This work was supported by the China Scholarship Council (CSC, to R. Xia), the German Centre for Cardiovascular Research (DZHK; 81X2600255 to S. Clauss, 81Z0600206 to S. K&#228;&#228;b), the Corona Foundation (S199/10079/2019 to S. Clauss), the SFB 914 (project Z01 to H. Ishikawa-Ankerhold and S. Massberg and project A10 to C. Schulz), the ERA-NET on Cardiovascular Diseases (ERA-CVD; 01KL1910 to S. Clauss) and the Heinrich-and-Lotte-M&#252;hlfenzl Stiftung (to S. Clauss). The funders had no role in manuscript preparation.

Animal care and all experimental procedures were conducted in accordance with the guidelines of the Animal Care and Ethics committee of the University of Munich, and all the procedures undertaken on mice were approved by the Government of Bavaria, Munich, Germany (ROB-55.2-2532.Vet_02-16-106, ROB-55.2-2532.Vet_02-19-86). C57BL6/J mice were purchased from Jackson Laboratory.
NOTE: Figure 1 shows the instruments needed for the experiment. Figure 2 shows an illustration of the gross cardiac anatomy. Figure 3 shows the location of SAN and AVN in an adult mouse heart. Figure 4 shows the prepared sample loaded on the confocal microscopy.
1. Preparations
<ol>
	<li>Prepare a 4% formaldehyde solution (4% PFA) by diluting 10 mL of 16% formaldehyde solution in 30 mL of PBS.</li>
	<li>Prepare a 15% sucrose solution and 30% sucrose solution by dissolving 15 g and 30 g of sucrose powder into 100 mL PBS, respectively. After the sucrose powder is fully dissolved, filter the solution using 0.2 &#181;m syringe filter before storing at 4 &#176;C.</li>
	<li>Prepare a 3%-4% agarose gel, add 3 g or 4 g agarose powder and 100 mL of 1x TAE (diluted from 50x TAE stock solution) into a beaker. Place the beaker on a magnetic stirrer and boil it until the agarose is completely dissolved.</li>
	<li>Prepare the dissection dish by gently pouring 30 mL of agarose gel (3%-4%) in a 100 mm diameter Petri dish. Leave the Petri dish on the bench at room temperature to cool down and harden.</li>
	<li>Prepare blocking solution and washing solution according to the recipes provided in the Table 1. Prepare all of the solutions prepared at the day of use. Long term storage is not recommended.</li>
	<li>Prepare the antibody solutions by diluting them in cold (4 &#176;C) 1x PBS, protect the dilution from light (e.g., by wrapping aluminum foil around) and keep dilutions on ice until used. Dilute the antibodies shortly before incubation. Avoid leaving the diluted antibodies at room temperature. 
	NOTE: The appropriate antibody dilution should be tested with comparable tissue samples by performing a conventional immunofluorescent staining. Here, we tested the antibodies by staining frozen sections cut at a thickness of 10 &#181;m.</li>
</ol>
2. Organ harvest and tissue preparation
<ol>
	<li>Anesthetize the mouse by placing it into an incubation chamber connected to&#160;an isoflurane vaporizer.&#160;Set the vaporizer to deliver 4-5% isoflurane (96-95% oxygen, respectively). 
	NOTE: Full anesthesia is confirmed by the loss of the postural reaction and righting reflex by gently rolling the chamber until the mouse is placed on its back.</li>
	<li>Put the mouse in a supine position on the surgical table. Place the mouse nose into an anesthesia mask connected to a modified Bain circuit with its inner tube connected to the isoflurane vaporizer. To maintain anesthesia, use 1-2% isoflurane in oxygen at a flow rate of 1 L/min. Scavenge away excess anesthetic vapor from the mouse via the outer tube of the mask and draw through a canister of activated charcoal, which absorbs the excess anesthetic gas.</li>
	<li>When full anesthesia is achieved, inject fentanyl for analgesia (0.1&#160;&#181;g/25 g body weight i.p.).</li>
	<li>When the toe-pinch reflex is undetectable, make a clear cut from the jugulum to the symphysis using iris scissors to remove fur and skin. Make another cut from left to right underneath the ribs using iris scissors to carefully open the abdomen.</li>
	<li>Life the xiphoid a little bit using curved forceps to allow cutting the diaphragm from left to right without injuring any organs. Cut the rib cage in a medial axillary line on both sides using iris scissors to flip it cranially and to allow access to the heart.</li>
	<li>Cut the inferior vena cava and descending thoracic aorta at the level of the diaphragm using iris scissors. Puncture the heart with a 27 G needlein the area of the apex and then carefully push the needle into the left ventricle (LV). Gently inject 5-10 mL of ice-cold PBS into the LV to perfuse the heart. 
	NOTE: The color of the heart should turn from red to gray indicating successful perfusion with PBS.</li>
	<li>Carefully lift the apex of the heart using a tweezer allowing to cut the large vessels and to remove the heart.</li>
	<li>Excise the heart by cutting the large arteries and veins as far away from the heart as possible to avoid any damage to the superior vena cava (SVC). Conserve the SVC since it will serve as important landmark during later processing.</li>
	<li>After heart removal, turn off the isoflurane vaporizer.</li>
	<li>Put the heart into a dissection dish filled with ice cold PBS under the dissecting microscope. After determination of the left/right and front/back of the heart, turn the heart around with the front of the heart at the bottom of the dish (to expose the large vessels that are located posterior).</li>
	<li>Immobilize the heart by putting little pinsthrough the apex and the left atrial appendage (LAA) into the agarose at the bottom of the dissection dish (Figure 2A). Using fine tweezers and scissors, carefully remove non-cardiac tissue around the SVC and inferior vena cava (IVC) (e.g., lungs, fat, pericardium) to expose the inter-caval region (Figure 3A).</li>
	<li>Remove the majority of the ventricles by cutting parallel the groove between the ventricles and the atria with the micro scissor. Preserve a small part of the ventricular tissue for the later loading on the Plexiglas&#160;ring.</li>
	<li>For fixation and dehydration, put the sample (containing the atria, SVC and IVC) in 4% PFA overnight at 4 &#176;C.</li>
	<li>The next day, transfer the heart to 15% sucrose solution for 24 hours at 4 &#176;C.</li>
	<li>The next day, transfer the heart to 30% sucrose solution for 24 hours at 4 &#176;C. 
	NOTE: For the fixation and dehydration in step 2.13, 2.14 and 2.15, the samples are left at 4&#176;C without further stirring or rocking.</li>
</ol>
3. Whole-mount immunofluorescence staining
<ol>
	<li>Wash the heart in 1% Triton X-100 diluted in PBS and block and permeabilize in blocking solution (Table 1) overnight at 4 &#176;C.</li>
	<li>Place the heart in a 1.5 mL tube and incubate with rabbit anti-mouse connexin-43 (dilution of 1:200) and rat anti-mouse HCN4 (dilution of 1:200) antibodies diluted with blocking solution for 7 days at 4 &#176;C. 
	NOTE: The optimal concentration of primary antibodies should be tested before using the datasheets as orientation. Other antigens of interest could also be stained in this step, as long as the host species of the primary antigen is different from the other ones. A higher concentration of Triton X-100 may help obtain more efficient antibody staining as demonstrated before10, but concentration might be determined individually.</li>
	<li>After 7 days, remove the solution containing primary antibodies using a pipetteand wash the heart with 1%Triton X-100 solution 3 times (each time for 1 h at room temperature on the orbital shaker).</li>
	<li>After washing, incubate the heart = with Alexa Fluor 488 goat anti-rat IgG (dilution of 1:200) and Alexa Fluor 647 goat anti-rabbit IgG (dilution of 1:200) for 7 days at 4 &#176;C.</li>
	<li>After 7 days, remove all the solution containing the secondary antibodies using a pipette. Then wash the heart using washing solution (Table 1) 3 times (each time for 1 h at room temperature on the orbital shaker).</li>
	<li>To stain nuclei, incubate the heart in DAPI solution (10 &#181;g/mL) overnight at 4 &#176;C.</li>
	<li>On the next day, wash the heart with washing solution 3 times (each time for 1 h at room temperature on the orbital shaker). 
	NOTE: For the blocking, permeabilization, antibodies incubation and DAPI staining in step 3.1, 3.2, 3.4 and 3.6, leave the samples at steady state in the 4 &#176;C, no stirring is required. The stained tissue could be preserved fully covered with washing solution at 4 &#176;C and protected from light for a few days until imaging at the confocal microscope.</li>
</ol>
4. Confocal microscopy
<ol>
	<li>Prepare Plexiglas&#160;rings and fill with plasticine (Figure 1). In the center of the plasticine a little groove is formed in the center for loading the heart. Make a shallow groove, as this would be easier to acquire a flat imaging plane, and to adjust the space and avoid air bubbles between the sample and the coverslips in step 4.4.</li>
	<li>Use the same heart sample for the imaging of both SAN and AVN sequentially. For SAN imaging, directly load the heart whereas for AVN imaging, perform microdissection before.
	<ol>
		<li>SAN imaging
		<ol>
			<li>Identify the SAN by anatomical landmarks: it is located on the dorsal side of the heart within the inter-caval region (the region between the superior and inferior vena cava). The crista terminalis (CT) is the muscle streak between the SAN and RAA.</li>
			<li>Place the heart into the plasticine groove with the back of the heart facing up. Add PBS onto the heart to displace all air within the cavity until the heart is fully covered with PBS (usually a few drops of PBS are sufficient).</li>
			<li>Under the dissection microscope, gently press the RAA, LAA and remaining parts of the left ventricle into the plasticine to fix the heart. Make sure that the whole inter-caval region and the crista terminalis could be clearly seen (not covered by plasticine). The area that will be imaged is shown in Figure 3A.</li>
		</ol>
		</li>
		<li>AVN imaging
		<ol>
			<li>After finish the imaging of SAN, recollect the same sample and subsequently use for the AVN imaging. For AVN microdissection, place the heart = on a dissection dish under the dissection microscope.</li>
			<li>Orient the heart with the right side facing up (including the remaining parts of the right ventricle). Put pins through the remaining part of the LV free wall (which is now at the bottom) to immobilize the tissue (Figure 2B).</li>
			<li>Cut the remaining RV free wall upwards through the tricuspid valve and superior vena cava. Then flip the RV and RA away to expose interventricular and interatrial septum. (Figure 3B) 
			NOTE: Pay attention not to damage the coronary sinus (CS), as it is an important anatomical landmark to find the triangle of Koch which then allows to localize the AVN. The CS runs transversely in the left atrioventricular groove on the posterior side of the heart, and the CS orifice opening is located between IVC and the tricuspid valve at the inferior part of the interatrial septum (Figure 3A and B).</li>
			<li>Identify the triangle of Koch that can be found on the endocardial surface of the right atrium, bordered anteriorly by the hinge-line of the septal leaflet of the tricuspid valve (TV), and posteriorly by the tendon of Todaro. The base is formed by the orifice of the coronary sinus (Figure 3B). Since this is the target region for AVN imaging, it needs to be clearly visible.</li>
			<li>Transfer the heart into the plasticine groove within the Plexiglas&#160;ring with the triangle of Koch clearly being exposed (i.e. not covered by plasticine). Gently press the tissue around the Koch triangle into the plasticine to fix the sample. Add PBS onto the heart to displace all air within the cavity until the heart is fully covered with PBS (usually a few drops of PBS are sufficient).</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Apply silicone to the edges of the Plexiglas&#160;rings to allow covering the hearts loaded within the plasticine-filled Plexiglas&#160;rings with cover slips.</li>
	<li>Gently press the back side of the plasticine to squeeze out parts of the PBS and to attach the heart to the coverslip while avoiding any air bubbles within the imaging area. 
	NOTE: Make sure that the regions of interest are not folded and covered during pressing the back side of the plasticine. A flat imaging plane without sample overcompression is necessary for conserving the anatomy and for proper confocal imaging of the samples. For SAN, it is important to make sure the inter-caval region is clearly exposed. For AVN, the triangle of Koch should be fully exposed.</li>
	<li>Put the whole-mount staining samples up-side down on the platform of the confocal microscope. The exposed SAN/AVN attached to the cover slip is now on the bottom of the microscope platform (Figure 4). 
	NOTE: To take images, we use the Carl Zeiss LSM800 with Airyscan Unit and the software ZEN 2.3 SP1 black.</li>
	<li>Choose plate &#34;BP420-480 + LP605&#34; for the excitation of Alexa Fluor 647 conjugated anti-HCN4. Vary the master gain from 650-750.</li>
	<li>Take an overview image of the whole SAN and AVN region by using Tile Scan function. Then select the HCN4-positive region by clicking and adding a square on the overview image around the area of interest that will be scanned.</li>
	<li>Check the parameters, including plates and master gain for the remaining channels (Alexa Fluor 488 and DAPI) of the confocal microscope and set as described in step 4.6.</li>
	<li>Slowly adjust the focus from top to bottom of the sample to preview the whole sample and to set the First and Last for the Z-stack range. Set an optimal interval for the Z-stack based on the thickness of the optical sample. We use an 20x objective, and 0.8-1 &#181;m as the interval for Z-stack.</li>
	<li>After all the parameters are properly set, scan the whole area of the SAN and AVN.</li>
	<li>Perform 3D reconstruction of the images using software (e.g., Imaris version 8.4.2).
	<ol>
		<li>Select Images Processing | Baseline Subtraction to remove background staining.</li>
		<li>Select surface creation onto setting for selected channel and region of interest for processing.</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no conflicts of interest.

Cardiac anatomy has traditionally been studied using thin histological sections11. However, these methods do not preserve the three-dimensional structure of the conduction system and thus, only provides 2D information. The whole-mount immunofluorescence staining protocol described here allows to overcome these limitations and can be routinely used for SAN and AVN imaging.
In comparison to standard methods such as conventional immunohistochemistry that require paraffin-e...

An erratum was issued for:&#160;<a href="https://www.jove.com/t/62058">Whole-Mount Immunofluorescence Staining, Confocal Imaging and 3D Reconstruction of the Sinoatrial and Atrioventricular Node in the Mouse</a>. The Authors section was updated from:
Ruibing Xia1,2,3 
Julia Vlcek1,2 
Julia Bauer1,2,3 
Stefan K&#228;&#228;b1,3 
Hellen Ishikawa-Ankerhold1,2 
Dominic Adam van den Heuvel1,2 
Christian Schulz1,2,3 
Steffen Massberg1,2,3 
Sebastian Clauss1,2,3 
1University Hospital Munich, Department of Medicine I, Ludwig Maximilian University Munich 
2Walter Brendel Center of Experimental Medicine, Ludwig Maximilian University Munich 
3German Center for Cardiovascular Research (DZHK), Partner Site Munich, Munich Heart Alliance
to:
Ruibing Xia1,2,3 
Julia Vlcek1,2 
Julia Bauer1,2,3 
Stefan K&#228;&#228;b1,3 
Hellen Ishikawa-Ankerhold1,2 
Dominic Adam van den Heuvel1,2 
Christian Schulz1,2,3 
Steffen Massberg1,2,3 
Sebastian Clauss1,2,3 
1University Hospital Munich, Department of Medicine I, Ludwig Maximilians University (LMU) Munich 
2Institute of Surgical Research at the Walter Brendel Center of Experimental Medicine, University Hospital Munich, Ludwig Maximilians University (LMU) Munich 
3German Center for Cardiovascular Research (DZHK), Partner Site Munich, Munich Heart Alliance

An erratum was issued for:&#160;<a href="https://www.jove.com/t/62058">Whole-Mount Immunofluorescence Staining, Confocal Imaging and 3D Reconstruction of the Sinoatrial and Atrioventricular Node in the Mouse</a>. The Authors section was updated.

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

Arrhythmias are common diseases affecting millions of people, and are the cause of significant morbidity and mortality worldwide. Despite enormous advances in treatment and prevention, such as the development of cardiac pacemakers, treatment of arrhythmias remains challenging, primarily due to the very limited knowledge regarding underlying disease mechanisms1,2,3. A better understanding of both the normal electrophysiology and the pathophysiology of arrhythmias may help to develop novel, innovative and causal treatment strategies in the future. Additionally, to comprehensively study arrhythmogenesis, it is important to localize and visualize the specific cardiac conduction system in animal models such as the mouse, as mice are widely used in electrophysiology research.
The major parts of the cardiac conduction system are the sinoatrial node (SAN), where the electrical impulse is generated in specialized pacemaker cells, and the atrioventricular node (AVN), which is the only electrical connection between the atria and the ventricles4. Whenever the electrophysiological properties of SAN and AVN are altered, arrhythmias such as sick sinus syndrome or atrioventricular block can occur which may lead to hemodynamic deterioration, syncope and even death, and thus underline the essential role of both SAN and AVN in electrophysiology and arrhythmogenesis5.
Comprehensive studies on SAN or AVN require a precise localization and visualization of both structures, ideally within their physiologic environment. However, due to their small size and location within the working myocardium, without establishing a clear macroscopically visible structure, studying the anatomy and electrophysiology of SAN and AVN is challenging. Anatomical landmarks can be used to roughly identify the region that contains SAN and AVN6,7,8. In brief, SAN is located in the inter-caval region of the right atrium adjacent to the muscular crista terminalis (CT), AVN is located within the triangle of Koch established by the tricuspid valve, the ostium of the coronary sinus and the tendon of Todaro. Thus far, these anatomical landmarks were mainly used to localize, remove and then study SAN and AVN as individual structures (e.g., by conventional histology). To better understand the complex electrophysiology of SAN and AVN (e.g., regulatory effects of adjacent cells of the working myocardium), however, studying the conduction systems within the physiologic 3D environment is necessary.
Whole-mount immunofluorescence staining is a method that is used to study anatomical structures in situ while preserving the integrity of the surrounding tissue9. Taking advantage of confocal microscopy and image analysis software, SAN and AVN can be visualized with fluorescently labeled antibodies targeting ion channels specifically expressed in these regions.
This following protocol explains the necessary steps to perform a well-established whole-mount staining method for SAN and AVN microscope localization and visualization. Specifically, this protocol describes how (1) to localize SAN and AVN by anatomical landmarks to prepare these samples for staining and microscopy analysis (2) to perform whole-mount immunofluorescence staining of the reference markers HCN4 and Cx43 (3) to prepare SAN and AVN samples for confocal microscopy (4) to perform confocal imaging of SAN and AVN. We also describe how this protocol can be modified to include additional staining of surrounding working myocardium or non-myocyte cells such as autonomous nerve fibers which allows a thorough investigation of the cardiac conduction system within the heart.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="4">Anesthesia</td>
		</tr>
		<tr>
			<td>Isoflurane vaporizer system&#160;</td>
			<td>Hugo Sachs Elektronik</td>
			<td>34-0458, 34-1030, 73-4911, 34-0415, 73-4910</td>
			<td>Includes an induction chamber, a gas evacuation unit and charcoal filters</td>
		</tr>
		<tr>
			<td>Modified Bain circuit</td>
			<td>Hugo Sachs Elektronik</td>
			<td>73-4860</td>
			<td>Includes an anesthesia mask for mice</td>
		</tr>
		<tr>
			<td>Surgical Platform</td>
			<td>Kent Scientific</td>
			<td>SURGI-M</td>
			<td></td>
		</tr>
		<tr>
			<td colspan="4">In vivo instrumentation</td>
		</tr>
		<tr>
			<td>Fine forceps</td>
			<td>Fine Science Tools</td>
			<td>11295-51</td>
			<td></td>
		</tr>
		<tr>
			<td>Iris scissors</td>
			<td>Fine Science Tools</td>
			<td>14084-08</td>
			<td></td>
		</tr>
		<tr>
			<td>Spring scissors</td>
			<td>Fine Science Tools</td>
			<td>91500-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue forceps</td>
			<td>Fine Science Tools</td>
			<td>11051-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue pins</td>
			<td>Fine Science Tools</td>
			<td>26007-01</td>
			<td>Could use 27G needles as a substitute</td>
		</tr>
		<tr>
			<td colspan="4">General lab instruments</td>
		</tr>
		<tr>
			<td>Orbital shaker</td>
			<td>Sunlab</td>
			<td>D-8040</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnetic stirrer</td>
			<td>IKA&#160;</td>
			<td>RH basic</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette,volume 10 &#181;L, 100 &#181;L, 1000 &#181;L</td>
			<td>Eppendorf</td>
			<td>Z683884-1EA</td>
			<td></td>
		</tr>
		<tr>
			<td colspan="4">Microscopes</td>
		</tr>
		<tr>
			<td>Dissection stereo- zoom microscope&#160;</td>
			<td>VWR</td>
			<td>10836-004</td>
			<td></td>
		</tr>
		<tr>
			<td>Laser Scanning Confocal microscope</td>
			<td>Zeiss</td>
			<td>LSM 800</td>
			<td></td>
		</tr>
		<tr>
			<td colspan="4">Software</td>
		</tr>
		<tr>
			<td>Imaris 8.4.2</td>
			<td>Oxford instruments</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ZEN 2.3 SP1 black</td>
			<td>Zeiss</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td colspan="4">General Lab Material</td>
		</tr>
		<tr>
			<td>0.2 &#181;m syringe filter</td>
			<td>Sartorius</td>
			<td>17597</td>
			<td></td>
		</tr>
		<tr>
			<td>100 mm petri dish</td>
			<td>Falcon</td>
			<td>351029</td>
			<td></td>
		</tr>
		<tr>
			<td>27G needle</td>
			<td>BD Microlance 3</td>
			<td>300635</td>
			<td></td>
		</tr>
		<tr>
			<td>50 ml Polypropylene conical Tube</td>
			<td>Falcon</td>
			<td>352070</td>
			<td></td>
		</tr>
		<tr>
			<td>5ml Syringe</td>
			<td>Braun</td>
			<td>4606108V</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover slips</td>
			<td>Thermo Scientific</td>
			<td>7632160</td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf Tubes</td>
			<td>Eppendorf</td>
			<td>30121872</td>
			<td></td>
		</tr>
		<tr>
			<td colspan="4">Chemicals</td>
		</tr>
		<tr>
			<td>0.5 M EDTA</td>
			<td>Sigma</td>
			<td>20-158</td>
			<td>Components of TEA</td>
		</tr>
		<tr>
			<td>16% Formaldehyde Solution</td>
			<td>Thermo Scientific&#160;</td>
			<td>28908</td>
			<td>use as a 4% solution&#160;</td>
		</tr>
		<tr>
			<td>Acetic acid</td>
			<td>Merck</td>
			<td>100063</td>
			<td>Components of TEA</td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>Biozym</td>
			<td>850070</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>Sigma</td>
			<td>A2153-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>DPBS (1X) Dulbecco&#39;s Phosphate Buffered Saline</td>
			<td>Gibco</td>
			<td>14190-094</td>
			<td></td>
		</tr>
		<tr>
			<td>Normal goat serum</td>
			<td>Sigma</td>
			<td>NS02L</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma</td>
			<td>S1888-1kg</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris-base</td>
			<td>Roche</td>
			<td>TRIS-RO</td>
			<td>Components of TEA</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma</td>
			<td>T8787-250ml</td>
			<td>Diluted to 1% in PBS</td>
		</tr>
		<tr>
			<td>Tween 20</td>
			<td>Sigma</td>
			<td>P2287-500ml</td>
			<td></td>
		</tr>
		<tr>
			<td colspan="4">Drugs</td>
		</tr>
		<tr>
			<td>Fentanyl 0.5&#160;mg/10&#160;mL</td>
			<td>Braun Melsungen</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane 1 mL/mL</td>
			<td>Cp-pharma</td>
			<td>31303</td>
			<td></td>
		</tr>
		<tr>
			<td>Oxygen 5 L</td>
			<td>Linde</td>
			<td>2020175</td>
			<td>Includes a pressure regulator</td>
		</tr>
		<tr>
			<td colspan="4">Antibodies</td>
		</tr>
		<tr>
			<td>Goat anti-Rabbit IgG Alexa Fluor&#160;488&#160;</td>
			<td>Cell Signaling Technology</td>
			<td>#4412</td>
			<td>diluted to 1:200</td>
		</tr>
		<tr>
			<td>Goat anti-Rat IgG Alexa Fluor 647</td>
			<td>Invitrogen</td>
			<td>#A-21247</td>
			<td>diluted to 1:200</td>
		</tr>
		<tr>
			<td>Hoechst 33342, Trihydrochloride, Trihydrate (DAPI)</td>
			<td>Invitrogen</td>
			<td>H3570</td>
			<td>diluted to 1:1000</td>
		</tr>
		<tr>
			<td>Rabbit Anti-Connexin-43</td>
			<td>Sigma</td>
			<td>C6219</td>
			<td>diluted to 1:200</td>
		</tr>
		<tr>
			<td>Rat anti-HCN4 (SHG 1E5)</td>
			<td>Invitrogen</td>
			<td>MA3-903</td>
			<td>diluted to 1:200</td>
		</tr>
		<tr>
			<td colspan="4">Other</td>
		</tr>
		<tr>
			<td>Plastic ring</td>
			<td></td>
			<td></td>
			<td>Self-designed and 3D printed</td>
		</tr>
		<tr>
			<td>Plasticine</td>
			<td>Cernit</td>
			<td>49655005</td>
			<td></td>
		</tr>
		<tr>
			<td>Silikonpasten, Baysilone</td>
			<td>VWR</td>
			<td>291-1220</td>
			<td></td>
		</tr>
		<tr>
			<td colspan="4">Animals</td>
		</tr>
		<tr>
			<td>Mouse, C57BL/6</td>
			<td>The Jackson Laboratory</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

By using the protocol outlined above, confocal microscopy imaging of both SAN and AVN can be reliably performed. Specific staining of the conduction system using fluorescent antibodies targeting HCN4 and staining of the working myocardium using fluorescent antibodies targeting Cx43 allows the clear identification of SAN (Figure 5, Video 1) and AVN (Figure 6, Video 2)&#160;within the intact anatom...

Watch this Scientific Journal Video about Whole-Mount Immunofluorescence Staining, Confocal Imaging and 3D Reconstruction of the Sinoatrial and Atrioventricular Node in the Mouse at JoVE.com

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

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 ...

whole-mount-immunofluorescence-staining-confocal-imaging-3d

Research

JoVE Journal

Medicine

5.2K Views.  Ludwig Maximilians University (LMU) Munich. We provide a step-by-step protocol for whole-mount immunofluorescence staining of the sinoatrial node (SAN) and atrioventricular node (AVN) in murine hearts.

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

Technical Aspects of the Mouse Aortocaval Fistula

Technical aspects of creating an arteriovenous fistula in the mouse are discussed. Under general anesthesia, an abdominal incision is made, and the aorta and inferior vena cava (IVC) are exposed. The proximal infrarenal aorta and the distal aorta are dissected for clamp placement and needle puncture, respectively. Special attention is paid to avoid dissection between the aorta and the IVC. After clamping the aorta, a 25 G needle is used to puncture both walls of the aorta into the IVC. The surrounding connective tissue is used for hemostatic compression. Successful creation of the AVF will show pulsatile arterial blood flow in the IVC. Further confirmation of successful AVF can be achieved by post-operative Doppler ultrasound.

The goal is to produce an arteriovenous fistula that is simple and reproducible. This method does not use sutures or glue adhesive. Therefore the samples can be used with the least amount of foreign materials for analysis.

Technical aspects of creating an arteriovenous fistula in the mouse are discussed. Under general anesthesia, an abdominal incision is made, and the aorta ...

Cell Death Associated with Abnormal Mitosis Observed by Confocal Imaging in Live Cancer Cells

Phenanthrene derivatives acting as potent PARP1 inhibitors prevented the bi-focal clustering of supernumerary centrosomes in multi-centrosomal human cancer cells in mitosis. The phenanthridine PJ-34 was the most potent molecule. Declustering of extra-centrosomes causes mitotic failure and cell death in multi-centrosomal cells. Most solid human cancers have high occurrence of extra-centrosomes. The activity of PJ-34 was documented in real-time by confocal imaging of live human breast cancer MDA-MB-231 cells transfected with vectors encoding for fluorescent &gamma;-tubulin, which is highly abundant in the centrosomes and for fluorescent histone H2b present in the chromosomes. Aberrant chromosomes arrangements and de-clustered &gamma;-tubulin foci representing declustered centrosomes were detected in the transfected MDA-MB-231 cells after treatment with PJ-34. Un-clustered extra-centrosomes in the two spindle poles preceded their cell death. These results linked for the first time the recently detected exclusive cytotoxic activity of PJ-34 in human cancer cells with extra-centrosomes de-clustering in mitosis, and mitotic failure leading to cell death. According to previous findings observed by confocal imaging of fixed cells, PJ-34 exclusively eradicated cancer cells with multi-centrosomes without impairing normal cells undergoing mitosis with two centrosomes and bi-focal spindles. This cytotoxic activity of PJ-34 was not shared by other potent PARP1 inhibitors, and was observed in PARP1 deficient MEF harboring extracentrosomes, suggesting its independency of PARP1 inhibition. Live confocal imaging offered a useful tool for identifying new molecules eradicating cells during mitosis.

The cytotoxic activity of the phenanthridine PJ-34 in cancer cells undergoing mitosis was documented in real time by live confocal imaging. PJ-34 eradicated human breast cancer MDA-MB-231 cells harboring extra-centrosomes in mitosis. Unlike normal bi-focal mitosis, the extra-centrosomes were not clustered in the two spindle poles in the presence of PJ-34.

Phenanthrene derivatives acting as potent PARP1 inhibitors prevented the bi-focal clustering of supernumerary centrosomes in multi-centrosomal human ...

Diagnostic Ultrasound Imaging of Mouse Diaphragm Function

Function analysis of rodent respiratory skeletal muscles, particularly the diaphragm, is commonly performed by isolating muscle strips using invasive surgical procedures. Although this is an effective method of assessing in vitro diaphragm activity, it involves non-survival surgery. The application of non-invasive ultrasound imaging as an in vivo procedure is beneficial since it not only reduces the number of animals sacrificed, but is also suitable for monitoring disease progression in live mice. Thus, our ultrasound imaging method may likely assist in the development of novel therapies that alleviate muscle injury induced by various respiratory diseases. Particularly, in clinical diagnoses of obstructive lung diseases, ultrasound imaging has the potential to be used in conjunction with other standard tests to detect the early onset of diaphragm muscle fatigue. In the current protocol, we describe how to accurately evaluate diaphragm contractility in a mouse model using a diagnostic ultrasound imaging technique.

Diagnostic ultrasound imaging has proven to be effective in diagnosing various respiratory diseases in human and animal subjects. We demonstrate a comprehensive ultrasound protocol utilized by Dr. Zuo's lab to analyze diaphragm kinetics specifically in mouse models. This is also a non-invasive research technique which can provide quantitative information on mouse respiratory muscle function.

Function analysis of rodent respiratory skeletal muscles, particularly the diaphragm, is commonly performed by isolating muscle strips using invasive ...

Collection and Extraction of Saliva DNA for Next Generation Sequencing

The preferred source of DNA in human genetics research is blood, or cell lines derived from blood, as these sources yield large quantities of high quality DNA. However, DNA extraction from saliva can yield high quality DNA with little to no degradation/fragmentation that is suitable for a variety of DNA assays without the expense of a phlebotomist and can even be acquired through the mail. However, at present, no saliva DNA collection/extraction protocols for next generation sequencing have been presented in the literature. This protocol optimizes parameters of saliva collection/storage and DNA extraction to be of sufficient quality and quantity for DNA assays with the highest standards, including microarray genotyping and next generation sequencing.

DNA extraction from saliva can provide a readily available source of high molecular weight DNA, with little to no degradation/fragmentation. This protocol provides optimized parameters for saliva collection/storage and DNA extraction to be of sufficient quality and quantity for downstream DNA assays with high quality requirements.

The preferred source of DNA in human genetics research is blood, or cell lines derived from blood, as these sources yield large quantities of high quality ...

Rose Bengal Photothrombosis by Confocal Optical Imaging In Vivo: A Model of Single Vessel Stroke

In vivo imaging techniques have increased in utilization due to recent advances in imaging dyes and optical technologies, allowing for the ability to image cellular events in an intact animal. Additionally, the ability to induce physiological disease states such as stroke in vivo increases its utility. The technique described herein allows for physiological assessment of cellular responses within the CNS following a stroke and can be adapted for other pathological conditions being studied. The technique presented uses laser excitation of the photosensitive dye Rose Bengal in vivo to induce a focal ischemic event in a single blood vessel. 
The video protocol demonstrates the preparation of a thin-skulled cranial window over the somatosensory cortex in a mouse for the induction of a Rose Bengal photothrombotic event keeping injury to the underlying dura matter and brain at a minimum. Surgical preparation is initially performed under a dissecting microscope with a custom-made surgical/imaging platform, which is then transferred to a confocal microscope equipped with an inverted objective adaptor. Representative images acquired utilizing this protocol are presented as well as time-lapse sequences of stroke induction. This technique is powerful in that the same area can be imaged repeatedly on subsequent days facilitating longitudinal in vivo studies of pathological processes following stroke.

Rose Bengal Photothrombosis by Confocal Optical Imaging In Vivo:  A Model of Single Vessel Stroke

Here, we describe a semi-invasive optical microscopy approach for the induction of a Rose Bengal photothrombotic clot in the somatosensory cortex of a mouse in vivo. The technical aspects of the imaging procedure are described from induction of a photothrombotic event to application and data collection.

In vivo imaging techniques have increased in utilization due to recent advances in imaging dyes and optical technologies, allowing for the ability to ...

Imaging- and Flow Cytometry-based Analysis of Cell Position and the Cell Cycle in 3D Melanoma Spheroids

Three-dimensional (3D) tumor spheroids are utilized in cancer research as a more accurate model of the in vivo tumor microenvironment, compared to traditional two-dimensional (2D) cell culture. The spheroid model is able to mimic the effects of cell-cell interaction, hypoxia and nutrient deprivation, and drug penetration. One characteristic of this model is the development of a necrotic core, surrounded by a ring of G1 arrested cells, with proliferating cells on the outer layers of the spheroid. Of interest in the cancer field is how different regions of the spheroid respond to drug therapies as well as genetic or environmental manipulation. We describe here the use of the fluorescence ubiquitination cell cycle indicator (FUCCI) system along with cytometry and image analysis using commercial software to characterize the cell cycle status of cells with respect to their position inside melanoma spheroids. These methods may be used to track changes in cell cycle status, gene/protein expression or cell viability in different sub-regions of tumor spheroids over time and under different conditions.

We describe two complementary methods using the fluorescence ubiquitination cell cycle indicator (FUCCI) and image analysis or flow cytometry to identify and isolate cells in the inner G1 arrested and outer proliferating regions of 3D spheroids.

Three-dimensional (3D) tumor spheroids are utilized in cancer research as a more accurate model of the in vivo tumor microenvironment, compared to ...

A Postoperative Evaluation Guideline for Computer-Assisted Reconstruction of the Mandible

Valid comparisons of postoperative accuracy results in computer-assisted reconstruction of the mandible are difficult due to heterogeneity in imaging modalities, mandibular defect classification, and evaluation methodologies between studies. This guideline uses a step-by-step approach guiding the process of imaging, classification of mandibular defects and volume assessment of three-dimensional (3D) models, after which a legitimized quantitative accuracy evaluation method can be performed between the postoperative clinical situation and the preoperative virtual plan. The condyles and the vertical and horizontal corners of the mandible are used as bony landmarks to define virtual lines in the computer-assisted surgery (CAS) software. Between these lines the axial, coronal, and both sagittal mandibular angles are calculated on both pre- and postoperative 3D models of the (neo)mandible and subsequently the deviations are calculated. By superimposing the postoperative 3D model to the preoperative virtually planned 3D model, which is fixed to the XYZ axis, the deviation between pre- and postoperative virtually planned dental implant positions can be calculated. This protocol continues and specifies an earlier publication of this evaluation guideline.

Here, we propose a practical, feasible and reproducible evaluation guideline for computer-assisted reconstruction of the mandible in order to create uniformity between studies regarding postoperative accuracy evaluation. This protocol continues and specifies an earlier publication of this evaluation guideline.

Valid comparisons of postoperative accuracy results in computer-assisted reconstruction of the mandible are difficult due to heterogeneity in imaging ...

Blood Circuit Reconstruction in an Abdominal Mouse Heart Transplantation Model

The surgical technique of heterotopic abdominal heart transplantation in mice is a standard model for research in transplantation immunology. Here, the established technique for a modified blood circuit reconstruction in a heterotopic abdominal heart transplantation model is presented. This method uses the intrathoracic inferior vena cava (IIVC) instead of the pulmonary artery of the donor heart for the anastomosis to the inferior vena cava of the recipient. It is facilitating and improving success rates for abdominal heart transplantation in mice.

A novel technique for blood circuit reconstruction in a heterotopic abdominal mouse heart transplantation model is demonstrated.

The surgical technique of heterotopic abdominal heart transplantation in mice is a standard model for research in transplantation immunology. Here, the ...

Refined CLARITY-Based Tissue Clearing for Three-Dimensional Fibroblast Organization in Healthy and Injured Mouse Hearts

Cardiovascular disease is the most prevalent cause of mortality worldwide and is often marked by heightened cardiac fibrosis that can lead to increased ventricular stiffness with altered cardiac function. This increase in cardiac ventricular fibrosis is due to activation of resident fibroblasts, although how these cells operate within the 3-dimensional (3-D) heart, at baseline or after activation, is not well understood. To examine how fibroblasts contribute to heart disease and their dynamics in the 3-D heart, a refined CLARITY-based tissue clearing and imaging method was developed that shows fluorescently labeled cardiac fibroblasts within the entire mouse heart. Tissue resident fibroblasts were genetically labeled using Rosa26-loxP-eGFP florescent reporter mice crossed with the cardiac fibroblast expressing Tcf21-MerCreMer knock-in line. This technique was used to observe fibroblast localization dynamics throughout the entire adult left ventricle in healthy mice and in fibrotic mouse models of heart disease. Interestingly, in one injury model, unique patterns of cardiac fibroblasts were observed in the injured mouse heart that followed bands of wrapped fibers in the contractile direction. In ischemic injury models, fibroblast death occurred, followed by repopulation from the infarct border zone. Collectively, this refined cardiac tissue clarifying technique and digitized imaging system allows for 3-D visualization of cardiac fibroblasts in the heart without the limitations of antibody penetration failure or previous issues surrounding lost fluorescence due to tissue processing.

A refined method of tissue clearing was developed and applied to the adult mouse heart. This method was designed to clear dense, autofluorescent cardiac tissue, while maintaining labeled fibroblast fluorescence attributed to a genetic reporter strategy.

Cardiovascular disease is the most prevalent cause of mortality worldwide and is often marked by heightened cardiac fibrosis that can lead to increased ...

Isolation and Culture of Resident Cardiac Macrophages from the Murine Sinoatrial and Atrioventricular Node

Resident cardiac macrophages have been demonstrated to facilitate the electrical conduction in the heart. The physiologic heart rhythm is initiated by electrical impulses generated in sinoatrial node (SAN) and then conducted to ventricles via atrioventricular node (AVN). To further study the role of resident macrophages in cardiac conduction system, a proper isolation of resident macrophages from SAN and AVN is necessary, but it remains challenging. Here, we provide a protocol for the reliable microdissection of the SAN and AVN in murine hearts followed by the isolation and culture of resident macrophages.
Both, SAN which is located at the junction of the crista terminalis with the superior vena cava, and AVN which is located at the apex of the triangle of Koch, are identified and microdissected. Correct location is confirmed by histologic analysis of the tissue performed with Masson's trichrome stain and by anti-HCN4.
Microdissected tissues are then enzymatically digested to obtain single cell suspensions followed by the incubation with a specific panel of antibodies directed against cell-type specific surface markers. This allows to identify, count, or isolate different cell populations by fluorescent activated cell sorting. To differentiate cardiac resident macrophages from other immune cells in the myocardium, especially recruited monocyte-derived macrophages, a delicate devised gating strategy is needed. First, lymphoid lineage cells are detected and excluded from further analysis. Then, myeloid cells are identified with resident macrophages being determined by high expression of both CD45 and CD11b, and low expression of Ly6C. With cell sorting, isolated cardiac macrophages can then be cultivated in vitro over several days for further investigation. We, therefore, describe a protocol to isolate cardiac resident macrophages located within the cardiac conduction system. We discuss pitfalls in microdissecting and digesting SAN and AVN, and provide a gating strategy to reliably identify, count and sort cardiac macrophages by fluorescence-activated cell sorting.

The protocol presented here provides a step-by-step approach for the isolation of cardiac resident macrophages from the sinoatrial node (SAN) and atrioventricular node (AVN) region of mouse hearts.

Resident cardiac macrophages have been demonstrated to facilitate the electrical conduction in the heart. The physiologic heart rhythm is initiated by ...