This work is supported by research grants from the National Institutes of Health (HL068915; HL088206) and contributions to the Cardiac Conduction Fund at Children's Hospital Boston.

Part 1: Prepare solutions and the isolated perfused heart system
<ol>
<li>On the morning of the experiment, 4.0 L of Krebs-Henseleit solution is prepared as previously described[5, 6].</li>
<li>11 mMol/L 2,3-butanedione monoxime (BDM) is dissolved in 1.0 L of Krebs-Henseleit solution decanted from the perfusate prepared in step 1.1.</li>
<li>An additional 150 mL of Krebs-Henseleit is removed from the perfusate prepared in step 1.1 and mixed with 5 Mol/L di-8-ANEPPS (diluted from a 10 mMol/L stock dissolved in dimethyl sulfoxide (DMSO)).</li>
<li>These 3 solutions are transferred to water-jacketed glass reservoirs (Radnoti) where they are pre-warmed to 41.0&deg; C and oxygenated with a submerged bubbler (Radnoti) using 0.2 &mu;m-filtered 95% O2; 5% CO2 gas. Solutions from the 3 stock reservoirs are pumped to the wall-mounted Langendorff system with MasterFlex&trade; L/S peristaltic pumps and low-absorption silicone tubing (Cole-Parmer). </li>
<li>Prior to assembly,the Langendorff system glassware (Radnoti) is meticulously washed with E-TOXA-CLEAN reagent (Sigma-Aldrich) and thoroughly rinsed with 0.1 M/L HCl, 100% ethanol, and distilled water.</li>
<li>The non-recirculating Langendorff apparatus was constructed to deliver 70 mm Hg constant-pressure from 3 separate oxygenated reservoirs, which are separated from the heart with water-jacketed heating coils and bubble traps (Radnoti)[7]. The 3 independent perfusion lines converge above the heart at two 3-way stopcocks, allowing us to precisely control delivery of the solutions prepared in steps 1.1 to 1.3. Water-jacketed glassware is connected in series with MasterFlex PharMed BPT tubing (Cole-Parmer) and warmed to 39&deg; C with distilled water using two E100 circulators (Lauda). Heat loss through the platinum-cured silicone tubing (Cole-Parmer) that connects the pressure-head reservoirs to the heating coils and bubble traps resulted in a 37&deg; C perfusate reaching the heart. </li>
</ol>
Part 2: Harvest the rat heart and set-up Langendorff perfusion
<ol>
<li>To induce deep general anesthesia in 200-250 g Lewis rats, 100 mg/kg Ketamine and 10 mg/kg Xylazine is injected into the intraperitoneal cavity. To this mixture, we add 500 U/kg Heparin to prevent blood coagulation and myocardial ischemia during the explantation procedure.</li>
<li>For easy access to the heart and great vessels, the anterior chest wall is removed. After that, surrounding tissue is carefully dissected and the pericardial sack opened.</li>
<li>Following identification of the inferior vena cava, this vessel is ligated with 5-0 silk (Ethicon) and the entire heart-lung block is explanted. Tissue is immediately placed in ice-cold Krebs-Henseleit solution contained in a 50 mL beaker on ice.</li>
<li>The ascending aorta is quickly identified and dissected from the surrounding tissue. An appropriately-sized cannula (Harvard Apparatus) is inserted into the aorta, taking care to avoid interrupting obligatory perfusion of the coronary arteries by inserting the cannula too far into the aortic root. The cannula is secured to the ascending aorta with 5-0 silk (Ethicon).</li>
<li>The rat heart is then placed on the Langendorff apparatus without introducing air bubbles into the cannula. Retrograde coronary vascular perfusion is now established with the warm, oxygenated Krebs-Henseleit solution from the pressure-head described in step 1.6.</li>
<li>Extra tissue, including the lungs, is now removed and the heart is perfused for 20 minutes to permit recovery of function and stabilize the rhythm. During this time, a very thin thermocouple temperature probe (Cole-Parmer) is introduced into the left ventricular cavity and sutured in place with 5-0 Prolene suture (Ethicon). The probe is connected to a thermo-controller (Digi-Sense) to ensure the temperature of the heart is maintained at 37&deg; C by adjusting the settings of the water circulatory pumps. Motion from perfusate dripping from the cardiac apex is minimized by placing a piece of gauze in the effluent receptacle.</li>
</ol>
Part 3: Load heart with potentiometric dye and acquire electrographic and optical signals
<ol>
<li>Di-8-ANEPPS (Invitrogen) is loaded into the heart by switching to the perfusion line containing the Krebs-Henseleit mixed with the fluorescent dye using a stopcock. In addition, an 18G cannula is placed into the left and/or right atrium and additional 50 mL of dye solution is slowly administered into each of these chambers because they are not sufficiently perfused with dye introduced through the coronary arteries.</li>
<li>During the loading procedure, 3 ECG leads (Harvard Apparatus) are gently placed on the surface of the heart that is not facing the optics used for mapping. Electrode No 1 is positioned on the posterior apical portion of the left ventricle, No 2 on the left atrium, and No 3 as a reference electrode on the aortic root (Figure 1). The atrial and ventricular electrographic signals are subsequently amplified, digitized, and displayed alongside optical signals using the software (RedShirt Imaging) (Figure 2). An oscilloscope (Tektronix model TDS 1002) is also used to visualize the surface ECG s in real-time and to assure adequate pacing.</li>
<li>ECG amplifier (Hugo Sachs Elektronik) settings:
High Pass Filter: 0.1 Hz Low Pass Filter: 150 Hz</li>
<li>The CMOS camera (RedShirt Imaging) and macroscope is positioned using the X-Y-Z adjustments so that the surface of the heart is in focus and centered in the acquisition frame. The camera and optics are mounted on a vibration isolation table (Minus K Technology) to minimize resonant frequencies. At the same time, a coaxial pacing electrode (Harvard Apparatus) controlled with an isolated, S48 electrical-stimulation unit (Grass), is placed on the right atrium and the heart is paced at 300 beats per minute (Figure 1).</li>
<li>Electrical stimulation (Grass) settings: Rate: 5 pulses per second Delay: 0.2 ms Duration: 2 ms Volts: 6-12 V Mode: Repeat Pulse: Single</li>
<li>For optical recordings which lack motion artifacts from contraction, the heart needs to be electromechanically uncoupled. We do this by again switching perfusion lines to Krebs-Henseleit solution that contains 11 mMol/L BDM. Between acquisitions, the heart is perfused with unadulterated Krebs-Henseleit to help preserve viability of the preparation.</li>
<li>The recording parameters are set using the software (RedShirt Imaging) with the following acquisition settings: Configuration: 2,000 Hz; 128x128 pixel array Frame Interval: 0.5 ms Camera Amplifier Gain: 5x On-Chip Gain: 8x-12Me-well Shutter: 500 ms delay Number of Frames: 4000 Duration: 2000 ms</li>
<li>All room and equipment lights are turned off or shielded to eliminate background noise during recording. The LED light illuminates the heart only during the optical recording to reduce photo-bleaching and dye toxicity. The light source shutter is controlled with a 5 V pulse delivered through the control panel by way of a D-to-A board in the computer (RedShirt Imaging).</li>
</ol>
Part 4: Analyze acquisition information using RedShirt Imaging Software
<ol>
<li>Following acquisition, data is processed using different filter settings. We generally use the default settings except when adjusting the Band Stop/Pass filter, which is set with the left boundary at 44.0 and the right boundary at 98.0. Afterward the recorded information is processed and a movie is generated (RedShirt Imaging).</li>
<li>Data from one acquisition corresponds to the local electrical activation at 16,384 sites on the heart surface over a period of 2 seconds. The software allows these local signals to be directly compared with one another and with the atrial and ventricular electrographic recordings. Data is then visualized by mapping local electrical activation to color and rendering this information as an animation showing the spatiotemporal electrical activation on the cardiac surface. To create such an animation, we use the software to: 
<ul>
<li>temporally and/or spatially filter the data</li>
<li>select a start and end time for the animation</li>
<li>map optical signals to color based on resting light intensity of each pixel</li>
<li>overlay the resulting color data with a picture of the heart </li>
<li>generate the animation</li>
</ul>
</li>
</ol>
Part 5: Representative results
If the perfused heart preparation was motionless during recording, the optical signals show one distinct peak for every pixel involved in a change of the emission intensity of di-8-ANEPPS. The corresponding movies (Figures 3 and 4) demonstrate an excitation wave front propagating across the epicardial surface of the heart as well as the simultaneously acquired electrographic recordings (Figure 2).
<img src="/files/ftp_upload/1138/Figure3.jpg" alt="Figure 1" /> Figure 1. A photograph of a Langendorff-perfused heart preparation depicting the positions of the pacing electrode on the right atrium and the ECG leads as described in step 3.2.
<img src="/files/ftp_upload/1138/Figure4thumb.jpg" alt="Figure 2" border="0" /> Figure 2. Representative optical signals and electrographic recordings from a perfused Lewis rat heart. Panel A shows an image of the epicardial surface used for optical imaging. The position of the pixels selected to demonstrate the changes in fluorescence emission over time in Panel B are indicated by colored arrows. Electrographic signals are shown in Panel C with the red line indicating atrial activation and the light blue line corresponding to the ventricular signal. Please <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/1138/Figure4.jpg">click here</a> to see a larger version of this figure.

<ol>
	<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>Hucker, W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Images+in+cardiovascular+medicine.+Optical+mapping+of+the+human+atrioventricular+junction.">Images in cardiovascular medicine. Optical mapping of the human atrioventricular junction.</a> Circulation. 117 (11), 1474-147 (2008).</li><li>Entcheva, E., Bien, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Macroscopic+optical+mapping+of+excitation+in+cardiac+cell+networks+with+ultra-high+spatiotemporal+resolution.">Macroscopic optical mapping of excitation in cardiac cell networks with ultra-high spatiotemporal resolution.</a> Prog Biophys Mol Biol. 92 (2), 232-257 (2006).</li><li>Entcheva, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescence+imaging+of+electrical+activity+in+cardiac+cells+using+an+all-solid-state+system.">Fluorescence imaging of electrical activity in cardiac cells using an all-solid-state system.</a> IEEE Trans Biomed Eng. 51 (2), 333-341 (2004).</li><li>Stamm, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+endotoxin-induced+alterations+in+myocardial+calcium+handling:+obligatory+role+of+cardiac+TNF-alpha.">Rapid endotoxin-induced alterations in myocardial calcium handling: obligatory role of cardiac TNF-alpha.</a> Anesthesiology. 95 (6), 1396-1405 (2001).</li><li>Choi, Y. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+conduction+through+engineered+tissue.">Cardiac conduction through engineered tissue.</a> Am J Pathol. 169 (1), 72-85 (2006).</li><li>Skrzypiec-Spring, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolated+heart+perfusion+according+to+Langendorff---still+viable+in+the+new+millennium.">Isolated heart perfusion according to Langendorff---still viable in the new millennium.</a> J Pharmacol Toxicol Methods. 55 (2), 113-126 (2007).</li><li>Fedorov, V. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Application+of+blebbistatin+as+an+excitation-contraction+uncoupler+for+electrophysiologic+study+of+rat+and+rabbit+hearts.">Application of blebbistatin as an excitation-contraction uncoupler for electrophysiologic study of rat and rabbit hearts.</a> Heart Rhythm. 4 (5), 619-626 (2007).</li></ol>

Experiments on animals were performed in accordance with the guidelines and regulations set forth by the Institutional Animal Care and Use Committee at Children's Hospital Boston.

Removal of the heart from anesthetized rats must be performed rapidly to avoid myocardial ischemia. If ischemia or insufficient coronary perfusion occurs, the heart will likely develop arrhythmias and may become infarcted. Additionally, these hearts will show insufficient fluorescence emission for informative recordings and subsequent analyses. Before loading the dye, myocardial cells need to be adequately perfused with Krebs-Henseleit solution to establish and maintain a physiologic electrolyte milieu for the stability of electrical impulses. Accurate preparation of the perfusate is also necessary to maintain proper organ viability and function. Differences in the electrolyte concentration or insufficient filtering of the perfusate will probably lead to fatal myocardial dysfunction and heart rhythm disorders. For the optical recordings, the heart must be completely loaded with voltage-sensitive dye. This is especially important for the atrial myocardium as these chambers are not well-perfused by the coronary arteries. We have found that additional intra-cavitary perfusion of the atria will establish a good optical signal. Furthermore, acquisition of top-quality voltage tracings requires the perfused heart to be motionless; otherwise, changes in fluorescence emission cannot be reliably used to track changes in membrane potential with high fidelity due to artifacts caused by signal drift. This will result in multiple peaks for one pixel rather than a single peak. Other methods to eliminate motion artifacts from optical recordings include mechanical immobilization, treatment with other excitation-contraction uncouplers (e.g. cytochalasin D, blebbistatin), signal processing, and by mathematical modeling[1, 8]. Finally, the method described here merely provides information regarding action potential movement on the cardiac epicardial surface. Alternative tissue preparations and infrared potentiometric dyes may resolve the electrical propagation characteristics in other regions of the heart.

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		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
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<tr>
<td>CardioCMOS-SM128f</td>
<td>Equipment</td>
<td>RedShirt Imaging, Decatur, GA 30030 USA </td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>CardioPlex Software</td>
<td>Equipment</td>
<td>RedShirt Imaging, Decatur, GA 30030 USA</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>LUXEON LED Light Source 460-490 nm </td>
<td>Equipment</td>
<td>Lumileds Lighting, US, LLC, San Jose, CA 95131 USA</td>
<td>LXHL-PB02</td>
<td>&nbsp;</td>
<tr>
<td>ECG Amplifier Type 689 Hugo Sachs Elektronik</td>
<td>Equipment</td>
<td>Harvard Apparatus, Holliston, MA 01746 USA</td>
<td>730149</td>
<td>&nbsp;</td>
<tr>
<td>Dichroic Mirror 505 nm</td>
<td>Equipment</td>
<td>Semrock, Rochester, NY 14624 USA</td>
<td>FF505-SDi01-25x36</td>
<td>&nbsp;</td>
<tr>
<td>Emission Filter 605 nm Long Pass</td>
<td>Equipment</td>
<td>SciMedia, Costa Mesa, CA 92626 USA </td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>THT Sideways</td>
<td>Equipment</td>
<td>SciMedia, Costa Mesa, CA 92626 USA</td>
<td>25 BM-8</td>
<td>&nbsp;</td>
<tr>
<td>Mini Ball Joint Holder</td>
<td>Equipment</td>
<td>Harvard Apparatus, Holliston, MA 01746 USA</td>
<td>BS4 73-0177</td>
<td>&nbsp;</td>
<tr>
<td>Small Stimulation Electrode Set</td>
<td>Equipment</td>
<td>Harvard Apparatus, Holliston, MA 01746 USA</td>
<td>BS4 73-0160</td>
<td>&nbsp;</td>
<tr>
<td>BM-6 Benchtop Vibration Isolation Platform</td>
<td>Equipment</td>
<td>Technology Inc., Inglewood, CA 90301</td>
<td>25 BM-6</td>
<td>&nbsp;</td>
<tr>
<td>Monopolar ECG Electrode</td>
<td>Equipment</td>
<td>Harvard Apparatus, Holliston, MA 01746 USA</td>
<td>BS4 73-0200</td>
<td>&nbsp;</td>
<tr>
<td>Roller Pump SCI 400</td>
<td>Equipment</td>
<td>Watson-Marlow Bredel Inc., Wilmington, MA 01887 USA</td>
<td>401U/D1</td>
<td>&nbsp;</td>
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<td>Roller Pump MasterFlex Easy Load II</td>
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<td>Cole Parmer, Vernon Hills, Illinois 60061 USA</td>
<td>Model 77201-60</td>
<td>&nbsp;</td>
<tr>
<td>Tubing Marprene #14</td>
<td>Equipment</td>
<td>Watson-Marlow Bredel Inc., Wilmington, MA 01887 USA</td>
<td>902.0016.016</td>
<td>&nbsp;</td>
<tr>
<td>MasterFlex Tubing</td>
<td>Equipment</td>
<td>PharMed, Westlake, OH 44145 USA</td>
<td>06485-25</td>
<td>&nbsp;</td>
<tr>
<td>S48 Square Pulse Stimulator</td>
<td>Equipment</td>
<td>Grass Technologies, West Warwick, RI 02893 USA</td>
<td>Model S48</td>
<td>&nbsp;</td>
<tr>
<td>SIU5 RF TRANSFORMER ISOLATION UNIT</td>
<td>Equipment</td>
<td>Grass Technologies, West Warwick, RI 02893 USA</td>
<td>Model SIU5</td>
<td>&nbsp;</td>
<tr>
<td>5 Liter Water Jacketed Reservoir</td>
<td>Equipment</td>
<td>Radnoti Glass Technology Inc., Monrovia CA 91016 USA</td>
<td>120142-5</td>
<td>&nbsp;</td>
<tr>
<td>2 Liter Water Jacketed Reservoir</td>
<td>Equipment</td>
<td>Radnoti Glass Technology Inc., Monrovia CA 91016 USA</td>
<td>120142-2</td>
<td>&nbsp;</td>
<tr>
<td>0.5 Liter Water Jacketed Reservoir</td>
<td>Equipment</td>
<td>Radnoti Glass Technology Inc., Monrovia CA 91016 USA</td>
<td>120142-0</td>
<td>&nbsp;</td>
<tr>
<td>0.25 Liter Water Jacketed Reservoir</td>
<td>Equipment</td>
<td>Radnoti Glass Technology Inc., Monrovia CA 91016 USA </td>
<td>120142-025</td>
<td>&nbsp;</td>
<tr>
<td>10 ml Heating Coil</td>
<td>Equipment</td>
<td>Radnoti Glass Technology Inc., Monrovia CA 91016 USA</td>
<td>158822</td>
<td>&nbsp;</td>
<tr>
<td>Compliance Bubble Trap</td>
<td>Equipment</td>
<td>Radnoti Glass Technology Inc., Monrovia CA 91016 USA</td>
<td>130149</td>
<td>&nbsp;</td>
<tr>
<td>Luer Disconnect Cannula</td>
<td>Equipment</td>
<td>Harvard Apparatus, Holliston, MA 01746 USA</td>
<td>72-1444</td>
<td>&nbsp;</td>
<tr>
<td>3-Way stopcock, FLL to MLT, No Port Covers</td>
<td>Equipment</td>
<td>Harvard Apparatus, Holliston, MA 01746 USA</td>
<td>BS4 72-2630</td>
<td>&nbsp;</td>
<tr>
<td>Thermocouple Thermometer</td>
<td>Equipment</td>
<td>Cole Parmer, Vernon Hills, Illinois 60061 USA</td>
<td>WU-91100-40</td>
<td>&nbsp;</td>
<tr>
<td>Ultra Fine IT-Series Flexible Microprobe </td>
<td>Equipment</td>
<td>PhysiTemp Instruments Inc., Clifton, NJ 07013 USA</td>
<td>IT-24P</td>
<td>&nbsp;</td>
<tr>
<td>Oscilloscope Tektronix TDS 1002</td>
<td>Equipment</td>
<td>Tektronix Inc., Beaverton, OR 97005 USA</td>
<td>TDS 1002B</td>
<td>&nbsp;</td>
<tr>
<td>2,3-Butanedione monoxime </td>
<td>Reagent</td>
<td>Sigma, St. Louis, MO 63132 USA</td>
<td>B0753</td>
<td>&nbsp;</td>
<tr>
<td>Ketamine HCl</td>
<td>Reagent</td>
<td>Hospira Inc., Lake Forest, IL 60045 USA</td>
<td>RL-0065</td>
<td>&nbsp;</td>
<tr>
<td>Xylazine</td>
<td>Reagent</td>
<td>Lloyd Inc., Iowa 51601 USA</td>
<td>LB15705A</td>
<td>&nbsp;</td>
<tr>
<td>E-TOXA-CLEAN&reg;</td>
<td>Reagent</td>
<td>Sigma, St. Louis, MO 63132 USA </td>
<td>E9029</td>
<td>&nbsp;</td>
<tr>
<td>Di-8-ANEPPS</td>
<td>Reagent</td>
<td>Invitrogen, Carlsbad, CA 92008 USA</td>
<td>D-3167</td>
<td>&nbsp;</td>		</tbody>
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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.

Watch this Scientific Journal Video about Optical Mapping of Langendorff-perfused Rat Hearts at JoVE.com

Optical Mapping of Langendorff-perfused Rat Hearts

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-mapping-of-langendorff-perfused-rat-hearts

Nanyang Technological University

Research

JoVE Journal

Biology

20.7K Views.  Children's Hospital Boston and Harvard Medical School. 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).

Video: Optical Mapping of Langendorff-perfused Rat Hearts

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.

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Major Components of the Light Microscope

The light microscope is a basic tool for the cell biologist, who should have a thorough understanding of how it works, how it should be aligned for different applications, and how it should be maintained as required to obtain maximum image-forming capacity and resolution. The components of the microscope are described in detail here.

The light microscope is a basic tool for the cell biologist, who should have a thorough understanding of how it works, how it should be aligned for different applications, and how it should be maintained as required to obtain maximum image-forming capacity and resolution. The components of the microscope are described in detail here.

The light microscope is a basic tool for the cell biologist, who should have a thorough understanding of how it works, how it should be aligned for ...

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

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Semi-automated Optical Heartbeat Analysis of Small Hearts

We have developed a method for analyzing high speed optical recordings from Drosophila, zebrafish and embryonic mouse hearts (Fink, et. al., 2009). Our Semi-automatic Optical Heartbeat Analysis (SOHA) uses a novel movement detection algorithm that is able to detect cardiac movements associated with individual contractile and relaxation events. The program provides a host of physiologically relevant readouts including systolic and diastolic intervals, heart rate, as well as qualitative and quantitative measures of heartbeat arrhythmicity. The program also calculates heart diameter measurements during both diastole and systole from which fractional shortening and fractional area changes are calculated. Output is provided as a digital file compatible with most spreadsheet programs. Measurements are made for every heartbeat in a record increasing the statistical power of the output. We demonstrate each of the steps where user input is required and show the application of our methodology to the analysis of heart function in all three genetically tractable heart models.

We have developed a Semi-automated Optical Heartbeat Analysis method (SOHA) for analyzing high speed optical recordings from Drosophila, zebrafish and embryonic mouse hearts. We demonstrate the application of our methodology to the analysis of heart function in fruit fly and embryonic mouse hearts.

We have developed a method for analyzing high speed optical recordings from Drosophila, zebrafish and embryonic mouse hearts (Fink, et. al., 2009). Our ...

Single Cell Fate Mapping in Zebrafish

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, lymphatic, and blood vessel lineages arise in developing embryos requires fate mapping and lineage tracing of undifferentiated precursor cells. Recently, photoactivatable proteins which include: Eos1, 2, PAmCherry3, Kaede4-7, pKindling8, and KikGR9, 10 have received wide interest as cell tracing probes. The fluorescence spectrum of these photosensitive proteins can be easily converted with UV excitation, allowing a population of cells to be distinguished from adjacent ones. However, the photoefficiency of the activated protein may limit long-term cell tracking11. As an alternative to photoactivatable proteins, caged fluorescein-dextran has been widely used in embryo model systems7, 12-14. Traditionally, to uncage fluorescein-dextran, UV excitation from a fluorescence lamp house or a single photon UV laser has been used; however, such sources limit the spatial resolution of photoactivation. Here we report a protocol to fate map, lineage trace, and detect single labeled cells. Single cells in embryos injected with caged fluorescein-dextran are photoactivated with near-infrared laser pulses produced from a titanium sapphire femtosecond laser. This laser is customary in all two-photon confocal microscopes such as the LSM 510 META NLO microscope used in this paper. Since biological tissue is transparent to near-infrared irradiation15, the laser pulses can be focused deep within the embryo without uncaging cells above or below the selected focal plane. Therefore, non-linear two-photon absorption is induced only at the geometric focus to uncage fluorescein-dextran in a single cell. To detect the cell containing uncaged fluorescein-dextran, we describe a simple immunohistochemistry protocol16 to rapidly visualize the activated cell. The activation and detection protocol presented in this paper is versatile and can be applied to any model system. 
Note: The reagents used in this protocol can be found in the table appended at the end of the article.

A method is described to photoactivate single cells containing a caged fluorescent protein using two-photon absorption from a Ti:Sapphire femtosecond laser oscillator. To fate map the photoactivated cell, immunohistochemistry is used. This technique can be applied to any cell type.

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, ...

Cell Labeling and Injection in Developing Embryonic Mouse Hearts

Testing the fate of embryonic or pluripotent stem cell-derivatives in in vitro protocols has led to controversial outcomes that do not necessarily reflect their in vivo potential. Preferably, these cells should be placed in a proper embryonic environment in order to acquire their definite phenotype. Furthermore, cell lineage tracing studies in the mouse after labeling cells with dyes or retroviral vectors has remained mostly limited to early stage mouse embryos with still poorly developed organs. To overcome these limitations, we designed standard and ultrasound-mediated microinjection protocols to inject various agents in targeted regions of the heart in mouse embryos at E9.5 and later stages of development. &#160;Embryonic explant or embryos are then cultured or left to further develop in utero. These agents include fluorescent dyes, virus, shRNAs, or stem cell-derived progenitor cells. Our approaches allow for preservation of the function of the organ while monitoring migration and fate of labeled and/or injected cells. These technologies can be extended to other organs and will be very helpful to address key biological questions in biology of development.

We describe a series of methods to inject dyes, DNA vectors, virus, and cells in order to monitor both cell fate and phenotype of endogenous and grafted cells derived from embryonic or pluripotent cells within mouse embryos at embryonic day (E)9.5 and later stages of development.

Testing the fate of embryonic or pluripotent stem cell-derivatives in in vitro protocols has led to controversial outcomes that do not necessarily reflect ...

Recovery of Adult Zebrafish Hearts for High-throughput Applications

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and purification of RNA, DNA, and proteins. All of these applications demand the rapid recovery of significant numbers of zebrafish hearts to avoid gene regulatory, metabolic, and other changes that begin after death. Adult zebrafish hearts are also required for studying heart structure for a variety of mutants and for studying heart regeneration. However, the traditional zebrafish heart dissection is slow and difficult and requires specialized tools, making large-scale dissection of adult zebrafish hearts tedious. Traditional methods also harbor the risk of damaging the heart during the dissection. Here, we describe a method for dissection of adult zebrafish hearts that is fast, reproducible, and preserves heart architecture. Furthermore, this method does not require specialized tools, is painless for the zebrafish, can be performed on fresh or fixed specimens, and can be performed on zebrafish as young as one month old. The approach described expands the use of adult zebrafish for cardiovascular research.

Use of zebrafish for cardiovascular research is expanding towards research on adult hearts. For these applications, quick and simple isolation of cardiac tissues is key to avoid post-mortem changes and to obtain an adequate number of samples. Here, we describe a fast and reproducible method for dissecting adult zebrafish hearts.

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and ...

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.

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 ...

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 ...

High-resolution Optical Mapping of the Mouse Sino-atrial Node

Sino-atrial node (SAN) dysfunctions and associated complications constitute important causes of morbidity in patients with cardiac diseases. The development of novel pharmacological therapies to cure these patients relies on the thorough understanding of both normal physiology and pathophysiology of the SAN. Among the studies of cardiac pacemaking, the mouse SAN is widely used due to its feasibility for modifications in the expression of different genes that encode SAN ion channels or calcium handling proteins. Emerging evidence from electrophysiological and histological studies has also proved the representativeness and similarity of the mouse SAN structure and functions to larger mammals, including the presence of specialized conduction pathways from the SAN to the atrium and a complex pacemakers' hierarchy within the SAN. Recently, the technique of optical mapping has greatly facilitated the exploration and investigation of the origin of excitation and conduction within and from the mouse SAN, which in turn has extended the understanding of the SAN and benefited clinical treatments of SAN dysfunction associated diseases. In this manuscript, we have described in detail how to perform the optical mapping of the mouse SAN from the intact, Langendorff-perfused heart and from the isolated atrial preparation. This protocol is a useful tool to enhance the understanding of mouse SAN physiology and pathophysiology.

Here, we present a protocol for optical mapping of electrical activity from the mouse right atrium and especially the sino-atrial node, at a high spatial and temporal resolution.