We would like to thank Dr. Claycomb for providing HL-1 cells and detailed maintenance protocol. We would also like to thank Ms. Elena Carrillo and Dr. Seth Robia for their assistance with generating the cell movie and Mr. Pete Caron for his assistance with making some accessories for our dual channel optical mapping system.&#160;
This work was supported by American Heart Association (10GRNT3770030 &#38; 12GRNT12050478 to XA), National Institutes of Health (HL113640 to XA), and Loyola University Research Development Fund (to XA).&#160;

1. Solution Preparation
<ol>
	<li>Norepinephrine (NP) solution
	<ol>
		<li>Dissolve 40 mg of NP into 25 ml of 30 mM ascorbic acid (0.1475 g ascorbic acid in 25 ml of purified Milli-Q (MQ) water. Avoid direct light exposure during this solution preparation because norepinephrine (NP) is light sensitive.</li>
		<li>Filter the NP solution with a 0.22 &#181;m syringe filter. Aliquot the solution into 1 ml sterile microtubes and store at -20 &#176;C. Once frozen, NP is stable for one month.</li>
	</ol>
	</li>
	<li>Supplemented Wash and Final Claycomb media
	<ol>
		<li>Start with 43.5 ml of Claycomb medium and supplement with 5 ml of fetal bovine serum (FBS), 0.5 ml of penicillin/streptomycin, 0.5 ml of NP and 1:100 diluted L-Glutamine to make 50 ml of final supplemented Claycomb medium. Avoid direct light exposure during this solution preparation because Claycomb medium is light sensitive.</li>
		<li>Prepare the Wash Claycomb medium using the same recipe as the supplemented Claycomb medium except add 5% FBS and omit the NP and L-Glutamine components. 
		NOTE: The supplemented Final and Wash Claycomb media are good for two weeks when stored at 4 &#176;C without direct light exposure.</li>
	</ol>
	</li>
	<li>Soybean Trypsin inhibitory solution
	<ol>
		<li>Dissolve 25 mg of soybean trypsin inhibitor into 100 ml of Ca-free and Mg-free Dulbecco&#8217;s phosphate buffered saline (PBS).</li>
		<li>Filter the solution using a 0.22 &#181;m syringe filter. 
		NOTE: The sterilized solution is stable for one month when stored at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Gelatin/Fibronectin dish coating solution
	<ol>
		<li>Dissolve 0.1 g of gelatin into 500 ml of MQ water to make 0.02% gelatin.</li>
		<li>Autoclave, then dilute 1 ml of fibronectin into 199 ml of 0.02% gelatin and mix gently.</li>
		<li>Aliquot 6 ml of gelatin solution into separate tubes and store at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Hanks&#8217; Salt solution
	<ol>
		<li>Using a stir plate, dissolve the following salts in MQ water (mmol/L), NaCl (136.9), KCl (5.366), KH2PO4 (0.4409), NaH2PO4 (0.4000), MgSO4 (0.8142), D-Glc (5.051), NaHCO3 (4.162), HEPEs (5.042), CaCl2 (1.261).</li>
	</ol>
	</li>
	<li>Calcium sensitive dye (Rhod2) solution
	<ol>
		<li>Dilute 1 mg of Rhod2 dry powder in 400 &#956;l of 20% Pluronic F-127 in DMSO to make a Rhod2 dye stock solution and store at -20 &#176;C.</li>
		<li>Dilute this stock solution 1:1,000 with Hanks&#39; Salt solution to make a final working solution before starting the experiment.</li>
	</ol>
	</li>
	<li>Voltage sensitive dye (Rh237) solution
	<ol>
		<li>Dilute 5 mg of Rh237 with 2 ml of DMSO to make stock solution and store at -20 &#176;C. Dilute the stock solution 1:1,000 with Hanks&#39; Salt solution to make a final working solution before starting the experiment.</li>
	</ol>
	</li>
</ol>
&#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; NOTE: 1-4 solutions are based on the Claycomb HL-1 cell culture protocol with minor modification.
2. Passaging and Culturing HL-1 Atrial Myocytes onto Coverslips
NOTE: The HL-1 cells can be obtained from Dr. Claycomb (Louisiana State University).
<ol>
	<li>Grow HL-1 cells in T25 flasks and feed with 5 ml of supplemented Claycomb Medium daily. Passage the cells in T25 flasks every five days according to the Claycomb HL-1 cell culture protocol (described below)10.</li>
	<li>Place round coverslips (20 mm diameter) into 12-well culture plates 24 hr before the cells in the T25 flask are fully confluent and ready to be passaged, and then cover the coverslips with gelatin/fibronectin overnight at 37 &#176;C.</li>
	<li>The next day, aspirate and discard the gelatin/fibronectin from the culture plates and replace with 1.5 ml of supplemented Final Claycomb medium in each well.</li>
	<li>Split cells from a T25 culture flask after reaching full confluence at day 5. Aspirate the medium from the T25 flask containing the confluent culture and rinse the culture briefly with PBS at 37 &#176;C.</li>
	<li>Aspirate PBS and add 3 ml of 0.05% trypsin/EDTA at 37 &#176;C by pipetting the trypsin/EDTA onto the bottom of the T25 flask to avoid direct contact with the cells. Incubate at 37 &#176;C for 1 min.</li>
	<li>Remove trypsin/EDTA by aspiration and add another 1.3 ml of trypsin/EDTA per T25 flask. Incubate at room temperature for 1 min. Examine under a microscope and if cells are still attached to the flask, tap to fully dislodge remaining cells.</li>
	<li>Add 1.3 ml of soybean trypsin inhibitor to the cells and transfer cells from the flask into a 15 ml centrifuge tube. Rinse the empty flask with 5 ml of Wash Claycomb medium. Add the rinse to the cells in the 15 ml centrifuge tube. Centrifuge at 500 x g for 5 min.</li>
	<li>After centrifuging, aspirate the supernatant and gently re-suspend the cell pellet in 6 ml of supplemented Final Claycomb medium.</li>
	<li>Seed each well of a 12-well plate with 0.5 ml of cell suspension from cell passaging. Feed the cells with fresh supplemented Final Claycomb medium daily. 
	NOTE: Closely monitor the growth of the cells, typically the cell monolayers reach confluent status on day 4 and the spontaneous activities remain visible. Then, the cultured monolayer will be ready for optical mapping on day 4-5.</li>
</ol>
3. Loading Calcium and Voltage Sensitive Dyes to the Cells
<ol>
	<li>For optical mapping studies, gently remove the coverslip from the 12-well cell culture plate and rinse with Hanks&#39; Salt solution.</li>
	<li>To stain cells with Ca sensitive dye Rhod2, incubate the cell monolayer on the coverslip with 3 ml of the oxygenated (bubbling with 5% CO2/95% O2 gas) Rhod2 working solution at 37 &#176;C in a water bath for 30 min.</li>
	<li>To stain the cells with Vm sensitive dye Rh237, immerse the Rhod2 loaded cell monolayer into 3 ml of oxygenated Rh237 working solution at 37 &#176;C in a water bath for another 15 min.</li>
</ol>
4. Dual Channel Optical Mapping
<ol>
	<li>Place the cell monolayer, which is double-stained with Vm and Ca dyes, into a circulation chamber on an inverted fluorescence microscope. Use a heated circulation system to keep the temperature of the cell chamber at 35 &#177; 1 &#176;C. Carefully control the temperature of the perfusate by adjusting the speed of the circulating flow through the chamber to avoid a gradient of more than 0.3 &#176;C across the chamber.
	<ol>
		<li>Before recording Vm/Ca signals, wash the cell monolayer with fresh Hanks solution for approximately 5 min to remove the remaining dye from the previous staining steps.</li>
	</ol>
	</li>
	<li>Pace healthy cell monolayers using a bipolar electrode composed of a glass pipette filled with Hanks&#39; Salt solution and a silver wire coiled around the pipette tip as previously described12,17.</li>
	<li>Pace healthy cell monolayers at a cycle length of 200-500 msec and pulse width of 2 msec, expecting a pacing threshold at approximately 1 mV. 
	NOTE: For each batch of cultured monolayers, at least two non-treated monolayers should be used to test the condition of the cell batch. Healthy control monolayers should be able to be captured by electrical pacing at cycle lengths of 200-500 msec. If the control monolayers fail to be electrically paced, this suggests that the entire cell batch may not be suitable for optical mapping experiments.</li>
	<li>To excite both Vm and Ca sensitive dyes, use an LED light source (520 nm wavelength) coupled with an excitation filter (510 nm/40) and a dichroic mirror (561 nm).
	<ol>
		<li>Collect the fluorescent signals emitted by the cells via a 40X objective and split with a dichroic mirror (594 nm) into a Vm component and a Ca component. Filter the emitted light in the Vm channel with a long pass filter (700 nm) and the emitted light in the Ca channel with a band pass filter (585/40 nm).</li>
		<li>Collect the filtered fluorescent light using two CCD cameras (80 x 80 pixels, 1,000 fps; RedShirt Imaging).</li>
		<li>To avoid dye photobleaching, make recordings for less than 2 sec on each field of view to limit the time of light exposure. Here, typically image four different areas for each monolayer.</li>
	</ol>
	</li>
</ol>
5. Data Processing of Vm or Ca Recordings
<ol>
	<li>For both Vm and Ca signals (80 x 80 pixels), average the intensity values of twenty-five (5 x 5) neighboring pixels into one intensity value at different recording time points to reduce background noise. 
	NOTE: After the data averaging, each recording time point of an individual action potential or Ca transient yields a data matrix containing 16 x 16 averaged intensity values (averaged data channels) with a spatial resolution of 50 &#956;m.</li>
	<li>Define activation time (AT) of recorded Vm or Ca signals.
	<ol>
		<li>To define an activation time (AT) of an individual action potential or Ca transient, determine the time of 50% of peak amplitude of the Vm or Ca signals using a custom-made MATLAB-based algorithm as previously described.12</li>
		<li>Manually proofread the analyzed ATs to eliminate any errors that may occur during the program analysis.</li>
		<li>Store the defined AT matrix (16 x 16 averaged data channels) of recorded action potential and calcium transient, respectively, in the software used.</li>
	</ol>
	</li>
	<li>Calculate action potential or calcium transient conduction velocity (CV) vectors on the AT matrix using the vector field analysis method as previously described with minor modifications12,18.
	<ol>
		<li>Calculate the CV vector of a data channel relative to its surrounding AT data points (5 &#215; 5 neighboring averaged data channels) to simulate a smooth polynomial surface of activation times (AT surface) using a least-square algorithm within the MATLAB program12,18.</li>
		<li>Calculate conduction vector using the formulas below 
		Formula 1: 
		<img alt="Equation 1" fo:content-width="1.5in" src="/files/ftp_upload/52542/52542eq1.jpg" /> 
		Formula 2: 
		&#952; = arctan [(dy/dT)/(dx/dT)] 
		NOTE: Ve is a CV vector projected at x- and y-axes; dx/dT is the projection of the CV vector in x-axis; dy/dT is the projection of the CV vector in y-axis. T is the AT surface; x, y refer to the 2-D coordinates; Tx = &#8706;T/&#8706;x is the partial derivative of T with respect to x; Ty = &#8706;T/&#8706;y is the partial derivative of T with respect to y; &#952; is the final direction of a CV vector.</li>
		<li>Group all the vectors of the AT matrix into 36 pathways, each of which covers a ten-degree sector on the compass.</li>
		<li>Select the pathway containing the largest population of the CV vectors among the 36 conduction pathways. Calculate the average CV of this pathway to represent the CV of the monolayer.</li>
	</ol>
	</li>
</ol>
6. Visualization of the Wave Front Propagation
<ol>
	<li>Normalize the signal intensity of an individual action potential or Ca transient at different recording time points relative to its peak value using MATLAB software.</li>
	<li>Temporarily store the normalized intensity data of every recorded time point into a 16 x 16 intensity matrix in the MATLAB software used.</li>
	<li>Construct a color-coded intensity map from the stored intensity matrix of each recording time point using the surf.m function. 
	NOTE: In the intensity map, red represents high intensity of the Vm or Ca signals, and blue reflects low intensity of the fluorescence signals.</li>
	<li>Create an .avi file using the avifile.m function.</li>
	<li>Save all the intensity maps at different recording time points of the action potential or Ca transient using the addframe.m function. 
	NOTE: At this point, two separate avi files of Vm and Ca propagation will be generated.</li>
</ol>

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	<li>Efimov, I. R., Nikolski, V. P., Salama, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical+imaging+of+the+heart.">Optical imaging of the heart.</a> Circ Res. 95 (1), 21-33 (2004).</li><li>Salama, G., Hwang, S. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+optical+mapping+of+intracellular+free+calcium+and+action+potentials+from+Langendorff+perfused+hearts.">Simultaneous optical mapping of intracellular free calcium and action potentials from Langendorff perfused hearts.</a> Current protocols in cytometry / editorial board, J. Paul Robinson, managing editor ... [et al.]. 12 (Unit 12 17), (2009).</li><li>Laurita, K. R., Singal, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mapping+action+potentials+and+calcium+transients+simultaneously+from+the+intact+heart.">Mapping action potentials and calcium transients simultaneously from the intact heart.</a> Am J Physiol Heart Circ Physiol. 280 (5), H2053-H2060 (2001).</li><li>Fast, V. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+optical+imaging+of+membrane+potential+and+intracellular+calcium.">Simultaneous optical imaging of membrane potential and intracellular calcium.</a> J Electrocardiol. 38 (4 Suppl), 107-112 (2005).</li><li>Sowell, B., Fast, V. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ionic+mechanism+of+shock-induced+arrhythmias:+role+of+intracellular+calcium.">Ionic mechanism of shock-induced arrhythmias: role of intracellular calcium.</a> Heart Rhythm. 9 (1), 96-104 (2012).</li><li>Fast, V. G., Kleber, A. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microscopic+conduction+in+cultured+strands+of+neonatal+rat+heart+cells+measured+with+voltage-sensitive+dyes.">Microscopic conduction in cultured strands of neonatal rat heart cells measured with voltage-sensitive dyes.</a> Circ Res. 73 (5), 914-925 (1993).</li><li>Hou, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+major+role+for+HERG+in+determining+frequency+of+reentry+in+neonatal+rat+ventricular+myocyte+monolayer.">A major role for HERG in determining frequency of reentry in neonatal rat ventricular myocyte monolayer.</a> Circ Res. 107 (12), 1503-1511 (2010).</li><li>Beauchamp, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Relative+contributions+of+connexins+40+and+43+to+atrial+impulse+propagation+in+synthetic+strands+of+neonatal+and+fetal+murine+cardiomyocytes.">Relative contributions of connexins 40 and 43 to atrial impulse propagation in synthetic strands of neonatal and fetal murine cardiomyocytes.</a> Circ Res. 99 (11), 1216-1224 (2006).</li><li>Xia, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+expression+of+L-+and+T-type+Ca2++channels+in+murine+HL-1+cells.">Functional expression of L- and T-type Ca2+ channels in murine HL-1 cells.</a> J Mol Cell Cardiol. 36 (1), 111-119 (2004).</li><li>Claycomb, W. 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No conflicts of interest declared.

This article describes the key aspects of optical mapping in a cultured HL-1 atrial myocyte monolayer stained with calcium and voltage sensitive fluorescent dyes. It includes culturing an optimal HL-1 cell monolayer, setup of the mapping equipment, mapping a cultured monolayer, and data analysis.
To successfully map cultured cells, the key is to prepare a uniformly distributed cell monolayer. When seeding the cells on to the coverslips, be sure to evenly disperse the cells. Always map fully-gr...

A unique calcium (Ca) and voltage (Vm) dual channel optical mapping technique1-5 is emerging as an efficient tool to simultaneously record Vm and Ca signals in both intact hearts and cultured cell monolayers. This technique makes it possible to obtain powerful information regarding the relationship between calcium transients and action potentials to better understand the underlying electrophysiological mechanisms of cardiac arrhythmias.
Cultured cell monolayers have proven to be a useful cellular model to study cardiac electrophysiology and the underlying mechanism of arrhythmias.4,6-8 HL-1 cells are a well-characterized atrial myocyte culture line. HL-1 cells also maintain a uniquely differentiated genotype and phenotype that includes morphologic, electrophysiologic, and pharmacologic characteristics of adult atrial myocytes. These cells express cardiac genes and proteins, including important cardiac ion channels (i.e., L- and T-type calcium channels)9 and mature isoforms of sarcomeric contractile proteins normally found in adult atrial myocytes as others and we have previously reported.10-13 In addition, HL-1 cells can be cultured to form a 2-dimensional (2-D) myocyte monolayer. Thus, the advantages of using cultured HL-1 monolayers include: 1) relatively lower cost and easier to maintain a myocyte culture line than&#160;isolating and culturing primary&#160;neonatal myocytes; 2) a confluent monolayer of cells reduces the structural complexities that result from the 3-D structure of the heart; 3) a cell monolayer can eliminate the interference of interstitial fibrosis that occurs in the intact heart. This can be used to dissect specific electrophysiological functions of a group of myocytes without interference of fibroblasts and interstitial matrix; 4) assessment of functional consequences from pharmacological or genetic manipulation in cultured cell monolayers can be effectively achieved. Therefore, HL-1 cells have become a widely used cellular model for studying diverse aspects of myocyte physiology as well as pacing-induced abnormal electrical activities.13-16 However, special handling and care is required to culture healthy cell monolayers that respond to external electrical pacing for optical mapping studies. In addition, dual florescent dye staining procedure may easily damage the integrity of cultured confluent cell monolayers. Thus, performing Vm/CA&#160;dual channel optical mapping in cultured HL-1 monolayers has been a great challenge.
The goal of this method is to provide the key steps for successfully performing dual channel optical mapping in cultured HL-1 monolayers. Here we have provided extensive details on an optimized protocol for HL-1 cell monolayer preparation, dual channel optical mapping of a cultured cell monolayer, and mapping data processing.

<table><tbody><tr><td>Rhod2 dry powder</td><td>AAT Bioquest</td><td>21062</td><td></td></tr><tr><td>pluronic</td><td>AAT Bioquest</td><td>20050</td><td></td></tr><tr><td>DMSO</td><td>Sigma</td><td>276855</td><td></td></tr><tr><td>Rh237</td><td>Invitrogen</td><td>S-1109</td><td></td></tr><tr><td>NaCl</td><td>Sigma</td><td>S7653</td><td></td></tr><tr><td>KCl</td><td>Sigma</td><td>P3911</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>P0662</td><td></td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>S9638</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>D-Glucose</td><td>Sigma</td><td>G8270</td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Sigma</td><td>S6014</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma</td><td>C3881</td><td></td></tr><tr><td>HEPEs</td><td>Sigma</td><td>H3375</td><td></td></tr></tbody></table>

voltage and calcium dual channel optical mapping of cultured hl-1 atrial myocyte monolayer

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

A cultured confluent monolayer exhibits a regular intrinsic&#160;rhythm as demonstrated in Movie 1. We then performed Vm/Ca dual channel optical mapping in a fully confluent HL-1 monolayer. Figure 1A shows example traces of Vm and Ca signals from a recorded single beat. Representative isochronal maps of uniformly propagated Vm and Ca signals using the dual channel optical mapping system are shown in Figures 1B and 1C. Rep...

Watch this Scientific Journal Video about Voltage and Calcium Dual Channel Optical Mapping of Cultured HL-1 Atrial Myocyte Monolayer at JoVE.com

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

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

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

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10.2K Views.  Loyola University Chicago. This article describes the technique used to perform dual channel optical mapping in cultured HL-1 atrial cell monolayers. This unique protocol allows the simultaneous visualization of both calcium (Ca) and voltage (Vm) activity in the same area for the detailed detection and analysis of electrophysiological properties of culture monolayers.

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

Multiparametric Optical Mapping of the Langendorff-perfused Rabbit Heart

Optical imaging and fluorescent probes have significantly advanced research methodology in the field of cardiac electrophysiology in ways that could not have been accomplished by other approaches1. With the use of the calcium- and voltage-sensitive dyes, optical mapping allows measurement of transmembrane action potentials and calcium transients with high spatial resolution without the physical contact with the tissue. This makes measurements of the cardiac electrical activity possible under many conditions where the use of electrodes is inconvenient or impossible1. For example, optical recordings provide accurate morphological changes of membrane potential during and immediately after stimulation and defibrillation, while conventional electrode techniques suffer from stimulus-induced artifacts during and after stimuli due to electrode polarization1. 
The Langendorff-perfused rabbit heart is one of the most studied models of human heart physiology and pathophysiology. Many types of arrhythmias observed clinically could be recapitulated in the rabbit heart model. It was shown that wave patterns in the rabbit heart during ventricular arrhythmias, determined by effective size of the heart and the wavelength of reentry, are very similar to that in the human heart2. It was also shown that critical aspects of excitation-contraction (EC) coupling in rabbit myocardium, such as the relative contribution of sarcoplasmic reticulum (SR), is very similar to human EC coupling3. Here we present the basic procedures of optical mapping experiments in Langendorff-perfused rabbit hearts, including the Langendorff perfusion system setup, the optical mapping systems setup, the isolation and cannulation of the heart, perfusion and dye-staining of the heart, excitation-contraction uncoupling, and collection of optical signals. These methods could be also applied to the heart from species other than rabbit with adjustments to flow rates, optics, solutions, etc.
Two optical mapping systems are described. The panoramic mapping system is used to map the entire epicardium of the rabbit heart4-7. This system provides a global view of the evolution of reentrant circuits during arrhythmogenesis and defibrillation, and has been used to study the mechanisms of arrhythmias and antiarrhythmia therapy8,9. The dual mapping system is used to map the action potential (AP) and calcium transient (CaT) simultaneously from the same field of view10-13. This approach has enhanced our understanding of the important role of calcium in the electrical alternans and the induction of arrhythmia14-16.

This article describes the basic procedures for conducting optical mapping experiments in the Langendorff-perfused rabbit heart using the panoramic imaging system, and the dual (voltage and calcium) imaging modality.

Optical imaging and fluorescent probes have significantly advanced research methodology in the field of cardiac electrophysiology in ways that could not ...

Bioengineering

Optical Mapping of Action Potentials and Calcium Transients in the Mouse Heart

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. Cardiovascular physiological phenotyping of the mouse heart can be easily done using fluorescence imaging employing various probes for transmembrane potential (Vm), calcium transients (CaT), and other parameters. Excitation-contraction coupling is characterized by action potential and intracellular calcium dynamics; therefore, it is critically important to map both Vm and CaT simultaneously from the same location on the heart1-4. Simultaneous optical mapping from Langendorff perfused mouse hearts has the potential to elucidate mechanisms underlying heart failure, arrhythmias, metabolic disease, and other heart diseases. Visualization of activation, conduction velocity, action potential duration, and other parameters at a myriad of sites cannot be achieved from cellular level investigation but is well solved by optical mapping1,5,6. In this paper we present the instrumentation setup and experimental conditions for simultaneous optical mapping of Vm and CaT in mouse hearts with high spatio-temporal resolution using state-of-the-art CMOS imaging technology. Consistent optical recordings obtained with this method illustrate that simultaneous optical mapping of Langendorff perfused mouse hearts is both feasible and reliable.

This paper details the dissection procedure, instrumental setup, and experimental conditions during optical mapping of transmembrane potential (Vm) and intracellular calcium transient (CaT) in intact isolated Langendorff perfused mouse hearts.

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. ...

Optical Imaging of Isolated Murine Ventricular Myocytes

The ability to isolate adult cardiac myocytes has permitted researchers to study a variety of cardiac pathologies at the single cell level. While advances in calcium sensitive dyes have permitted the robust optical recording of single cell calcium dynamics, recording of robust transmembrane optical voltage signals has remained difficult. Arguably, this is because of the low single to noise ratio, phototoxicity, and photobleaching of traditional potentiometric dyes. Therefore, single cell voltage measurements have long been confined to the patch clamp technique which while the gold standard, is technically demanding and low throughput. However, with the development of novel potentiometric dyes, large, fast optical responses to changes in voltage can be obtained with little to no phototoxicity and photobleaching. This protocol describes in detail how to isolate adult murine myocytes which can be used for cellular shortening, calcium, and optical voltage measurements. Specifically, the protocol describes how to use a ratiometric calcium dye, a single-excitation calcium dye, and a single excitation voltage dye. This approach can be used to assess the cardiotoxicity and arrhythmogenicity of various chemical agents. While phototoxicity is still an issue at the single cell level, methodology is discussed on how to reduce it.

We present the methodology for the isolation of murine myocytes and how to obtain voltage or calcium traces simultaneously with sarcomere shortening traces using fluorescence photometry with simultaneous digital cell geometry measurements.

The ability to isolate adult cardiac myocytes has permitted researchers to study a variety of cardiac pathologies at the single cell level. While advances ...

Preclinical Cardiac Electrophysiology Assessment by Dual Voltage and Calcium Optical Mapping of Human Organotypic Cardiac Slices

Human cardiac slice preparations have recently been developed as a platform for human physiology studies and therapy testing to bridge the gap between animal and clinical trials. Numerous animal and cell models have been used to examine the effects of drugs, yet these responses often differ in humans. Human cardiac slices offer an advantage for drug testing in that they are directly derived from viable human hearts. In addition to having preserved multicellular structures, cell-cell coupling, and extracellular matrix environments, human cardiac tissue slices can be used to directly test the effect of innumerable drugs on adult human cardiac physiology. What distinguishes this model from other heart preparations, such as whole hearts or wedges, is that slices can be subjected to longer-term culture. As such, cardiac slices allow for studying the acute as well as chronic effects of drugs. Furthermore, the ability to collect several hundred to a thousand slices from a single heart makes this a high-throughput model to test several drugs at varying concentrations and combinations with other drugs at the same time. Slices can be prepared from any given region of the heart. In this protocol, we describe the preparation of left ventricular slices by isolating tissue cubes from the left ventricular free wall and sectioning them into slices using a high precision vibrating microtome. These slices can then either be subjected to acute experiments to measure baseline cardiac electrophysiological function or cultured for chronic drug studies. This protocol also describes dual optical mapping of cardiac slices for simultaneous recordings of transmembrane potentials and intracellular calcium dynamics to determine the effects of the drugs being investigated.

This protocol describes the procedure for sectioning and culturing human cardiac slices for preclinical drug testing and details the use of optical mapping for recording transmembrane voltage and intracellular calcium signals simultaneously from these slices.

Human cardiac slice preparations have recently been developed as a platform for human physiology studies and therapy testing to bridge the gap between ...

Measuring Fast Calcium Fluxes in Cardiomyocytes

Cardiomyocytes have multiple Ca2+ fluxes of varying duration that work together to optimize function 1,2. Changes in Ca2+ activity in response to extracellular agents is predominantly regulated by the phospholipase C&beta;- G&alpha;q pathway localized on the plasma membrane which is stimulated by agents such as acetylcholine 3,4. We have recently found that plasma membrane protein domains called caveolae5,6 can entrap activated G&alpha;q 7. This entrapment has the effect of stabilizing the activated state of G&alpha;q and resulting in prolonged Ca2+ signals in cardiomyocytes and other cell types8. We uncovered this surprising result by measuring dynamic calcium responses on a fast scale in living cardiomyocytes. Briefly, cells are loaded with a fluorescent Ca2+ indicator. In our studies, we used Ca2+ Green (Invitrogen, Inc.) which exhibits an increase in fluorescence emission intensity upon binding of calcium ions. The fluorescence intensity is then recorded for using a line-scan mode of a laser scanning confocal microscope. This method allows rapid acquisition of the time course of fluorescence intensity in pixels along a selected line, producing several hundreds of time traces on the microsecond time scale. These very fast traces are transferred into excel and then into Sigmaplot for analysis, and are compared to traces obtained for electronic noise, free dye, and other controls. To dissect Ca2+ responses of different flux rates, we performed a histogram analysis that binned pixel intensities with time. Binning allows us to group over 500 traces of scans and visualize the compiled results spatially and temporally on a single plot. Thus, the slow Ca2+ waves that are difficult to discern when the scans are overlaid due to different peak placement and noise, can be readily seen in the binned histograms. Very fast fluxes in the time scale of the measurement show a narrow distribution of intensities in the very short time bins whereas longer Ca2+ waves show binned data with a broad distribution over longer time bins. These different time distributions allow us to dissect the timing of Ca2+fluxes in the cells, and to determine their impact on various cellular events.

We present a method to isolate rapid (microsecond) calcium events from slower fluxes in living cells using laser scanning confocal microscopy. The method measures fluorescence intensity fluctuations of calcium indicators by recording line scans of several hundred pixels in a cell. Histogram analysis allows us to isolate the time scales of different calcium fluxes.

Cardiomyocytes have multiple Ca2+ fluxes of varying duration that work together to optimize function 1,2. Changes in Ca2+ activity in response to ...

Single-Cell Optical Action Potential Measurement in Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes

Conventional intracellular microelectrode techniques to quantify cardiomyocyte electrophysiology are extremely complex, labor intensive, and typically carried out in low throughput. Rapid and ongoing expansion of induced pluripotent stem cell (iPSC) technology presents a new standard in cardiovascular research and alternate methods are now necessary to increase throughput of electrophysiological data at a single cell level. VF2.1Cl is a recently derived voltage sensitive dye which provides a rapid single channel, high magnitude response to fluctuations in membrane potential. It possesses kinetics superior to those of other existing voltage indicators and makes available functional data equivalent to that of traditional microelectrode techniques. Here, we demonstrate simplified, non-invasive action potential characterization in externally paced human iPSC derived cardiomyocytes using a modular and highly affordable photometry system.

Here we describe optical acquisition and characterization of action potentials from induced pluripotent stem cell derived cardiomyocytes using a high-speed modular photometry system.

Conventional intracellular microelectrode techniques to quantify cardiomyocyte electrophysiology are extremely complex, labor intensive, and typically ...

Electrophysiological Assessment of Murine Atria with High-Resolution Optical Mapping

Recent genome-wide association studies targeting atrial fibrillation (AF) have indicated a strong association between the genotype and electrophysiological phenotype in the atria. That encourages us to utilize a genetically-engineered mouse model to elucidate the mechanism of AF. However, it is difficult to evaluate the electrophysiological properties in murine atria due to their small size. This protocol describes the electrophysiological evaluation of atria using an optical mapping system with a high temporal and spatial resolution in Langendorff perfused murine hearts. The optical mapping system is assembled with dual high-speed complementary metal oxide semiconductor cameras and high magnification objective lenses, to detect the fluorescence of a voltage-sensitive dye and Ca2+ indicator. To focus on the assessment of murine atria, optical mapping is performed with an area of 2 mm &#215; 2 mm or 10 mm x 10 mm, with a 100 &#215; 100 resolution (20 &#181;m/pixel or 100 &#181;m/pixel) and sampling rate of up to 10 kHz (0.1 ms) at maximum. A 1-French size quadripolar electrode pacing catheter is placed into the right atrium through the superior vena cava avoiding any mechanical damage to the atrium, and pacing stimulation is delivered through the catheter. An electrophysiological study is performed with programmed stimulation including constant pacing, burst pacing, and up to triple extrastimuli pacing. Under a spontaneous or pacing rhythm, the optical mapping recorded the action potential duration, activation map, conduction velocity, and Ca2+ transient individually in the right and left atria. In addition, the programmed stimulation also determines the inducibility of atrial tachyarrhythmias. Precise activation mapping is performed to identify the propagation of the excitation in the atrium during an induced atrial tachyarrhythmia. Optical mapping with a specialized setting enables a thorough electrophysiological evaluation of the atrium in murine pathological models.

This protocol describes the electrophysiological evaluation of murine atria utilizing an optical mapping system with a high temporal and spatial resolution, including dual recordings of the membrane voltage and Ca2+ transient under programmed stimulation through a specialized electrode catheter.

Recent genome-wide association studies targeting atrial fibrillation (AF) have indicated a strong association between the genotype and ...

Medicine

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

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

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

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

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

Optical Mapping of Intra-Sarcoplasmic Reticulum Ca2+ and Transmembrane Potential in the Langendorff-perfused Rabbit Heart

Sarcoplasmic reticulum (SR) Ca2+ handling plays a key role in normal excitation-contraction coupling and aberrant SR Ca2+ handling is known to play a significant role in certain types of arrhythmia. Because arrhythmias are spatially distinct, emergent phenomena, they must be investigated at the tissue level. However, methods for directly probing SR Ca2+ in the intact heart remain limited. This article describes the protocol for dual optical mapping of transmembrane potential (Vm) and free intra-SR [Ca2+] ([Ca2+]SR) in the Langendorff-perfused rabbit heart. This approach takes advantage of the low-affinity Ca2+ indicator Fluo-5N, which has minimal fluorescence in the cytosol where intracellular [Ca2+] ([Ca2+]i) is relatively low but exhibits significant fluorescence in the SR lumen where [Ca2+]SR is in the millimolar range. In addition to revealing SR Ca2+ characteristics spatially across the epicardial surface of the heart, this approach has the distinct advantage of simultaneous monitoring of Vm, allowing for investigations into the bidirectional relationship between Vm and SR Ca2+ and the role of SR Ca2+ in arrhythmogenic phenomena.

Optical Mapping of Intra-Sarcoplasmic Reticulum Ca2+ and Transmembrane Potential in the Langendorff-perfused Rabbit Heart

This article describes the detailed protocol and equipment necessary for dual optical mapping of transmembrane potential (Vm) and free intra-sarcoplasmic reticulum (SR) Ca2+ in the Langendorff-perfused rabbit heart. This method allows for direct observation and quantification of Vm and SR Ca2+ dynamics in the intact heart.

Sarcoplasmic reticulum (SR) Ca2+ handling plays a key role in normal excitation-contraction coupling and aberrant SR Ca2+ handling is known to play a ...

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 ...

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

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