In addition, we thank the cardiac surgeons at Heidelberg University for the provision of human atrial tissue and Claudia Liebetrau, Katrin Kupser and Ramona Nagel for their excellent technical support. Special thanks also to Andy W. Trafford for his helpful suggestions and advice during the establishment of the Ca2+ transient measurements. The authors wish to express their deepest gratitude to the members of the Department of Pharmacology and Toxicology (head: Ursula Ravens) of the Dresden University of Technology for the opportunity they offered us to learn basic techniques and skills in cellular electrophysiology and cardiomyocyte isolation.
The authors' research is supported by the German Research Foundation (Do769/1-1-3), the German Federal Ministry of Education and Research through the Atrial Fibrillation Competence Network (01Gi0204) and the German Centre for Cardiovascular Research, the European Union through the European Network for Translational Research in Atrial Fibrillation (EUTRAF, FP7-HEALTH-2010, large-scale integrating project, Proposal No. 261057) and the European-North American Atrial Fibrillation Research Alliance (ENAFRA) grant of Fondation Leducq (07CVD03).
The schematic overview shown in the video file was produced using Servier medical art.

Experimental protocols need to be approved by the local ethics committee and all patients need to give written informed consent. Our research was approved by the ethics committee of the Medical Faculty Mannheim, University of Heidelberg (#2011-216N-MA) and was performed in compliance with all institutional, national and international guidelines for human welfare. All patients gave written informed consent.
0. Obtaining Human Atrial Tissue
During routine cannulation procedures in patients undergoing open-heart surgery for cardiopulmonary bypass grafting, the tip of the right atrial appendage is usually removed and can be used for isolation of atrial cardiomyocytes. After excision the tissue sample is transferred immediately into a 50 ml Falcon tube with sterile Ca2+-free transport solution containing 2,3-butanedione monoxime (Table I; BDM, contractile inhibitor, preventing myocyte contracture). With transport times between 30 and 45 min, we did not recognize a clear advantage of cooling or oxygenation with respect to both number and quality of isolated atrial cardiomyocytes. In general transport to the lab should be as quick as possible. However, if longer transportation time cannot be avoided, transport at 4 &deg;C in oxygenated solution might be advantageous.
1. Prearrangements
<ol>
	<li>Switch on the thermocirculator, which controls the temperature of the jacketed beaker (Table II). Make sure the temperature within the beaker equals 37 &deg;C. Cover the beaker with a glass lid to maintain a constant temperature within the beaker throughout the whole isolation process.</li>
	<li>Weigh the Collagenase I and Protease XXIV into three glass beakers and store it at room temperature. The enzymes will be diluted just before use.</li>
	<li>Store 20 ml of Ca2+ free solution in a Petri dish at 4 &deg;C.</li>
	<li>Store 250 ml of Ca2+ free solution in a water bath at 37 &deg;C.</li>
</ol>
2. Cleaning of the Tissue
<ol>
	<li>Transfer the tissue sample together with the transport solution into a Petri dish (~10 cm diameter) and remove fatty tissue roughly using scissors.</li>
	<li>Weigh the tissue sample. Between 200 and 600 mg are used for cell isolation. The remaining tissue can be frozen in liquid nitrogen for later biochemical analysis.</li>
	<li>Transfer the tissue sample into the Petri dish containing 20 ml Ca2+ free solution at 4 &deg;C (see step 1.3) and chop the tissue sample into small chunks of approximately 1 mm3 in size.</li>
	<li>The following steps should be executed at 37 &deg;C and under continuous gassing with 100% O2. A pipette tip can be placed onto the oxygen tube to avoid strong bubbling and generate continuous air flow.</li>
	<li>Transfer the tissue chunks together with the Ca2+-free solution into the jacketed beaker and stir carefully for about 3 min with a magnetic stirring bar.</li>
	<li>Stop stirring and allow the tissue chunks to settle down for a few seconds. Carefully strain the supernatant through a nylon mesh (200 &mu;m, Table II). Return the restrained tissue chunks from the mesh into the beaker using forceps.</li>
	<li>Refill the beaker with Ca2+ free solution and repeat the washing step 2.4 and 2.5 two times.</li>
</ol>
3. First Enzymatic Digestion
<ol>
	<li>Re-suspend the tissue chunks with 20 ml Enzyme solution E1 (Table I) containing collagenase and protease and stir carefully for 10 min.</li>
	<li>Add 40 &mu;l of 10 mM CaCl2-solution to obtain a final concentration of 20 &mu;M Ca2+.</li>
	<li>After 35 min strain the supernatant carefully through a nylon mesh (200 &mu;m, Table II) in a way that most of the tissue chunks remain in the beaker. Return restrained tissue chunks from the mesh into the beaker.</li>
</ol>
4. Second Enzymatic Digestion
<ol>
	<li>Resuspend the tissue chunks again with 20 ml enzyme solution E2 containing collagenase I only (Table I). Add 40 &mu;l of 10 mM CaCl2-solution immediately to obtain a final concentration of 20 &mu;M Ca2+.</li>
	<li>During the second digestion use scissors to further chop tissue clots occasionally.</li>
	<li>After 5 min take a sample using a Pasteur pipette to check the dissociation of cells. Repeat this every 2-3 min until rod-shaped, striated cardiomyocytes appear.</li>
	<li>Stop stirring and allow the tissue chunks to settle down for about 20-30 sec.</li>
	<li>Strain the supernatant carefully through a nylon mesh (200 &mu;m, Table II) into a 50 ml Falcon tube (Tube A). Return the restrained tissue chunks from the mesh into the beaker.</li>
	<li>Re-suspend the tissue chunks in the beaker with 20 ml storage solution (Table I)10 and further dissociate the cells by gentle mechanical trituration using a 20 ml serological pipette with dispenser. Try to avoid bubble formation since bubbles are detrimental to cell survival and quality.</li>
	<li>Strain the supernatant carefully through a nylon mesh (200 &mu;m, Table II) into a 50 ml Falcon Tube (Tube B).</li>
</ol>
5. Final Preparation and Adjustment of Final Ca2+ Concentration
<ol>
	<li>Centrifuge both Falcon tubes A and B (see step 4.5 and 4.7) at 95 x g for 10 min.</li>
	<li>Remove the supernatant from both tubes carefully by aspiration. Make sure not to disturb the pellet. Discard the supernatant.</li>
	<li>Re-suspend both pellets in 1.5 ml storage solution (Table I) each (room temperature).</li>
	<li>Add two times 7.5 &mu;l of 10 mM CaCl2 solution to each Falcon and stir carefully. Incubate for 10 min after each step.</li>
	<li>Add 15 &mu;l of 10 mM CaCl2 solution to obtain a final Ca2+ concentration of 0.2 mM.</li>
</ol>
6. Loading of Myocytes with the Fluorescent Ca2+-indicator Fluo-3 AM (Figure 1)
<ol>
	<li>Transfer 1.5 ml of cell suspension (tube A and/or tube B) into a 2 ml microcentrifuge tube (Eppendorf tube).</li>
	<li>The following steps should be executed under consideration of the light sensitivity of the fluorescent Ca2+ Indicator Fluo-3.</li>
	<li>Dissolve 50 &mu;g of the membrane permeable acetoxymethyl ester derivative of Fluo-3 (Fluo-3 AM, Table III) in 44 &mu;l of the Pluronic F-127 (Table III) stock solution (20% w/v in anhydrous DMSO) to get a 1 mM Fluo-3 AM stock solution, which can be stored at -20 &deg;C for at most 1 week.</li>
	<li>Add 15 &mu;l of the Fluo-3 AM stock solution to the microcentrifuge tube containing 1.5 ml of cell suspension (see step 6.1) and agitate carefully.</li>
	<li>Incubate the cell suspension for 10 min in an optically opaque box.</li>
	<li>Briefly centrifuge at about 6,000 rpm.</li>
	<li>Discard the supernatant and re-suspend the pellet in 1.5 ml bath solution (Table IV).</li>
	<li>Leave the cell suspension for about 30 min for de-esterification before beginning with experiments.</li>
</ol>
7. Simultaneous Patch-clamp and Epifluorescent Ca2+ Measurements
Since patch-clamp measurements are not the major topic of this review, we refer the interested reader to other publications providing a more in depth description of this technique.11-14 For the sake of completeness we provide a brief summary of a protocol to measure action potentials or L-type Ca2+ currents, both together with simultaneous Ca2+-transient recordings.
During experiments myocytes are superfused at 37 &deg;C with bath solution (Table IV) using a rapid perfusion system (Octaflow IITM, ALA Scientific Instruments, NY). For voltage-clamp experiments, K+ currents are blocked by adding 4-aminopyridine (5 mmol/L) and BaCl2 (0.1 mmol/L) to the bath solution. Borosilicate glass microelectrodes are used and should have tip resistances of 2-5 M&Omega; when filled with pipette solution (Table V). In addition to the Fluo-3 AM loading of the myocytes (see step 6), Fluo-3 is also included in the pipette solution (Table V). Fluorescence is excited at 488 nm and emitted light (&lt;520 nm) converted to [Ca2+]i assuming
<img alt="Equation 1" fo:content-width="1.540in" fo:src="/files/ftp_upload/50235/50235eq1highres.jpg" src="/files/ftp_upload/50235/50235eq1.jpg" />
 
where kd=dissociation constant of Fluo-3 (864 nM), F=Fluo-3 fluorescence; Fmax=Ca2+-saturated fluorescence obtained at the end of each experiment.12 Both electrical signals and epifluorescent Ca2+ signals are recorded simultaneously. Action potentials are stimulated at 0.5 Hz in current-clamp mode using 1 msec current pulses of 1.2x threshold strength. L-type Ca2+-currents are measured in voltage-clamp mode using a holding potential of -80 mV and a 100-msec ramp-pulse to -40 mV to inactivate the fast Na+-current, followed by a 100-msec test-pulse to +10 mV at 0.5 Hz.

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Worthington Biochemical Corp. supported this publication by covering production costs of video file. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Here we describe a method for isolation of human atrial myocytes from right atrial appendages obtained from patients undergoing open heart surgery. In order to use these myocytes for measurements of cytosolic Ca2+ we adapted a previously described method4-11 by omitting EGTA from the storage solution.
Already in 1970 it was observed that although myocytes dissociate in the presence of Ca2+ during digestion, all of them were in contracture and non-viable.16,17 Therefore, cell isolation is performed in Ca2+-free solution. However, the re-introduction of physiological concentrations of Ca2+ resulted in rapid Ca2+ influx and cell death. This has been described as the Ca2+ paradox phenomenon which was originally observed in perfused hearts by Zimmerman and Hulsman.18 Modifications of the isolation media including reduction of the pH to 7.0,19 addition of taurin20 or of small amounts of Ca2+ (see step 3.2 and 4.1),21 as well as storage of isolated myocytes in EGTA containing storage-solution22 have been suggested to prevent the Ca2+ paradox.17 However, it is well known that Ca2+ buffering through EGTA reduces the amplitude of L-type Ca2+ current-induced Ca2+ transient amplitudes and results in a biphasic decay of the Ca2+ transients.23 Therefore, we omitted EGTA throughout the whole isolation process in order to obtain Ca2+ transients with typical properties and monophasic decays. To protect the cells from the Ca2+ paradox we increased the final Ca2+ concentration of the storage solution in a stepwise manner until 0.2 mM.
The choice of collagenase is probably the most critical step for successful myocyte isolation. Conventional collagenases are crude preparations obtained from Clostridium histolyticum and contain collagenase in addition to a number of other proteinases, polysaccharidases and lipases. Based on their general composition collagenases are subdivided into different types.24 Worthington collagenase Types I and II have been successfully used for isolation of human atrial myocytes.4-10,15,25-30 In our presently described protocol we recommend the use of collagenase Type I, although we were also able to obtain acceptable amounts of viable cells using collagenase Type II. However, even within a single collagenase type there is a significant batch-to-batch variation regarding the enzyme activities. These variations require careful batch selection and testing of various batches to optimize isolation procedure. The online available batch-selection tool from Worthington Biochemical Corp. (<a href="http://www.worthington-biochem.com/cls/match.php" target="_blank">http://www.worthington-biochem.com/cls/match.php</a>) may be used to find available batches with a composition that has been shown to be suitable for the isolation of human atrial myocytes. Currently we use collagenase type I with 250 U/mg collagenase activity, 345 U/mg caseinase activity, 2.16 U/mg clostripain activity and 0.48 U/mg tryptic activity (lot# 49H11338).
 
 
The cells obtained using the procedure described in this manuscript may be used within 8 hr for patch-clamp studies, Ca2+ transient measurements and a combination of both.15 In addition, these cells allow measurements of cellular contraction in response to electric field stimulation or electric stimulation using the patch-clamp pipette (unpublished observations).

Studying electrophysiological properties of cardiac ion channels with the patch-clamp technique and the exploration of cellular Ca2+ handling abnormalities require isolated cardiomyocytes. These are usually obtained following the in vitro exposure of cardiac tissue samples to digestive enzymes (collagenase, hyaluronidase, peptidase etc.). Since the first report of isolation of viable cardiac myocytes in 19552 a large quantity of protocols has been developed in order to harvest single atrial and ventricular cardiomyocytes from different species including mouse, rat, rabbit, dog, guinea pig and human. In this review we focus on isolation of human atrial myocytes. Regarding procedures for isolation of myocytes from other species we refer to the "Worthington Tissue Dissociation guide" provided by Worthington Biochemical Corp., USA (<a href="http://www.tissuedissociation.com" target="_blank">www.tissuedissociation.com</a>).
Human atrial myocyte isolation protocols are generally derived from the method described by Bustamante, et al.3 Here we provide a step-by-step description of a technique, which is adapted from a previously published method, in order to obtain atrial myocytes suitable not only for patch-clamp experiments but also for simultaneous intracellular Ca2+ measurements.4-11

<table border="1">
 <tbody>
 <tr>
 <td> &nbsp;</td>
 <td> &nbsp;</td>
 <td> &nbsp;</td>
 <td>Transport solution</td>
 </tr>
 <tr>
 <td>Albumin</td>
 <td>Sigma-Aldrich</td>
 <td>A3059</td>
 <td>Storage solution: 1%</td>
 </tr>
 <tr>
 <td>BDM</td>
 <td>Sigma-Aldrich</td>
 <td>31550</td>
 <td>Transport solution: 30</td>
 </tr>
 <tr>
 <td>DL-b-Hydroxy-butyric acid</td>
 <td>Sigma-Aldrich</td>
 <td>H6501</td>
 <td>Storage solution: 10</td>
 </tr>
 <tr>
 <td>Glucose</td>
 <td>Sigma-Aldrich</td>
 <td>G8270</td>
 <td>Transport solution: 20 
 Ca2+-free solution: 20 
 Enzyme solution E1 and E2: 20 
 Storage solution: 10</td>
 </tr>
 <tr>
 <td>L-Glutamic acid</td>
 <td>Sigma-Aldrich</td>
 <td>G1251</td>
 <td>Storage solution: 70</td>
 </tr>
 <tr>
 <td>KCl</td>
 <td>Merck</td>
 <td>1049360250</td>
 <td>Transport solution: 10 
 Ca2+-free solution: 10 
 Enzyme solution E1 and E2: 10 
 Storage solution: 20</td>
 </tr>
 <tr>
 <td>KH2PO4</td>
 <td>Sigma-Aldrich</td>
 <td>P5655</td>
 <td>Transport solution: 1.2 
 Ca2+-free solution: 1.2 
 Enzyme solution E1 and E2: 1.2 
 Storage solution: 10</td>
 </tr>
 <tr>
 <td>MgSO4</td>
 <td>Sigma-Aldrich</td>
 <td>M9397</td>
 <td>Transport solution: 5 
 Ca2+-free solution: 5 
 Enzyme solution E1 and E2: 5</td>
 </tr>
 <tr>
 <td>MOPS</td>
 <td>Sigma-Aldrich</td>
 <td>M1254</td>
 <td>Transport solution: 5 
 Ca2+-free solution: 5 
 Enzyme solution E1 and E2: 5</td>
 </tr>
 <tr>
 <td>NaCl</td>
 <td>Sigma-Aldrich</td>
 <td>S3014</td>
 <td>Transport solution: 100 
 Ca2+-free solution: 100 
 Enzyme solution E1 and E2: 100</td>
 </tr>
 <tr>
 <td>Taurin</td>
 <td>Sigma-Aldrich</td>
 <td>86330</td>
 <td>Transport solution: 50 
 Ca2+-free solution: 50 
 Enzyme solution E1 and E2: 50 
 Storage solution: 10</td>
 </tr>
 <tr>
 <td>Collagenase I</td>
 <td>Worthington</td>
 <td>4196</td>
 <td>Enzyme solution E1 and E2: 286 U/ml</td>
 </tr>
 <tr>
 <td>Protease XXIV</td>
 <td>Sigma-Aldrich</td>
 <td>P8038</td>
 <td>Enzyme solution E1 and E2: 5 U/ml*</td>
 </tr>
 <tr>
 <td>pH</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Transport solution: 7.00 
 Ca2+-free solution: 7.00 
 Enzyme solution E1 and E2: 7.00 
 Storage solution: 7.40</td>
 </tr>
 <tr>
 <td>adjusted with</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Transport solution: 1 M NaOH 
 Ca2+-free solution: 1 M NaOH 
 Enzyme solution E1 and E2: 1 M NaOH 
 Storage solution: 1 M KOH</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Concentrations in mM unless otherwise stated. BDM, 2,3-Butanedione monoxime. *Protease XXIV is included in Enzyme solution E1 only.</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Table I. Solutions.</td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Nylon mesh (200 &mu;m)</td>
 <td>VWR-Germany</td>
 <td>510-9527</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Jacketed reaction beaker</td>
 <td>VWR</td>
 <td>KT317000-0050</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Table II. Specific equipment.</td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dimethyl-sulphoxide</td>
 <td>Sigma-Aldrich</td>
 <td>D2650</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Fluo-3 AM (special packaging)</td>
 <td>Invitrogen</td>
 <td>F-1242</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Pluronic F-127</td>
 <td>Invitrogen</td>
 <td>P6867</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Table III. Substances for loading of myocytes with Fluo-3 AM.</td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Bath solution</td>
 </tr>
 <tr>
 <td>4-aminopyridine*</td>
 <td>Sigma-Aldrich</td>
 <td>A78403</td>
 <td>Bath solution: 5</td>
 </tr>
 <tr>
 <td>BaCl2*</td>
 <td>Sigma-Aldrich</td>
 <td>342920</td>
 <td>Bath solution: 0.1</td>
 </tr>
 <tr>
 <td>CaCl2 &times; 2H2O</td>
 <td>Sigma-Aldrich</td>
 <td>C5080</td>
 <td>Bath solution: 2</td>
 </tr>
 <tr>
 <td>Glucose</td>
 <td>Sigma-Aldrich</td>
 <td>G8270</td>
 <td>Bath solution: 10</td>
 </tr>
 <tr>
 <td>HEPES</td>
 <td>Sigma-Aldrich</td>
 <td>H9136</td>
 <td>Bath solution: 10</td>
 </tr>
 <tr>
 <td>KCl</td>
 <td>Merck</td>
 <td>1049360250</td>
 <td>Bath solution: 4</td>
 </tr>
 <tr>
 <td>MgCl &times; 6H2O</td>
 <td>Sigma-Aldrich</td>
 <td>M0250</td>
 <td>Bath solution: 1</td>
 </tr>
 <tr>
 <td>NaCl</td>
 <td>Sigma-Aldrich</td>
 <td>S3014</td>
 <td>Bath solution: 140</td>
 </tr>
 <tr>
 <td>Probenecid</td>
 <td>Sigma-Aldrich</td>
 <td>P8761</td>
 <td>Bath solution: 2</td>
 </tr>
 <tr>
 <td>pH</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Bath solution: 7.35</td>
 </tr>
 <tr>
 <td>adjusted with</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Bath solution: 1 M HCl</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Table IV. Bath solution for patch-clamp. *4-aminopyridine and BaCl were included for voltage-clamp experiments only.</td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Pipette solution</td>
 </tr>
 <tr>
 <td>DL-aspartat K+-salt</td>
 <td>Sigma-Aldrich</td>
 <td>A2025</td>
 <td>Pipette solution: 92</td>
 </tr>
 <tr>
 <td>EGTA</td>
 <td>Sigma-Aldrich</td>
 <td>E4378</td>
 <td>Pipette solution: 0.02</td>
 </tr>
 <tr>
 <td>GTP-Tris</td>
 <td>Sigma-Aldrich</td>
 <td>G9002</td>
 <td>Pipette solution: 0.1</td>
 </tr>
 <tr>
 <td>HEPES</td>
 <td>Sigma-Aldrich</td>
 <td>H9136</td>
 <td>Pipette solution: 10</td>
 </tr>
 <tr>
 <td>KCl</td>
 <td>Merck</td>
 <td>1049360250</td>
 <td>Pipette solution: 48</td>
 </tr>
 <tr>
 <td>MgATP</td>
 <td>Sigma-Aldrich</td>
 <td>A9187</td>
 <td>Pipette solution: 1</td>
 </tr>
 <tr>
 <td>Na2ATP</td>
 <td>Sigma-Aldrich</td>
 <td>A2383</td>
 <td>Pipette solution: 4</td>
 </tr>
 <tr>
 <td>Fluo-3**</td>
 <td>Invitrogen</td>
 <td>F3715</td>
 <td>Pipette solution: 0.1</td>
 </tr>
 <tr>
 <td>pH</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>7.20</td>
 </tr>
 <tr>
 <td>adjusted with</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>1 M KOH</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Table V. Pipette solution for patch-clamp*. *On experimental days pipette solution is stored on ice until use. **Fluo-3 is added from a 1 mM stock solution on experimental days.</td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 </tbody>
</table>

isolation of human atrial myocytes for simultaneous measurements of ca2+ transients and membrane currents

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

Figure 2A shows three representative examples from isolated human right atrial myocytes. To quantify the cell yield we pipette 10 &mu;l of cell suspension (step 5.5) on a CellFinder microscope slide (<a href="http://www.antenna.nl/microlab/index-uk.html" target="_blank">http://www.antenna.nl/microlab/index-uk.html</a>). Averaged cell yields in Figure 2B clearly indicate that there is a tendency to lower cell yields in chronic AF (cAF) patient samples (tube A: 16.5&plusmn;3.1 cells/10 &mu;l (n=29) vs. 5.1&plusmn;2.3 cells/10 &mu;l (n=10) in SR and cAF, respectively, p&lt;0.05; tube B: 17.9&plusmn;3.9 cells/10 &mu;l (n=29) vs. 5.9&plusmn;2.0 cells/10 &mu;l (n=9) in SR and cAF, respectively, p=0.107).
Representative examples of action-potential measurements and simultaneous recordings of cytosolic Ca2+ transients are given in Figure 3. In about 90% of the investigated cells, the action-potential-triggered Ca2+ release causes clear and regular cell contractions. As reported previously, the resting membrane potential, which is an accepted indicator for cell integrity, averaged about -73.9&plusmn;2.7 mV (n = 23/10 myocytes/patients) and -77.7&plusmn;1.8 mV (n = 19/8 myocytes/patients) in SR and cAF respectively (p&gt;0.05).15 Figure 4 shows representative simultaneous recordings of voltage-gated L-type Ca2+ currents and cytosolic Ca2+ transients. Application of the non-selective &beta;-adrenoceptor agonist isoprenaline (1 &mu;M) increases amplitudes of both ICa,L and cytosolic Ca2+ transients, suggesting intact &beta;-adrenergic signal transduction cascade.
<img alt="Figure 1" fo:content-width="4in" fo:src="/files/ftp_upload/50235/50235fig1highres.jpg" src="/files/ftp_upload/50235/50235fig1.jpg" /> 
Figure 1. Flow chart of the myocytes Fluo-3 AM loading protocol (see step 6.1-6.5). m/v, mass/volume.
<img alt="Figure 2" fo:content-width="4in" fo:src="/files/ftp_upload/50235/50235fig2highres.jpg" src="/files/ftp_upload/50235/50235fig2.jpg" /> 
Figure 2. A, Isolated human right atrial myocytes after one hour in storage solution. B, Mean&plusmn;SEM of the cell yield counted in 10 &mu;l of cells in storage solution (see step 5.5). n refers to the number of preparations within each group. *p&lt;0.05.
<img alt="Figure 3" fo:content-width="4in" fo:src="/files/ftp_upload/50235/50235fig3highres.jpg" src="/files/ftp_upload/50235/50235fig3.jpg" /> 
Figure 3. Representative recordings of action-potential-triggered Ca2+-transients (CaT) in an atrial myocyte from a sinus rhythm and a chronic atrial fibrillation patient. Top: Injected membrane current (IM) used for stimulation (0.5 Hz). Below: Simultaneous recording of membrane potential (VM), and triggered CaT (bottom). (Replotted with permission from Voigt et al. 2012)15
<img alt="Figure 4" fo:content-width="4in" fo:src="/files/ftp_upload/50235/50235fig4highres.jpg" src="/files/ftp_upload/50235/50235fig4.jpg" /> 
Figure 4. Representative recordings of the isoprenaline (1 &mu;M) effect on L-type Ca2+ current-triggered Ca2+-transients (CaT) in an atrial myocyte from a sinus rhythm and a chronic atrial fibrillation patient. Top: Voltage-clamp protocol (0.5 Hz). Below: Simultaneous recording of total net membrane current (IM), predominantly reflecting L-type Ca2+ current (middle) and triggered CaT (bottom). (Replotted with permission from Voigt, et al. 2012)15

<img src="/files/ftp_upload/50235/50235table1.jpg" alt="Table 1" fo:content-width="4in" fo:src="/files/ftp_upload/50235/50235table1highres.jpg" /> 
Table I. Solutions. 
<img src="/files/ftp_upload/50235/50235table2.jpg" alt="Table 2" fo:content-width="2in" fo:src="/files/ftp_upload/50235/50235table2highres.jpg"/> 
Table II. Specific equipment. 
<img src="/files/ftp_upload/50235/50235table3.jpg" alt="Table 3" fo:content-width="2in" fo:src="/files/ftp_upload/50235/50235table3highres.jpg"/> 
Table III. Substances for loading of myocytes with Fluo-3 AM. 
<img src="/files/ftp_upload/50235/50235table4.jpg" alt="Table 4" fo:content-width="2.5in" fo:src="/files/ftp_upload/50235/50235table4highres.jpg"/> 
Table IV. Bath solution for patch-clamp. 
<img src="/files/ftp_upload/50235/50235table5.jpg" alt="Table 5" fo:content-width="2.5in" fo:src="/files/ftp_upload/50235/50235table5highres.jpg"/> 
Table V. Pipette solution for patch-clamp*.

Watch this Scientific Journal Video about Isolation of Human Atrial Myocytes for Simultaneous Measurements of Ca2+ Transients and Membrane Currents at JoVE.com

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

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

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

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

isolation-human-atrial-myocytes-for-simultaneous-measurements-ca2

Research

JoVE Journal

Biology

20.3K Views.  University of Duisburg-Essen. Descriviamo l'isolamento di miociti atriali umani, che possono essere utilizzati per Ca intracellulare 2 + Misure in combinazione con studi elettrofisiologici di patch-clamp.

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

Primary Culture of Adult Rat Heart Myocytes

Cultured primary adult rodent heart cells are an important model system for cardiovascular research. Nevertheless, establishment of robust, viable cultured adult myocytes can be a technically challenging, rate-limiting step for many researchers. Here we described a protocol to obtain a high yield of adult rat heart myocytes that remain viable in culture for several days. The heart is isolated and perfused with collagenase and protease under low Ca2+ conditions to recover single myocytes. Ca2+-tolerant cells are obtained by stepwise increases in extracellular Ca2+ concentration in three subsequent wash steps. Cells are filtered, resuspended in culture medium, and plated on laminin coated slips. Cultured myocytes obtained using this protocol are viable for up to four days and are suitable for most experiments including electrophysiology, biochemistry, imaging and molecular biology.

In this paper, we described a typical way to isolate and culture adult rat heart myocytes. Collagenase and protease are used to digest and isolate single myocytes. Myocytes cultured follow this protocol meet most experiment requirements.

Cultured primary adult rodent heart cells are an important model system for cardiovascular research.  Nevertheless, establishment of robust, viable ...

Isolation and Genetic Manipulation of Adult Cardiac Myocytes for Confocal Imaging

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

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

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

Techniques for Imaging Ca2+ Signaling in Human Sperm

Fluorescence microscopy of cells loaded with fluorescent, Ca2+-sensitive dyes is used for measurement of spatial and temporal aspects of Ca2+ signaling in live cells. Here we describe the method used in our laboratories for loading suspensions of human sperm with Ca2+-reporting dyes and measuring the fluorescence signal during physiological stimulation. Motile cells are isolated by direct swim-up and incubated under capacitating conditions for 0-24 h, depending upon the experiment. The cell-permeant AM (acetoxy methyl ester) ester form of the Ca2+-reporting dye is then added to a cell aliquot and a period of 1 h is allowed for loading of the dye into the cytoplasm. We use visible wavelength dyes to minimize photo-damage to the cells, but this means that ratiometric recording is not possible. Advantages and disadvantages of this approach are discussed. During the loading period cells are introduced into an imaging chamber and allowed to adhere to a poly-D-lysine coated coverslip. At the end of the loading period excess dye and loose cells are removed by connection of the chamber to the perfusion apparatus. The chamber is perfused continuously, stimuli and modified salines are then added to the perfusion header. Experiments are recorded by time-lapse acquisition of fluorescence images and analyzed in detail offline, by manually drawing regions of interest. Data are normalized to pre-stimulus levels such that, for each cell (or part of a cell), a graph showing the Ca2+ response as % change in fluorescence is obtained.

Techniques for Imaging Ca2+ Signaling in Human Sperm

Stimulus-evoked [Ca2+]i signals of individual human sperm are assessed. Motile cells are loaded with Ca2+-sensitive fluorescent dye (AM-ester method) and immobilised in a perfusable chamber. Cells are imaged by time-lapse fluorescence microscopy and stimulated via the perfusing medium. Responses of single cells (or regions) are analysed offline using Excel.

Fluorescence microscopy of cells loaded with fluorescent, Ca2+-sensitive dyes is used for measurement of spatial and temporal aspects of Ca2+ signaling in ...

Isolation and Kv Channel Recordings in Murine Atrial and Ventricular Cardiomyocytes

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

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

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

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

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

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

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

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

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

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

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

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

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.

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). ...

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

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

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

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

Simultaneous Mapping and Quantitation of Ribonucleotides in Human Mitochondrial DNA

Established approaches to estimate the number of ribonucleotides present in a genome are limited to the quantitation of incorporated ribonucleotides using short synthetic DNA fragments or plasmids as templates and then extrapolating the results to the whole genome. Alternatively, the number of ribonucleotides present in a genome may be estimated using alkaline gels or Southern blots. More recent in vivo approaches employ Next-generation sequencing allowing genome-wide mapping of ribonucleotides, providing the position and identity of embedded ribonucleotides. However, they do not allow quantitation of the number of ribonucleotides which are incorporated into a genome. Here we describe how to simultaneously map and quantitate the number of ribonucleotides which are incorporated into human mitochondrial DNA in vivo by Next-generation sequencing. We use highly intact DNA and introduce sequence specific double strand breaks by digesting it with an endonuclease, subsequently hydrolyzing incorporated ribonucleotides with alkali. The generated ends are ligated with adapters and these ends are sequenced on a Next-generation sequencing machine. The absolute number of ribonucleotides can be calculated as the number of reads outside the recognition site per average number of reads at the recognition site for the sequence specific endonuclease. This protocol may also be utilized to map and quantitate free nicks in DNA and allows adaption to map other DNA lesions that can be processed to 5&#180;-OH ends or 5&#180;-phosphate ends. Furthermore, this method can be applied to any organism, given that a suitable reference genome is available. This protocol therefore provides an important tool to study DNA replication, 5&#180;-end processing, DNA damage, and DNA repair.

Here we describe a method amenable to simultaneously quantitate and genome-wide map ribonucleotides in highly intact DNA at single-nucleotide resolution, combining enzymatic cleavage of genomic DNA with its alkaline hydrolysis and subsequent 5&#180;-end sequencing.

Established approaches to estimate the number of ribonucleotides present in a genome are limited to the quantitation of incorporated ribonucleotides using ...

Simultaneous Measurements of Intracellular Calcium and Membrane Potential in Freshly Isolated and Intact Mouse Cerebral Endothelium

Cerebral arteries and their respective microcirculation deliver oxygen and nutrients to the brain via blood flow regulation. Endothelial cells line the lumen of blood vessels and command changes in vascular diameter as needed to meet the metabolic demand of neurons. Primary endothelial-dependent signaling pathways of hyperpolarization of membrane potential (Vm) and nitric oxide typically operate in parallel to mediate vasodilation and thereby increase blood flow. Although integral to coordinating vasodilation over several millimeters of vascular length, components of endothelium-derived hyperpolarization (EDH) have been historically difficult to measure. These components of EDH entail intracellular Ca2+ [Ca2+]i increases and subsequent activation of small- and intermediate conductance Ca2+-activated K+ (SKCa/IKCa) channels.
Here, we present a simplified illustration of the isolation of fresh endothelium from mouse cerebral arteries; simultaneous measurements of endothelial [Ca2+]i and Vm using Fura-2 photometry and intracellular sharp electrodes, respectively; and a continuous superfusion of salt solutions and pharmacological agents under physiological conditions (pH 7.4, 37 &#176;C). Posterior cerebral arteries from the Circle of Willis are removed free of the posterior communicating and the basilar arteries. Enzymatic digestion of cleaned posterior cerebral arterial segments and subsequent trituration facilitates removal of adventitia, perivascular nerves, and smooth muscle cells. Resulting posterior cerebral arterial endothelial &#34;tubes&#34; are then secured under a microscope and examined using a camera, photomultiplier tube, and one to two electrometers while under continuous superfusion. Collectively, this method can simultaneously measure changes in endothelial [Ca2+]i and Vm in discrete cellular locations, in addition to the spreading of EDH through gap junctions up to millimeter distances along the intact endothelium. This method is expected to yield a high-throughput analysis of the cerebral endothelial functions underlying mechanisms of blood flow regulation in the normal and diseased brain.

Demonstrated here are protocols for (1) freshly isolating intact cerebral endothelial "tubes" and (2) simultaneous measurements of endothelial calcium and membrane potential during endothelium-derived hyperpolarization. Further, these methods allow for pharmacological tuning of endothelial cell calcium and electrical signaling as individual or interactive experimental variables.

Cerebral arteries and their respective microcirculation deliver oxygen and nutrients to the brain via blood flow regulation. Endothelial cells line the ...