Analysis of Tubular Membrane Networks in Cardiac Myocytes from Atria and Ventricles

NOTE: All animal procedures were reviewed and approved by the Institutional Animal Care and Use Committee of the University Medical Center Goettingen in compliance with the humane care and use of laboratory animals.

1. Isolation of Atrial and Ventricular Myocytes from the Mouse Heart

1. Handle animals as gently as possible and according to approved protocols to minimize stress in general and specifically to avoid potential powerful inadvertent effects from neurohormonal excesses on isolated cardiac cells. In addition, inject each mouse with heparin (500 IU/kg body weight s.c.) at least 20 min prior to heart extraction to prevent blood clotting and micro emboli which can significantly compromise the yield and integrity of cardiac cells during isolation.
2. Anesthetize mice aged 12 weeks or older by isoflurane inhalation, confirm absence of pain withdrawal reflexes, and euthanize animals by cervical dislocation.
   1. Extract the heart swiftly following previously established expert protocols (e.g., see Kaestner et al.²⁶ and Louch et al.²⁷). Avoid any inadvertent damage to the atria by unnecessary squeezing or stretching.
   2. Carefully preserve the tissue of the proximal ascending aorta using a pair of stump forceps and straight scissors to establish a continuous transversal cutting edge through the aortic vessel wall, which is important for successful cannulation and perfusion of the heart.
3. Transfer the excised heart immediately into ice-cold nominally Ca²⁺ free perfusion buffer (for solutions see Table 1). Keep the great vessels clamped during transfer through air into buffer to avoid inadvertent air embolism until the heart is fully submerged. Use BDM in the buffer and ice-cooled solutions to inhibit cardiac contraction.
4. Use a binocular zoom microscope with sufficient 3D illumination.
   1. Cannulate the aorta under panoramic stereovision of the heart with a smooth, surface polished 21 G cannula (outer diameter 0.81 mm; for normal mouse heart weight) which must be completely filled with buffer. Ensure that there are no air bubbles in the cannula by connecting a proximal solution reservoir (e.g., a syringe) through a 2-way Luer valve allowing for rapid solution flow control.
   2. Confirm under binocular magnification that the cannula is positioned correctly inside the aorta, which is approximately 1 mm above the aortic valves and coronary artery branches. Absolutely avoid any passage through or inadvertent perforation of the aortic valves with the cannula (this will permanently disrupt aortic valve closure and consequently disrupt cardiac perfusion).
5. Tie the aorta gently to custom-made, circumferentially oriented anti-slip grooves near the end of the cannula using two silk sutures. Do not flush the coronary arteries forcefully at any point. Connect the solution filled cannula tied to the aorta and the heart to a tightly fitting outflow connector of a customized and pre-calibrated perfusion system, a.k.a. the modified Langendorff setup (either using constant pressure or constant flow; see also discussion section below).
6. Perfuse the heart as soon as possible for 4 min using oxygenated perfusion buffer at 37 °C (target perfusion rate: 4 ml/min). Start the digestion by switching the perfusion to collagenase containing digestion buffer (600 U/ml collagenase type II) for 8-10 min at 37 °C. Monitor the progress of tissue digestion by confirming similar tissue changes including increasing opaqueness, softness, and flaccidity throughout the apparent heart surface.
7. Dissect the cardiac chambers as needed following digestion. Place the cannulated heart under a binocular microscope and visualize the posterior heart wall. Dissect residual non-cardiac tissue e.g., lung and vessel parts to avoid cell contamination in the digestion buffer using micro scissors (e.g., spring scissors with 8 mm straight blades) as shown in Figure 1.
8. Follow a check list which particular chambers, regions, and/or cells of the collagenase digested heart should be harvested (Figure 1): left and/or right atrium, free left and/or right ventricular wall, and/or the ventricular septum.
   1. For dissection of specific cardiac tissues, use a relatively wide and flat dissection bath coated with a several mm thick layer of silicone plastic elastomer. Fix the apex of the heart with a fine insect steel pin to the bottom elastomer layer.
   2. Deflect the right atrial appendage and dissect the right atrium just above the atrioventricular valves. Continue the dissection with the left atrium. Dissect and discard the fibrous valve apparatus. Finally, dissect the left and right free ventricular walls and the septum and/or smaller tissue parts as needed.
   3. Note only for trainees: to gain practice, start with non-digested mouse hearts. For ease of anatomical orientation, practice the tissue handling including all consecutive dissection steps under binocular vision as shown in Figure 1. Once the 3D anatomy, manual handling under binocular vision, and dissection steps are sufficiently familiarized, continue with the collagenase digested mouse hearts as outlined above.
9. For ventricular myocyte (VM) cell isolation: transfer the ventricular tissue into 2.5 ml of fresh digestion buffer. If simultaneous cell isolations from several organ parts, e.g., atria and ventricles is attempted, a second person may take over one leg of the cell dissociation procedure, both to minimize and optimize use of mice through coordinated handling of multiple cardiac tissues. Continue with steps 1.9.1-1.9.4, thereafter 1.10.
   1. Dissect either the entire ventricular tissue or specific parts thereof (e.g., LV, RV, free walls, and/or septum) into approximately 1 mm³ pieces in 2.5 ml digestion buffer using sharp scissors (e.g., spring scissors with 8 mm straight blades) in a 60 mm Petri dish.
   2. Gently dissociate VMs into cell suspension by slow trituration of the tissue pieces with a transfer pipette. Avoid any air bubbling in cell suspension.
   3. Add 8 ml of stop buffer to the VM cell suspension and transfer the cell suspension into a 15 ml conical tube. Allow the remaining tissue pieces to settle at the bottom for approximately 15 sec, but short enough for isolated cells to remain in suspension. Next, harvest the VM suspension via transfer of the supernatant volume to a new 15 ml tube. If excessive tissue pieces are present, alternatively use a minimally 200 µm spaced nylon mesh to separate the tissue pieces from the cell suspension.
   4. Let the VM cell suspension settle to the bottom of a 15 ml conical tube by gravity for 8 min.
   5. Wash step: remove the supernatant and gently resuspend the remaining VM pellet in 10 ml of perfusion buffer. Repeat wash step 1.9.5 (optional: add additional wash steps to gradually increase the Ca²⁺ concentration as needed).
   6. Resuspend the settled VM pellet in 10 ml perfusion buffer and distribute the remaining cell suspension volume in 1.5 ml microcentrifuge tubes (approximately 50,000 VM cells per tube).
10. For atrial myocyte (AM) cell isolation: transfer the digested/dissected atrial tissue into 1 ml of fresh digestion buffer.
    1. Cut the partially digested atrial tissue into approximately 1 mm³ pieces in 1 ml digestion buffer using micro scissors in a small Petri dish (e.g., 60 mm diameter). Gently dissociate AM cells out of the digested tissue pieces into cell suspension using trituration with a 1 ml plastic pipette with a cut tip to avoid damaging fluid jets. During trituration strictly avoid any air bubbling in cell suspension. Following mechanical agitation, add 4 ml of stop buffer (50 µM CaCl₂; 10% BCS) to arrest any remaining collagenase activity in cell suspension.
    2. Transfer the AM cell suspension into a 15 ml conical tube. Allow the remaining tissue pieces to settle at the bottom for approximately 15 sec, but short enough for the isolated cells to remain in suspension. Harvest the supernatant volume containing the free AM cells via solution transfer to a new 15 ml tube.
    3. Centrifuge the AM cell suspension, e.g., 2 min at 20 x g at RT or – preferable for membrane studies – let the cells settle down slowly by gravity for 20 min in a 15 ml conical tube.
    4. Wash step: discard the supernatant and gently resuspend the AM pellet in 5 ml perfusion buffer. Repeat 1.10.4.
    5. Resuspend AM cells gently in 5 ml perfusion buffer. Distribute the cell suspension volume in 1.5 ml microcentrifuge tubes (approximately 1,000 AM cells per tube).
11. Analyze and document the isolated cell population quality for each heart including the cell yield using trypan blue staining.
    1. For this, dilute 500 µl of the cell suspension as 1:1 vol/vol with trypan blue solution (final concentration 0.02%) using 1 ml cut pipette tips. Mix the cells and the trypan blue gently by very slow up/down pipetting. Immediately apply the trypan blue containing cell suspension to a Neubauer-type improved cytometer and count the intact myocytes using an inverted microscope.
    2. Exclude any cells with apparent damage, membrane blebs, disrupted striations, contractures, and cells accumulating intracellular trypan blue (Figure 2). Also exclude spontaneously contracting cells, which are susceptible to subsequent cell death. To assess the count of intact cells in suspension, use only cardiac myocytes with regular striations which exclude trypan blue throughout the cell volume.
12. Judge the integrity of individual atrial and ventricular myocytes by transmitted light microscopy. Save the bright field image as a Tif file for documentation and further analysis.
    1. Use the following criteria for analysis of cardiac cell integrity:
       1. confirm the presence of regular striations throughout the visible cell volume;
       2. confirm the continuous integrity of the lateral surface membrane on both cell sides parallel to the myofilaments;
       3. visualize sharp serrations at the intercalated discs on both cell sides which reflect the integrity of specific surface membrane structures; and
       4. visualize the fluorescent signals of TATS membranes (or immunolabeled caveolin-3 protein or other membrane markers) as described in section 3 for localization correlation next to the underlying cell-specific bright field image morphology (e.g., combine both images with ImageJ as overlay compound image).
       5. Determine the sarcomere length from the bright field images. For mean sarcomere length, measure the distance of sequentially aligned sarcomere striations, and divide the distance by the number of sarcomeres. Measure at least two locations per cell. Perform the analysis with commercial software or offline with ImageJ.
          NOTE: If treated with uncoupling agents, intact relaxed VMs from mouse heart show a mean sarcomere length of ~1.9 µm²⁸.
13. Quantify the morphology and dimensions of individual atrial and ventricular myocytes from the transmitted light microscopy image. Consider that AM and VM cells differ significantly in size. Measure the cell length, width, and area, and calculate the length:width ratio.
    1. Analyze the 2D cell dimensions from the transmitted light image in ImageJ using the commands polygon selection tool and add to ROI manager, analogous to Figure 3. If morphological changes are expected within specific study contexts, further document for all cells the specific mouse strain, age, sex, heart size, and any interventions for subsequent data classification.

2. Staining of TATS Membranes in Living Atrial and Ventricular Myocytes

1. Load the imaging chamber (e.g., POC-R2) with a 42 mm glass coverslip. For stable myocyte attachment to the coverslip, prepare 20 µl of laminin solution by 1:10 dilution of the laminin stock in physiological perfusion buffer (final concentration 0.2 mg/ml). Spread 20 µl of the laminin solution evenly on the glass coverslip.
2. Prepare 800 µl of a 50 µM di-8-ANEPPS solution in perfusion buffer. For this, dilute 20 µl of 2 mM di-8-ANEPPS stock solution in 780 µl physiological buffer.
3. To stain VMs, let the cells settle by gravity for 8 min in a 1.5 ml reaction tube. To stain AMs, either use gravity sedimentation or spin the cell suspension for 2 min (refer to 1.10.3). For both AMs and VMs, carefully remove the supernatant while avoiding unnecessary agitation of the cell pellet and gently resuspend the cell pellet in 800 µl of di-8-ANEPPS containing solution (50 µM). Immediately transfer the di-8-ANEPPS/myocyte suspension onto the laminin-coated coverslip in the imaging chamber.
4. Stain the VM suspension for 15 min at RT in the dark.
5. Slowly remove excess volume via the upward fluid meniscus at the sides of the imaging chamber with a manual pipette. Confirm that the majority of di-8-ANEPPS stained myocytes remains firmly attached to the laminin-coated coverslip and do not become exposed to air. Next, wash the attached myocyte suspension once by slowly adding 1 ml of perfusion buffer followed by removing any excess fluid including non-adherent cells.
6. Carefully overlay the stained and surface attached myocytes with 1 ml of perfusion buffer slowly from the side of the imaging chamber. Place the imaging chamber on the microscope stage.

3. Imaging of TATS Membrane Structures in Living Atrial and Ventricular Myocytes

1. In general, carefully choose the best possible fluorescent microscope option(s) available for TATS membrane imaging. For confocal imaging, consider recent generation, modern fluorescence microscopes with optimized PMT array detectors and photon recycling pathways that maximize fluorescent signal intensity. For confocal imaging of smaller details of TATS membrane structures use a 63X 1.4 NA oil objective or – depending on availability – use a STED superresolution microscope for smallest TATS details as reviewed for myocyte-specific applications by Kohl et al.³⁴ For general principles of high resolution fluorescence microscopy, refer to the discussion section.
2. Set the imaging parameters to detect most, ideally all di-8-ANEPPS stained intracellular membranes inside a given myocyte imaging plane. Use the following parameters as starting point for confocal laser scanning microscopy: excitation 458 nm e.g., at 3% of the maximal laser power; detect the emitted signal between 550 nm and 740 nm; detector gain (e.g., master command 800); and pinhole 1 AU for an optical slice thickness of 900 nm. Adjust these parameters to optimize the signal-to-noise.
3. Use the bright field mode to select an intact AM or VM cell as appropriate (Figures 4A and 4B). Refer to point 1.12 for relevant criteria how to judge cell integrity to select cells as summarized: cell-wide regular striations and equal sarcomere spacing, sharp surface edges and serrations on all four cell sides, continuous integrity of the lateral surface membrane, and absence of any membrane blebs.
4. Take a sample image of a central intracellular myocyte section. To adjust the ROI use the “crop” function →adjust the crop window →the final pixel size measures 100 nm x 100 nm. Adjust the x-axis of the crop window to correspond with the major (axial / longitudinal) axis of the myocyte.
5. Select the final imaging plane. Use single image frames to manually select the appropriate imaging plane in the z-direction. Confirm that the TATS membranes, including T-tubule and A-tubule components, are visually apparent in the focal plane. Note that a typical intracellular imaging plane may include a nucleus as intracellular reference point. Refer to the examples in Figures 4A and 4B.
   NOTE: In general, keep cell exposure to laser light as short as possible. If possible, use single image frames to determine the optimal focal plane in cardiac myocytes.
6. Adjust the pixel dwell time to approximately 0.5 µsec. Select 16x averaging and record the image as snapshot. Repeat the image snapshot step to establish the appropriate imaging plane as outlined for TATS membrane structures in 3.5 as needed.
7. Save the final image and confirm that the file was saved in its target folder. In general, save all image files in the same format (e.g., lsm) for uniform application of analysis software. Prior to any image analysis, once again confirm sufficient cell integrity off-line (consider criteria listed under step 1.12.1) and exclude any damaged cells from analysis. Refer to the examples in Figures 4C and 4D.

4. Analysis of the TATS Membrane Network and its Components

The following image processing steps for direct analysis of TATS membrane components are summarized as top-down workflow diagram in Figures 5A and 5B.

1. Open the image file of a di-8-ANEPPS stained myocyte in Fiji (http://fiji.sc/), a freely available variant of ImageJ which contains analysis plugins essential for image processing. For further information please refer to Schindelin et al²⁹.
2. Save the image →File →Save As →Tif.
3. To analyze the TATS membrane components, select the appropriate ROI excluding the outer surface membrane signal, then use the “polygon selection” tool to delineate the ROI border excluding the outer surface membrane (sarcolemma) and including the intracellular portions of TATS membranes as shown in Figure 5A (ROI). Add the selected ROI to the “ROI Manager” by applying Analyze →Tools →ROI Manager.
   1. To select a specific ROI for the orientation analysis of TATS components, align the main longitudinal cell axis and the image x-axis in parallel. If the cell is slightly curved, select several ROIs and align each ROI individually. Exclude any nuclei from the analysis. Exclude any excessively curved cells from the analysis because precise alignment of ROIs will become increasingly difficult and increase orientation errors during analysis.
4. Delete any unwanted signal information from the outer surface membrane: Edit →Clear Outside to generate the “selected ROI” (Figure 5A). Ensure that the selected ROI only contains the intracellular membrane portions, which correspond with the TATS network.
5. Perform the following chain of image processing steps (commands) prior to the subsequent quantitative analysis as documented in Figure 5A.
   1. Click on →Process →Subtract Background. Set the Rolling ball radius to 5 pixels.
      NOTE: Set the rolling ball radius to 5 pixels if the image to be analyzed has a pixel size of 100 nm x 100 nm. For other pixel sizes set the rolling ball radius to the number of pixels which approximately correspond to a physical radius of 500 nm.
   2. Click on →Process →Enhance Local Contrast (CLAHE). Set the block size to 49, the histogram bins to 256, the maximum slope to 3, and the mask to “none”.
   3. Click on →Process →Smooth.
   4. Click on →Plugins →Segmentation →Statistical Region Merging. Set the parameters to Q100 →click on Show Averages.
   5. Confirm complete processing of the statistical region merging indicated by automatic presentation of a new image frame and confirm that the label “SRM Q=100” appears. Continue the following steps with this image file. Click on →Image →Type →8-bit.
   6. Click on →Image →Adjust →Threshold. Choose the threshold sufficiently low to detect most, ideally all TATS structures, in particular avoid exclusion of TATS components with a low signal intensity (use a threshold of 40 as starting point). Consult the “Representative Results” section for detailed data output and the examples in Figure 6 (upper threshold of 255). Document the final choice of threshold parameters. Note that the correct choice of threshold should produce only specific TATS membrane structures but not false positive signals due to background noise.
   7. Confirm correct superimposition of TATS image details versus extracted skeleton data, in particular for intermediate and high fluorescence signal levels, which should match the (dis-) continuity of the skeleton structure. Once an appropriate threshold has been identified, apply this same threshold to all images during analysis and repeat the overlay comparison of the original signal versus skeleton data to consistently minimize this potential source of bias.
   8. Click on →Apply: the image data becomes binary as shown in Figure 5 under “Threshold”.
   9. Click on →Plugins →Skeleton →Skeletonize (2D/3D). Save the skeletonized 2D image (as shown in Figure 5) as Tif file by clicking on →File →Save as →Tif. Analyze the skeletonized image file for quantitative data output by clicking on →Plugins →Analyze Skeleton (2D/3D). Choose Prune cycle method: none. Confirm automatic generation of the resulting data table.
   10. Save the automatically generated data table as txt file. Select the relevant quantitative parameters e.g., the total number of branch points or the average branch length as shown by exemplary data in Figure 7. Consider further data analysis with complementary/supporting software tools like Excel and as appropriate.
6. Consider to use automated image processing routines whenever possible including all necessary steps described under 4.5 to harmonize the analysis between individual images and/or image batches. Use the example Supplementary Code File containing an image processing Macro programmed for Fiji (adjust the programming as needed).
7. Analyze the individual orientations of all or select TATS network components from the skeletonized image data by the Fiji plugin “Directionality”. Generate a directionality histogram where the respective axially oriented A-tubule or transversally oriented T-tubule components are represented by the 0° or 90° bins. Note the correct reference of image orientation and that it is important for the x-axis of the image to correspond closely with the main (longitudinal) 0° axis of the VM cell as shown in Figure 8 and Representative Results.
   1. Click on →Analyze →Directionality →specify Method: Fourier components, Nbins 180, Histogram start -45 →click on Display table.
   2. Save the newly generated and displayed results table including associated histogram data as txt file. Consider further analysis of the txt file data by complimentary software tools like Excel and as needed.
8. To generate subclasses of data as grouped datasets e.g., for all cells treated under the same condition (and potentially other conditions), repeat the analysis steps 4.1-4.7 for all relevant images as appropriate. Import the skeletonized data parameters from all images into one combined Excel file in order to derive average values. In addition, import all directionality histogram data from the same grouped dataset into a combined Excel file to calculate and generate the average directionality histogram. Consider further processing of grouped datasets to analyze additional TATS network parameters of interest for individual or between different treatment groups as needed.