The following histological technique for preparing paraffin embedded and frozen sections of drosophila thoracic muscles allows a powerful approach to analysis of muscle morphology and protein expression. This is accomplished by first immobilizing animals in collars then embed the tissue. In paraffin or OCT medium, the embedded oph thax are sectioned by microtome.
Finally, sections are stained to visualize muscle tissue structures or detect different molecular components of the muscles. The resulting specimens when used in conjunction with classical histological staining methods, fluorescent dyes or immuno staining can show muscle tissue morphology and localization of muscle specific components. The main advantage of this technique is the ability to recover thin muscle sucs in a configuration that is found naturally within the animal.
This method can easily be adapted to other systems such as the Drosophila header abdomen. It was developed in the laboratory of Helena Sheta, the laboratory of gene expression and signaling, and will be demonstrated today by Maria Rinko. Keep all solutions for submerging collars in glass staining jars to start.
Prepare fresh car noise fixative by combining absolute ethanol chloroform and glacial acetic acid in the proportions of six to three to one respectively. Create a stack of aluminum foil, pockets of the correct size for the collars. Also prepare the ethanol, methylate, and paraffin solutions in staining jars.
Now place the methyl benzoate and paraffin containers in an incubator set to 60 to 65 degrees Celsius. In addition, warm more paraffin to 60 to 65 degrees Celsius. To pour into foil pockets, attach the collar under the microscope using tape in a vertical position so that the entry point for the fly is visible.
Anesthetize the flies either by using carbon dioxide or via hypothermia using an ice block. Be careful not to freeze the flies using forceps. Pick up individual flies by the wings and place each fly into the collar oriented, so its head and thorax are on top of the blades and its abdomen is below the blades.
Place 10 to 20 flies into each collar when analyzing several genotypes. Note the collar number and corresponding genotype. Transfer the collar to Carnoy solution and fix the tissue at four degrees Celsius overnight.
After fixation, dehydrate the sample by submerging the collar into increasing concentrations of ethanol baths for 10 minutes each at room temperature. Next, incubate the collar in methyl benzoate and methyl benzoate plus paraffin solution for 30 minutes each, and then infiltrate the collar in two changes of paraffin for 60 minutes each. At 60 to 65 degrees Celsius rapidly transfer the collar into a foil pocket.
At this point, the collar should be positioned in the foil pockets to facilitate the plant orientation of the sections longitudinal or transverse. Then fill the pockets with melted paraffin in order to allow the paraffin to solidify. Leave the foil pockets at room temperature overnight.
Unwrap the dry paraffin blocks with collars from the foil and gently separate the collar from the paraffin block with the help of a sharp later scalpel. Carefully carve out the extra paraffin from around the fly tissue on a rotation microtome. Cut the paraffin block with seven to 10 micron section steps.
Allow the cut tissue to float flat in a 30 degree Celsius water bath. Finally, place the section tissue on polar slides and dry overnight. These slides are now ready for staining with hematin and eosin.
An hour before starting the experiment, put the bottle of cryo embedding medium upside down in a four degrees Celsius fridge in order to cool it down and to minimize the formation of air bubbles. In addition, set up a freezing cooler of minus 60 degrees Celsius with the help of ethanol and dry ice. As with paraffin sections, immobilize the flies in a collar, transfer the collar with flies to a pre-cool foil pocket and rapidly fill with OCT the cryo embedding compound.
Let the OCT get solid. Carefully unwrap the formed block inside the cooler. Gently separate the collar from the embedding block and put it at minus 20 degrees Celsius for at least one day on a cryo microtome, cut the frozen muscles between minus 15 and minus 18 degrees Celsius with a section thickness of 10 to 15 microns.
Then place the samples on polarized slides. Store samples at minus 20 degrees Celsius until ready for further processing. Lipid dropouts can be detected with oil red o stain on cryo sections.
Using a protocol adopted from seber and tum, remember that immediately before staining, the frozen sections have to be fixed in 4%formaldehyde for 10 minutes after fixation. Wash the slides with water twice for five minutes each. Then equilibrate in propylene glycol for 10 minutes.
Now incubate for three hours in oil. Red o stain at room temperature. Then wash samples two times for five minutes in polypropylene glycol, followed by 30 minutes in PBS mount samples in 30%glycerol, hematin and eosin stain transverse in longitudinal sections of indirect flight muscles differentiate abnormal size and morphology from normal structured muscles.
Paraffin embedded sections can also be stained effectively with antibodies as indicated in this section of drosophila intestinal tract illuminated for the nuclear envelope marker. Lamin C and the nuclear stain dpi. Here, transverse sections of drosophila thax are stained with anti lamb C and DPI to show an enlarged view of normal and deteriorated muscles.
Similarly, longitudinal frozen sections of indirect flight muscles can be stained with anti distro, glycan muscle, sarcolemma marker, and DPI when stained with oil red O transverse frozen sections of drosophila thorax label lipid droplets. A combination of these techniques with other methods like HNE staining and immunohistochemical detection offers a powerful approach to determine tissue structure and protein localization in muscles.
