Transmission Electron Microscopy to Quantify Glycogen Distribution in Human Skeletal Muscles

Isolate a small specimen from the muscle biopsy or whole muscle, which has a maximum diameter of 1 millimeter in any direction, and is longer longitudinally than cross-sectionally.

Place the specimen in the tube containing the cold primary fixation solution. Store it at 5 degrees Celsius for 24 hours. Then, wash the specimen 4 times in 0.1 molar sodium cacodylate buffer. Using transfer pipettes, remove the used buffer from the tube, leaving the specimen untouched, and add the fresh buffer. Post-fix the specimen with 1% osmium tetroxide, and 1.5% potassium ferrocyanide in 0.1 molar sodium cacodylate buffer for 120 minutes at 4 degrees Celsius.

Rinse the specimen twice in double-distilled water at room temperature. Dehydrate by submerging in a graded series of ethanol at room temperature. Infiltrate the specimen with propylene oxide and epossidic resin-graded mixtures at room temperature using the mentioned volume ratios.

The following day, embed specimens in 100% fresh epossidic resin in molds, and polymerize at 60 degrees Celsius for 48 hours.

Mount the block of a specimen on the ultramicrotome holder. Trim the block on the surface with a razor blade to reach the tissue level. Mount a diamond knife in front of the sample, and align the sample surface parallel to the knife.

Produce a semi-thin section of 1-micrometer thickness with the diamond knife to check the orientation of the sample. Stain the semi-thin section with toluidine blue for observation with light microscopy. Then, trim the block further to reduce the area of interest to get proper ultra-thin sections.

Cut 60 to 70-nanometer thickness ultra-thin sections with a second diamond knife. Collect 1 to 2 sections on one-hole copper grids using a Perfect Loop. For contrasting the sections, immerse the grids in uranyl acetate solution for 20 minutes, and wash the grids in double-distilled water, and then, immerse the grids in lead citrate for 15 minutes, and re-wash the grids in double-distilled water.

Turn on the transmission electron microscope, computer, and image recording software. Record digital images with a digital slow-scan CCD camera, and the associated imaging software. After inserting the grid with multiple sections in the microscope stage, screen the grid initially at low magnification to choose the best quality sections, and determine the direction of the muscle fibers.

Next, focus the image at magnification above 30,000, with the beam centered on a peripheral fiber in the section, and record images with 1 second exposure time at the desired magnification. Ensure that the images are distributed across the length and width of the fiber in a randomized, but systematic order to obtain unbiased results, and repeat the imaging until 6 to 10 fibers are imaged.

Import images to imageJ by clicking on File, followed by Open. Set global scale to match the original size of the image by clicking on Analyze, and then, Set Scale. To zoom in 100%, click on Image, go to Zoom, and click on In.

Measure the thickness of one Z-disc per image of the myofibrillar space, using the Straight Line tool from the Tools menu, and calculate the average Z-disc thickness of each of the 6 to 10 fibers. Use the Segmented Line tool to measure the length of the outermost myofibril, visible just below the subsarcolemmal region.

To insert a grid, click on Analyze, select Tools, and then, select Grid. Now set the Area Per Point at 32,400 square nanometers. Count the number of hits within the available length, in the 12 subsarcolemmal images where a cross hits the subsarcolemmal glycogen.

Insert a grid by clicking on Analyze, then, select Tools, followed by Grid, and set Area Per Point at 160,000 square nanometers. Count the number of hits in the 12 myofibrillar images, where a cross hits the intramyofibrillar space.

Then again, to insert a grid, click on Analyze, select Tools, and, then, select Grid. Now, set the Area Per Point at 3,600 square nanometers. Count the number of hits in the 12 myofibrillar images where a cross hits the intramyofibrillar glycogen.

Insert a grid by clicking on Analyze, then, select Tools, followed by Grid, and set Area Per Point at 32,400 square nanometers. Count the number of hits in the 12 myofibrillar images where a cross hits the intermyofibrillar glycogen.

Using the 32,400 square nanometers grid to randomly choose the glycogen particles, start with the upper left square and move left if necessary. Using Straight Line tool, measure the diameter of 5 randomly-chosen glycogen particles of each pool for each of the 12 images to obtain an average of 60 particles per pool per fiber.