Direct Cryosectioning of Drosophila Heads for Enhanced Brain Fluorescence Staining and Immunostaining

Our team studies metabolic, cardiac, muscle sleep and aging disorder using Drosophila model. This Jove paper detailed simplified protocol for brain tissue processing, including decapitation, fixation, cryosectioning, staining, and imaging. My research group has pioneered the development of Drosophila model to study several human disease and aging.

We have also investigated intervention like time restricted feeding and exercise. We use machine learning, omics and molecular approach to study factor like circadian rhythm and genetics to reveal their impact on cellular integrity, physiology, and behavior. This simplified Drosophila brain research protocol avoids complex dissections, requires only one handed execution, and eliminates the need for costly confocal microscopy, increasing accessibility and reducing equipment dependency.

To begin, obtain the flies in aging vials, then open the valve on the carbon dioxide mat, and quickly dump the flies onto the mat to prevent escape. Once the flies become mostly unconscious, position the flies underneath the SZ61 microscope by moving the mat under the objective lens. Adjust the magnification and focus until the flies are clearly visible and comfortable to decapitate.

Using spring scissors, position the blades between the thorax and head of the fly and squeeze firmly to decapitate five to 10 fly heads per experimental group. Place any and used flies back into their aging vial. Using a brush, gently transfer the decapitated fly heads into labeled 1.5 milliliter tubes placed on ice.

After removing the tubes from the ice, pipette 100 microliters of 4%paraformaldehyde and PBS into each tube ensuring all fly heads are fully submerged. If heads adhere to the tube walls, gently push them downward using a brush or tap the tube lightly against a horizontal surface to ensure proper contact with the solution. Then incubate the tubes for 15 minutes on an orbital shaker at a medium setting.

Discard the paraformaldehyde solution and replace it with PBS, ensuring all fly heads are submerged. After the final wash, transfer the fly heads into a 10%sucrose solution in PBS, ensuring they're submerged within the tube. Fill a labeled mold to approximately 50%capacity with optimal cutting temperature or OCT compound, allowing it to spread to all four corners of the mold.

Using a brush, carefully place the collected fixed fly heads onto the surface of the OCT compound within the mold. Using the tip of forceps, gently push each head to the bottom of the mold, ensuring the eyes are oriented downward for a cut through the coronal plane. Carefully align all heads in the X, Y, and Z dimensions to ensure that all sections will contain all experimental groups simultaneously.

After aligning the heads properly, carefully placed the molds directly into a freezer set at minus 20 degrees Celsius to freeze. Once the mold has mostly frozen, fill the remaining space with OCT compound. For mold cryosectioning, apply a generous amount of OCT compound to the top of the mold to attach the chuck bit to the mold.

Place the bit flat on top and let it freeze completely inside the cryostat, which typically takes five minutes. Then release the bit and the OCT block from the mold. Place the block into the chuck, ensuring the proper top to bottom orientation is maintained and tighten the chuck key until the block is securely in place.

Using the adjustment knobs and chuck depth controls, align the block with the blade. Set the section width on the cryostat to 20 micrometers and cut each slice using a slow consistent motion while allowing the anti-roll glass to capture each slice. Then capture the sections using warm slides by touching the slide to the closer edge of the section and allowing the section to rise onto the slide.

Allow the slides to dry at room temperature for at least 30 minutes, but no more than one hour. Take the cryosectioned Drosophila fly heads and use a razor blade to remove any residual OCT compound on the edges of the slide, leaving space for a hydrophobic border. Then use a hydrophobic marker to draw a border around each slide and allow it to dry for five minutes.

Wash all slides three times for five minutes each using PBS by gently pipetting on the slide. After washing the slides, pipette 3%BSA in Tris Buffered Saline blocking solution onto the slides and incubate for 30 minutes to one hour. Then discard the blocking solution and pipette the primary antibody solution onto the slides.

Incubate the antibody overnight at four degrees Celsius or for one hour at room temperature using wet tissue wipes to prevent drying. Discard the primary antibody solution and wash the slides three times for five minutes each using PBS. Then add the secondary antibody solution to the slides and incubate for one hour at room temperature.

After washing the slides three times with PBS, leave only a small amount of PBS on the slide. Next, using a 1000 microliter pipette, add three to five drops of hardening mounting media containing DAPI evenly across the slide. Place a cover slip onto the slide ensuring no air bubbles form during placement.

Carefully handle the freshly mounted slides and store them flat until the mounting media is completely dry. If long-term storage is needed, seal the edges of the slides. Capture images of the slides as soon as possible to minimize fluorescent decay.

During image acquisition, focus on each unique subject in the same channel for consistency across all experimental groups. Calibrate exposures for each channel to avoid overexposure in all selected channels. Ensure that the exposures selected remain constant and are appropriate for all subjects, especially between different experimental groups.

Expression of the ApoE antibody tag was clearly localized in brain regions of Elav+and Elav-ApoE4 specimens. Lipid accumulation was indicated by Nile red staining, which was visible in the brain regions in both genotypes.