We thank Dr. Christian Rickert for critical reading of the manuscript. This work was supported by a grant from the National Heart Lung and Blood Institute (R01-HL088427) to CP. EJS was supported by 5T32-AG000279 from the National Institute on Aging. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

All animal procedures were performed in accordance with protocols approved by the Institutional Animal Care and Use Committee of the University of Colorado Anschutz Medical Campus. The standard protocol below has been optimized using male C57BL/6J mice of 2-3 months of age.
1. Prepare Solution Stocks and Supplies in Advance of Experiments
NOTE: Refer to Materials Table for necessary equipment and supplies.
<ol>
	<li>Prepare 1 L each of the following solutions as indicated in Table 1: Complete Tyrode&#39;s, Low Ca2+/Mg2+ Tyrode&#39;s, Modified Kraft-Br&#252;he (KB) Solution, and Bovine Serum Albumin (BSA) Solution. Use ultrapure filtered deionized water for all solutions. Divide each solution into 50 ml aliquots and store at 20 &#176;C for up to six months. Thaw individual aliquots immediately before experiments, and store for up to one week at 4 &#176;C.</li>
	<li>Prepare 50 ml of 10 mM NaCl and 1.8 mM CaCl2 Adaptation Solution by dissolving NaCl and CaCl2 in ultrapure filtered deionized water (Table 1). Store at room temperature for up to six months.</li>
	<li>Prepare 4.75 enzyme activity unit (U) aliquots of elastase by pipetting into microfuge tubes. Store at 4 &#176;C for up to three months.</li>
	<li>Prepare 375 &#956;g aliquots (in ultrapure H2O) of collagenase-protease enzyme blend by pipetting into microfuge tubes. Store at -20 &#176;C for up to three months.</li>
	<li>Prepare two fire-polished Pasteur pipettes, one for tissue transfer (~1.5 mm final opening diameter; Figure 1Aiv and Figure 1C) and one for dissociation (~2 mm diameter; Figure 1Aiii and Figure 1C).
	<ol>
		<li>Score Pasteur pipettes with a glass cutter and snap along the score to produce an opening slightly larger than the desired size. Fire-polish the cut end of each pipette for ~30-60 sec over a low flame on a Bunsen burner to produce a thick polished wall and an opening of the desired diameter. Ensure the fire-polished opening is free of any cracks or rough edges.</li>
	</ol>
	</li>
	<li>Prepare two dissection dishes by adding ~25 ml silicone elastomer mixed according to manufacturer&#39;s directions to each 100 mm Petri dish (Figure 1Ai and Figure 1B). Allow to cure at room temperature for 48 hr.</li>
	<li>For culture only: prepare 25-50 ml each Plating Medium and Culture Medium as per Table 2. Store for up to two weeks at 4 &#176;C.</li>
</ol>
2. Prepare Solutions to Be Used on Day of Cell Isolation
NOTE: The following amounts are for isolation of sinoatrial myocytes from one mouse.
<ol>
	<li>Add 2.5 ml of Low Ca2+/Mg2+ Tyrode&#39;s (pH 6.9) to each of three small, round-bottomed culture tubes. Place tubes in 35 &#177; 1 &#176;C water bath.</li>
	<li>Add 2.5 ml of Low Ca2+/Mg2+ Tyrode&#39;s to one large round-bottomed culture tube. To this tube, add 1 aliquot (4.75 U) elastase and one aliquot (375 &#956;g) collagenase-protease enzyme blend. Swirl to mix. Place tube in 35 &#177; 1 &#176;C water bath.</li>
	<li>Add 2.5 ml of KB Medium to three additional small, round-bottomed culture tubes and one additional large, round-bottomed culture tube. Place tubes in 35 &#177; 1 &#176;C water bath.</li>
	<li>Add ~7 ml of BSA solution to one large bottomed culture tube. Keep this tube at room temperature.</li>
	<li>Place 20-40 ml of Complete Tyrode&#39;s solution in a 50 ml beaker (Figure 1Aii). Add 10 USP/ml heparin, swirl to mix, and place the beaker in the 35 &#177; 1 &#176;C water bath.</li>
</ol>
3. Prepare Additional Solutions and Materials for Cultured Cells (Skip These Steps for Acutely Isolated Cells)
<ol>
	<li>In a sterile tissue culture hood, place two 12 mm round glass coverslips per mouse into individual wells of a 24-well plate. 
	NOTE: The 24 well plate is used because the wells are a convenient size, which serves to limit the volume for subsequent viral infections.</li>
	<li>Pipette approximately 200 &#181;l of a 100 ng/ml solution of mouse laminin diluted in sterile phosphate buffered saline (PBS) onto each coverslip.</li>
	<li>Incubate coverslips with laminin for the duration of the isolation (at least 1 hr) in an incubator at 37 &#176;C.</li>
	<li>Pre-warm the Plating Medium and Culture Medium (from Step 1.7 and Table 3) to 37 &#176;C.</li>
</ol>
4. Sinoatrial Node Isolation
<ol>
	<li>In a chemical fume hood, place a mouse in one chamber of a two-chamber box and anesthetize with ~200 &#956;l liquid isoflurane introduced via a cotton swab into the other chamber. Confirm anesthesia (usually within ~30-60 sec) with a toe pinch. Euthanize mouse by cervical dislocation. 
	NOTE: An empty 1 ml pipette tip box can be used to form the two-chamber box; turn the rack upside down in the box to create the separate compartment for the isoflurane to prevent the mouse from contacting it directly.</li>
	<li>Remove fur from chest with scissors and transect rib cage to expose the chest cavity using external tissue forceps (Figure 1Aviii) and dissection scissors (Figure 1Avix). Bathe the chest cavity with ~2 ml warmed Complete Tyrode&#39;s with Heparin using a transfer pipette. 
	NOTE: Continue bathing chest cavity when necessary, do not allow the preparation to dry out.</li>
	<li>Under a dissecting microscope, carefully remove the lungs and thymus using internal scissors (Figure 1Avi) and dissection forceps (Figure 1Avii).</li>
	<li>While gently holding the apex of the heart with the internal dissection forceps, carefully cut the inferior vena cava and the aorta with the internal scissors to remove the heart from the chest cavity. Transfer the heart to one of the silicone dissection dishes and bathe with ~4 ml warmed heparinized Complete Tyrode&#39;s using the transfer pipette.</li>
	<li>Orient the heart such that the posterior vessels are visible and facing up, with the animal&#39;s right atrium on the experimenter&#39;s right and left atrium on the experimenter&#39;s left. Once oriented, immobilize the heart by pinning through the apex into the silicone dissection dish.</li>
	<li>Locate the groove between the ventricles and the atria (clear ring above the ventricles). </li>
	<li>Using the internal dissection scissors, make an incision at the groove, keeping closer to the ventricles than the atria. Flush the groove and incision with additional warmed heparinized Complete Tyrode&#39;s as needed to allow a clear view of the atria and the valves. Continue to cut along the groove to separate the atria from the ventricles.</li>
	<li>Transfer the atrial tissue to the second silicone dissection dish and bathe with ~3 ml warmed heparinized Complete Tyrode&#39;s.</li>
	<li>Orient the tissue so that the animal&#39;s right atrium is now on the experimenter&#39;s left, and the left atrium is on the right. 
	NOTE: The right atrium is more transparent, while the left atrium has more of a dark red tone.</li>
	<li>Pin the tissue through the inferior and superior vena cavae and the right and left atrial appendages, stretching the preparation gently. Remove any remaining fat or other tissue to allow a clear view of the preparation (be careful to not cut into the atrial wall, as the sinoatrial node is quite delicate and can be easily damaged).</li>
	<li>Open the anterior wall of the atria by cutting through the venae cavae. Re-position the pins as necessary to visualize the interatrial septum.</li>
	<li>Cut along the interatrial septum to remove the left atrium. Re-pin the preparation, stretching gently.</li>
	<li>Remove the right atrial appendage and free the sinoatrial node by cutting along the cristae terminalis, which appears as a dark orange streak bordering the atrial appendage.</li>
	<li>Re-pin the nodal tissue and cut it laterally (perpendicular to the crista terminalis) to produce three equally sized strips.</li>
</ol>
5. Sinoatrial Node Digestion
<ol>
	<li>Using the narrow fire-polished pipette (Figure 1Aiv), transfer the three strips of sinoatrial node tissue into the first of three small, round-bottomed tube containing 2.5 ml of low Ca2+/Mg2+ Tyrode&#39;s in the 35 &#177; 1 &#176;C water bath. Incubate for 5 min.</li>
	<li>Transfer tissue strips to the second small, round bottom tube containing 2.5 ml low Ca2+/Mg2+ Tyrode&#39;s in the 35 &#177; 1 &#176;C water bath, using the same narrow pipette. Wash the tissue strips by gentle swirling the tube or by gently pipetting with the narrow pipette. Do not invert the tube.</li>
	<li>Transfer tissue strips to the third small, round bottom tube containing 2.5 ml low Ca2+/Mg2+ Tyrode&#39;s, and repeat the washing step described in step 5.2.</li>
	<li>Transfer strips into the large round-bottomed tube containing 2.5 ml of low Ca2+/Mg2+ Tyrode&#39;s with enzymes (elastase plus collagenase-protease blend) in the 35 &#176;C water bath. Ensure that all three tissue strips are present. Incubate for 10-15 min at 35 &#177; 1 &#176;C. Mix every 5 min by gently swirling the tube. Do not invert the tube.</li>
</ol>
6. Sinoatrial Node Myocyte Dissociation
<ol>
	<li>Following the enzyme digestion, use the narrow fire-polished pipette to gently transfer the tissue strips to the first small, round-bottomed tube containing 2.5 ml KB solution at 35 &#177; 1 &#176;C. Wash tissue by gently swirling the tube. 
	Note: Tissue strips will appear somewhat translucent and may clump together at this point. Handle very gently after enzymatic digestion to avoid losing cells.</li>
	<li>Transfer the tissue to the second small round-bottomed tube containing 2.5 ml KB at 35 &#177; 1 &#176;C. Swirl gently to wash.</li>
	<li>Transfer the tissue to the third small round-bottomed tube containing 2.5 ml KB at 35 &#177; 1 &#176;C. Swirl gently to wash.</li>
	<li>Transfer the tissue to the large, round-bottomed tube containing 2.5 ml KB at 35 &#177; 1 &#176;C.</li>
	<li>Using the larger fire-polished pipette (Figure 1Aiii and Figure 1C), dissociate the cells in the large round-bottomed tube by constant trituration at approximately 0.5-1 Hz for 5-10 min, taking care to keep the dissociation tube submerged in the 35 &#177; 1 &#176;C water bath and to avoid introducing bubbles into the solution. 
	Note: Trituration time varies with diameter of the dissociation pipette and force of pipetting. Time should be adjusted so that the tissue pieces remaining at the end of the dissociation appear thin, transparent and wispy. If tissue retains any color, the dissociation is likely to be incomplete. Frequency (0.5-1 Hz) is determined by hand.</li>
	<li>Remove round-bottomed tube containing dissociated SAMs from the water bath and equilibrate at room temperature for 5 min.</li>
</ol>
7. Sinoatrial Node Calcium Re-adaptation (Performed at Room Temperature)
NOTE: For SAMs destined for culture experiments, the procedures in the following section should be performed in a sterile tissue culture hood. If SAMs are to be used for acute experiments, there is no need to perform these steps in a sterile environment.
<ol>
	<li>Add 75 &#181;l of NaCl/CaCl2 adaptation solution (Table 2). Swirl gently to mix and incubate for 5 min.</li>
	<li>Add 160 &#181;l of NaCl/CaCl2 adaptation solution. Swirl gently to mix and incubate for 5 min.</li>
	<li>Add 390 &#181;l of BSA solution (Table 2). Swirl gently to mix and incubate for 4 min.</li>
	<li>Add 1.25 ml of BSA solution. Swirl gently to mix and incubate for 4 min.</li>
	<li>Add 4.37 ml of BSA solution. Swirl gently to mix and incubate for 4 min. 
	Note: The final concentration of calcium will be 1.8 mM.</li>
	<li>Following calcium re-adaptation, collect SAMs by allowing to settle by gravity for ~10 min or by centrifugation at ~2,000 x g for 3 min.
	<ol>
		<li>For acutely isolated cells, gently remove and discard about 5 ml of the supernatant using a glass Pasteur pipette, leaving cells in a volume of ~2 ml. Store these cells at room temperature for up to ~8 hr for patch clamp recordings.</li>
		<li>For cultured cells, remove as much of the supernatant as possible using a sterile glass Pasteur pipette. Resuspend cell pellet in 1 ml pre-warmed (37 &#176;C) Plating Medium (Table 2).</li>
	</ol>
	</li>
</ol>
8. Plating and Culture of Sinoatrial Myocytes (Skip for Acutely Isolated Cells)
<ol>
	<li>Remove laminin solution from coverslips from Step 3.3 with a Pasteur pipette. Immediately seed 500 &#181;l (~50-100 cells) onto each laminin-coated coverslip (from step 3). 
	NOTE: The contractile inhibitor 2,3-butanedione monoxime (BDM) is included in the plating and culture media to prevent contraction, which causes attrition of cells9.</li>
	<li>Return the 24-well plate containing newly seeded SAMs to the incubator and maintain at 37 &#176;C in an atmosphere of 95% air/5% CO2. Allow cells to adhere to coverslips for 4-6 hr in plating media (Table 2).</li>
	<li>Gently remove Plating Medium using a sterile Pasteur pipette. Replace with 500 &#181;l per well of pre-warmed (37 &#176;C) Culture Medium (Table 2).</li>
</ol>
9. Adenoviral Transduction of Adult Sinoatrial Myocyte Cultures (Skip for Acutely Isolated Cells)
<ol>
	<li>Estimate number of cells per coverslip immediately before applying adenovirus by counting the cells in a field of view under a microscope, adjusting for magnification. Count all cells, not just SAMs.</li>
	<li>Dilute adenovirus in Medium 199 and adjust the dilution for the viral titer so that application of 1-10 &#956;l is required to achieve a final multiplicity of infection (MOI) of 100 viral units per cell. Add adenoviral solution in a dropwise fashion directly onto plated SAMs.</li>
	<li>Incubate cells with the virus-containing medium overnight (~12-14 hr). Then exchange with fresh culture medium. Maintain cells in incubator, changing culture medium every 48 hr, until desired protein expression is obtained.</li>
</ol>
10. Functional Evaluation of Acutely Isolated or Cultured SAMs
NOTE: The following protocol is an example of functional assessments of isolated SAMs using the amphotericin perforated-patch technique to record both spontaneous APs and If from the same cell (see reference9).
<ol>
	<li>Prepare recording solutions as described in Table 3.</li>
	<li>Prepare a stock solution of 20 mg/ml amphotericin-B in DMSO fresh on the day of recording. Keep stock at room temperature and protect from light. Dilute amphotericin-B stock in intracellular solution to a final concentration of 200 ug/ml just before use. Maintain the final pipette solution on ice and protect from light. 
	NOTE: The amphotericin-containing pipette solution for experiments should be prepared fresh hourly by diluting an aliquot of the stock solution into the intracellular solution and vortexing for at least 1 min.</li>
	<li>Transfer an aliquot of the acutely dissociated SAM cell suspension or a fragment of glass coverslip bearing cultured SAMs to a recording chamber containing Tyrode&#39;s solution at 35 &#177; 1 &#176;C. Perfuse cells with Tyrode&#39;s solution for at least 2 min prior to electrophysiological recordings to remove any residual BDM remaining from culture medium. 
	NOTE: Spontaneous contractions should be evident immediately upon transfer to the Tyrode&#39;s solution. SAMs for recording can be identified by a combination of features including spontaneous contractile activity, characteristic morphology (e.g., Figure 2), lack of striations, expression of HCN4 protein, presence of If current, membrane capacitance &#60;50 pF, and spontaneous APs with waveforms that include a diastolic depolarization phase and a slow upstroke.</li>
	<li>Using borosilicate glass pipettes with resistances of 1.5-3.0 M&#937;, fill the tip with intracellular solution lacking amphotericin by dipping for 10-30 sec. Then back-fill the pipette with the amphotericin-containing solution. A G&#937; cell-attached seal should be obtained as quickly as possible. If seal formation is difficult, increase the tip-fill time. 
	NOTE: The access resistance should be continuously monitored following formation of the cell-attached seal, and recordings should only be commenced after obtaining a stable access resistance of &#60;10 M&#8486;.</li>
	<li>To record spontaneous APs, switch the amplifier to fast current-clamp mode with no current injection. 
	NOTE: 1 nM isoproterenol is included in the extracellular Tyrode&#39;s solution when recording APs to stabilize the firing rate, as previously reported10.</li>
</ol>

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This paper presents detailed protocols for the isolation and culture of fully differentiated sinoatrial node myocytes from adult mice. The isolation protocol reliably produces spontaneously active mouse SAMs suitable for either immediate electrophysiological analysis or subsequent culture. Similar protocols have been reported by many other groups (for example, see references11,12,10,13-17). However, our protocol for maintaining adult mouse SAMs in vitro preserves the characteristic morphology, spontan...

Pacemaker myocytes in the sinoatrial node of the heart (sinoatrial myocytes, &#34;SAMs&#34;) generate spontaneous, rhythmic action potentials (APs) that propagate through the myocardium to initiate each heartbeat. Experiments using acutely isolated SAMs from many species have been essential for elucidation of mechanisms that contribute to the generation of pacemaker activity. SAMs are highly specialized cardiomyocytes that differ substantially from their counterparts in the atrial and ventricular myocardium in terms of morphology, function, and protein expression. The hallmark of spontaneous APs in SAMs is a spontaneous depolarization during diastole that drives the membrane potential to threshold to trigger the next AP1,2. This &#34;pacemaker potential&#34; depends on the coordinated activity of many different membrane currents including the &#34;funny current&#34; (If), T- and L-type calcium currents, and the sodium-calcium exchanger current (INCX), which is driven by Ca2+ release from the sarcoplasmic reticulum3,4.
While acutely isolated mouse SAMs are an essential experimental preparation for the study of pacemaking, the isolation of SAMs from mice can be a challenging method to adopt because the indistinct anatomy and small size of the mouse SAN requires a nuanced microdissection and the combined enzymatic and mechanical dissociation of the cells requires careful optimization.
We provide here a detailed video demonstration of a protocol that has been successfully used to isolate SAMs from adult mice for patch clamp recordings5-8. To our knowledge, there is no such visual demonstration available from any other source. In addition, a new method is demonstrated in which isolated SAMs from adult mice can be maintained in vitro for several days, thereby permitting the introduction of proteins, genetically encoded reporter molecules or RNAi via adenoviral infection9.

<table><tbody><tr><td>Sylgruard/Elastomer Kit</td><td>Dow Corning</td><td>184 SIL ELAST KIT 0.5KG</td><td></td></tr><tr><td>Borosilicate 9&quot; pasteur pipettes</td><td>Fisher Scientific</td><td>13-678-20C</td><td></td></tr><tr><td>Small, round bottomed culture tubes</td><td>Fisher Scientific</td><td>352059</td><td></td></tr><tr><td>Large, round bottomed culture tubes</td><td>Corning</td><td>14-959-11B</td><td></td></tr><tr><td>Elastase</td><td>Worthington Biochemical</td><td>LS002279</td><td></td></tr><tr><td>Liberase TM</td><td>Roche</td><td>5401119001</td><td>Tissue dissociation solution</td></tr><tr><td>Heparin</td><td>SAGENT Pharmaceuticals&amp;nbsp;</td><td>NDC 25021-400-10</td><td></td></tr><tr><td>Mouse Laminin</td><td>Corning</td><td>CB-354232</td><td></td></tr><tr><td>12 mm round glass coverslips</td><td>Fisher&amp;nbsp;</td><td>12-545-80</td><td></td></tr><tr><td>24-well culture plate</td><td>Fisher</td><td>08-772-1</td><td></td></tr><tr><td>Ad-mCherry</td><td>Vector Biolabs</td><td>1767</td><td></td></tr><tr><td>Ad-eGFP</td><td>Vector Biolabs</td><td>1060</td><td></td></tr><tr><td>Plastic, disposable transfer pipette</td><td>Fisher Scientific</td><td></td><td></td></tr><tr><td>Micro scissors</td><td>Fisher Scientific</td><td>17-467-496</td><td></td></tr><tr><td>Dumont #4 Forceps</td><td>Roboz Instruments</td><td>RS-4904</td><td></td></tr><tr><td>Tissue Forceps</td><td>Roboz Instruments</td><td>RS-8164</td><td></td></tr><tr><td>Dissecting Iris Scissors</td><td>WPI, Inc.</td><td>501264</td><td></td></tr><tr><td>Dissecting Pins</td><td>Fine Science Tools</td><td>26002-20</td><td></td></tr><tr><td>NaCl</td><td>Sigma</td><td>71376</td><td></td></tr><tr><td>KCl</td><td>Sigma</td><td>60128</td><td></td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>60353</td><td></td></tr><tr><td>HEPES</td><td>Sigma</td><td>54457</td><td></td></tr><tr><td>glucose</td><td>Sigma</td><td>G0350500</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma</td><td>M8266</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma</td><td>C1016</td><td></td></tr><tr><td>taurine</td><td>Sigma</td><td>T0625</td><td></td></tr><tr><td>BSA</td><td>Sigma</td><td>A2153</td><td></td></tr><tr><td>K-glutamate</td><td>Sigma</td><td>G1501</td><td></td></tr><tr><td>K-aspartate</td><td>Sigma</td><td>A6558</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>M7506</td><td></td></tr><tr><td>creatine</td><td>Sigma</td><td>C0780</td><td></td></tr><tr><td>EGTA</td><td>Sigma</td><td>E3889</td><td></td></tr><tr><td>Mg-ATP</td><td>Sigma</td><td>A9187</td><td></td></tr><tr><td>Amphotericin-B</td><td>Fisher Scientific</td><td>1397-89-3</td><td></td></tr><tr><td>Isoproterenol</td><td>Calbiochem</td><td>420355</td><td></td></tr><tr><td>Media199</td><td>Sigma</td><td>M4530</td><td></td></tr><tr><td>2,3-butanedione monoxime (BDM)</td><td>Sigma</td><td>B0753</td><td></td></tr><tr><td>Fetal Bovine Serum (FBS)</td><td>Sigma</td><td>SH30071</td><td></td></tr><tr><td>Bovine Serum Albumin (BSA)</td><td>Sigma</td><td>A5611</td><td></td></tr><tr><td>Insulin&amp;nbsp;&amp;nbsp;</td><td>Sigma</td><td>I3146</td><td></td></tr><tr><td>Transferrin</td><td>Sigma</td><td>I3146</td><td></td></tr><tr><td>Selenium</td><td>Sigma</td><td>I3146</td><td></td></tr><tr><td>Penicillin</td><td>GE Healthcare</td><td>SV30010</td><td></td></tr><tr><td>Streptomycin</td><td>Hyclone</td><td>SV30010</td><td></td></tr></tbody></table>

methods for the isolation, culture, and functional characterization of sinoatrial node myocytes from adult mice

Sinoatrial node myocytes (SAMs) act as the natural pacemakers of the heart, initiating each heart beat by generating spontaneous action potentials (APs). These pacemaker APs reflect the coordinated activity of numerous membrane currents and intracellular calcium cycling. However the precise mechanisms that drive spontaneous pacemaker activity in SAMs remain elusive. Acutely isolated SAMs are an essential preparation for experiments to dissect the molecular basis of cardiac pacemaking. However, the indistinct anatomy, complex microdissection, and finicky enzymatic digestion conditions have prevented widespread use of acutely isolated SAMs. In addition, methods were not available until recently to permit longer-term culture of SAMs for protein expression studies. Here we provide a step-by-step protocol and video demonstration for the isolation of SAMs from adult mice. A method is also demonstrated for maintaining adult mouse SAMs in vitro and for expression of exogenous proteins via adenoviral infection. Acutely isolated and cultured SAMs prepared via these methods are suitable for a variety of electrophysiological and imaging studies.

The protocols described here have been previously employed to isolate spontaneously active SAMs from adult mice that are suitable for a variety of different patch clamp studies5-8. In addition, the protocols allow for isolated SAMs that can be maintained in culture for up to one week. Gene transfer into the cultured cells can be accomplished via adenoviral infection9. The results presented in this section derive from our previous work and are shown here as examples o...

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Methods for the Isolation, Culture, and Functional Characterization of Sinoatrial Node Myocytes from Adult Mice

Methods are demonstrated for the isolation of sinoatrial node myocytes (SAMs) from adult mice for patch clamp electrophysiology or imaging studies. Isolated cells can be used directly or can be maintained in culture to permit expression of proteins of interest, such as genetically encoded reporters.

Sinoatrial node myocytes (SAMs) act as the natural pacemakers of the heart, initiating each heart beat by generating spontaneous action potentials (APs). ...

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JoVE Journal

Biology

13.1K Views.  University of Colorado Anschutz Medical Campus. Methods are demonstrated for the isolation of sinoatrial node myocytes (SAMs) from adult mice for patch clamp electrophysiology or imaging studies. Isolated cells can be used directly or can be maintained in culture to permit expression of proteins of interest, such as genetically encoded reporters.

Video: Methods for the Isolation, Culture, and Functional Characterization of Sinoatrial Node Myocytes from Adult Mice

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 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 ...

Medicine

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 and Culture of Resident Cardiac Macrophages from the Murine Sinoatrial and Atrioventricular Node

Resident cardiac macrophages have been demonstrated to facilitate the electrical conduction in the heart. The physiologic heart rhythm is initiated by electrical impulses generated in sinoatrial node (SAN) and then conducted to ventricles via atrioventricular node (AVN). To further study the role of resident macrophages in cardiac conduction system, a proper isolation of resident macrophages from SAN and AVN is necessary, but it remains challenging. Here, we provide a protocol for the reliable microdissection of the SAN and AVN in murine hearts followed by the isolation and culture of resident macrophages.
Both, SAN which is located at the junction of the crista terminalis with the superior vena cava, and AVN which is located at the apex of the triangle of Koch, are identified and microdissected. Correct location is confirmed by histologic analysis of the tissue performed with Masson's trichrome stain and by anti-HCN4.
Microdissected tissues are then enzymatically digested to obtain single cell suspensions followed by the incubation with a specific panel of antibodies directed against cell-type specific surface markers. This allows to identify, count, or isolate different cell populations by fluorescent activated cell sorting. To differentiate cardiac resident macrophages from other immune cells in the myocardium, especially recruited monocyte-derived macrophages, a delicate devised gating strategy is needed. First, lymphoid lineage cells are detected and excluded from further analysis. Then, myeloid cells are identified with resident macrophages being determined by high expression of both CD45 and CD11b, and low expression of Ly6C. With cell sorting, isolated cardiac macrophages can then be cultivated in vitro over several days for further investigation. We, therefore, describe a protocol to isolate cardiac resident macrophages located within the cardiac conduction system. We discuss pitfalls in microdissecting and digesting SAN and AVN, and provide a gating strategy to reliably identify, count and sort cardiac macrophages by fluorescence-activated cell sorting.

The protocol presented here provides a step-by-step approach for the isolation of cardiac resident macrophages from the sinoatrial node (SAN) and atrioventricular node (AVN) region of mouse hearts.

Resident cardiac macrophages have been demonstrated to facilitate the electrical conduction in the heart. The physiologic heart rhythm is initiated by ...

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 ...

Microelectrode Array Recording of Sinoatrial Node Firing Rate to Identify Intrinsic Cardiac Pacemaking Defects in Mice

The sinoatrial node (SAN), located in the right atrium, contains the pacemaker cells of the heart, and dysfunction of this region can cause tachycardia or bradycardia. Reliable identification of cardiac pacemaking defects requires the measurement of intrinsic heart rates by largely preventing the influence of the autonomic nervous system, which can mask rate deficits. Traditional methods to analyze intrinsic cardiac pacemaker function include drug-induced autonomic blockade to measure in vivo heart rates, isolated heart recordings to measure intrinsic heart rates, and sinoatrial strip or single-cell patch-clamp recordings of sinoatrial pacemaker cells to measure spontaneous action potential firing rates. However, these more traditional techniques can be technically challenging and difficult to perform. Here, we present a new methodology to measure intrinsic cardiac firing rate by performing microelectrode array (MEA) recordings of whole-mount sinoatrial node preparations from mice. MEAs are composed of multiple microelectrodes arranged in a grid-like pattern for recording in vitro extracellular field potentials. The method described herein has the combined advantage of being relatively faster, simpler, and more precise than previous approaches for recording intrinsic heart rates, while also allowing easy pharmacological interrogation.

This protocol aims to describe a new methodology to measure intrinsic cardiac firing rate using microelectrode array recording of the whole sinoatrial node tissue to identify pacemaking defects in mice. Pharmacological agents can also be introduced in this method to study their effects on intrinsic pacemaking.

The sinoatrial node (SAN), located in the right atrium, contains the pacemaker cells of the heart, and dysfunction of this region can cause tachycardia or ...

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 ...

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.

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

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

Mouse models play a crucial role in arrhythmia research and allow studying key mechanisms of arrhythmogenesis including altered ion channel function and calcium handling. For this purpose, atrial or ventricular cardiomyocytes of high quality are necessary to perform patch-clamp measurements or to explore calcium handling abnormalities. However, the limited yield of high-quality cardiomyocytes obtained by current isolation protocols does not allow both measurements in the same mouse. This article describes a method to isolate high-quality murine atrial and ventricular myocytes via retrograde enzyme-based Langendorff perfusion, for subsequent simultaneous measurements of calcium transients and L-type calcium current from one animal. Mouse hearts are obtained, and the aorta is rapidly cannulated to remove blood. Hearts are then initially perfused with a calcium-free solution (37 &#176;C) to dissociate the tissue at the level of intercalated discs and afterwards with an enzyme solution containing little calcium to disrupt extracellular matrix (37 &#176;C). The digested heart is subsequently dissected into atria and ventricles. Tissue samples are chopped into small pieces and dissolved by carefully pipetting up and down. The enzymatic digestion is stopped, and cells are stepwise reintroduced to physiologic calcium concentrations. After loading with a fluorescent Ca2+-indicator, isolated cardiomyocytes are prepared for simultaneous measurement of calcium currents and transients. Additionally, isolation pitfalls are discussed and patch-clamp protocols and representative traces of L-type calcium currents with simultaneous calcium transient measurements in atrial and ventricular murine myocytes isolated as described above are provided.

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

Mouse models allow studying key mechanisms of arrhythmogenesis. For this purpose, high quality cardiomyocytes are necessary to perform patch-clamp measurements. Here, a method to isolate murine atrial and ventricular myocytes via retrograde enzyme-based Langendorff perfusion, which allows simultaneous measurements of calcium-transients and L-type calcium current, is described.

Mouse models play a crucial role in arrhythmia research and allow studying key mechanisms of arrhythmogenesis including altered ion channel function and ...

Methods for the Isolation, Culture, and Functional Characterization of Sinoatrial Node Myocytes from Adult Mice

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Isolation, Culture, and Functional Characterization of Adult Mouse Cardiomyoctyes

Isolation of Atrial Myocytes from Adult Mice

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

Isolation and Culture of Resident Cardiac Macrophages from the Murine Sinoatrial and Atrioventricular Node

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

Microelectrode Array Recording of Sinoatrial Node Firing Rate to Identify Intrinsic Cardiac Pacemaking Defects in Mice

Isolation, Culture and Transduction of Adult Mouse Cardiomyocytes

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

An Antegrade Perfusion Method for Cardiomyocyte Isolation from Mice

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