So my research entails exploring glial roles in experience-dependent critical period synapse remodeling. I'm determining the role of glia in synaptic pruning, specifically during critical periods, and the neuron-to-glia communication enabling pruning driven by experience. Our protocol has the proven power of Drosophila genetics to define molecular mechanisms.
To study gene-environment interactions, we can precisely manipulate the odor concentration, duration, and timing in a short, well-defined critical period. This protocol is incredibly versatile in inducing synapse remodeling. Our results established an excellent model amenable to genetic and pharmaceutical manipulations that could potentially reopen critical period-like plasticity during adulthood, which is a topic of enormous interest for the treatment of injury, trauma, and neurological disorders.
To begin, take the vial the Drosophila larvae. Using a fine paintbrush, sort 40 to 50 very dark pupae in the polystyrene Drosophila vials containing standard cornmeal molasses food. Place fine stainless-steel wire mesh over the end of the Drosophila vials to contain the flies while allowing good airflow.
Secure the wire-mesh caps with a tape transparent film onto the side of the vials. Then add one milliliter of 100%mineral oil or dissolved EB in mineral oil in 1.5-milliliter tubes. Place these tubes upright within a 3, 700-milliliter Glasslock container.
Using tape, anchor the tubes in the center of the odorant chambers along with the Drosophila vials. Place the sealed vehicle control and EB odorant chambers containing the pupae vials in a humidified incubator on a 12-hour light and 12-hour dark cycle. After four hours of odorant chamber exposure, rapidly transfer 20 to 25 newly enclosed flies to fresh Drosophila vials in chambers with freshly made odorant.
Keep the flies in their sealed odorant chambers in the incubators for a total of 24 hours. To begin, take Drosophila flies exposed to mineral oil and EB for 24 hours. Label vials as a vehicle control or EB exposed and maintain these designations throughout brain dissection and processing.
Using forceps, transfer a single anesthetized fly into a small dish of freshly made PBS. Fully submerge the fly in PBS for full brain dissection. Then position the fly ventral side up.
Take two fine sharpened forceps to grasp the upper thorax with one forcep and the head under the proboscis with the other. Pull the head and the rest of the body in opposite directions to remove the head. Ensure the head easily detaches from the thorax and leaves the isolated head for dissection.
Slide the forceps previously used to grasp the thorax under the opposite side of the proboscis. Gently pull the exoskeleton cuticle in opposite directions to tear it between the eyes and reveal the brain. Continue to remove the exoskeleton cuticle from the head, ensuring all parts are removed.
Now rinse a P20 pipette tip with PBS and 0.2%Triton X-100. Transfer the brains into the fixing solution in the capped tube immediately after dissection. To begin, take a slide and add two thin strips of double-sided adhesive tape to it to prepare for imaging.
Pipette approximately 10 to 15 microliters of mounting medium between the two strips of tape for brain mounting. Transfer the labeled Drosophila brains onto the microscope slide with a P20 pipette tip pre-rinsed with PBS and 0.2%Triton X-100. Once the brains are transferred, use a fine paintbrush to align them, ensuring the antenna lobes face upwards.
After orienting the brains, cover them with a glass coverslip. Then fill in the sides of the coverslip with additional mounting medium. Seal the edges of the coverslip with clear nail polish.
After drying the slide thoroughly, store it in the refrigerator for subsequent imaging. Use a laser scanning confocal microscope equipped with a 63x oil-immersion objective for imaging. Employ an argon 488 and helium-neon 543 laser for imaging the antenna lobe synaptic glomeruli and Or42a olfactory sensory neurons.
Determine the optimal gain and offset for both channels. Position the imaging to the center of the brain and focus on the VM7 glomeruli located approximately to the hole in the middle of the brain. Select the imaging resolution of 1024-by-1024.
Capture an entire confocal Z-stack projection through the antenna lobe to ensure the full Or42a neuron innervation of the VM7 glomeruli is captured. Load the genotype or condition-blinded image into Fiji. To split the laser line channels, go to image, click Color, and select Split Channels.
In the Or42a olfactory sensory channel, scroll through the Z-stack to determine which slices contain the Or42a neuron innervation, identifying the beginning and end of fluorescence. To create a sum slices projection that includes only slices with Or42a neuron innervation, click Image and go to Stacks. Then click Z Project, select some slices, and enter the desired range.
Use the lasso tool in Fiji to trace the outline of the Or42a neuron innervation in the VM7 glomerulus. Multiply the circumference of the traced area by the number of Z-stack slices and the thickness of each slice to calculate the VM7 synaptic glomerulus innervation volume. After 24-hour exposure to a control odor and vehicle, dense Or42a mCD8-GFP innervation persists in VM7 glomeruli of both antenna lobes.
In contrast, a 24-hour exposure to 25%EB odorant causes significant pruning and loss of synaptic glomerular volume. Increasing EB odorant concentration causes progressively greater synaptic glomeruli pruning. Representative quantifications for this range of EB odorant concentrations showed similar results for fluorescence intensity and innervation volume.
