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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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. 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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 border="0" cellpadding="0" cellspacing="0" width="756">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:367px;">Name of Material/ Equipment</td>
			<td style="width:103px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Rhod2 dry powder</td>
			<td style="width:103px;">&#160;AAT Bioquest</td>
			<td style="width:119px;">21062</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">pluronic</td>
			<td style="width:103px;">AAT Bioquest</td>
			<td style="width:119px;">20050</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:367px;">DMSO</td>
			<td style="width:103px;">sigma</td>
			<td style="width:119px;">276855</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Rh237</td>
			<td style="width:103px;">Invitrogen</td>
			<td style="width:119px;">&#160;S-1109</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">NaCl</td>
			<td style="width:103px;">sigma</td>
			<td style="width:119px;">S7653</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">KCl</td>
			<td style="width:103px;">sigma</td>
			<td style="width:119px;">P3911</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">KH2PO4&#160;</td>
			<td style="width:103px;">sigma</td>
			<td style="width:119px;">P0662</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">NaH2PO4</td>
			<td style="width:103px;">sigma</td>
			<td style="width:119px;">S9638</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">MgSO4&#160;</td>
			<td style="width:103px;">sigma</td>
			<td style="width:119px;">M7506</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">D-Glucose</td>
			<td style="width:103px;">sigma</td>
			<td style="width:119px;">G8270</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">NaHCO3</td>
			<td style="width:103px;">sigma</td>
			<td style="width:119px;">S6014</td>
			<td></td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;">CaCl2</td>
			<td style="width:103px;">sigma</td>
			<td style="width:119px;">C3881</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">HEPEs</td>
			<td style="width:103px;">sigma</td>
			<td style="width:119px;">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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Biology

10.1K 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

Proper Care and Cleaning of the Microscope

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a microscope causes reduction in contrast and resolution. DIC is especially sensitive to contamination and scratches on the lens surfaces. This protocol details the procedure for keeping the microscope clean.

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a microscope causes reduction in contrast and resolution. DIC is especially sensitive to contamination and scratches on the lens surfaces. This protocol details the procedure for keeping the microscope clean.

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a ...

Phase Contrast and Differential Interference Contrast (DIC) Microscopy

Phase-contrast microscopy is often used to produce contrast for transparent, non light-absorbing, biological specimens. The technique was discovered by Zernike, in 1942, who received the Nobel prize for his achievement. DIC microscopy, introduced in the late 1960s, has been popular in biomedical research because it highlights edges of specimen structural detail, provides high-resolution optical sections of thick specimens including tissue cells, eggs, and embryos and does not suffer from the  phase halos  typical of phase-contrast images. This protocol highlights the principles and practical applications of these microscopy techniques.

Phase Contrast and Differential Interference Contrast DIC Microscopy

This protocol highlights the principles and practical applications of Phase and Differential Interference Contrast (DIC) Microscopy

Phase-contrast microscopy is often used to produce contrast for transparent, non light-absorbing, biological specimens. The technique was discovered by ...

Optical Mapping of Langendorff-perfused Rat Hearts

Optical mapping of the cardiac surface with voltage-sensitive fluorescent dyes has become an important tool to investigate electrical excitation in experimental models that range in scale from cell cultures to whole-organs[1, 2]. Using state-of-the-art optical imaging systems, generation and propagation of action potentials during normal cardiac rhythm or throughout initiation and maintenance of arrhythmias can be visualized almost instantly[1]. The latest commercially-available systems can provide information at exceedingly high spatiotemporal resolutions and were based on custom-built equipment initially developed to overcome the obstacles imposed by more conventional electrophysiological methods[1]. Advancements in high-resolution and high-speed complementary metal-oxide-semiconductor (CMOS) cameras and intensely-bright, light-emitting diodes (LEDs) as well as voltage-sensitive dyes, optics, and filters have begun to make electrical signal acquisition practical for cardiovascular cell biologists who are more accustomed to working with microscopes. Although the newest generation of CMOS cameras can acquire 10,000 frames per second on a 16,384 pixel array, depending on the type of sample preparation, long-established fluorescence acquisition technologies such as photodiode arrays, laser scanning systems, and cooled charged-coupled device (CCD) cameras still have some distinct advantages with respect to dynamic range, signal-to-noise ratio, and quantum efficiency[1, 3]. In the present study, Lewis rat hearts were perfused ex vivo with a crystalloid perfusate (Krebs-Henseleit solution) at 37&deg;C on a modified Langendorff apparatus. After a 20 minute stabilization period, the hearts were intermittently perfused with 11 mMol/L 2,3-butanedione monoxime to eliminate contraction-associated motion during image acquisition. For optical mapping, we loaded hearts with the fast-response potentiometric probe di-8-ANEPPS[4] (5 &mu;Mol/L) and briefly illuminated the preparation with 475&plusmn;15 nm excitation light. During a typical 2 second period of illumination, &gt;605 nm light emitted from the cardiac preparation was imaged with a high-speed CMOS camera connected to a horizontal macroscope. For this demonstration, hearts were paced at 300 beats per minute with a coaxial electrode connected to an isolated electrical stimulation unit. Simultaneous bipolar electrographic recordings were acquired and analyzed along with the voltage signals using readily-available software. In this manner, action potentials on the surface of Langendorff-perfused rat hearts can be visualized and registered with electrographic signals.

This article describes a high temporal and spatial resolution technique to optically image action potential movement on the surface of Langendorff-perfused rat hearts using a potentiometric dye (di-8-ANEPPS).

Optical mapping of the cardiac surface with voltage-sensitive fluorescent dyes has become an important tool to investigate electrical excitation in ...

Optical Recording of Electrical Activity in Guinea-pig Enteric Networks using Voltage-sensitive Dyes

The enteric nervous system (ENS) is a self-contained network with identified functions, capable of performing complex behaviors in isolation. Its neurons (10 to 25 &mu;m in diameter) are arranged in plexuses that are confined to distinct planes of the gut wall 1; the myenteric plexus can be found between the longitudinal and circular muscle layers, and the submucous plexus between the circular muscle layer and the mucosa. Since the effector systems for these plexuses (transporting epithelium, endocrine cells, immune elements, blood vessels and smooth muscle) are also contained within the gut wall, semi-intact preparations can be dissected that preserve individual components of different reflex pathways. The behavior of the effector systems is controlled by the submucous and myenteric plexuses acting in concert. Therefore, detailed knowledge of synaptic interactions within and between ganglia, and of communication between the plexuses, is essential for understanding normal gastrointestinal function. The ENS, as an intact nervous system, is a unique experimental model in which one can correlate molecular and cellular events with the electrical behavior of the neuronal network and its physiological outputs. Because of the quasi-two-dimensional organization of its plexuses, the ENS is particularly well suited for the study of neural networks using multiple site optical recording techniques that employ voltage-sensitive dyes 2,7,8,9. We will illustrate here the use of a relatively new naphthylstyryl-pyridinium dye (di-4-ANEPPDHQ) 3 that offers multiple advantages over its predecessors, including very low phototoxicity, slow rate of internalization, and remarkable chemical stability. When used in conjunction with a camera that permits sub-millisecond time resolution, this dye allows us to monitor the electrical activity of all the neurons in the field of view with a maximal spatial resolution of ~ 2.5 &mu;m at 100X magnification. At lower magnification (10X or 20X), the sacrifice of single-cell resolution is compensated by a gain in perspective, revealing the intricacies of the inter-ganglionic circuitry.

This protocol illustrates how voltage-sensitive dyes enable optical recording of electrical activity from intact neural networks such as the plexuses of the guinea-pig enteric nervous system, with an adjustable resolution that ranges from single-cells to multi-ganglionic circuitry.

The enteric nervous system (ENS) is a self-contained network with identified functions, capable of performing complex behaviors in isolation. Its neurons ...

Photoconversion of Purified Fluorescent Proteins and Dual-probe Optical Highlighting in Live Cells

Photoconvertible fluorescent proteins (pc-FPs) are a class of fluorescent proteins with "optical highlighter" capability, meaning that the color of fluorescence can be changed by exposure to light of a specific wavelength. Optical highlighting allows noninvasive marking of a subpopulation of fluorescent molecules, and is therefore ideal for tracking single cells or organelles.
Critical parameters for efficient photoconversion are the intensity and the exposure time of the photoconversion light. If the intensity is too low, photoconversion will be slow or not occur at all. On the other hand, too much intensity or too long exposure can photobleach the protein and thereby reduce the efficiency of photoconversion.
This protocol describes a general approach how to set up a confocal laser scanning microscope for pc-FP photoconversion applications. First, we describe a procedure for preparing purified protein droplet samples. This sample format is very convenient for studying the photophysical behavior of fluorescent proteins under the microscope. Second, we will use the protein droplet sample to show how to configure the microscope for photoconversion. And finally, we will show how to perform optical highlighting in live cells, including dual-probe optical highlighting with mOrange2 and Dronpa.

This protocol describes a general approach to perform photoconversion of fluorescent proteins on a confocal laser scanning microscope. We describe procedures for the photoconversion of puried protein samples, as well as for dual-probe optical highlighting in live cells with mOrange2 and Dronpa.

Photoconvertible fluorescent proteins (pc-FPs) are a class of fluorescent proteins with "optical highlighter" capability, meaning that the color of ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

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 ...

Reconstitution of a Transmembrane Protein, the Voltage-gated Ion Channel, KvAP, into Giant Unilamellar Vesicles for Microscopy and Patch Clamp Studies

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow GUVs must be modified in order to form GUVs containing functional transmembrane proteins. This article describes two dehydration-rehydration methods &#8212; electroformation and gel-assisted swelling &#8212; to form GUVs containing the voltage-gated potassium channel, KvAP. In both methods, a solution of protein-containing small unilamellar vesicles is partially dehydrated to form a stack of membranes, which is then allowed to swell in a rehydration buffer. For the electroformation method, the film is deposited on platinum electrodes so that an AC field can be applied during film rehydration. In contrast, the gel-assisted swelling method uses an agarose gel substrate to enhance film rehydration. Both methods can produce GUVs in low (e.g., 5 mM) and physiological (e.g., 100 mM) salt concentrations. The resulting GUVs are characterized via fluorescence microscopy, and the function of reconstituted channels measured using the inside-out patch-clamp configuration. While swelling in the presence of an alternating electric field (electroformation) gives a high yield of defect-free GUVs, the gel-assisted swelling method produces a more homogeneous protein distribution and requires no special equipment.

The reconstitution of the transmembrane protein, KvAP, into giant unilamellar vesicles (GUVs) is demonstrated for two dehydration-rehydration methods &#8212; electroformation, and gel-assisted swelling. In both methods, small unilamellar vesicles containing the protein are fused together to form GUVs that can then be studied by fluorescence microscopy and patch-clamp electrophysiology.

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow ...

Single-channel Analysis and Calcium Imaging in the Podocytes of the Freshly Isolated Glomeruli

Podocytes (renal glomerular epithelial cells) are known to regulate glomerular permeability and maintain glomerular structure; a key role for these cells in the pathogenesis of various renal diseases has been established since podocyte injury leads to proteinuria and foot process effacement. It was previously reported that various endogenous agents may cause a dramatic overload in intracellular Ca2+ concentration in podocytes, presumably leading to albuminuria, and this likely occurs via calcium-conducting ion channels. Therefore, it appeared important to study calcium handling in the podocytes both under normal conditions and in various pathological states. However, available experimental approaches have remained somewhat limited to cultured and transfected cells. Although they represent a good basic model for such studies, they are essentially extracted from the native environment of the glomerulus. Here we describe the methodology of studying podocytes as a part of the freshly isolated whole glomerulus. This preparation retains the functional potential of the podocytes, which are still attached to the capillaries; therefore, podocytes remain in the environment that conserves the major parts of the glomeruli filtration apparatus. The present manuscript elaborates on two experimental approaches that allow 1) real-time detection of calcium concentration changes with the help of ratiometric confocal fluorescence microscopy, and 2) the recording of the single ion channels activity in the podocytes of the freshly isolated glomeruli. These methodologies utilize the advantages of the native environment of the glomerulus that enable researchers to resolve acute changes in the intracellular calcium handling in response to applications of various agents, measure basal concentration of calcium within the cells (for instance, to evaluate disease progression), and assess and manipulate calcium conductance at the level of single ion channels.

Changes in the intracellular calcium levels in the podocytes are one of the most important means to control the filtration function of glomeruli. Here we explain a high-throughput approach that allows detection of real-time calcium handling and single ion channels activity in the podocytes of the freshly isolated glomeruli.

Podocytes (renal glomerular epithelial cells) are known to regulate glomerular permeability and maintain glomerular structure; a key role for these cells ...

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 ...