We are supported by the University of Wisconsin-Madison Medical School start-up (A.V.G.).

All experiments were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals (NIH Pub. No. 80-23). All methods and protocols used in these studies have been approved by The University of Wisconsin Animal Care and Use Protocol Committee following the Guidelines for Care and Use of Laboratory Animals published by NIH (publication No. 85-23, revised 1996). All animals used in this study received humane care in compliance with the Guide for the Care and Use of Laboratory Animals.
1. Heart Removal and Langendorff Perfusion
<ol>
	<li>Prior to heart isolation, warm Tyrode solution to 37 &#176;C by using a water bath and a water jacket.</li>
	<li>Anesthetize the mouse with 5% isoflurane/95% O2.</li>
	<li>Ensure appropriate level of anesthesia by checking the loss of pain reflex.</li>
	<li>Perform thoracotomy. Open the chest by using curved 5.5&#34; Mayo scissors and 5.5&#34; Kelly hemostatic forceps to make a 1 cm cut on the front of the thorax.
	<ol>
		<li>Quickly extract the heart from the chest (within 30 sec). Use 4&#34; curved Iris forceps to grab the lung tissue and cut out the lung, thymus and heart together with the pericardium using 4.3&#34; Iris scissors. Do not grab the heart directly.</li>
		<li>Wash it in oxygenated (95% O2 / 5% CO2), constant-temperature (36.8 &#177; 0.4&#176;C) modified Tyrode&#39;s solution. Use the following solution composition (in mM): 128.2 NaCl, 4.7 KCl, 1.19 NaH2PO4, 1.05 MgCl2, 1.3 CaCl2, 20.0 NaHCO3, and 11.1 glucose (pH 7.35 &#177; 0.05).</li>
	</ol>
	</li>
	<li>While bathed in the same solution, identify the lung, thymus, and fat tissue. Carefully dissect them from the heart. Use curved 3&#34; Vannas-Tubingen scissors and 4.3&#34; #5 forceps.</li>
	<li>Then identify the aorta and cannulate it onto a custom-made 21 G cannula. Use two curved 4.3&#34; #5B forceps.</li>
	<li>After cannulation, simultaneously perfuse (through the cannula at a flow rate of ~5 ml/min; this should be adjusted based on the aortic pressure, see below) and superfuse (in the bath at a flow rate of ~50 ml/min) the heart with warmed and filtered Tyrode solution throughout the entire experiment. Pass the perfusion solution through an in-line 11 &#181;m nylon filter to prevent clogging of coronary circulation.</li>
	<li>Using a pressure transducer (or a pressure gauge) connected via a 3-way stopcock Luer lock to the perfusion line, monitor the aortic pressure and maintain it between 60 and 80 mmHg. Adjust perfusion speed if required to keep the aortic pressure within this range.</li>
</ol>
2. Optical Mapping of the SAN from the Langendorff-perfused Whole Heart
<ol>
	<li>Instrumentation
	<ol>
		<li>Place the heart horizontally and pin the ventricular apex to the silicon-coated bottom of the chamber using 0.1 mm diameter pins to prevent stream-induced movement during the experiment.12</li>
		<li>Insert a small (.012&#34; I.D. x .005&#34;) silicon tube into the left ventricle (LV) via the pulmonary vein, the left atria (LA) and the mitral valve (MV). Fix the tube by a silk suture (4-0) to the sounding connective tissue.</li>
		<li>Position the heart with its posterior side facing up (Figure 1A, left panel).</li>
		<li>Pin the edge of the RA appendage (RAA) to the silicon bottom of the chamber using 0.1 mm diameter minutien pins. Adjust its level in order to make the posterior surface of the RA flat and allow it to be located the camera&#39;s focal plane. This will enable optical measurements from the maximal surface area of the atrium.</li>
		<li>Noose superior (SVC) and inferior (IVC) vena cava by a silk suture (4-0), stretch and pin the other end of the suture to the bottom of a silicon-coated chamber (see Figure 3A). Make sure the sutures do not block the optical field of view.</li>
		<li>Place the custom-made pacing electrode on the edge of the RAA. To make electrodes, use silicon coated 0.25 mm diameter silver wires, with 0.5 mm inter-electrode distance and devoid of silicon for a length of 1 mm at the pacing end. Then place two ECG 12 mm needle (29-gauge) monopolar electrodes near the base of the right and left ventricles. Place the ground ECG electrode near the apex of the ventricles .</li>
	</ol>
	</li>
	<li>Staining
	<ol>
		<li>Since both fluorescent dyes and electro-mechanical uncoupler blebbistatin are light-sensitive, perform all the procedures described below in a dark room. First prepare voltage-sensitive dye RH-237 or di-4-ANNEPS as a stock solution, 1.25 mg/ml in dimethyl sulfoxide (2.5 mM), aliquot it (30 &#181;l each) and store the aliquots at -20 &#176;C.</li>
		<li>Dilute 5-10 &#181;l of dye stock solution in 1 ml of warmed (37 &#176;C) Tyrode solution and then inject it into the coronary perfusion line over a period of 5-7 min using an in-line Luer injection port.</li>
		<li>Prepare blebbistatin as a stock solution (2 mg/ml in dimethyl sulfoxide, 6.8 mM) in advance and store it at 4&#176;C.</li>
		<li>After 20 min of stabilization, add 0.5 ml of warmed (37 &#176;C) blebbistatin in the perfusate and dilute 0.1 ml of blebbistatin in 1 ml of warmed (37 &#176;C) Tyrode solution. Then inject into the coronary perfusion line over a period of 5-7 min using an in-line Luer injection port.</li>
	</ol>
	</li>
</ol>
3. Optical mapping of the SAN from the Isolated Atrial Preparation
<ol>
	<li>Instrumentation
	<ol>
		<li>For the isolated atrial preparation, isolate and cannulate the heart as described above in steps 1.1-1.5 for whole-heart preparation.</li>
		<li>Dissect the ventricles away from the anterior side. See Figure 1A, right panel, cut #1 for details.</li>
		<li>Open the RA by cutting through the tricuspid valve (TV) along the TV-SVC axis. See Figure 1A, right panel, cut #2 for details.</li>
		<li>Cut the medial limb of the crista terminalis to open the RAA. See Figure 1A, right panel, cut #3.</li>
		<li>Open the anterior atrial free wall by performing a cut from the midline of the previous cut#3 to the edge of the right bottom corner of the RAA, flatten and pin the atrial free wall to the bottom of a silicon-coated chamber. See the direction shown by the hollow arrow in Figure 1A, cut #4. Preserve a rim of ventricular tissue for pinning the preparation to prevent damage to the atria.</li>
		<li>Similarly, open LA by cutting through the MV along the MV-upper corner of the LA appendage (LAA).</li>
		<li>To open the LAA, cut from the middle of the opened LAA, through the anterior atrial free wall until near a middle rim of the LAA.</li>
		<li>Open the anterior atrial free wall along the same direction, then flatten and pin it to the bottom of a Sylgard-coated chamber.</li>
		<li>Partially remove the interatrial septal tissue. This will reduce scattering of the optical signal from tissue that is not in focus. Therefore, both the LA and RA as well as the SAN and atrio-ventricular junction (AVJ) are accessible in this preparation (Figure 1B).</li>
		<li>Lift up the final preparation by about 0.5 mm from the bottom of the silicon-coated chamber in order to allow superfusion from both epicardial and endocardial surfaces.</li>
		<li>Superfuse the preparation with warmed (37 &#176;C) Tyrode solution at a constant rate of ~12 ml/min for a focused flow located near the SVC and ~30 ml/min for a bath superfusion.</li>
		<li>Place the custom-made pacing Ag/AgCl2 electrode (0.25 mm diameter) on the edge of the dissected RAA. Then place two ECG 12 mm needle (29G) electrodes (monopolar) near the RAA and LAA, respectively. Place the ground ECG electrode near the AVJ.</li>
	</ol>
	</li>
	<li>Staining
	<ol>
		<li>Perform a direct application of the voltage-sensitive dye (RH-237 or di-4-ANNEPS). For this, dilute 1-2 &#181;l of the dye stock solution in 1 mm of warmed (37 &#176;C) Tyrode solution and slowly release the diluted dye on the surface of the preparation by using a 1 mm pipette.</li>
		<li>Alternatively, perform atrial staining with the voltage-sensitive dye through a coronary perfusion of the Langendorff-perfused heart similar to that described above for the whole-heart preparation (steps 2.2.1-2.2.2).
		<ol>
			<li>After the coronary perfusion staining, isolate the atrial preparation as described. Perform additional surface staining when needed to reach a satisfactory fluorescence level. Quick atrial isolation does not bleach the dye and does not affect the quality of staining.</li>
		</ol>
		</li>
		<li>Immobilize the preparation with the electro-mechanical uncoupler, blebbistatin. Dilute 3-5 &#181;l blebbistatin stock solution in the warmed (37 &#176;C) Tyrode solution and slowly release the diluted blebbistatin on the surface of the preparation and surrounding solution. Since blebbistatin has been shown to be light sensitive,13,14 avoid long exposure to light during preparation and staining.</li>
		<li>Perform additional blebbistatin application as much as needed to suppress contraction. Normally up to 3 additional applications (3-5 &#181;l per each application) can be used to suppress the motion artifact sufficiently in order to accurately reconstruct SAN and atrial activation.</li>
		<li>Add additional 0.5 ml of warmed blebbistatin in the perfusate. During this procedure, pause the superfusion for 30 sec to allow the dye/blebbistatin to stain the atrial tissue.</li>
	</ol>
	</li>
</ol>
4. Optical Mapping Set Up
NOTE: A detailed description of the optical mapping system is provided elsewhere.12
<ol>
	<li>Use a camera with a temporal resolution of 2,000 frames/sec or higher, and a series of condenser lenses to reach spatial resolution of 100 &#181;m/pixel or higher. This is required to reconstruct SAN activation and intranodal propagation.</li>
	<li>To reduce motion artifact from the vibrating solution, fix a small cover glass on the surface of the solution over the heart/isolated atrial preparation.</li>
	<li>Use excitation light (520 &#177; 45 nm wavelength) provided by a constant-current, low-noise halogen lamp. Direct the filtered light beam onto the preparation by using a flexible bifurcated light guide.</li>
	<li>Filter the emitted fluorescence by a long-pass filter (&#62;715 nm). Collect, digitize and save the acquired fluorescent signal on a computer using software provided by the camera manufacturer.</li>
</ol>
5. Data Processing
NOTE: Optical mapping data is collected and stored as a series of matrices of fluorescent intensity at different time points. Each pixel represents a measure of fluorescent intensity collected from a specific location on the tissue preparation at a specific time point. Background fluorescence is automatically removed per pixel to allow for better visualization of fluorescence changes produced by membrane voltage changes and inverted to correspond with action potentials (APs) measured by microelectrode systems. Details of the different steps in processing optical imaging data, including image segmentation, spatial filtering, temporal filtering, and baseline drift removal, are provided in the focused review.15
<ol>
	<li>Manually or by using the special algorithm (for example, a thresholding or edge detection)15 to determine tissue boundaries, create a binary mask of 1 (included pixels) and 0 (excluded pixels) and apply the mask to each frame in the sequence. This process creates a binary image that highlights areas of interest for further processing.</li>
	<li>To improve signal-to-noise ratio of the optical data, filter the signals within the selected areas of interest. For this, use spatial (i.e. averaging of neighboring fluorescent pixels as defined by a desired convolution bin or kernel, for instance 3 x 3 or, for noisy data, 5 x 5) and/or temporal filtering (for example, Butterworth, Chebyshev type 1, Chebyshev type 2, Elliptic etc.) (Figure 2). Keep in mind the possibility of filtered-induced artifacts when interpreting the data. For more details, see review.15</li>
	<li>Remove drift of baseline in optical recordings when needed (by using high-pass filtering or polynomial fitting to the original signal) and then normalize each pixel signal from 0 (minimum fluorescence) to 1 (maximum fluorescence).</li>
	<li>Select a time window that encompasses the activation times of all the pixels for a single AP propagation. Assign each pixel its activation time as the time of maximum upstroke derivative (dF/dtmax, where F is fluorescence intensity). Using all pixel activation times, reconstruct the isochronal activation map. Each isochron will thus show the pixels activated at the same time.</li>
	<li>To reconstruct the AP duration (APD) distribution map, for each pixel calculate the duration between the activation time and the time at a specific level of repolarization (for example, at 80% of repolarization, APD80). Using all pixel APD values, reconstruct the APD distribution isochronal map. Each isochron will thus show the pixels with the same APD.</li>
	<li>To calculate conduction velocity for AP propagation, fit a surface of the preparation to the activation time data calculated in 5.4. For this, use polynomial surface fitting or local kernel surface smoothing and then calculate local conduction velocity vectors from the gradient of the fitted surface.</li>
	<li>To create the repolarization map, assign each pixel its repolarization time, defined as the maximum second derivative (d2F/dt2) of the optical signal at the end of the OAP, or the time of 90% repolarization.</li>
</ol>

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</ol>

No conflicts of interest are declared.

Here, we presented two types of mouse SAN preparations: 1) intact SAN in the Langendorff-perfused whole heart, and 2) SAN in the isolated, opened atrial preparation. These two types of preparation serve different experimental purposes. In the Langendorff-perfused whole heart preparation, the intact atrial structure is preserved which makes it possible to study complex atrial arrhythmias such as atrial fibrillation as well as interactions between the SAN and atrium during reentrant tachyarrhythmias.6 In contras...

Novel scientific breakthroughs that lead to leaps in the understanding of human physiology are often preceded by technological advances. Fluorescent optical mapping, for example, enables investigation of multiple physiological parameters in both cells and tissues.1,2 It significantly improved our understanding of how the anatomical structure is associated with electrophysiological functions and dysfunctions. In the heart, the natural pacemaker and conduction systems such as the sinoatrial node (SAN) and the atrioventricular junction consist of nodal myocytes that are insulated by the surrounding atrial myocytes.3-5 Such organization creates complex three-dimensional structures with specialized electrical properties. Both structural and functional remodeling of these pacemaker structures has been recognized to form significant electrophysiological heterogeneities.6,7 Understanding the mechanism underlying how such heterogeneities result in SAN dysfunctions and atrial arrhythmogenesis will dramatically benefit the clinical treatment of these diseases. It requires a technique to visualize the propagation of electrical signals at the tissue level, such as optical mapping. 
Recently, accumulating evidence has proven the advance of optical mapping in studies of atrial electrophysiology and pathology.2 However, novel and rigorous studies are dependent on the accurate interpretation of experimental data, whose validity and stability rely on careful experiment protocols. Genetically modified mice are extensively used for research as animal models of human diseases including sick sinus syndrome, pacemaker abnormalities and atrial arrhythmias.6,8-11 Thus, a combination of fluorescent optical mapping with transgenic mouse models provides a powerful tool to study cardiac electrical abnormalities associated with various pathologies. In this paper, we present a protocol for high-resolution optical mapping of the mouse SAN and atrium. Specifically, we discuss and compare different dye loading approaches, time effects on dye bleaching and heart rate stability during the experiment.

<table><tbody><tr><td>water jacket</td><td>Radnoti</td><td>1660 Series Tissue Bath for Large Organ or Single Cell Isolation Procedures</td><td></td></tr><tr><td>water bath / circulator</td><td>Fisher Scientific</td><td>1016S</td><td></td></tr><tr><td>pressure amplifier</td><td>AD Instruments</td><td>MLT0670</td><td></td></tr><tr><td>EMD Millipore Nylon Net Filters</td><td>Fisher Scientific</td><td>NY1102500</td><td></td></tr><tr><td>Pressure transducer</td><td>AD Instruments</td><td>MLT0670</td><td></td></tr><tr><td>Stainless Steel Minutien Pins - 0.1 mm Diameter</td><td>Fine Science Tools</td><td>26002-10&amp;nbsp;</td><td></td></tr><tr><td>Perfusion pump</td><td>World Precision Instruments</td><td>PERIPRO-4LS</td><td></td></tr><tr><td>Superfusion pump</td><td>World Precision Instruments</td><td>PERIPRO-4HS</td><td></td></tr><tr><td>Vannas Tubingen scissors&amp;nbsp;</td><td>World Precision Instruments</td><td>503379</td><td></td></tr><tr><td>Dumont forceps</td><td>World Precision Instruments</td><td>501201, 500085</td><td></td></tr><tr><td>Mayo scissors</td><td>World Precision Instruments</td><td>501750</td><td></td></tr><tr><td>Kelly hemostatic forceps</td><td>World Precision Instruments</td><td>501241</td><td></td></tr><tr><td>Iris forceps</td><td>World Precision Instruments</td><td>15917</td><td></td></tr><tr><td>Iris scissors</td><td>World Precision Instruments</td><td>501263</td><td></td></tr><tr><td>ECG 12 mm needle (29-gauge) electrodes (monopolar)&amp;nbsp;</td><td>AD Instruments</td><td>MLA1203</td><td></td></tr><tr><td>in-line Luer injection port</td><td>Ibidi</td><td>10820</td><td></td></tr><tr><td>Ultima-L CMOS camera&amp;nbsp;</td><td>SciMedia</td><td>MiCAM-05&amp;nbsp;</td><td></td></tr><tr><td>halogen lamp</td><td>Moritex USA Inc</td><td>MHAB-150W</td><td></td></tr><tr><td>NaCl</td><td>Fisher Scientific</td><td>S271-1</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt; (2H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Fisher Scientific</td><td>C79-500</td><td></td></tr><tr><td>KCl</td><td>Fisher Scientific</td><td>S217-500</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt; (6H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Fisher Scientific</td><td>M33-500</td><td></td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt; (H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Fisher Scientific</td><td>S369-500</td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Fisher Scientific</td><td>S233-3</td><td></td></tr><tr><td>D-Glucose</td><td>Fisher Scientific</td><td>D16-1</td><td></td></tr><tr><td>Blebbistatin</td><td>Tocris Bioscience</td><td>1760</td><td></td></tr><tr><td>RH237</td><td>ThermoFisher Scientific</td><td>S1109</td><td></td></tr><tr><td>Dimethyl sulphoxide (DMSO)</td><td>Sigma-Aldrich</td><td>D2650</td><td></td></tr></tbody></table>

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.

Optical Mapping of the Intact SAN from the Langendorff-perfused Heart 
A typical example of an RA activation contour map reconstructed for spontaneous sinus rhythm is shown in Figure 3 for a Langendorff-perfused mouse heart. The early activation point is located within the intercaval region near the SVC where the SAN is anatomically defined.3,16 Two RA activation contour maps acquired at 1.0 and 0.5 msec sampling rate are shown in Fi...

Watch this Scientific Journal Video about High-resolution Optical Mapping of the Mouse Sino-atrial Node at JoVE.com

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

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.

Sino-atrial node (SAN) dysfunctions and associated complications constitute important causes of morbidity in patients with cardiac diseases. The ...

high-resolution-optical-mapping-of-the-mouse-sino-atrial-node

Research

JoVE Journal

Biology

15.6K Views.  University of Wisconsin-Madison School of Medicine and Public Health.  Hier presenteren we een protocol voor optische kaart brengen van de elektrische activiteit van de muis rechteratrium en vooral de sinusknoop, bij een hoge ruimtelijke en temporele resolutie. 

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

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. ...

High-Resolution Endocardial and Epicardial Optical Mapping in a Sheep Model of Stretch-Induced Atrial Fibrillation

Atrial fibrillation (AF) is a complex cardiac arrhythmia with high morbidity and mortality.1,2 It is the most common sustained cardiac rhythm disturbance seen in clinical practice and its prevalence is expected to increase in the coming years.3 Increased intra-atrial pressure and dilatation have been long recognized to lead to AF,1,4 which highlights the relevance of using animal models and stretch to study AF dynamics. Understanding the mechanisms underlying AF requires visualization of the cardiac electrical waves with high spatial and temporal resolution. While high-temporal resolution can be achieved by conventional electrical mapping traditionally used in human electrophysiological studies, the small number of intra-atrial electrodes that can be used simultaneously limits the spatial resolution and precludes any detailed tracking of the electrical waves during the arrhythmia. The introduction of optical mapping in the early 90's enabled wide-field characterization of fibrillatory activity together with sub-millimeter spatial resolution in animal models5,6 and led to the identification of rapidly spinning electrical wave patterns (rotors) as the sources of the fibrillatory activity that may occur in the ventricles or the atria.7-9 Using combined time- and frequency-domain analyses of optical mapping it is possible to demonstrate discrete sites of high frequency periodic activity during AF, along with frequency gradients between left and right atrium. The region with fastest rotors activates at the highest frequency and drives the overall arrhythmia.10,11 The waves emanating from such rotor interact with either functional or anatomic obstacles in their path, resulting in the phenomenon of fibrillatory conduction.12 Mapping the endocardial surface of the posterior left atrium (PLA) allows the tracking of AF wave dynamics in the region with the highest rotor frequency. Importantly, the PLA is the region where intracavitary catheter-based ablative procedures are most successful terminating AF in patients,13 which underscores the relevance of studying AF dynamics from the interior of the left atrium. Here we describe a sheep model of acute stretch-induced AF, which resembles some of the characteristics of human paroxysmal AF. Epicardial mapping on the left atrium is complemented with endocardial mapping of the PLA using a dual-channel rigid borescope c-mounted to a CCD camera, which represents the most direct approach to visualize the patterns of activation in the most relevant region for AF maintenance.

This report provides a detailed description of the methodology and results of simultaneous endocardial and epicardial optical mapping of electrical excitation in the intact left atrium of a Langendorff-perfused sheep heart during stretch-induced atrial fibrillation.

 Atrial fibrillation (AF) is a complex cardiac arrhythmia with high morbidity and mortality.1,2 It is the most common sustained cardiac rhythm disturbance ...

Medicine

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 ...

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 ...

Light-Sheet Microscopy and Tissue Clearing for Comprehensive Cardiac Imaging

Light-Sheet Imaging to Reveal Cardiac Structure in Rodent Hearts

Light-sheet microscopy (LSM) plays a pivotal role in comprehending the intricate three-dimensional (3D) structure of the heart, providing crucial insights into fundamental cardiac physiology and pathologic responses. We hereby delve into the development and implementation of the LSM technique to elucidate the micro-architecture of the heart in mouse models. The methodology integrates a customized LSM system with tissue clearing techniques, mitigating light scattering within cardiac tissues for volumetric imaging. The combination of conventional LSM with image stitching and multiview deconvolution approaches allows for the capture of the entire heart. To address the inherent trade-off between axial resolution and field of view (FOV), we further introduce an axially swept light-sheet microscopy (ASLM) method to minimize out-of-focus light and uniformly illuminate the heart across the propagation direction. In the meanwhile, tissue clearing methods such as iDISCO enhance light penetration, facilitating the visualization of deep structures and ensuring a comprehensive examination of the myocardium throughout the entire heart. The combination of the proposed LSM and tissue clearing methods presents a promising platform for researchers in resolving cardiac structures in rodent hearts, holding great potential for the understanding of cardiac morphogenesis and remodeling.

Author Spotlight: Customized Light-Sheet Imaging for Investigating Myocardial Structures in Rodent Hearts

The protocol utilizes advanced light-sheet microscopy along with adapted tissue clearing methods to investigate intricate cardiac structures in rodent hearts, holding great potential for the understanding of cardiac morphogenesis and remodeling.

Light-sheet microscopy (LSM) plays a pivotal role in comprehending the intricate three-dimensional (3D) structure of the heart, providing crucial insights ...

Whole-Mount Immunofluorescence Staining, Confocal Imaging and 3D Reconstruction of the Sinoatrial and Atrioventricular Node in the Mouse

The electrical signal physiologically generated by pacemaker cells in the sinoatrial node (SAN) is conducted through the conduction system, which includes the atrioventricular node (AVN), to allow excitation and contraction of the whole heart. Any dysfunction of either SAN or AVN results in arrhythmias, indicating their fundamental role in electrophysiology and arrhythmogenesis. Mouse models are widely used in arrhythmia research, but the specific investigation of SAN and AVN remains challenging.
The SAN is located at the junction of the crista terminalis with the superior vena cava and AVN is located at the apex of the triangle of Koch, formed by the orifice of the coronary sinus, the tricuspid annulus, and the tendon of Todaro. However, due to the small size, visualization by conventional histology remains challenging and it does not allow the study of SAN and AVN within their 3D environment.
Here we describe a whole-mount immunofluorescence approach that allows the local visualization of labelled mouse SAN and AVN. Whole-mount immunofluorescence staining is intended for smaller sections of tissue without the need for manual sectioning. To this purpose, the mouse heart is dissected, with unwanted tissue removed, followed by fixation, permeabilization and blocking. Cells of the conduction system within SAN and AVN are then stained with an anti-HCN4 antibody. Confocal laser scanning microscopy and image processing allow differentiation between nodal cells and working cardiomyocytes, and to clearly localize SAN and AVN. Furthermore, additional antibodies can be combined to label other cell types as well, such as nerve fibers.
Compared to conventional immunohistology, whole-mount immunofluorescence staining preserves the anatomical integrity of the cardiac conduction system, thus allowing the investigation of AVN; especially so into their anatomy and interactions with the surrounding working myocardium and non-myocyte cells.

We provide a step-by-step protocol for whole-mount immunofluorescence staining of the sinoatrial node (SAN) and atrioventricular node (AVN) in murine hearts.

The electrical signal physiologically generated by pacemaker cells in the sinoatrial node (SAN) is conducted through the conduction system, which includes ...

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

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

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

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

Mesoscopic Optical Imaging of Whole Mouse Heart

Both genetic and non-genetic cardiac diseases can cause severe remodeling processes in the heart. Structural remodeling, such as collagen deposition (fibrosis) and cellular misalignment, can affect electrical conduction, introduce electromechanical dysfunctions and, eventually, lead to arrhythmia. Current predictive models of these functional alterations are based on non-integrated and low-resolution structural information. Placing this framework on a different order of magnitude is challenging due to the inefficacy of standard imaging methods in performing high-resolution imaging in massive tissue. In this work, we describe a methodological framework that allows imaging of whole mouse hearts with micrometric resolution. The achievement of this goal has required a technological effort where advances in tissue transformation and imaging methods have been combined. First, we describe an optimized CLARITY protocol capable of transforming an intact heart into a nanoporous, hydrogel-hybridized, lipid-free form that allows high transparency and deep staining. Then, a fluorescence light-sheet microscope able to rapidly acquire images of a mesoscopic field of view (mm-scale) with the micron-scale resolution is described. Following the mesoSPIM project, the conceived microscope allows the reconstruction of the whole mouse heart with micrometric resolution in a single tomographic scan. We believe that this methodological framework will allow clarifying the involvement of the cytoarchitecture disarray in the electrical dysfunctions and pave the way for a comprehensive model that considers both the functional and structural data, thus enabling a unified investigation of the structural causes that lead to electrical and mechanical alterations after the tissue remodeling.

We report a method for mesoscopic reconstruction of the whole mouse heart by combining new advancements in tissue transformation and staining with the development of an axially scanned light-sheet microscope.

Both genetic and non-genetic cardiac diseases can cause severe remodeling processes in the heart. Structural remodeling, such as collagen deposition ...

Dual-Dye Optical Mapping of Hearts from RyR2R2474S Knock-In Mice of Catecholaminergic Polymorphic Ventricular Tachycardia

The pro-arrhythmic cardiac disorder catecholaminergic polymorphic ventricular tachycardia (CPVT) manifests as polymorphic ventricular tachycardia episodes following physical activity, stress, or catecholamine challenge, which can deteriorate into potentially fatal ventricular fibrillation. The mouse heart is a widespread species for modeling inherited cardiac arrhythmic diseases, including CPVT. Simultaneous optical mapping of transmembrane potential (Vm) and calcium transients (CaT) from Langendorff-perfused mouse hearts has the potential to elucidate mechanisms underlying arrhythmogenesis. Compared with the cellular level investigation, the optical mapping technique can test some electrophysiological parameters, such as the determination of activation, conduction velocity, action potential duration, and CaT duration. This paper presents the instrumentation setup and experimental procedure for high-throughput optical mapping of CaT and Vm in murine wild-type and heterozygous RyR2-R2474S/+ hearts, combined with programmed electrical pacing before and during the isoproterenol challenge. This approach has demonstrated a feasible and reliable method for mechanistically studying CPVT disease in an ex vivo mouse heart preparation.

Dual-Dye Optical Mapping of Hearts from RyR2R2474S Knock-In Mice of Catecholaminergic Polymorphic Ventricular Tachycardia

This protocol introduces dual-dye optical mapping of mouse hearts obtained from wild-type and knock-in animals affected by catecholaminergic polymorphic ventricular tachycardia, including electrophysiological measurements of transmembrane voltage and intracellular Ca2+ transients with high temporal and spatial resolution.

The pro-arrhythmic cardiac disorder catecholaminergic polymorphic ventricular tachycardia (CPVT) manifests as polymorphic ventricular tachycardia episodes ...

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

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