na+, Ca2+ and K+ currents (as illustrated above), permits the investigator to rigorously interrogate atrial myocyte electrophysiology in a plethora of experimental conditions. Our laboratory has routinely employed these techniques to study atrial myocyte electrophysiology in normal mice, in mouse models of heart disease, and in genetically modified mice6,14,17,18,19,20.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59588/59588fig1.jpg" /> 
Figure 1: Flowchart for the atrial myocyte isolation protocol. Summary of the steps used to isolate atrial myocytes using this protocol. <a href="https://www.jove.com/files/ftp_upload/59588/59588fig1large.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/59588/59588fig2.jpg" /> 
Figure 2: Experimental setup and dissection tools for atrial myocyte isolation. (A). A small bore fire-polished pipette with an opening 1 mm in diameter (left) is used for tissue transfer following dissection, a medium bore fire-polished pipette with an opening 3 mm in diameter (middle) is used to transfer tissue strips during the isolation, and a large bore fire-polished pipette with an opening 5 mm in diameter (right) is used for trituration of digested atrial tissue. (B). Experimental setup for atrial myocyte isolation. <a href="https://www.jove.com/files/ftp_upload/59588/59588fig2large.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/59588/59588fig3.jpg" /> 
Figure 3: Image of the atrial appendage dissection. (A). Representative bright field image of an excised atrial appendage cut open and pinned out. (B). Representative bright field image of the atrial appendage cut into tissue strips of approximately 0.7 mm in width. Scale bar = 1 mm. <a href="https://www.jove.com/files/ftp_upload/59588/59588fig3large.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/59588/59588fig4.jpg" /> 
Figure 4: Images of isolated atrial myocytes. (A). Brightfield image of isolated atrial myocytes immediately after isolation. Scale bar = 50 &#956;m. (B). Brightfield image of a single isolated atrial myocyte. Scale bar = 100 &#956;m. <a href="https://www.jove.com/files/ftp_upload/59588/59588fig4large.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/59588/59588fig5.jpg" /> 
Figure 5: Representative patch-clamp data obtained from isolated atrial myocytes. (A). Representative stimulated AP recording from an isolated atrial myocyte. Summary of AP parameters is presented in Table 6. Amphotericin B (200 &#956;g/mL) was added to the pipette solution to permeabilize the cellular membrane. (B). Representative INa recordings (left) and summary INa IV curve (right) from an isolated atrial myocyte. Nifedipine (10 &#956;M) was added to the modified Tyrode&#8217;s solution to block ICa,L&#173; when recording INa&#173;. C. Representative ICa,L recordings (left) and summary ICa,L IV curve (right) from an isolated atrial myocyte. (D). Representative IK recordings (left) and summary IK IV curve (right) from an isolated atrial myocyte. The solutions used to record each of these currents are listed in Table 7 and Table 8. Summary IV curves are averaged measurements from 10 atrial myocytes isolated from a 15 week-old male wildtype C57Bl/6 mouse. <a href="https://www.jove.com/files/ftp_upload/59588/59588fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>Stock Tyrode&#39;s pH 6.9</td>
			<td>Stock Tyrode&#39;s pH 7.4</td>
		</tr>
		<tr>
			<td>Chemical</td>
			<td>in mM</td>
			<td>in mM</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>140</td>
			<td>140</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>5.4</td>
			<td>5.4</td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>1.2</td>
			<td>1.2</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>5</td>
			<td>5</td>
		</tr>
		<tr>
			<td>Final volume</td>
			<td>500 mL</td>
			<td>1 L</td>
		</tr>
		<tr>
			<td>Final pH with NaOH</td>
			<td>6.9</td>
			<td>7.4</td>
		</tr>
	</tbody>
</table>
Table 1: Stock Tyrode&#8217;s pH 7.4 and stock Tyrode&#8217;s pH 6.9 solutions. Composition of stock Tyrode&#8217;s solutions (pH 7.4 and pH 6.9) that can be made in advance and stored at 4 &#176;C for up to 2 months.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Chemical</td>
			<td>in mM</td>
		</tr>
		<tr>
			<td>K-glutamate</td>
			<td>100</td>
		</tr>
		<tr>
			<td>K-aspartate</td>
			<td>10</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>25</td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>10</td>
		</tr>
		<tr>
			<td>MgSO4</td>
			<td>2</td>
		</tr>
		<tr>
			<td>Taurine</td>
			<td>20</td>
		</tr>
		<tr>
			<td>Creatine</td>
			<td>5</td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>0.5</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>20</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>5</td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>0.10%</td>
		</tr>
		<tr>
			<td>Final volume&#160;</td>
			<td>1 L</td>
		</tr>
		<tr>
			<td>Final pH with KOH</td>
			<td>7.2</td>
		</tr>
	</tbody>
</table>
Table 2: Modified KB solution. Recipe for modified KB solution that can be made in advance, aliquoted, and stored at -20 &#176;C for up to 2 months.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Chemical</td>
			<td>Amount</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>5.55 mM</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>1.8 mM</td>
		</tr>
		<tr>
			<td>Stock Tyrode&#39;s pH 7.4</td>
			<td>50 mL</td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>250 &#956;L</td>
		</tr>
	</tbody>
</table>
Table 3: Modified Tyrode&#8217;s pH 7.4 solution with glucose, magnesium, calcium, and heparin. Composition of modified Tyrode&#8217;s pH 7.4 solution used for the atrial tissue dissection. This solution should be made fresh and kept in a 35 &#176;C water bath until use.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Chemical</td>
			<td>Amount</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>18.5 mM</td>
		</tr>
		<tr>
			<td>Taurine</td>
			<td>49.96 mM</td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>15 mg</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>0.066 mM</td>
		</tr>
		<tr>
			<td>Stock Tyrode&#39;s pH 6.9</td>
			<td>15 mL</td>
		</tr>
	</tbody>
</table>
Table 4: Modified Tyrode&#39;s pH 6.9 solution containing glucose, taurine, BSA, and low calcium. Composition of the modified Tyrode&#8217;s pH 6.9 solution used for the atrial myocyte isolation. This solution should be made fresh and kept in a 35 &#176;C water bath until use.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Chemical</td>
			<td>Amount</td>
		</tr>
		<tr>
			<td>Collagenase</td>
			<td>1,064 U</td>
		</tr>
		<tr>
			<td>Elastase</td>
			<td>9 U</td>
		</tr>
		<tr>
			<td>Protease solution</td>
			<td>65.2 &#956;L&#160;</td>
		</tr>
		<tr>
			<td>Modified Tyrode&#39;s pH 6.9</td>
			<td>5 mL</td>
		</tr>
	</tbody>
</table>
Table 5: Enzymatic solution. Composition of the enzymatic solution used to enzymatically digest atrial tissue strips. This solution should be made fresh and kept in a 35 &#176;C water bath until use.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Parameter</td>
			<td>Average</td>
		</tr>
		<tr>
			<td>RMP (mV)</td>
			<td>-74.2 &#177; 0.7</td>
		</tr>
		<tr>
			<td>Vmax (V/s)</td>
			<td>144.6 &#177; 5.8</td>
		</tr>
		<tr>
			<td>OS (mV)</td>
			<td>71.9 &#177; 3.0</td>
		</tr>
		<tr>
			<td>APD50 (ms)</td>
			<td>11.1 &#177; 1.7</td>
		</tr>
		<tr>
			<td>APD70 (ms)</td>
			<td>23.0 &#177; 4.6</td>
		</tr>
		<tr>
			<td>APD90 (ms)</td>
			<td>54.7 &#177; 7.8</td>
		</tr>
	</tbody>
</table>
Table 6: Summary of AP parameters from isolated atrial myocytes. Data are presented as mean &#177; SEM, n = 10 atrial myocytes isolated from a 15 week male wildtype C57Bl/6 mouse.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>Potassium currents and APs</td>
			<td>Sodium currents</td>
			<td>Calcium currents</td>
		</tr>
		<tr>
			<td>Chemical</td>
			<td>in mM</td>
			<td>in mM</td>
			<td>in mM</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>140</td>
			<td>5</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>5.4</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>1</td>
			<td>1</td>
			<td>1</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>1</td>
			<td>1</td>
			<td>2</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>10</td>
			<td>10</td>
			<td>10</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>5.5</td>
			<td>5.5</td>
			<td>5.5</td>
		</tr>
		<tr>
			<td>CsCl</td>
			<td></td>
			<td>130</td>
			<td></td>
		</tr>
		<tr>
			<td>TEA-Cl</td>
			<td></td>
			<td>5.4</td>
			<td>145.5</td>
		</tr>
		<tr>
			<td>pH</td>
			<td>7.4 with NaOH</td>
			<td>7.4 with CsOH</td>
			<td>7.4 with CsOH</td>
		</tr>
	</tbody>
</table>
Table 7: Composition of Tyrode&#8217;s solutions used during patch-clamp experiments. Composition of the Tyrode&#8217;s solutions used to record APs, INa&#173;, ICa,L, and IK from isolated atrial myocytes.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>Potassium currents and APs</td>
			<td>Sodium currents</td>
			<td>Calcium currents</td>
		</tr>
		<tr>
			<td>Chemical</td>
			<td>in mM</td>
			<td>in mM</td>
			<td>in mM</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>5</td>
			<td>5</td>
			<td>5</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>140</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>1</td>
			<td>1</td>
			<td>1</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>0.2</td>
			<td>0.2</td>
			<td>0.2</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>10</td>
			<td>10</td>
			<td>10</td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>5</td>
			<td></td>
			<td>5</td>
		</tr>
		<tr>
			<td>Mg-ATP</td>
			<td>4</td>
			<td>5</td>
			<td>4</td>
		</tr>
		<tr>
			<td>Na-GTP</td>
			<td>0.3</td>
			<td>0.3</td>
			<td>0.3</td>
		</tr>
		<tr>
			<td>Na-phosphocreatine</td>
			<td>6.6</td>
			<td></td>
			<td>6.6</td>
		</tr>
		<tr>
			<td>CsCl</td>
			<td></td>
			<td>130</td>
			<td>135</td>
		</tr>
		<tr>
			<td>BAPTA</td>
			<td></td>
			<td>5</td>
			<td></td>
		</tr>
		<tr>
			<td>pH</td>
			<td>7.2 with KOH</td>
			<td>7.2 with CsOH</td>
			<td>7.2 with CsOH</td>
		</tr>
	</tbody>
</table>
Table 8: Composition of internal pipette solution used during patch-clamp experiments. Composition of the pipette filling solutions used to record APs, INa&#173;, ICa,L, and IK from isolated atrial myocytes.

Watch this Scientific Journal Video about Isolation of Atrial Myocytes from Adult Mice at JoVE.com

Isolation of Atrial Myocytes from Adult Mice

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 ...

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Research

JoVE Journal

Medicine

10.2K Views.  University of Calgary. 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.

Video: Isolation of Atrial Myocytes from Adult Mice

The WATCHMAN Left Atrial Appendage Closure Device for Atrial Fibrillation

Atrial fibrillation (AF) is the most common cardiac arrhythmia, affecting an estimated 6 million people in the United States 1. Since AF affects primarily elderly people, its prevalence increases parallel with age. As such, it is expected that 15.9 million Americans will be affected by the year 2050 2. Ischemic stroke occurs in 5% of non-anticoagulated AF patients each year. Current treatments for AF include rate control, rhythm control and prevention of stroke 3.
The American College of Cardiology, American Heart Association, and European Society of Cardiology currently recommended rate control as the first course of therapy for AF 3. Rate control is achieved by administration of pharmacological agents, such as &beta;-blockers, that lower the heart rate until it reaches a less symptomatic state 3. Rhythm control aims to return the heart to its normal sinus rhythm and is typically achieved through administration of antiarrhythmic drugs such as amiodarone, electrical cardioversion or ablation therapy. Rhythm control methods, however, have not been demonstrated to be superior to rate-control methods 4-6. In fact, certain antiarrhythmic drugs have been shown to be associated with higher hospitalization rates, serious adverse effects 3, or even increases in mortality in patients with structural heart defects 7. Thus, treatment with antiarrhythmics is more often used when rate-control drugs are ineffective or contraindicated. Rate-control and antiarrhythmic agents relieve the symptoms of AF, including palpitations, shortness of breath, and fatigue 8, but don't reliably prevent thromboembolic events 6.
Treatment with the anticoagulant drug warfarin significantly reduces the rate of stroke or embolism 9,10. However, because of problems associated with its use, fewer than 50% of patients are treated with it. The therapeutic dose is affected by drug, dietary, and metabolic interactions, and thus requires detailed monitoring. In addition, warfarin has the potential to cause severe, sometimes lethal, bleeding 2. As an alternative, aspirin is commonly prescribed. While aspirin is typically well tolerated, it is far less effective at preventing stroke 10. Other alternatives to warfarin, such as dabigatran 11 or rivaroxaban 12 demonstrate non-inferiority to warfarin with respect to thromboembolic events (in fact, dabigatran given as a high dose of 150 mg twice a day has shown superiority). While these drugs have the advantage of eliminating dietary concerns and eliminating the need for regular blood monitoring, major bleeding and associated complications, while somewhat less so than with warfarin, remain an issue 13-15.
Since 90% of AF-associated strokes result from emboli that arise from the left atrial appendage (LAA) 2, one alternative approach to warfarin therapy has been to exclude the LAA using an implanted device to trap blood clots before they exit. Here, we demonstrate a procedure for implanting the WATCHMAN Left Atrial Appendage Closure Device. A transseptal cannula is inserted through the femoral vein, and under fluoroscopic guidance, inter-atrial septum is crossed. Once access to the left atrium has been achieved, a guidewire is placed in the upper pulmonary vein and the WATCHMAN Access Sheath and dilator are advanced over the wire into the left atrium. The guidewire is removed, and the access sheath is carefully advanced into the distal portion of the LAA over a pigtail catheter. The WATCHMAN Delivery System is prepped, inserted into the access sheath, and slowly advanced. The WATCHMAN device is then deployed into the LAA. The device release criteria are confirmed via fluoroscopy and transesophageal echocardiography (TEE) and the device is released.

The accompanying video describes a procedure for percutaneous placement of the WATCHMAN Left Atrial Appendage (LAA) Device. The WATCHMAN is a nitinol device designed to be permanently implanted at, or slightly distal to, the opening of the left atrial appendage (LAA) to trap blood clots before they exit the LAA, preventing thromboembolic stroke.

Atrial fibrillation (AF) is the most common cardiac arrhythmia, affecting an estimated 6 million people in the United States 1. Since AF affects primarily ...

Catheter Ablation in Combination With Left Atrial Appendage Closure for Atrial Fibrillation

Atrial fibrillation (AF) is the most common sustained cardiac arrhythmia, affecting millions of individuals worldwide 1-3. The rapid, irregular, and disordered electrical activity in the atria gives rise to palpitations, fatigue, dyspnea, chest pain and dizziness with or without syncope 4, 5. Patients with AF have a five-fold higher risk of stroke 6.
Oral anticoagulation (OAC) with warfarin is commonly used for stroke prevention in patients with AF and has been shown to reduce the risk of stroke by 64% 7. Warfarin therapy has several major disadvantages, however, including bleeding, non-tolerance, interactions with other medications and foods, non-compliance and a narrow therapeutic range 8-11. These issues, together with poor appreciation of the risk-benefit ratio, unawareness of guidelines, or absence of an OAC monitoring outpatient clinic may explain why only 30-60% of patients with AF are prescribed this drug 8.
The problems associated with warfarin, combined with the limited efficacy and/or serious side effects associated with other medications used for AF 12,13, highlight the need for effective non-pharmacological approaches to treatment. One such approach is catheter ablation (CA), a procedure in which a radiofrequency electrical current is applied to regions of the heart to create small ablation lesions that electrically isolate potential AF triggers 4. CA is a well-established treatment for AF symptoms 14, 15, that may also decrease the risk of stroke. Recent data showed a significant decrease in the relative risk of stroke and transient ischemic attack events among patients who underwent ablation compared with those undergoing antiarrhythmic drug therapy 16.
Since the left atrial appendage (LAA) is the source of thrombi in more than 90% of patients with non-valvular atrial fibrillation 17, another approach to stroke prevention is to physically block clots from exiting the LAA. One method for occluding the LAA is via percutaneous placement of the WATCHMAN LAA closure device. The WATCHMAN device resembles a small parachute. It consists of a nitinol frame covered by fabric polyethyl terephthalate that prevents emboli, but not blood, from exiting during the healing process. Fixation anchors around the perimeter secure the device in the LAA (Figure 1). To date, the WATCHMAN is the only implanted percutaneous device for which a randomized clinical trial has been reported. In this study, implantation of the WATCHMAN was found to be at least as effective as warfarin in preventing stroke (all-causes) and death (all-causes) 18. This device received the Conformit&eacute; Europ&eacute;enne (CE) mark for use in the European Union for warfarin eligible patients and in those who have a contraindication to anticoagulation therapy 19.
Given the proven effectiveness of CA to alleviate AF symptoms and the promising data with regard to reduction of thromboembolic events with both CA and WATCHMAN implantation, combining the two procedures is hoped to further reduce the incidence of stroke in high-risk patients while simultaneously relieving symptoms. The combined procedure may eventually enable patients to undergo implantation of the WATCHMAN device without subsequent warfarin treatment, since the CA procedure itself reduces thromboembolic events. This would present an avenue of treatment previously unavailable to patients ineligible for warfarin treatment because of recurrent bleeding 20 or other warfarin-associated problems.
The combined procedure is performed under general anesthesia with biplane fluoroscopy and TEE guidance. Catheter ablation is followed by implantation of the WATCHMAN LAA closure device. Data from a non-randomized trial with 10 patients demonstrates that this procedure can be safely performed in patients with a CHADS2 score of greater than 1 21. Further studies to examine the effectiveness of the combined procedure in reducing symptoms from AF and associated stroke are therefore warranted.

Catheter ablation is combined with placement of the WATCHMAN Left Atrial Appendage Closure Device to prevent ischemic stroke in a patient with non-valvular atrial fibrillation.

Atrial fibrillation (AF) is the most common sustained cardiac arrhythmia, affecting millions of individuals worldwide 1-3. The rapid, irregular, and ...

Implanting Glass Spinal Cord Windows in Adult Mice with Experimental Autoimmune Encephalomyelitis

Experimental autoimmune encephalomyelitis (EAE) in adult rodents is the standard experimental model for studying autonomic demyelinating diseases such as multiple sclerosis. Here we present a low-cost and reproducible glass window implantation protocol that is suitable for intravital microscopy and studying the dynamics of spinal cord cytoarchitecture with subcellular resolution in live adult mice with EAE. Briefly, we surgically expose the vertebrae T12-L2 and construct a chamber around the exposed vertebrae using a combination of cyanoacrylate and dental cement. A laminectomy is performed from T13 to L1, and a thin layer of transparent silicone elastomer is applied to the dorsal surface of the exposed spinal cord. A modified glass cover slip is implanted over the exposed spinal cord taking care that the glass does not directly contact the spinal cord. To reduce the infiltration of inflammatory cells between the window and spinal cord, anti-inflammatory treatment is administered every 2 days (as recommended by ethics committee) for the first 10 days after implantation. EAE is induced only 2-3 weeks after the cessation of anti-inflammatory treatment.
Using this approach we successfully induced EAE in 87% of animals with implanted windows and, using Thy1-CFP-23 mice (blue axons in dorsal spinal cord), quantified axonal loss throughout EAE progression. Taken together, this protocol may be useful for studying the recruitment of various cell populations as well as their interaction dynamics, with subcellular resolution and for extended periods of time. This intravital imaging modality represents a valuable tool for developing therapeutic strategies to treat autoimmune demyelinating diseases such as EAE.

We describe a method for implanting and maintaining glass windows over the exposed spinal cords of adult mice for studying experimental autoimmune encephalomyelitis. These windows provide chronic optical access to the spinal cord for monitoring cell dynamics at the subcellular level in live animals using two-photon microscopy.

Experimental autoimmune encephalomyelitis (EAE) in adult rodents is the standard experimental model for studying autonomic demyelinating diseases such as ...

Robotic Ablation of Atrial Fibrillation

Background: Pulmonary vein isolation (PVI) is an established treatment for atrial fibrillation (AF). During PVI an electrical conduction block between pulmonary vein (PV) and left atrium (LA) is created. This conduction block prevents AF, which is triggered by irregular electric activity originating from the PV. However, transmural atrial lesions are required which can be challenging. Re-conduction and AF recurrence occur in 20 - 40% of the cases. Robotic catheter systems aim to improve catheter steerability. Here, a procedure with a new remote catheter system (RCS), is presented. Objective of this article is to show feasibility of robotic AF ablation with a novel system. Materials and Methods: After interatrial trans-septal puncture is performed using a long sheath and needle under fluoroscopic guidance. The needle is removed and a guide wire is placed in the left superior PV. Then an ablation catheter is positioned in the LA, using the sheath and wire as guide to the LA. LA angiography is performed over the sheath. A circular mapping catheter is positioned via the long sheath into the LA and a three-dimensional (3-D) anatomical reconstruction of the LA is performed. The handle of the ablation catheter is positioned in the robotic arm of the Amigo system and the ablation procedure begins. During the ablation procedure, the operator manipulates the ablation catheter via the robotic arm with the use of a remote control. The ablation is performed by creating point-by-point lesions around the left and right PV ostia. Contact force is measured at the catheter tip to provide feedback of catheter-tissue contact. Conduction block is confirmed by recording the PV potentials on the circular mapping catheter and by pacing maneuvers. The operator stays out of the radiationfield during ablation. Conclusion: The novel catheter system allows ablation with high stability on low operator fluoroscopy exposure.

Pulmonary vein isolation (PVI) with an ablation catheter is a curative treatment for atrial fibrillation (AF). Robotic catheter systems aim to improve catheter steerability. Here, a procedure with a new robotic catheter system is presented. The goal of the procedure is electrical block between pulmonary vein and left atrium.

Background: Pulmonary vein isolation (PVI) is an established treatment for atrial fibrillation (AF). During PVI an electrical conduction block between ...

Isolation and Cultivation of Adult Rat Cardiomyocytes

In an intact heart, adjacent cells influence adult cardiomyocytes. With the method of isolation and cultivation of adult cardiomyocytes, a precise investigation of the behavior of these cells under specific treatments and environments is possible. This manuscript presents a protocol for successful isolation and cultivation of adult rat ventricular cardiomyocytes (ARVC).
The rat is sacrificed by cervical dislocation under deep anesthesia. Then, the heart is extracted and the aorta is uncovered. Subsequently, perfusion on the Langendorff perfusion system with calcium depletion and collagenase treatment is performed. Afterwards, ventricular tissue gets minced, re-circulated, and filtered, followed by three centrifugation steps with gradual addition of CaCl2 until physiological calcium concentration is reached. ARVC are plated on cell culture dishes. After refreshing the cell culture medium, ARVC can be cultivated for up to six days without changing the serum-containing culture medium. Isolation of ARVC is a calcium sensitive process. Small changes in the intracellular calcium concentration cause a decrease in the quality and viability of the isolated cells.
Freshly isolated ARVC are rod shaped. Within the first days of cultivation they lose the rod-shaped morphology and form pseudopodia-like structures (spreading). During this morphological formation ARVC initially degrade their contractile elements followed by a reformation through actin stress fibers and de novo sarcomerogenesis. After one week of cultivation, most ARVC show a widespread appearance with a clearly detectable cross striation. This process is sensitive to intracellular calcium concentration, as treatment with ionomycin attenuates spreading. Key markers in this process of de- and re-differentiation are &#946;-myosin heavy chain (&#946;-MHC), oncostatin M (OSM), and swiprosin-1 (EFHD2). Recent studies have suggested that cardiac re- and de-differentiation occurring under culture conditions mimics features seen in vivo during cardiac remodeling. Therefore, isolation and cultivation of ARVC play a key role in understanding the biology of cardiomyocytes.

Here, we present a protocol for the isolation and cultivation of adult rat ventricular cardiomyocytes (ARVC). Isolated ARVC can be used for short and long-term cultivation. The isolation and cultivation of ARVC can play a key role in developing new treatment regimens for cardiac diseases.

In an intact heart, adjacent cells influence adult cardiomyocytes. With the method of isolation and cultivation of adult cardiomyocytes, a precise ...

Isolation of Atrial Cardiomyocytes from a Rat Model of Metabolic Syndrome-related Heart Failure with Preserved Ejection Fraction

In this article, we describe an optimized, Langendorff-based procedure for the isolation of single-cell atrial cardiomyocytes (ACMs) from a rat model of metabolic syndrome (MetS)-related heart failure with preserved ejection fraction (HFpEF). The prevalence of MetS-related HFpEF is rising, and atrial cardiomyopathies associated with atrial remodeling and atrial fibrillation are clinically highly relevant as atrial remodeling is an independent predictor of mortality. Studies with isolated single-cell cardiomyocytes are frequently used to corroborate and complement in vivo findings. Circulatory vessel rarefication and interstitial tissue fibrosis pose a potentially limiting factor for the successful single-cell isolation of ACMs from animal models of this disease.
We have addressed this issue by employing a device capable of manually regulating the intraluminal pressure of cardiac cavities during the isolation procedure, substantially increasing the yield of morphologically and functionally intact ACMs. The acquired cells can be used in a variety of different experiments, such as cell culture and functional Calcium imaging (i.e., excitation-contraction-coupling).
We provide the researcher with a step-by-step protocol, a list of optimized solutions, thorough instructions to prepare the necessary equipment, and a comprehensive troubleshooting guide. While the initial implementation of the procedure might be rather difficult, a successful adaptation will allow the reader to perform state-of-the-art ACM isolations in a rat model of MetS-related HFpEF for a broad spectrum of experiments.

Here, we describe an optimized, Langendorff-based procedure for the isolation of single-cell atrial cardiomyocytes from a rat model of metabolic syndrome-related heart failure with preserved ejection fraction. A manual regulation of intraluminal pressure of cardiac cavities is implemented to yield functionally intact myocytes suitable for excitation-contraction-coupling studies.

In this article, we describe an optimized, Langendorff-based procedure for the isolation of single-cell atrial cardiomyocytes (ACMs) from a rat model of ...

Ultrasonic-augmented Primary Adult Fibroblast Isolation

Primary adult fibroblasts have become an important tool to study fibrosis, fibroblast interactions and inflammation in all body tissues. Since primary fibroblasts cannot divide indefinitely due to myofibroblast differentiation or senescence induction, new cultures must be established regularly. However, there are several obstacles to overcome during the processes of developing a reliable isolation protocol and primary fibroblast isolation itself: the method&#8217;s degree of difficulty (especially for beginners), the risk of bacterial contamination, the required time until primary fibroblasts can be used for experiments, and subsequent cell quality and viability. In this study, a fast, reliable and easy-to-learn protocol to isolate and culture primary adult fibroblasts from mouse heart, lung, liver and kidney combining enzymatic digestion and ultrasonic agitation is provided.

We present a protocol to isolate primary adult fibroblasts in an easy, fast and reliable way, performable by beginners (e.g., students). The procedure combines enzymatic tissue digestion and mechanical agitation with ultrasonic waves to obtain primary fibroblasts. The protocol can easily be adapted to specific experimental requirements (e.g., human tissue).

Primary adult fibroblasts have become an important tool to study fibrosis, fibroblast interactions and inflammation in all body tissues. Since primary ...

Echocardiographic Evaluation of Atrial Communications before Transcatheter Closure

Transthoracic (TTE) and transesophageal echocardiography (TEE) is the standard imaging method for atrial septal defect (ASD) and patent foramen ovale (PFO) detection, for patient selection for transcatheter ASD/PFO closure,&#160;for intraoperative guidance and for long-term follow-up. The size, shape, location and the number of the atrial communications schould be determined. The accuracy of PFO detection can be improved by using agitated saline together with maneuvers to transiently increase the right atrial (RA) pressure. The appearance of microbubbles in the left atrium (LA) within 3 cardiac cycles after opacification of the RA is considered positive for the presence of an intracardiac shunt. Three dimensional TEE identifies further septal fenestrations and describes the dynamic morphology of ASD/PFO and atrial septal aneurysm. Follow-up evaluations with TTE is recommended at 1, 6, and 12 months after the procedure, with a subsequent evaluation every year. Previous studies showed an increased incidence of atrial arrhythmias early after device closure. Speckle tracking analysis may help to understand functional left atrial remodeling following percutaneous closure and its impact on atrial arrhythmias.

Transthoracic (TTE) and transesophageal (TEE) echocardiography represent the basic imaging tools for interatrial septum examination. Three dimensional (3D) TEE provides incremental information in the assessment of the interatrial septum. Further advanced echocardiography techniques using speckle tracking echocardiography are applied for sensitive volumetric and functional assessment of the heart chambers.&#160;

Transthoracic (TTE) and transesophageal echocardiography (TEE) is the standard imaging method for atrial septal defect (ASD) and patent foramen ovale ...

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 ...

Sterile Pericarditis in Aachener Minipigs As a Model for Atrial Myopathy and Atrial Fibrillation

Atrial fibrillation (AF) is the most common arrhythmia caused by structural remodeling of the atria, also called atrial myopathy. Current therapies only target the electrical abnormalities and not the underlying atrial myopathy. For the development of novel therapies, a reproducible large animal model of atrial myopathy is necessary. This paper presents a model of sterile pericarditis-induced atrial myopathy in Aachener minipigs. Sterile pericarditis was induced by spraying sterile talcum and leaving a layer of sterile gauze over the atrial epicardial surface. This led to inflammation and fibrosis, two crucial components of the pathophysiology of atrial myopathy, making the atria susceptible to the induction of AF. Two pacemaker electrodes were positioned epicardially on each atrium and connected to two pacemakers from different manufacturers. This strategy allowed for repeated non-invasive atrial programmed stimulation to determine the inducibility of AF at specified time points after surgery. Different protocols to test AF inducibility were used. The advantages of this model are its clinical relevance, with AF inducibility and the rapid induction of inflammation and fibrosis-both present in atrial myopathy-and its reproducibility. The model will be useful in the development of novel therapies targeting atrial myopathy and AF.

We describe a sterile pericarditis model in minipigs to study atrial myopathy and atrial fibrillation (AF). We present surgical and anesthetic techniques, strategies for vascular access, and a protocol to study the inducibility of AF.

Atrial fibrillation (AF) is the most common arrhythmia caused by structural remodeling of the atria, also called atrial myopathy. Current therapies only ...