Microdissection and Immunofluorescence Staining of Myocardial Sleeves in Murine Pulmonary Veins

Summary

This protocol demonstrates microscopy-guided isolation and immunofluorescence staining of murine pulmonary veins. We prepare tissue samples containing the left atrium, pulmonary veins, and the corresponding lungs and stain them for cardiac Troponin T and Connexin 43.

Abstract

Pulmonary veins (PVs) are the major source of ectopic beats in atrial arrhythmias and play a crucial role in the development and progression of atrial fibrillation (AF). PVs contain myocardial sleeves (MS) composed of cardiomyocytes. MS are implicated in the initiation and maintenance of AF, as they preserve similarities to the cardiac working myocardium, including the ability to generate ectopic electrical impulses. Rodents are widely used and may represent excellent animal models to study the pulmonary vein myocardium since cardiomyocytes are widely present all over the vessel wall. However, precise microdissection and preparation of murine PVs is challenging due to the small organ size and intricate anatomy.

We demonstrate a microscopy-guided microdissection protocol for isolating the murine left atrium (LA) together with the PVs. Immunofluorescence staining using cardiac Troponin-T (cTNT) and connexin 43 (Cx43) antibodies is performed to visualize the LA and PVs in full length. Imaging at 10x and 40x magnification provides a comprehensive view of the PV structure as well as detailed insights into the myocardial architecture, particularly highlighting the presence of connexin 43 within the MS.

Introduction

Atrial fibrillation (AF) is the most common sustained arrhythmia¹. The prevalence of AF is increasing even further with an expected number of ~17.9 million patients in Europe in 2060¹. AF is clinically highly important since it is an essential risk factor for the development of myocardial infarction, heart failure, or stroke, resulting in an enormous individual, social, and socioeconomic burden¹. Even though AF has been known for decades, the pathophysiology of AF is still not fully understood².

Already in the late 1990s, studies demonstrated the great impact of pulmonary veins (PVs) in initiating and maintaining AF, as they are the main source of AF-triggering ectopic beats³. It has been demonstrated that PVs structurally differ from other blood vessels. While typical blood vessels contain smooth muscle cells, the tunica media of PVs also contains cardiomyocytes⁴. In rodents, this cardiac musculature is ubiquitously present throughout the whole PVs, including intra- and extrapulmonary parts, as well as the orifice region⁵. In humans, PVs also contain cardiomyocytes, which can be observed within extensions of the left atrial (LA) myocardium-so-called myocardial sleeves (MS)^6,7.

MS have morphological similarities to the atrial myocardium⁸. The shape and size of atrial and PV cardiomyocytes do not vary significantly between each other and show comparable electrophysiological properties⁸. Electrophysiologic recordings within the PV have proven the electrical activity of MS, and angiographic imaging has revealed contractions synchronized with the heartbeat^9,10.

Gap junctions are pore-forming protein complexes composed of six connexin subunits, which allow the passage of ions and small molecules¹¹. Gap junctions exist in the cell-to-cell appositions, interconnect neighboring cardiomyocytes, and enable an intercellular electrical coupling between cardiomyocytes^12,13. Several connexin isoforms are expressed in the heart with connexin 43 (Cx43) being the most common isoform expressed in all regions of the heart¹⁴. Previous studies provide evidence for the expression of Cx43 in cardiomyocytes of the PVs^15,16.

It remains challenging to investigate MS within intact PVs due to their delicate structure, especially in small animal models. Here, we demonstrate how to identify and isolate PVs together with LA and lung lobes in mice using microscopy-guided microdissection. Additionally, we demonstrate immunofluorescence (IF) staining of PVs to visualize cardiomyocytes and their interconnections within the PVs.

Protocol

Animal care and all experimental procedures were conducted following the guidelines of the Animal Care and Ethics Committee of the Ludwig-Maximilians-University of Munich, and all the procedures using mice were approved by the Regierung von Oberbayern (ROB 55.2-2532. Vet_02-20-215, ROB 55.2-2532. Vet_02-18-46, ROB 55.2-2532. Vet_02-19-86, ROB 55.2-2532. Vet_02-21-178, ROB 55.2-2532. Vet_02-22-170). C57BL6/N mice were commercially obtained.

1. Preparation

1. Prepare a 3% agarose gel by mixing 3 mg of agarose and 100 mL of 1x Tris-buffered saline in a 500 mL Erlenmeyer flask. Heat the solution in a microwave at 600 W for 2-3 min until the gel becomes homogeneous.
2. Prepare the dissection dish by gently pouring the gel into a 100 mm diameter Petri dish. Fill the Petri dish half with gel. Clear bubbles from the gel to ensure a homogeneous consistency. Leave the dissection dish to cool down until the gel becomes solid.
3. Prepare all the buffers and solutions needed, including fixing solution, permeabilization solution, blocking buffer, and washing buffer according to the recipes provided in Table 1.

2. Organ harvest and tissue preparation

NOTE: An extensive protocol detailing the procedure of mouse anesthesia and harvesting the heart has been previously published^17,18. Thus, we present only a brief description of that part. Experiments were performed on 12 to 16 weeks old C57BL6 mice (six male, four female). The male's body weight extended from 26 g to 28 g and the female's body weight from 19 g to 22 g. The following steps were performed without prior systemic heparin injection.

1. Anesthetize the mouse with isoflurane (5%, 95% oxygen, Flow: 1 L/min) and ensure sufficient anesthesia depth.
2. Transfer the mouse to the operation desk positioning it in a supine position. Maintain the narcosis with isoflurane inhalation (2%, 98% oxygen, Flow: 1 L/min) through an anesthesia mask and fix the mouse with tape at its extremities. Inject fentanyl for analgesia (0.1 mg/25 g body weight i.p.).
3. Lift the fur at the xiphoidal process and set a transversal 2 mm cut. Disconnect the fur from the subcutaneous fascia using blunt dissection (backside of scissors) and extend the cut cranially and laterally to expose the thorax.
4. Carefully open the abdomen caudal to the costal arch. Lift the xiphoidal process slightly to incise the diaphragm from caudal and make the lungs collapse. Then, cut the diaphragm from left to right without injuring any organs and open the thorax bilaterally to expose the heart.
5. Cut the inferior vena cava (IVC) while the heart is still beating to exsanguinate the mouse and proceed to perfuse the heart by penetrating the left ventricle with a 27 G needle and injecting 10 mL of ice-cold 1x phosphate-buffered saline (PBS).
6. After clearing the blood from the mouse heart, locate the aortic arch, superior vena cava (SVC), and the trachea. Cut them ~3 mm above the heart base and harvest the heart along with the connected lungs.
7. Submerge the harvested organs in a 10 mL conical tube containing 2 mL of sterilized 30% sucrose solution. Allow the samples to dehydrate for 24 h at 4 °C.

3. Microscopy-guided preparation of LA and PVs

1. Place the dehydrated heart and lungs into a dissection dish containing 40-50 mL of 1x PBS. Put it under a bright field microscope with 10x magnification and set up the lighting. Place the heart with its dorsal surface on the dissection dish. Identify the transition between the ventricular and atrial wall, and carefully separate the ventricles from the atria using surgical scissors.
   NOTE: Cut approximately 1 mm inferior to the atria to keep some ventricular tissue. This will be needed later for pinning the atria to the dissection dish without damaging structures at the heart base. We suggest keeping the ventricles since they can be used as a positive control sample. To embed the ventricular tissue, follow the same procedure outlined for embedding of the PVs, as described in Section 4.
2. Place the separated atria with the ventricular cutting surface (step 3.1) on the dissection dish so that the heart base is facing up. Orientate the atria so that the lung lobes are posterior, the LA and the left atrial appendage (LAA) are on the right, and the right atrium (RA) and right atrial appendage (RAA) are on the left. Fix the preparation by pinning the remaining left ventricular tissue to the dissection dish. Gently spread out the lung lobes without applying excessive force and pin them to the dissection dish (Figure 1A).
3. Get an overview of the anatomic landmarks at the heart base.
   1. Find the aorta with the aortic arch in the center of the heart base.
   2. Detect the pulmonary trunk (PT) directly to the right of the aorta and follow its course towards the posterior to distinguish the pulmonary arteries (PAs). Verify their location by following their course up to the left lobe and right superior/middle lobes to find the corresponding lung hilum (LH).
   3. Determine the position of the trachea and/or the left and right main bronchi (L Br, R Br), which are located posterior to the aorta, and verify their location by following their course towards the LH (Figure 1A).
   4. Find the SVC to the left of the aorta and verify by following its course into the RA next to the RAA.
4. Remove connective and fat tissue around the heart base while preserving all the identified landmarks mentioned above. Additionally, remove the aortic arch and trachea to have a free view of the heart base (Figure 1B).
5. Gently separate the PAs from the upper surface of the atrium by blunt dissection using fine forceps. Afterward, cut them off at LH and flip them to the side to expose the left atrial free wall (LAFW) and the PVs in their full dimension. Cut off the main bronchi from the LH and remove the bronchial tissue (Figure 1C).
   NOTE: The LH serves both as the common entry point for PAs and airways into the lungs, as well as the exit point for the PVs. It is important to perform every procedure at the LH carefully to avoid any damage to the PVs.
6. To separate the LA/RA and left and right ventricle (LV, RV), perform a cut starting from the anterior part of the remaining RV upwards through the tricuspid valve (TV) till the anterior side of the SVC to open the RA from the frontal side. Cut the posterior RA free wall (RAFW) next to the interatrial septum to separate the RA and LA. Cut the posterior right ventricular wall next to the interventricular septum to completely sever the LV and RV.
   NOTE: An extensive protocol describing the procedure of the preparation and isolation of the right atrium in detail has been published previously¹⁹. The separation of the RA is useful to remove unwanted tissue from the LA-PV tissue complex and to collect tissue for additional experiments.
7. Reduce the size of the lung lobes by cutting off some of the basal lung tissue so that approximately 3-4 mm of lung tissue is left over.
   ​NOTE: Preserve the apices of the lung lobes as they serve as floats during the subsequent embedding steps and allow embedding of the PVs horizontally in the O.C.T compound.

4. Tissue embedding

1. Transfer the preparation, including LA and PVs, into a cryomold, ensuring that the heart base and lung apex are facing upwards. Arrange them in a physiological configuration-ensure that the PVs are not twisted or flipped.
2. Fill the cryomold with O.C.T compound and gently compress the PVs with fine forceps to remove any remaining air. Maintain their physiological configuration unchanged throughout the process.
3. Put the cryomold with the embedded tissue on dry ice to freeze the O.C.T. compound. Store the samples at -80 °C for future use.
   ​NOTE: It is crucial to keep the PVs in a horizontal position to ensure obtaining sections that contain PVs in full length for subsequent procedures.

5. Cutting and collection of frozen tissue sections

NOTE: To cut the tissue blocks, the machine settings were adjusted to a specimen temperature of -18 °C, and blade temperature of -25 °C.

1. Install a tissue block on the specimen holder by applying a layer of O.C.T. compound between the tissue block and the specimen holder. Freeze them for 3-4 min.
2. Block the specimen head and load the specimen holder with the tissue block on the specimen head by closing the specimen chuck release lever. Finely adjust the position of the tissue block using the fine adjustment knobs.
3. Remove the cryomold from the specimen. Unblock the specimen head and the electrical brake of the cryostat.
4. Set the cryostat to trim mode with a trim size between 30 µm and 50 µm. Trim the tissue specimen until the PVs become visible.
5. Switch to the fine section-cutting mode and set the thickness to 10 µm. Start collecting sections, including PVs and connected lung and atrial tissue. Collect sections using an anti-roll plate or by fixing the free end of each section to the blade holder with a fine cold brush and spread them out.
6. Position a slide (kept at room temperature) adjacent to the section. Carefully release the section from the brush, allowing the section to naturally adhere to the slide, as the frozen section and O.C.T. compound melt upon touching the warmer slide surface.
   NOTE: Maintain a stable and gentle condition during the sectioning process to prevent any ruptures or folds in the sections. Replace the blade with a new blade if necessary.
7. Collect up to three sections on each slide. Label the slides and store them at -20 °C.
   ​NOTE: We recommend collecting at least five sections (equivalent to two slides) for each PV.

6. Immunofluorescence staining of cryosections from the PVs

1. Arrange the frozen slides on a staining system and allow the sections to thaw for approximately 20 min at room temperature.
   NOTE: Fill the staining system with water to keep the samples humidified. Drying of the tissue can damage antigens, causing non-specific antibody binding and overstaining artifacts during the staining procedures.
2. After 20 min of thawing, cover the sections with a few drops of fixing solution (4% paraformaldehyde [PFA], Table 1) to fix the tissue on the slides for 10 min at room temperature.
3. After fixation, transfer the slides to the vertical slide rack and immerse them in a container filled with 1x PBS. Place the container on a shaker at low speed and wash the slides for 5 min. Repeat this wash cycle two more times, using fresh 1x PBS each time.
4. After washing, circle every individual section on each slide with a xylol-containing liquid blocker pen. Rearrange the slides on the staining system and apply 1 or 2 drops of permeabilization solution (0.1% Triton X-100, Table 1) onto each sample with a Pasteur pipette. Allow the sections to incubate at room temperature for 10 min to permeabilize the cell and cell organelle membranes.
5. After permeabilization, remove the 0.1% Triton X-100 solution by washing the slides for 3 x 5 min in 1x PBS, following the same procedure as described in Step 6.3.
6. After washing, rearrange the slides on the staining system and apply 1 or 2 drops of the Blocking Buffer (Table 1) to each section, ensuring they are fully covered with the blocking buffer. Allow the slides to incubate for 1 h at room temperature.
7. Prepare 1 mL of the primary antibody mix by pipetting 5 µL of a mouse anti-cTNT antibody (diluted 1:200) and 1 µL of a rabbit anti-Cx43 antibody (diluted 1:1,000) into a 1.5 mL microcentrifuge tube filled with 994 µL of Blocking Buffer. Pipette the mixture up and down to ensure thorough mixing.
   NOTE: Adjust the amount of antibody mixture applied according to the size of the section. Ensure that the sections are fully covered by the antibody mixture. The concentration, incubation time, and optimal incubation temperature of antibodies can vary depending on factors such as antibody type, antibody clones, and tissue characteristics. We recommend testing and optimizing the staining conditions individually for each antibody.
8. After the blocking step (step 6.6), remove the remaining blocking buffer and directly apply 2 or 3 drops of the primary antibody mix onto each section. Allow the slides to incubate overnight at 4 °C.
9. After the primary antibody incubation, wash the slides for 3 x 5 min with Washing Buffer under gentle shaking (Table 1).
10. Prepare 1 mL of the secondary antibody mix by pipetting 1 µL of an AF488-conjugated goat antimouse secondary antibody (diluted 1:1,000) and 1 µL of an AF568-conjugated goat antirabbit secondary antibody (diluted 1:1,000) into a 1.5 mL microcentrifuge tube filled with 998 µL of Washing Buffer. Pipette the mixture up and down to mix well.
11. After the washing step (step 6.9), rearrange the slides in the staining system and apply 2 or 3 drops of the secondary antibody mixture onto each section with a pipette. Allow the antibodies to incubate for 45 min at room temperature while protecting the samples from light to prevent fluorophore quenching. After the secondary antibody incubation, wash the slides for 3 x 5 min with Washing Buffer under gentle shaking (Table 1).
12. Rearrange the slides on the staining system. Counterstain the sections with Hoechst-33342 (in a dilution of 1:1,000 in 1x PBS) by applying 2 to 3 drops of the Hoechst-33342 solution onto each section. Allow the slides to incubate for 10 min at room temperature.
13. After the incubation, wash the slides 3 x 5 min with Washing Buffer under gentle shaking (Table 1). Mount the slides by applying 1 or 2 drops of fluorescent mounting medium onto each slide. Cover the slides with coverslips.
    NOTE: Make sure the coverslip is flat when placing it. If any air bubbles are present, gently apply pressure on the side of the bubble to remove them.
14. Store the mounted slides in a slide saver at 4 °C.
    ​NOTE: It is recommended to image the slides as early as possible. This protocol is not exclusively valid for the mentioned antibodies. Target antigens and antibodies can be replaced or modified according to the individual experiment requirements or alternative antigen detection strategies.

7. Imaging of the immunofluorescence staining slices

1. Set up the microscope.
   1. Start the microscope with the laser light source, the connected computer, and the accompanying software following the manufacturer's manual.
   2. Enter the Configuration menu. Click to select the suitable camera type for fluorescence imaging (e.g., DFC365FX monochrome camera).
   3. Enter the Acquire menu and create three channels.
   4. Select channel FCr1 for the detection of Hoechst-33342, label it, and select the DAP filter cube in the selection section. Set the exposure time to 200 ms, the electronic gain to 2, and the light intensity to 17%.
   5. Select channel FCr2 for the detection of cTNT (stained with AF488-conjugated antibody), label it, and select the L5 filter cube in the selection section. Set the exposure time to 200-300 ms, the electronic gain to 1.8, and the light intensity to 100%.
   6. Select channel FCr3 for the detection of Cx43 (stained with AF568-conjugated antibody), label it, and select the TXR filter cube in the selection section. Set the exposure time to 150 ms, the electronic gain to 2, and the light intensity to 100%.
      NOTE: The spectral characteristics of the filter cubes are listed in Table 2.
2. Load the slide on the microscope stage.
3. Perform an overview imaging at 10x magnification:
   1. Select the 10x objective and open the laser shutter by clicking Live. With the FCr1 channel selected, use the navigator function to navigate through the slide until a signal is detected. Roughly focus the sample until the cell nuclei are distinguishable.
   2. Initiate the spiral scanning mode to capture a complete preview of the sample. Deactivate live by clicking the button again to protect the specimen. Identify the LA, PVs, and lung tissue.
   3. Define the area of interest for the subsequent tile scan; select multiple focus points within the area of interest. Click Live in the FCr1 channel and adjust the focus for each focus point. Save the corresponding Z-positions, which represent the exact focus position for each focus point. Start the tile scan at 10x magnification to capture a complete overview image of the sample.
   4. After the scan is complete, open the tile scan and merge the individual images. To enhance the brightness and contrast of single channels, select them in the object bar and adjust their signal range with the sliders as desired.
4. Obtain zoomed-in images at 40x magnification:
   1. Select the 40x objective. Adjust the exposure time and gain settings for each channel as follows: FCr1: DAP filter cube: exposure time: 50 ms, gain: 1.5; FCr2: L5 filter cube: exposure time: 80 ms, gain: 1.8; FCr3: TXR filter cube: exposure time: 20 ms, gain: 2.
   2. Use the overview image of step 7.3 to locate the PV orifice (PVO), as well as extrapulmonary and intrapulmonary PV (PV_ex, PV_in) regions. Define this area to be scanned.
   3. Open the submenu Image and click on z to activate Z-Stack.
      1. Select the FCr1 channel and open the laser shutter to obtain a live preview.
      2. Go to the Z-Stack submenu and determine the Z-Stack starting and end points by rotating the focus fine adjustment knob clockwise and counterclockwise until the overall signal begins to lose focus.
      3. After setting up the focus range, select the System Optimized Z-Stack mode, and activate the calculate extended depth of field images (EDF) option. Depending on how flat the tissue is, expect approximately 20 layers with a step interval of approximately 0.5 µm.
   4. Begin the tile scan. After scanning, open the tile scan and merge only the EDF image. To enhance the brightness and contrast of single channels, select them in the object bar and adjust their signal range with the sliders as desired (see step 7.3.4).
5. After generating the images, save the project and export both, the merged version and the raw channels of each image as a TIFF file.
   ​NOTE: Saving the files as ImageJ-readable TIFF allows ImageJ to import the original tile size in µm instead of pixels. Always close the laser shutter whenever a laser signal is not needed to prevent photobleaching of the secondary antibodies.

8. Image editing with ImageJ

1. Load the corresponding raw channel files in ImageJ. Click on Image | Color | Merge and choose the desired color for each channel.
2. Create a scale bar by selecting Analyze | Tools | Scale Bar and paste it in the bottom right corner. Proceed with further processing and analysis according to the image requirements. Save the merged images when finished.

Results

We performed the microdissection, staining, and imaging of the PVs in 10 12-16-week-old mice. Following the protocol, we successfully microdissected PVs together with the LA in all experimental mice and obtained sections with a comprehensive view of the PVs in eight mice. Overview images were taken at 10x magnification to identify the PV orifice (PVO) region at the LA-PV junction, the extrapulmonary PVs (PV_ex) (PVs in between the lung hilum and the LA-PV junction), and the intrapulmonary PVs (PV_in) ...

Discussion

With this protocol, we share a method to distinguish and isolate the PVs of the mouse heart and perform immunofluorescence staining on them. After the organ harvest, the heart and lungs were dehydrated in sterilized sucrose solution, followed by separating the ventricles from the atrium and lung lobes under microscopic guidance. Afterwards, the heart base was prepared to visualize the PVs followed by cutting them from the lungs at the hilum. The subsequent immunofluorescence staining was performed using a cryotechnique b...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Adhesion slides	Epredia	10149870
AF568-secondary antibody	Invitrogen	A11036	Host: Goat, Reactivity: Rabbit
Agarose	Biozym LE	840104
Alexa Fluor 488-secondary antibody	Cell Signaling Technology	4408S	Host: Goat, Reactivity: Mouse
Anti-Connexin 43 /GJA1 antibody	Abcam	ab11370	Polyclonal Antibody, Clone: GJA1, Host: Rabbit
Anti-cTNT antibody	Invitrogen	MA5-12960	Monoclonal Antibody, Clone: 13-11, Host: Mouse
Bovine serum albumin	Sigma-Aldrich	A2153
Brush	Lukas	5486	size 6
Cover slips	Epredia		24 mm x 50 mm
Cryotome Cryo Star NX70	Epredia		Settings: Specimen temperature: -18 °C, Blade Temperature: -25 °C
DFC365FX camera	Leica
DM6 B fluorescence microscope	Leica
Dry ice
Dubecco's phosphate-buffered saline (DPBS) 1x conc.	Gibco	14040133	500 mL
Dumont #5FS Forceps	F.S.T.	91150-20	2 pieces needed
Fine Scissors	F.S.T.	14090-09
Fluorescence mounting medium	DAKO	S3023
Graefe Forceps	F.S.T.	11052-10
Hoechst 33342	Invitrogen	H3570	Cell nuclei counterstaining
ImageJ	FIJI		analysis and processing software
LAS X	Leica		Imaging software for Leica DM6 B
Microtome blades S35	Feather	207500000
Microwave
Normal goat serum	Sigma-Aldrich	S26-M
O.C.T. compound	Tissue-Tek	4583
Paraformaldehyde 16%	Pierce	28908	methanol-free
Pasteur pipettes	VWR	612-1681
Petri dish	TPP	93100	100 mm diameter
Rocker 3D digital	IKA Schüttler	00040010000
Slide staining jars	EasyDip	M900-12
Specimen Molds	Tissue-Tek Cryomold	4557	25 mm x 20 mm x 5 mm
StainTray M920 staining system	StainTray	631-1923	Staining system for 20 slides
Sterican Needle	Braun	4657705	G 27 - used for injection (step 2) and pinning (step 3 and 4) in the protocol
Student Vannas Spring Scissors	F.S.T.	91500-09
Super PAP Pen Liquid Blocker	Super PAP Pen	N71310-N
Syringes	Braun	4606108V	10 mL
Tris base	Roche	TRIS-RO	component for 1x Tris-Buffered Saline (TBS)
Triton X-100	Sigma-Aldrich	T8787
Tween 20	Sigma-Aldrich	P2287