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

The overall goal of this procedure is to analyze the tubular membrane networks in atrial and ventricular myocytes. This is accomplished by first isolating atrial and ventricular myocytes from an adult mouse heart. In the second step, the transverse axial tubule system or tats is stained in the intact living isolated cardiac myocytes.

Next, the tats membrane network is visualized by confocal or stead fluorescence microscopy, and then the components of the network are quantitatively analyzed. Ultimately, the differences between the membrane networks of different cell types or between sham and treatment groups can be revealed by the quantitative tats image analysis. The main advantage of this cell isolation and analysis technique is that a comprehensive workflow of the isolation process leads to direct fluorescence imaging of the myocytes, allowing quantitative data analysis of the touch membrane network.

This method can provide insight into the organization of complex membrane networks in living healthy cardiomyocytes, but it can also be applied to myocytes from diseased or transgenic animals. Newcomers to this protocol will benefit from additional introductory training to achieve consistent isolation of high quality cardiac myocytes and important prerequisite for performing the technique To isolate the myocytes begin by perfusing the heart of a 12 week or older mouse with 37 degrees Celsius oxygenated perfusion buffer for four minutes as soon after the organ extraction is possible. Next, peruse the heart with 37 degrees Celsius digestion buffer for eight to 10 minutes, monitoring the progress of the tissue digestion for increasing opaqueness, softness and acidity throughout the apparent heart surface.

After the digestion, deflect the right atrial appendage and dissect the right atrium just above the atrial ventricular valves. Continue the dissection with the left atrium and then dissect and discard the fibrous valve apparatus. Finish the dissection by freeing the left and right free ventricular walls and the septum and smaller tissue parts as needed.

Next, transfer the entire ventricular tissue to a 60 millimeter Petri dish containing 2.5 milliliters of digestion buffer, and then cut the tissue into approximately one millimeter cubed pieces. Slowly ate the tissue pieces with a transfer pipette to gently dissociate the ventricular myocytes into a single cell suspension, taking care to avoid bubbles or fluid jet damage. Then add 7.5 milliliters of stop buffer to the cell suspension.

In parallel, cut the digested atrial tissue into approximately one millimeter cubed pieces in one milliliter of digestion, buffer in a 60 millimeter Petri dish, and then gently tritrate the atrial myocytes as just demonstrated. Following the mechanical agitation, add four milliliters of stop buffer to arrest the residual collagenase activity in the atrial myocyte suspension. Then transfer the cell suspension to a 15 milliliter conical tube and allow the remaining tissue pieces to sediment for 15 seconds prior to harvesting the isolated myocytes with the supinate fraction.

After one washing step, gently Resus suspend the atrial myocytes in five milliliters of perfusion buffer and divide the cell suspension into 1.5 milliliter micro centrifuge tubes in approximately one times 10 to the three cell aliquots to stain the tat membranes of the ventricular myocytes. First, let the cells settle by gravity for eight minutes and a 1.5 milliliter reaction tube to stain in the atrial myocytes send, refuse the cells for two minutes at 20 times G at room temperature for either cell type. Next, carefully remove the supinate taking care to avoid any unnecessary agitation of the cell pellet.

Then gently resuspend the cells in 800 microliters of the membrane specific tyle dye dye eight and eps immediately transfer the myocytes onto laminin coated cover slips in an imaging chamber after a 15 minute incubation in the dark wash. The cover slips once and then cover them with one milliliter of perfusion buffer. Take a sample image of a central intracellular myocyte section and then use the crop function to adjust the crop window and the final pixel size to 100 by 100 nanometers.

To select the final imaging plane, use single image frames to manually choose the most appropriate imaging plane in the Z direction. Confirm that the TAs membrane network includes both t tubial and a tubial components and is visually apparent in the focal plane. Note that a typical intracellular imaging plane may include a nucleus as a central intracellular reference point.

To analyze the tats membrane network and its components, now open the image file using the appropriate image analysis software. Use the polygon selection tool to delineate the region of interest while excluding the outer surface membrane and including intracellular portions of the tats membrane network as demonstrated in this image. Next, use the edit and clear outside commands to generate the selected ROI such that the selected intracellular area contains only the intracellular membrane portions of the tats network, but not any portions of the surface membrane.

Now click on process and subtract background and set the rolling ball radius to five pixels. Then select process and enhance local contrast. And set the block size to 49, the histogram bins to 256, the maximum slope to three and the mask to none.

Click process and smooth under plugins, select segmentation and statistical region merging and set the parameter to Q 100. Then click on show averages. Next, select on image type and eight bit, and then select image adjust and threshold, starting with a threshold of 40.

Note that the correct choice of threshold should produce only specific tats membrane structures and not false positive signals due to background noise. Then under plugins, select skeleton and skeletonized 2D 3D and save the skeletonized 2D image as a adoptive file. Now select plugins again and analyze skeleton 2D 3D to analyze the skeletonized image file for quantitative data output.

Then select none for the prune cycle method and confirm the automatic generation of the resulting data table. Now use the Fiji plugin directionality to analyze the individual orientations of the appropriate tax network components from the skeletonized image data. Under method, choose FURIER components, end bins 180 and a histogram start of minus 45 degrees.

Finally, check the display table and generate a directionality histogram. In this first figure, a representative skeletonized image of the tats in a healthy ventricular myocyte composed of transverse elements that are perpendicular to the X axis of the image and of axial components that are in parallel to the x axis is shown application of the directionality plugin results in an orientation histogram for the ventricular tats with comparable peaks at zero degrees, representing the axial proportion and at 90 degrees representing the transverse. In contrast, a representative skeletonized image of the tats in an atrial myocyte is primarily composed of axial components that are parallel to the X axis of the image.

Thus, the corresponding orientation histogram exhibits a dominant peak around the zero degree bin representing the axial tats proportion and only a small peak at 90 degrees, representing the transverse tats proportion While attempting this procedure, it's important to check for the cell quality in every step after the cell isolation, during imaging, and most importantly, just prior to the image analysis. After watching this video, you should have a good understanding of how complex membrane structures, tes networks, and potentially membrane associated proteins are fully differentiated. Adult cardiac myocytes can be imaged by confocal microscopy and quantitatively analyzed.

Please don't forget that working with lipophilic membrane eyes, digestion enzymes, and potentially infectious cardiac patient, em can be a hazard, and that precautions such as iron skin protection should always be taken while performing these procedures.