Atrial fibrillation is the most common arrhythmia, but the complex pathophysiology is still not fully understood. With our research, we contribute to a better understanding of the mechanisms that lead to the initiation and maintenance of atrial fibrillation. Translating medical research from bench to bedside is a major challenge.
To address this, we established a translational pipeline from cellular models over mouse models to large animal models and pigs, allowing us to identify and validate novel therapeutic targets. Mice are often used for studying arrhythmia. However, it is challenging to study the pulmonary veins in mice.
The pulmonary veins are the main triggers for atrial fibrillation in patients. Our protocol provides an effective guideline for identifying and microdissecting pulmonary veins to conduct further investigations. To begin, place the dehydrated heart and lungs isolated from a mouse into a dissection dish containing 40 to 50 milliliters of PBS.
Position it under a brightfield microscope with 10X magnification and set up the lighting. Then place the heart's dorsal surface on the dissection dish. Using surgical scissors, carefully separate the ventricles from the atria after identifying the transition between the ventricular and atrial wall.
Place the separated atria on the dissection dish with the heart base facing up. Orient the atria with the lung lobes positioned posteriorly, the left atrium and its appendage situated on the right, and the right atrium and its appendage situated on the left. Pin the remaining left ventricular tissue to the dissection dish to secure the preparation.
Spread the long lobes gently without exerting excessive force and pin them to the dissection dish. Next, find the aorta with the aortic arch in the center of the heart base. Remove connective and fat tissue around the heart base, while preserving all identified landmarks.
Also, remove the aortic arch to clear the view of the heart base. Detect the pulmonary trunk directly to the right of the aorta and trace its path toward the back to identify the pulmonary arteries. To confirm their position, track their root to the left lobe and right superior or middle lobes to pinpoint the lung hilum.
Determine the location of the left and right main bronchi behind the aorta. Confirm their position by tracing their path towards the lung hilum. Then identify the superior vena cava to the left of the aorta and trace its path into the right atrium near the right atrial appendage.
Using blunt dissection with fine forceps, carefully detach the pulmonary arteries from the atrium's top surface. Then sever them in the lung hilum and move them aside to unveil the left atrial free wall and the pulmonary veins in full view. Remove the main bronchi from the lung hilum and discard the bronchial tissue.
Examine the lung hilum to identify the ostium of the pulmonary artery and the main bronchus. To separate the atria and ventricles, make a cut from the front part of the remaining right ventricle and move upward through the tricuspid valve to the front side of the vena cava superior, opening the right atrium from the frontal side. Then cut the posterior right atrial free wall adjacent to the interatrial septum to create a division between the right and left atrium.
Cut the posterior right ventricular wall next to the interventricular septum to completely disconnect the left and right ventricles. Remove some of the basal lung tissue to trim the size of the lung lobes, leaving three to four millimeters of the lung tissue. To begin, isolate the left atrium and pulmonary veins or PVs from the mouse's heart.
Place the isolated tissue into a cryo mold with the heart base and lung apex facing up. Untwist and untangle the PVs in case they are convoluted. Then fill the cryo mold with the OCT compound.
Using fine forceps, apply gentle compression on the PVs without moving them to remove residual air. Freeze the tissue in the OCT compound on dry ice. For immunostaining, thaw the frozen mouse PV slides on a staining system for 20 minutes.
Then administer several drops of a 4%paraformaldehyde fixing solution onto the sections to secure the tissue to the slides. After incubation, move the slides to a vertical rack and submerge them in a container filled with PBS. Set the container on a slow shaker for five minutes to rinse the slides.
Using a liquid blocker pen containing xylol, encircle each section on every slide and reorganize the slides on the staining system. Using a Pasteur pipette, apply one or two drops of permeabilization solution onto each sample. Then clean the Triton X-100 solution by washing the slides for five minutes in PBS.
After rearranging the slides on the staining system, add one or two drops of the blocking buffer to each section, ensuring complete coverage. Next, prepare one milliliter of the primary antibody mix consisting of the primary antibodies and blocking buffer. Discard the leftover blocking buffer from the slide.
Administer two or three drops of the primary antibody mixture to each section. After incubation, place the slides in a vertical rack and rinse the slides for five minutes in wash buffer while maintaining gentle agitation. Then prepare one milliliter of the secondary antibody mix consisting of the secondary antibodies and washing buffer.
Reposition the slides on the staining system and use a pipette to dispense two or three drops of the secondary antibody blend onto each section. Subsequently, clean the slides three times for five minutes each in wash buffer while shaking. Next, add two or three drops of the Hoechst 33342 solution to each section arranged on the staining system.
Place the slides in a vertical rack and rinse them for five minutes in wash buffer with gentle agitation. Add one or two drops of fluorescent mounting medium to each slide and seal the slides with coverslips. The pulmonary vein orifice region at the left atrium and pulmonary vein junction, extrapulmonary veins and intrapulmonary veins were observed under 10X magnification.
Specific cTnT signals with a typical muscular striation in the pulmonary vein orifice, extrapulmonary veins and intrapulmonary veins were observed at 40X magnification. CX43 was found in all three pulmonary regions and was primarily projected between neighboring cardiomyocytes. CX43-related signals were observed on the polar side and the lateral side of the cardiomyocytes in the myocardial sleeves.
