Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

1. Drosophila larvae collection

NOTE: Prepare the yeast plate, grape paste, and the Fly condo before starting:

1. Yeast paste: In a small container, mix 5 g of active dry yeast with 10 mL of water to form a paste that has the consistency of peanut butter. Cover the yeast paste with plastic wrap and use a rubber band to firmly attach it to the container.
   NOTE: Fresh yeast paste will expand in its container and will pop off the lid unless firmly attached. Yeast paste will last for several days at room temperature (RT).
2. Grape plates: Follow the recipe for making grape plates (Table 1). If using plates stored at 4°C, make sure to pre-warm plates before use by placing them at RT for 1 h.
   1. Mix water (750 mL) and agar (18.75 g) in a 4 L flask, swirl, and autoclave for 20 min (liquid cycle).
   2. Mix the grape juice (250 mL) and sucrose (25 g) in a 1 L flask with a large stir bar on a heated plate (low heat). When the sucrose is dissolved, turn off the heat, wait until the flask can be touched before adding Tegosept (10%, 4 mL) and Propionic acid (5 mL). Keep the stir bar on.
   3. When the autoclaving is complete, let it cool until the flask can be touched (~60 °C), then mix in the grape juice mix.
   4. Combine all solutions in one flask and let stir on the plate.
   5. Pipette the solution into lids of small-sized Petri dishes (35 mm). Pipette roughly 9 mL per lid or until a convex dome is obtained.
   6. OPTIONAL: Flame the lids to get rid of any bubbles.
   7. When the plates solidify, stack the grape plates into a box with an airtight lid and place the box at 4 °C. Plates can be stored for up to 1 month.
3. Fly condo: Punch ~20 holes into a 6-ounce square bottom polypropylene Drosophila bottle using an 18 G needle.
4. Transfer adult flies (~100 OregonR or any genotype) to a fly condo and cap the condo with a grape agar plate topped with a dab of yeast paste. Place the dab towards the center of the plate and affix the plate to the condo with lab tape.
5. Invert the container so that the grape agar plate is on the bottom and place it in a 25 °C incubator for 24 h (Figure 2).

Figure 2: Visual representation of inverted fly bottle (condo) with male and female Drosophila adults. The plastic bottle has small punctures, generated with an 18 G needle, for oxygen exchange. The mouth of the bottle is sealed with an agar grape juice cap and is inverted and stored in a 25 °C incubator. Please click here to view a larger version of this figure.

6. After 24 h, change the grape agar plate and replace it with a new plate topped with yeast paste. Quickly switch the two plates while continuously tapping the condo lightly on the bench so that the adult flies do not escape.
7. Examine the plate by eye and assess the number of embryos on the plate. Drosophila embryos are oblong and white with two string-like appendages.
8. If there are very few embryos on the plate (less than 100), discard the plate (scrape out the agar into the fly trash and save the plastic lid for reuse). In many cases, adult females will not lay many embryos on the first night in a new condo. If this is the case, give the adult flies another 24 h to acclimate.
9. If there is a large number of embryos on the plate (at least 100), keep it, and carefully remove the yeast paste using a flat bottom spatula.
10. Once the yeast paste is removed, use a metal pick to manually remove all larvae from the grape plate under a dissecting microscope. When looking at the plate under a dissecting microscope, crawling larvae should be observed as well as embryos.
11. Remove all larvae by brushing the metal pick towards the side of one larva. Larvae are sticky and will stick to the pick. Once one larva is on the pick, additional larvae can be easily picked up by using the larva on the tool to attach more.
    NOTE: Larvae like to stick to each other. At this point, it does not matter if larvae get damaged. These larvae will be discarded.
12. After picking and removing all larvae, place the plate back into the 25 °C incubator. Ensure to place the plate in a larger container that can be sealed. Place wet paper towels in the bottom of the larger container to keep in moisture.
13. After 30-60 min, take the plate back to the dissecting microscope and now, carefully pick ~20-25 larvae from the same grape agar plate to ensure that the picked larvae are freshly hatched within a 30-60 min window of time.
14. Submerge the tip of the tool with the 20-25 freshly hatched larvae into a Petri dish (60 mm) filled with 1-2 mL of 1x Phosphate Buffered Saline (PBS) for 2 min.
15. After 2 min, tip the dish at an angle to pool the liquid at the bottom. Using a small paintbrush, brush the larvae out of the liquid up the bottom of the Petri dish.
16. Collect all larvae on the paintbrush and transfer the larvae to a new Petri dish (60 mm) containing 1-2 mL of 70% ethanol. Repeat steps 1.15 to collect larvae with a paintbrush and transfer them to a new Petri dish with 1-2 mL of 1x PBS.

2. Culture media and tool preparation

1. Spray the bench and work area with 70% ethanol and let dry.
2. Spray the dissection tools, forceps, and two glass watch dishes, with 70% ethanol and let them dry on the bench.
3. Make the supplemented Schneider's media (SSM, Table 2) and place it on ice.
4. Pipette 1 mL of SSM into each of the glass watch dishes.
5. 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. Using a micropipette with a sterile tip, transfer the freshly hatched larvae to the SSM in the second glass watch dish.

3. Dissections and brain cultures

1. Once the larvae are in the second glass watch dish with SSM, dissect the brains out of the larvae using forceps and a dissecting microscope. Adjust the magnification as needed.
2. Use one forceps to grab the mouth hooks and with the other, gently grab the body halfway down and pull in the opposite direction (Figure 3) to split the larva into two pieces.
   NOTE: The brain will be located right behind the mouth hooks. Note that there may be other tissues surrounding the brain. Be very careful when removing these tissues as it can result in damaging the brain

Figure 3: Drosophila larvae in a glass watch dish with SSM. Forceps are properly positioned for dissection. The location of the larval brain (dark grey) is posterior to the mouth hooks (black), and both are shown inside the larva. Please click here to view a larger version of this figure.

3. Once 15-20 brains have been dissected, add 1 mL of SSM into one well of a sterile 12-well culture tray. Transfer the freshly dissected brains into the SSM using a micropipette and a sterile tip (Figure 4A).
4. Place the brains in the SSM media in the 12-well culture tray into an incubator at 25 °C for 24 h (Figure 4A).

Figure 4: Brain culture and immunostaining. (A) Whole brains in a 12-well culture dish containing 1 mL of SSM. The culture dish is then placed in a 25 °C incubator for 24 h. (B) 72-well mini tray that holds brain explants during immunostaining. Brains are washed and solutions transferred using a P20 micropipette set to 10 μL. Please click here to view a larger version of this figure.

4. Proliferation assay, brain fixation, and antibody staining

1. The following day, make 1 mL of EdU SSM solution. Pipette 10 μL of a 10 mM stock of 5-Ethynyl-2′-deoxyuridine (EdU) with 990 μL of SSM (final EdU concentration equals 0.1 mM) into a sterile microcentrifuge tube and mix. After the 24 h incubation is complete, pipette the 1 mL of EdU SSM into one well of the sterile 12-well culture tray.
2. Transfer the brains using a micropipette with a sterile tip from the well containing SSM only to the new well containing the EdU SSM solution. Incubate for 1 h at 25 °C.
3. Next, transfer the EdU-labeled brains to another well in the same culture tray containing 1 mL of fixative (4% paraformaldehyde, see Table 3 for recipe) for 20 min.
   CAUTION: Paraformaldehyde is a biological hazard and must be disposed of properly.
4. After fixation, quickly transfer the brains to the wells of a 72-well mini tray using a micropipette. Each well can hold 10 brains and 10-15 µL of liquid (Figure 4B). Once the brains are transferred to the mini tray (no more than 10 brains per well), remove the fix and rinse the brains 3 times in 10 µL of 1x PBT (phosphate buffer, pH 7.4 containing 0.1% Triton-X 100).
   NOTE: Rinsing means pipetting 10 µL of 1x PBT on the brains, removing, and repeating 3 times.
5. Next, wash the brains 3 times for 10 min each, again in 10 µL 1x PBT. Ensure that the brains are covered with some liquid at all times.
6. After the washes are complete, pipette 10 µL of blocking solution (1x PBT with 10% normal goat serum) onto the brains. Cover the tray and seal it using a strip of parafilm around the edge.
7. Once sealed, place the mini tray into a sealed box with wet towels to provide a moist environment to prevent evaporation. Place the box containing the tray at 4 °C overnight.
8. The following day, make a primary antibody solution.
   NOTE: In this protocol, rabbit anti-scribble was used to label cell membranes and rat anti-deadpan to label neuroblasts, although any number of other primary antibodies could be used.
   1. To make the primary antibody solution, first, make dilutions of primary antibodies in the blocking solution. For example, rabbit anti-scribble is used at a final concentration of 1:1000. Therefore, first, dilute the rabbit anti-scribble antibody at 1:100 (1 µL of antibody plus 99 µL of blocking solution). Rat-deadpan is used at a final concentration of 1:100. Therefore, first, dilute the rat-deadpan antibody at 1:10 (1 µL of antibody plus 9 µL of blocking solution).
      NOTE: These dilutions can be stored long term at 4 °C if sodium azide (0.05%) is also added to inhibit bacterial growth.
   2. Next, count the number of wells that contain brains. The number of wells determines the volume of primary antibody solution to make. For example, if there are 2 wells of brains, prepare 20 µL of primary antibody solution (for 10 wells, 100 µL, etc.). To make a 20 µL primary antibody solution, add 2 µL of each primary antibody dilution and 16 µL of the blocking solution.
      NOTE: The final concentration of each of the primary antibodies is 1:1000 and 1:100, respectively. In short, make the first dilution of primary antibodies at a concentration so that the second dilution is always 1:10 to arrive at the respective final concentrations. In this case, 1:1000 for rabbit anti-scribble and 1:100 for rat anti-deadpan.
9. Remove the blocking solution with the micropipette set to 10 µL and pipette 10 µL of primary antibody solution into each well.
10. Cover and seal the tray using parafilm and place it back into the sealed box with wet towels. Incubate overnight at 4 °C.
    NOTE: Shaking is not required and is strongly discouraged. Antibodies will penetrate the brains without shaking or mixing.
11. The following day, remove the primary antibody solution using a micropipette and rinse the brains 3 times with 10 µL of 1x PBT. Next, wash the brains 4 times with 10 µL of 1x PBT for 10 min each. During the 10 min washes, prepare the secondary antibody solution.
12. To make the secondary antibody solution, choose secondary antibodies that recognize the primary antibodies. In this protocol, goat anti-rabbit Alexa Fluor 488 and goat anti-rat Alexa 555 were used.
    1. Pipette 1 µL of each of the secondary antibodies into a microcentrifuge tube with 298 µL of the blocking solution to make the final concentration 1:300 for each secondary antibody.
13. After the last 10 min wash, remove the 1x PBT and pipette 10 µL of the secondary antibody solution into each well. Seal the tray using parafilm and place it back into the box with moist towels. Incubate overnight at 4 °C.
    NOTE: Do not worry about removing every last µL in the wells between rinses, washes, or when adding primary and secondary antibody solutions. The brains will always remain submerged in a few µL of liquid, which is just fine.
14. The following day, remove the secondary antibody solution using a micropipette and rinse the brains 3 times with 10 µL each of 1x PBT. Next, wash the brains 4 times with 10 µL each of 1x PBT for 10 min each.
15. During the 10 min washes, prepare the EdU reaction mix to detect EdU incorporation. Prepare the EdU reaction mix according to the manufacturers' guidelines.
16. After the final wash, remove the 1x PBT and pipette 10 µL of the EdU reaction mix into each well with brains. Seal the microwell plate with parafilm and cover with aluminum foil. Leave plate on the bench for 30 min.
17. After 30 min, rinse the brains 3 times with 10 µL each of 1x PBT and wash the brains 3 times with 10 µL each of 1x PBT for 5 min each.
18. After the last wash, remove the 1x PBT and pipette 10 µL of a glycerol-based mounting media solution. Seal the plate and place at 4 °C overnight.

5. Mounting and imaging the brains

1. The following day, prepare microscope slides (25 mm x 75 mm x 1 mm): Glue (e.g., superglue) one 22 mm x 22 mm x 1 mm square cover glass to each end of the microscope slide to create a 'bridge' over which the larger 22 mm x 50 mm x 1 mm coverslip will be placed to make a space between the slide and the larger coverslip (Figure 5A). This space will allow the brain just enough movement to be correctly oriented while preventing it from being crushed.

Figure 5: Schematic showing microscope slide, orientation, and cell types in the larval brain. (A) Visual representation of microscope slide upon which a larval brain is mounted and is ready to be imaged. (B) A guideline is also shown to use for tissue orientation. (C) Microscope slide ready for imaging on a confocal microscope. (D) Cartoon showing some of the cell types in the larval brain. Please click here to view a larger version of this figure.

2. After gluing the 22 mm x 22 mm x 1 mm cover glasses to the microscope slide, pipette 9.3 µL of the glycerol-based mounting media solution containing one brain from a well of the microwell plate and place it onto the center of the slide (Figure 5A).
   NOTE: Larval brains may adhere to the pipette tip, so be careful. To avoid adhesion, begin by aspirating the antifade and then aspirating the single brain towards the end of the 9.3 µL volume.
3. Once the brain is on the slide, gently place the 22 mm x 50 mm x 1 mm coverslip on top. Position the brain as seen in Figure 5B. Gently move the coverslip around to orient the brain. The sample is then ready for imaging.
4. Use a confocal microscope equipped with high magnification and a high numerical aperture objective (Figure 5C) to acquire the best images. For example: 60x or 63x, 1.4 NA oil-immersion lens.
   1. Image the brains with the dorsal surface closest to the coverslip (and objective). Acquire the Z stacks through the entire brain hemisphere starting at the ventral surface (furthest away from the objective) at 1 µm intervals or Z step size.
      NOTE: Lasers used depend on the secondary antibodies. In this protocol, the laser lines used were 488 nm to detect Scribble staining, 555 nm to detect Deadpan, and 633 nm to detect EdU.

6. Data analysis

1. Use Fiji open-source software to analyze brain hemispheres and use the Fiji cell counter plugin to count the cells.