The overall goal of this Drosophila brain dissection protocol is to obtain intact brains from adult flies that can be used in a wide range of applications, such as immunofluorescence, GFP tagged protein localization, and ex-vivo culturing experiments. This method can help us answer key questions in the fields of developmental biology and neuroscience, such as the roles of specific proteins and signaling pathways and establishing patterns of neuronal connectivity during brain development. The main advantage of this technique is that once mastered, it provides a rapid and effective method for immunologically identifying morphological defects in specific regions of the adult brain.
This procedure can be difficult to learn, because removing the exoskeleton from the fly head can take a great deal of care and practice. Your movements must be slow and deliberate, and the dissector's arms must be well supported in order to stay steady. It's a good idea to practice on red-eyed flies and then white-eyed flies before actually dissecting genotypes of interest for actual experiments.
To set up for the dissection, place the anesthetized flies of interest on a cold metal pad or in a Petri dish sitting on ice. Alternatively, use a fly pad that emits carbon dioxide. Next, place a small amount of PTN in the center of the dissection dish to create a bubble of PTN.
Then, under a stereo microscope, fill the field of view with the PTN bubble under uniform illumination. Now, manipulate flies so that they are belly-up, and transfer one by the abdomen into the PTN. Completely submerge the animal in the PTN, and perform the entire dissection submerged in PTN.
With a second pair of forceps, grasp the proboscis and pull the head off the body. Occasionally, the proboscis will be removed, but the head will remain attached to the body. If this occurs, grab the medial edge of one retina and apply lateral force to remove the head.
Discard the abdomen and thorax. It is critical to keep the head in the PTN during this step. Connections from the central brain to VNC clearly cannot be analyzed using this method.
Next, grasp the medial edge of the left retina at the edge of the central hole in the cuticle and slowly pull the forceps directly apart from each other to prevent tearing of the optic lobe, which is the opaque structure covered by white, stringy trachea. As the retina dissociates, there will be a slight decrease in tension. In order to prevent the brain from tearing, it is critically important to grasp the medial edge of each eye and very slowly pull the forceps apart.
Moving too quickly can result in altered brain morphology or damage to brain tissue. As shown here, grasp the cuticle with each pair of forceps and very slowly pull them in opposite directions. If done correctly, the cuticle should separate from the brain tissue, leaving the brain intact.
Very sharp forceps are needed for this step. Next, carefully remove the surrounding cuticle piece by piece. While removing stubbornly attached cuticle, it can help to secure the brain by gripping the remaining VNC to avoid crushing the brain.
Finally, use a P200 pipette to transfer the dissected brains into a well of a plate filled with PTN for fixation and immunostaining. Under the microscope, use a P200 pipette to transfer 10 to 15 dissected brains of the same genotype into a 0.5 milliliter micro-centrifuge 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 four 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 four degrees Celsius for two days with gentle agitation. Remove the primary antibody using the five 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 four degrees Celsius for several days, but ideally, proceed directly with mounting. To mount the brains, build a bridge slide.
Position two base cover slips roughly 1 centimeter apart on a positively charged slide. Make certain that the positively charged side is face up. Then, adhere the cover slips to the slide with fingernail polish, and let the polish dry completely before proceeding.
Next, place the slide under a stereo microscope, and pipette brains and medium into the space between the cover slips. 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 cover slip over the brains, and use fingernail polish to seal the edges of the top cover slip that are attached to the base cover slips. Now, load the center cavity with fresh mounting media drop-wise, allowing the media to be drawn under the cover slip by capillary action. When the cavity is filled, seal it in completely using clear fingernail polish.
The described method can be used to visualize almost any structure within the adult brain, including the mushroom bodies. This region of the brain is required for learning and memory. The mushroom bodies can be easily visualized using antibodies that recognize the fasciculin 2 protein.
This protein is highly expressed in the alpha and beta lobes, as well as the centrally located ellipsoid body. This technique has allowed for the identification of multiple cellular processes required for proper axon guidance of mushroom body neurons. Disruption of these processes causes a diverse array of mutant phenotypes, such as inappropriate axon projection across the brain mid-line region and missing alpha lobes.
These mutant phenotypes are often incompletely penetrant, and therefore, require the examination of multiple brains. In order to examine the autonomous role of a protein in axon pathfinding, the Markham technique can also be used to visualize axon guidance decisions of individual GFP positive mutant or wild-type neurons in a non-fluorescent wild-type background. The method described here allows for the reliable and reproducible visualization of virtually region of the adult Drosophila brain.
Here, we focused on the mushroom body neurons. In the text version of this protocol, we have also demonstrated visualization of the optic lobes. Other studies have used similar techniques to visualize the pars intercerebralis, clock neurons, and antennal lobe projection neurons, among many others.
Once mastered, each brain can be dissected in 3 to 5 minutes, if it is performed properly. While attempting this procedure, it's important to remember to stabilize your arms near the microscope. It's also important to move slowly, and to remove small pieces of cuticle one at a time.
Moving too rapidly could cause unwanted damage to underlying brain tissue. In addition to using dissected brains for immunofluorescence-based microscopy experiments, brain tissue can also be used for additional experiments, as well.
