Visualizing Mushroom Body Neurons in Drosophila Brains via Immunostaining

Place wild-type and mutant Drosophila brains in a solution containing a fixative and a detergent. Incubate with gentle rocking. 

The fixative preserves the tissue morphology, while the detergent permeabilizes cell membranes.

Allow the brains to settle, then remove the solution.

Wash the tissues with a buffer.

Add a blocking solution to prevent non-specific antibody binding.

Remove the solution and add primary antibodies targeting a specific protein that regulates axonal development in the brain's mushroom body, or MB, neurons.

Incubate to promote antibody binding.

Wash with buffer to remove unbound antibodies.

Incubate with fluorophore-conjugated secondary antibodies targeting the primary antibodies.

Wash again to remove unbound antibodies, then resuspend the brains in a mounting medium.

Mount the brains on a bridge slide and visualize them under a fluorescence microscope.

Compared to the wild-type, the mutant brains exhibit a defective phenotype, with axonal mis-projections and missing MB lobes.

Under the microscope, use a P200 pipette to transfer 10 to 15 dissected brains at the same genotype into a 0.5-milliliter microcentrifuge tube filled with 4% paraformaldehyde diluted in PTN. Then, incubate the brains for 20 minutes with slow rocking at room temperature. Following the fixation, allow brains to settle to the bottom of the tube, then remove the fixative.

Next, perform two quick washes with 500 microliters of PTN per wash. Exchange the PTN as soon as the brains settle in the tube. Next, perform three long washes with PTN using gentle agitation at room temperature.

Following these washes, the brains may be stored overnight at 4 degrees Celsius in PTN. After removing the last PTN wash, incubate the brains in 0.5 milliliters of blocking solution for at least 30 minutes at room temperature, and with gentle agitation. After removing the blocking solution, add the primary antibodies diluted in blocking solution, and incubate the brains at 4 degrees Celsius for two days with gentle agitation.

Remove the primary antibody using the 5-wash PTN washing regiment, and then apply the secondary antibodies. Allow these antibodies to incubate with the tissues for three hours at room temperature while shielded from light with rocking.

After incubating brains in the secondary antibody solution, apply the PTN wash regiment, covering the samples in foil after each wash, and finish by removing as much buffer as possible. Next, add 75 microliters of fluorescent anti-fade mounting medium and flow the brains and medium in and out of the pipette tip just once. The tube may be then wrapped in foil and stored at 4 degrees Celsius for several days, but ideally, proceed directly with mounting.

To mount the brains, build a bridge slide. Position two base coverslips roughly 1 centimeter apart on a positively-charged slide. Make certain that the positively-charged slide is face up. Then, adhere the coverslips to the slide with finger nail polish, and let the polish dry completely before proceeding.

Next, place the slide under a stereomicroscope, and pipette brains and medium into the space between the coverslips. Be sure to adjust the lighting to improve visualization of the brains.

Next, aspirate the extra mounting media from the slide being careful to avoid the brains. Then, wick away remaining excess mounting media. This will allow the brains to be positioned more precisely.

Now using forceps, orient the brains into a grid pattern with their antennal lobes facing up. Then, place a coverslip over the brains, and use finger nail polish to seal the edges of the top coverslip that are attached to the base coverslips. Now, load the center cavity with fresh mounting media dropwise, allowing the media to be drawn under the coverslip by capillary action. When the cavity is filled, seal it in completely using clear finger nail polish.