The overall goal of this method is to visualize the structure of the cardiac matrix in the absence of cells or other non-matrix materials. This method can answer key questions in the field of fibrosis, especially how differences in the content of the matrix can affect the organization of the matrix, and how these changes might affect different cardiomyopathies or other fibrotic illnesses. The main advantage of this technique is that it provides a qualitative view of the matrix that can be used to augment quantitative measures.

Dr.Galindo and Dr.Sawyer first had the idea to use this technique after they found that regular treatment of post-infarct swine had changes in gene expression that were similar to changes in matrix content after treatment. Individuals new to this method may struggle if they don't have previous SEM sample preparation experience, due to the fine motor skills required. The tissue sample should be large enough to be easily seen, and stored in four percent glutaraldehyde solution at four degrees Celsius.

Remove the tissue from cold storage and rinse it off in distilled water. Then, immerse the tissue in a solution of 10 percent sodium hydroxide, made fresh from pellets in water. Be sure to take proper precautions.

Keep the tissue in the sodium hydroxide solution for six to 10 days at room temperature. Proceed when the color of the tissue becomes off-white or white. Then, rinse the tissue off with distilled water until it becomes transparent.

Next, put on gloves and immerse the tissue in one percent tannic acid for four hours at room temperature. After the tannic acid bath, rinse the tissue in distilled water overnight. In preparation, in a fume hood wearing gloves, make a 0.2 molar stock solution of sodium cacodylate buffer and a two percent aqueous stock solution of osmium tetroxide.

Use a splash guard, as osmium tetroxide is a severe inhalation hazard. Now, make a 0.1 molar dilution of the sodium cacodylate buffer, and incubate the tissue in it for five minutes with gentle agitation. Change the buffer twice to wash the tissue a total of three times with sodium cacodylate.

Next, make a one-to-one mixture of the stock sodium cacodylate and stock osmium tetroxide. Incubate the tissue in the new mixture on a rotator for an hour. Later, return the tissue to 0.1 molar sodium cacodylate and incubate it with gentle agitation for five minutes.

Change this bath solution twice over 15 minutes. Then, dehydrate the tissue using a series of ethanol baths with increasing concentrations. Let the tissue soak for 15 minutes in each bath with gentle agitation.

Next, transfer tissue and solution to a shallow Petri dish filled with 100 percent ethanol. Next, with care, hold two very sharp razor blades such that the flats of the sides are in contact with one another, and the cutting edges cross to form two sides of an equilateral triangle above the specimen. Now, use your dominant hand to place the second blade on the far left of the sample, and slice rightward.

Thus, slide the blades against each other from opposite directions to make a single smooth cut with minimal distortion or tearing force. The goal is to expose as large a surface area as possible without damaging the specimen. Repeat the process until enough satisfactory cross-sections are prepared for examination by SEM.

Using a spatula or tweezers, transfer the tissue to the sample holder of the CPD, while keeping the tissue immersed in 100 percent ethanol. The tissue cannot be exposed to the air for more than a few seconds. Use the CPD to replace the ethanol with liquid carbon dioxide, and perform the drying cycle as indicated by the manufacturer.

Then, prepare an SEM sample stub for each specimen. Adhere a carbon adhesive tab to the top surface of the aluminum stub. Under a stereoscope, carefully adhere the specimen to the adhesive tab with the cross-section surface of interest facing up, away from the tab, and maximally planar with the stub surface.

Do not probe or touch the surface of interest. For the smallest, most difficult specimens, break the wooden applicator stick to use as a spatula. Also, work over a large piece of filter paper to avoid losing a dropped specimen.

For improved adherence, use a broken applicator stick to make a tapered brush, and apply silver or carbon paint to the base and sides of the specimen. Extend a very thin line of paint up to an edge of the surface of interest. This will make a charge path from the surface of interest to the ground.

Then, apply two or three small dabs of paint around the perimeter of the carbon tab to provide a conductive path from the tab's surface to the metal stub. This functions as the ground. Set the specimen aside and allow the conductive paint to dry for two hours.

Once dried, operate a sputter coater to apply a relatively heavy coat of gold palladium alloy or gold onto the specimen. A 30 to 40 nanometer thick coating is desired. Perform the scanning electron microscopy at a relatively low accelerating voltage to minimize problems associated with poor charge dissipation in the sample.

Use the variable pressure mode if it is available. First, improve the working distance to provide enough depth of field to focus on fibers in the Z dimension. To do so, open the Navigation tab at upper right.

Then, access the Coordinates tab from the stage menu. Therein, enter a larger value for the Z coordinate, such as 20 millimeters. Finally, press Go To tab to execute the change.

Next, position the surface of interest orthogonally to the electron beam. Click and hold the right mouse button, then slide it left or right to focus the specimen near the periphery of the prepared surface plane of interest. Take note of the focused working distance.

Then, using the manual user interface joystick, move the view to the opposing edge and repeat the focusing method. If the working distance has changed, tilt the specimen to achieve an approximate agreement at both locations. Use the Coordinates tab of the Stage menu and enter a tilt value in the T coordinate.

Now, rotate the specimen 90 degrees by entering this value into the R coordinate field. Then, repeat the tilting process until all the positions are focused at approximately the same working distance. The described technique was applied to cardiac tissues from an unused human heart transplant donor.

The human cardiac matrix is an intricate weave of cross-linked proteins that has a honeycomb-like pattern in a cross-section. Each honeycomb structure is approximately 40 microns wide, normally circumventing a single myocyte. When connected by intercalated discs, several cardio myocytes can be envisioned as a rod running lengthwise through a tunnel-like shape.

In addition to providing structural information, an SEM of decellularized tissues can allow meaningful qualitative assessment in response to injury or non-injurious forms of cardiomyopathy, such as infarcted cardiac tissues from a swine. In a separate study, the GGF2 isoform of NRG-1 beta was shown to be a potential therapeutic treatment for heart failure. After watching this video, you should have a good idea of how to process tissue, heart tissue, for the scanning electron microscope, including the technique for exposing the surface of interest and for mounting the specimen.

Following this procedure, other techniques such as mass spec, RNA sequencing, and various other assays can be used in order to measure extracellular matrix proteins and other responsible for signaling and other organizational observational phenomena. Also, don't forget that osmium tetroxide is a dangerous chemical, due to the potential for vapor fixation of the mucous membranes. Always remember to work in a fume hood with nitrile gloves, using a splash guard or hood sash.