Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

We developed a method to reactivate quiescent neural stem cells in cultured drosophila brain explants. This method can supply exogenous factors to the culture media and can assay neuroblast reactivation. Better stem cell therapies could be developed by a better understanding of how quiescent stem cells respond to extrinsic cues and enter the cell cycle.

Demonstrating the procedure will be Susan Doyle, a research scientist from my laboratory. To begin carefully pick approximately 20 to 25 freshly hatched larvae from the grape agar plate under a dissecting microscope. Fill a Petri dish with about two milliliters of PBS and submerge the tool’s tip containing larvae in PBS for two minutes.

After two minutes, tip the dish at an angle to pool the liquid at the bottom, and using a small paint brush, brush the larvae out of the liquid up the bottom of the Petri dish. Collect all larvae on the paint brush and rinse the larvae briefly in 70%ethanol before transferring them back to the Petri dish containing PBS. Spray the work area, dissection tools, forceps, and two glass watch dishes with 70%ethanol and let them dry on the bench.

Make the supplemented Schneider’s culture medium, or SSM, and place it on ice. Pipette one milliliter of the medium into each glass watch dish. Using a micropipette with a sterile tip, transfer the freshly hatched larvae from the plate of PBS to the SSM in the first glass watch dish.

Dissect the brains out of the larvae placed in the second glass watch dish with SSM using forceps under a dissecting microscope and adjusting the magnification as needed. Use one forceps to grab the mouth hooks, and with the other, gently grab the body halfway down and pull in the opposite direction to split the larva into two pieces. After dissecting 15 to 20 brains, add one milliliter of SSM into one well of a sterile 24-well culture tray.

Using a micropipette and a sterile tip, transfer the freshly dissected brains into the SSM, followed by incubation of the media with brains for 24 hours at 25 degrees Celsius. Pipette 10 microliters of a 10-millimolar stock of 5-ethynyl-2’deoxyuridine, or EDU, with 990 microliters of SSM into a sterile microcentrifuge tube, mix, then pipette one milliliter of EDU SSM into one well of the sterile 12-well culture tray. Transfer the brains using a micropipette with a sterile tip from the well containing SSM to the new well containing the EDU SSM solution and incubate for one hour at 25 degrees Celsius.

Next, transfer the EDU-labeled brains to another well in the same culture tray containing one milliliter of fixative and allow the brains to be fixed for 20 minutes. After fixation, quickly transfer the brains to a 72-well minitray wells using a micropipette. Rinse the brains three times in 10 microliters of PBT and repeat three washes for 10 minutes each, ensuring the brains are always covered with some liquid.

Pipette 10 microliters of blocking solution onto the brains. Cover the tray and seal it with a strip of parafilm around the edge. The figure shows large-sized EDU-positive and deadpan-positive neuroblast cells after 24 hours of culturing in SSM with insulin and staining.

After 24 hours in supplemented Schneider’s media without insulin, the freshly hatched Oregon R wild type brains had no large-sized EDU-positive and deadpan-positive neuroblast cells, other than the four-mushroom-body neuroblast cells and one ventrolateral neuroblast cell. During confocal imaging, some brain hemispheres with damage, like small to large sized holes in the explanted brain tissue, were occasionally observed. Any brains with holes were not used for analysis.

Dissect tissues carefully and pipette gently to avoid any tissue damage. Also try to be as sterile as possible and not contaminate your cultures. Because the culture media can be easily manipulated by adding different factors, This technique can be used to address future hypotheses regarding extrinsic signaling and neuroblast quiescence entry and exit.