We acknowledge the LSAMP Bridges to Doctorate program for funding (CNK) as well as NIH/NIGMS (R01-GM120421 and R35-GM141886). We are grateful to Dr. Conor Sipe for Figure 1. We also thank all Siegrist lab members for their continued support and mentorship. We especially thank Chhavi Sood and Gary Teeters for their careful reading of the manuscript and for providing comments.

1. Drosophila larvae collection
NOTE: Prepare the yeast plate, grape paste, and the Fly condo before starting:
<ol>
	<li>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).</li>
	<li>Grape plates: Follow the recipe for making grape plates (Table 1). If using plates stored at 4&#176;C, make sure to pre-warm plates before use by placing them at RT for 1 h.
	<ol>
		<li>Mix water (750 mL) and agar (18.75 g) in a 4 L flask, swirl, and autoclave for 20 min (liquid cycle).</li>
		<li>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.</li>
		<li>When the autoclaving is complete, let it cool until the flask can be touched (~60 &#176;C), then mix in the grape juice mix.</li>
		<li>Combine all solutions in one flask and let stir on the plate.</li>
		<li>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.</li>
		<li>OPTIONAL: Flame the lids to get rid of any bubbles.</li>
		<li>When the plates solidify, stack the grape plates into a box with an airtight lid and place the box at 4 &#176;C. Plates can be stored for up to 1 month.</li>
	</ol>
	</li>
	<li>Fly condo: Punch ~20 holes into a 6-ounce square bottom polypropylene Drosophila bottle using an 18 G needle.</li>
	<li>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.</li>
	<li>Invert the container so that the grape agar plate is on the bottom and place it in a 25 &#176;C incubator for 24 h (Figure 2).</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63189/63189fig02.jpg" /> 
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 &#176;C incubator. <a href="https://www.jove.com/files/ftp_upload/63189/63189fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="6">
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>After picking and removing all larvae, place the plate back into the 25 &#176;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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
2. Culture media and tool preparation
<ol>
	<li>Spray the bench and work area with 70% ethanol and let dry.</li>
	<li>Spray the dissection tools, forceps, and two glass watch dishes, with 70% ethanol and let them dry on the bench.</li>
	<li>Make the supplemented Schneider&#39;s media (SSM, Table 2) and place it on ice.</li>
	<li>Pipette 1 mL of SSM into each of the glass watch dishes.</li>
	<li>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.</li>
</ol>
3. Dissections and brain cultures
<ol>
	<li>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.</li>
	<li>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&#160;removing these tissues as it can result in damaging the brain</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63189/63189fig03.jpg" /> 
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. <a href="https://www.jove.com/files/ftp_upload/63189/63189fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="3">
	<li>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).</li>
	<li>Place the brains in the SSM media in the 12-well culture tray into an incubator at 25 &#176;C for 24 h (Figure 4A).</li>
</ol>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63189/63189fig04.jpg" /> 
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 &#176;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 &#956;L. <a href="https://www.jove.com/files/ftp_upload/63189/63189fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Proliferation assay, brain fixation, and antibody staining
<ol>
	<li>The following day, make 1 mL of EdU SSM solution. Pipette 10 &#956;L of a 10 mM stock of 5-Ethynyl-2&#8242;-deoxyuridine (EdU) with 990 &#956;L 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.</li>
	<li>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 &#176;C.</li>
	<li>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.</li>
	<li>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 &#181;L 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 &#181;L of 1x PBT (phosphate buffer, pH 7.4 containing 0.1% Triton-X 100). 
	NOTE: Rinsing means pipetting 10 &#181;L of 1x PBT on the brains, removing, and repeating 3 times.</li>
	<li>Next, wash the brains 3 times for 10 min each, again in 10 &#181;L 1x PBT. Ensure that the brains are covered with some liquid at all times.</li>
	<li>After the washes are complete, pipette 10 &#181;L 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.</li>
	<li>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 &#176;C overnight.</li>
	<li>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.
	<ol>
		<li>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 &#181;L of antibody plus 99 &#181;L 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 &#181;L of antibody plus 9 &#181;L of blocking solution). 
		NOTE: These dilutions can be stored long term at 4 &#176;C if sodium azide (0.05%) is also added to inhibit bacterial growth.</li>
		<li>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 &#181;L of primary antibody solution (for 10 wells, 100 &#181;L, etc.). To make a 20 &#181;L primary antibody solution, add 2 &#181;L of each primary antibody dilution and 16 &#181;L 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.</li>
	</ol>
	</li>
	<li>Remove the blocking solution with the micropipette set to 10 &#181;L and pipette 10 &#181;L of primary antibody solution into each well.</li>
	<li>Cover and seal the tray using parafilm and place it back into the sealed box with wet towels. Incubate overnight at 4 &#176;C. 
	NOTE: Shaking is not required and is strongly discouraged. Antibodies will penetrate the brains without shaking or mixing.</li>
	<li>The following day, remove the primary antibody solution using a micropipette and rinse the brains 3 times with 10 &#181;L of 1x PBT. Next, wash the brains 4 times with 10 &#181;L of 1x PBT for 10 min each. During the 10 min washes, prepare the secondary antibody solution.</li>
	<li>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.
	<ol>
		<li>Pipette 1 &#181;L of each of the secondary antibodies into a microcentrifuge tube with 298 &#181;L of the blocking solution to make the final concentration 1:300 for each secondary antibody.</li>
	</ol>
	</li>
	<li>After the last 10 min wash, remove the 1x PBT and pipette 10 &#181;L 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 &#176;C. 
	NOTE: Do not worry about removing every last &#181;L in the wells between rinses, washes, or when adding primary and secondary antibody solutions. The brains will always remain submerged in a few &#181;L of liquid, which is just fine.</li>
	<li>The following day, remove the secondary antibody solution using a micropipette and rinse the brains 3 times with 10 &#181;L each of 1x PBT. Next, wash the brains 4 times with 10 &#181;L each of 1x PBT for 10 min each.</li>
	<li>During the 10 min washes, prepare the EdU reaction mix to detect EdU incorporation. Prepare the EdU reaction mix according to the manufacturers&#39; guidelines.</li>
	<li>After the final wash, remove the 1x PBT and pipette 10 &#181;L 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.</li>
	<li>After 30 min, rinse the brains 3 times with 10 &#181;L each of 1x PBT and wash the brains 3 times with 10 &#181;L each of 1x PBT for 5 min each.</li>
	<li>After the last wash, remove the 1x PBT and pipette 10 &#181;L of a glycerol-based mounting media solution. Seal the plate and place at 4 &#176;C overnight.</li>
</ol>
5. Mounting and imaging the brains
<ol>
	<li>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 &#39;bridge&#39; 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.</li>
</ol>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63189/63189fig05.jpg" /> 
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. <a href="https://www.jove.com/files/ftp_upload/63189/63189fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="2">
	<li>After gluing the 22 mm x 22 mm x 1 mm cover glasses to the microscope slide, pipette 9.3 &#181;L 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 &#181;L volume.</li>
	<li>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.</li>
	<li>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.
	<ol>
		<li>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 &#181;m 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.</li>
	</ol>
	</li>
</ol>
6. Data analysis
<ol>
	<li>Use Fiji open-source software to analyze brain hemispheres and use the Fiji cell counter plugin to count the cells.</li>
</ol>

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The authors have no competing interests.

The method described here to culture brain explants can be carried out in most lab environments. The tools required, as well as the procedure and data collection, are simple and straightforward. With this method, one can test a variety of hypotheses, including those related to the cell signaling cascades and extrinsic factors that regulate NB reactivation and proliferation. Here, using wild-type OregonR animals, we found that exogenous insulin was sufficient to reactivate NBs from quiescence independent of other animal-s...

Stem cells are of great interest because of their potential for use in regenerative medicine1,2. Many animals, especially those that are long-lived, maintain stem cells within their adult tissues. These resident stem cells function to maintain tissue homeostasis and are utilized for repair following physical injury or disease3,4. Most stem cells in adult animals are quiescent, a relatively dormant state characterized by cell cycle arrest and inactivation of growth signaling5. In response to extrinsic cues, stem cells exit from quiescence, enter the cell cycle and begin generating daughter progeny specific to their tissue type. For example, in order to mount an effective immune response, antigen-presenting cells induce quiescent naive T cells to enter cell cycle and clonally expand6. In response to skeletal muscle damage, muscle satellite stem cells enter the cell cycle and generate daughter myoblasts to replace damaged myofibrils5,7. While it is clear that quiescent stem cells respond to extrinsic signals, in many cases, the nature of the extrinsic cue remains unclear as well as the mechanism of cue-induced stem cell activation. Gaining a better understanding of how quiescent stem cells respond to extrinsic cues and enter the cell cycle will aid in the development of better stem cell therapies in the clinic and increase scientific knowledge.
For decades now, model organisms have been used to uncover the genes and cell signaling pathways that regulate stem cell proliferation during development and in adulthood. In Drosophila, neural stem cells (NSCs), known as neuroblasts (NBs), divide throughout development to generate all neurons and glia that ultimately integrate, forming the neural circuity required for brain function8,9. Like other stem cells, NBs divide asymmetrically to self-renew and, in some cases, symmetrically to expand the stem cell pool. NBs are specified during embryogenesis and most enter quiescence towards the end, coincident with declining maternal nutrient stores (Figure 1). After embryogenesis is complete, larvae hatch and begin feeding. In response to animal feeding, NBs reactivate from quiescence and resume cell divisions10,11,12,13,14,15,16. Because the Drosophila CNS is relatively simple and because NBs enter and exit quiescence at defined times, using Drosophila to investigate the regulation of quiescence, entry and exit, proves ideal.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63189/63189fig01.jpg" /> 
Figure 1: Relative proliferation of CB NBs (central brain neuroblasts, red) and MB NBs (mushroom body neuroblasts, blue) over developmental time. At the end of embryogenesis, most NBs (red line) cease proliferation and enter quiescence. Quiescence continues until freshly hatched larvae consume their first complete meal. The time points of focus for this methodology are denoted in red circles (1, quiescence and 2, reactivation). MB NBs (blue) are a subset of central brain NBs that divide continually throughout development (4 per brain hemisphere). <a href="https://www.jove.com/files/ftp_upload/63189/63189fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
In response to animal feeding, PI3-kinase and TOR growth signaling pathways become active in NBs and in their glial and tracheal niche10,11,15,16. When dietary nutrients are withdrawn or when levels of PI3-kinase are reduced, NBs fail to reactivate and growth of glia and trachea are also reduced10,11,15,16. The current model posits that NB reactivation is coupled to larval growth by the fat body, which releases a systemic signal in response to animal feeding12,17,18. This signal, which remains elusive, likely promotes the expression and release of Drosophila insulin-like peptide (Dilps) in the brain, which leads to the downstream activation of PI3-kinase in NBs and their glial and tracheal niche. To better understand the nature of the systemic cue(s), we developed a method to reactivate quiescent NBs in cultured brain explants. With this method, reactivation of NBs can be assayed in the absence of whole animal systemic cues. Exogenous factors can be resupplied to the culture media and NB reactivation assayed based on the incorporation of the thymidine analog, EdU. Using this method, we determined that exogenous insulin is sufficient to reactivate quiescent NBs in brain explants. Future work will be aimed at identifying additional factors that, when added back, either positively or negatively regulate NB quiescence in brain explants.

<table><tbody><tr><td>10 &amp;micro;L Pipette tips</td><td>Denville Sci</td><td>P2102</td><td></td></tr><tr><td>1000 &amp;micro;L Pipette tips</td><td>Denville Sci</td><td>P2103-N</td><td></td></tr><tr><td>1000 &amp;micro;L Pipettor</td><td>Gilson</td><td>P1000</td><td></td></tr><tr><td>16% paraformaldehyde (10 x 10 mL)</td><td>Electron Microscopy Sciences</td><td>2912.60.0000</td><td>Used for Fixation of Larval Brains</td></tr><tr><td>20 &amp;micro;L Pipette</td><td>Gilson</td><td>P20</td><td></td></tr><tr><td>200 &amp;micro;L Pipette tips</td><td>Denville Sci</td><td>1158U56</td><td></td></tr><tr><td>24-well multiwell culture plates</td><td>Fisher Scientific</td><td>50-197-4477</td><td></td></tr><tr><td>35 mm Petri dishes</td><td>Fisher Scientific</td><td>08-757-100A</td><td>Grape Plate Ingredients</td></tr><tr><td>4 &amp;deg;C refrigerator</td><td>Fisher Scientific</td><td></td><td>Provides an ideal temperature for &gt;24 h incubations in antibody solution</td></tr><tr><td>63x Objective</td><td>Lecia</td><td></td><td></td></tr><tr><td>Active dry yeast</td><td>Most supermarkets</td><td></td><td></td></tr><tr><td>Agarose</td><td>Fisher Scientific</td><td>214010</td><td>Grape Plate Ingredients</td></tr><tr><td>Click-iT EdU Cell Proliferation Kit for Imaging, Alexa Fluor 647 dye</td><td>Thermo Fisher Scientific</td><td>C10340</td><td>to label proliferating cells</td></tr><tr><td>Confocal Microscope</td><td>Leica</td><td>SP8</td><td></td></tr><tr><td>Coverslips 22 mm x 22 mm x 1 mm , 10 pack of 4 oz</td><td>Fisher Scientific</td><td>12-544-10</td><td>Two Coverslips are super glued to the ends of the microscope slide. This creates a space that allows for the brains to float in antifade while being imaged.</td></tr><tr><td>Coverslips, 22 mm x 50 mm x 1 mm</td><td>Fisher Scientific</td><td>12-545E</td><td>The coverslip is placed on two square coverslips on the microscope slide ensuring that the brain in the antifade does not move while imaging.</td></tr><tr><td>Dissecting microscope</td><td>Zeiss</td><td>Stemi 2000</td><td></td></tr><tr><td>Ethanol 200 proof (100%), Decon Labs, 1 gallon bottle</td><td>Fisher Scientific</td><td>2701</td><td>Used to wash off the larvae before the 24 hr hold in culture medium</td></tr><tr><td>Fetal Bovine Serum (10%)</td><td>Sigma</td><td>F4135-100ML</td><td>Supplement for cell culture media.</td></tr><tr><td>Fine forceps for dissection</td><td>Fine Science Tools</td><td>11295-20</td><td>Forcepts used in disections. They work best when sharpened.</td></tr><tr><td>Fly Bottles for Crossing</td><td>Genessee Scientific</td><td>32-130</td><td>This bottle is used as a container that lets the flies lay eggs on the grape plate.</td></tr><tr><td>Glass Dissection Dish (3 well)</td><td>These are no longer available</td><td></td><td></td></tr><tr><td>Glutathione</td><td>Sigma</td><td>G6013</td><td>Provides oxidative protection during cell culture.</td></tr><tr><td>Goat Serum</td><td>Sigma</td><td>G9023- 10ML</td><td>Blocking Agent</td></tr><tr><td>Grape Plates</td><td>Made in house</td><td>Made in house</td><td>Grape juice/agarose plates for collecting freshly hatched eggs</td></tr><tr><td>Image J</td><td>Imagej.net/fiji/downloads</td><td>Free Download:&amp;nbsp; https://fiji.sc</td><td>Imaging platform that is used to count cells and Edu reactivation</td></tr><tr><td>Incubator</td><td>Thermo Fisher Scientific</td><td></td><td>Ensures that the temperature, humidity, and light exposure is exactly the same throughout experiment.</td></tr><tr><td>Insulin</td><td>Sigma</td><td>I0516</td><td>Independant variable of the experiment</td></tr><tr><td>Laminar flow hood</td><td></td><td></td><td>For aliquoting culture media</td></tr><tr><td>L-Glutamine</td><td>Sigma</td><td>G7513</td><td>Provides support during cell culture</td></tr><tr><td>Nunc 72-well Microwell Mini Trays</td><td>Fisher Scientific</td><td>12-565-154</td><td>Immunostaining steps are performed in this tray</td></tr><tr><td>Parafilm</td><td>Fisher Scientific</td><td>S37440</td><td>Film used to seal plates in order to prevent evaporation</td></tr><tr><td>Pen-Strep</td><td>Sigma</td><td>P4458-100ml</td><td>Antibiodics used to prevent bacterial contamination of cells during culture.</td></tr><tr><td>Phosphate Buffer, pH7.4</td><td>Made in house</td><td>Made in house</td><td>Solvent used to wash the brains after fixing and staining steps</td></tr><tr><td>Pick</td><td>Fine Science Tools</td><td>10140-01</td><td>Used to pick larvae off of the grape plate</td></tr><tr><td>Propionic acid</td><td>Fisher Scientific</td><td>A-258</td><td>Grape Plate Ingredients</td></tr><tr><td>Rabbit 405</td><td>Abcam</td><td>ab175653</td><td>Antibodies used for immunostaining</td></tr><tr><td>Rat 555</td><td>Abcam</td><td>ab150166</td><td>Antibodies used for immunostaining</td></tr><tr><td>Rb Scribble</td><td>A Gift from Chris Doe</td><td></td><td>Antibodies used for immunostaining</td></tr><tr><td>Rt Deadpan</td><td>Abcam</td><td>ab195173</td><td>Antibodies used for immunostaining</td></tr><tr><td>Schneiders Culture Medium</td><td>Life Tech</td><td>21720024</td><td>Contains nutrients that help the cells grow and proliferate</td></tr><tr><td>SlowFade Diamond Antifade (5 x 2 mL)</td><td>Life Tech</td><td>S36963</td><td>Reagent that provides protection against fading fluorophores</td></tr><tr><td>Sterile Water</td><td>Autoclave Milli-Q water made in house</td><td></td><td>Needed for Solutions</td></tr><tr><td>Sucrose</td><td>Fisher</td><td>S2-12</td><td>Grape Plate Ingredients</td></tr><tr><td>Superfrost Microscope Slides</td><td>Fisher Scientific</td><td>12-544-7</td><td></td></tr><tr><td>Superglue</td><td>Most supermarkets</td><td></td><td></td></tr><tr><td>Tegosept</td><td>Genesee Scientific</td><td>20-259</td><td>Grape Plate Ingredients</td></tr><tr><td>Triton-X 100</td><td>Sigma</td><td>T9284-100ML</td><td>PBT</td></tr><tr><td>Welch&#039;s 100% grape grape juice</td><td>Most supermarkets</td><td></td><td>Grape Plate Ingredients</td></tr></tbody></table>

neural stem cell reactivation in cultured drosophila brain explants

Neural stem cells (NSCs) have the ability to proliferate, differentiate, undergo apoptosis, and even enter and exit quiescence. Many of these processes are controlled by the complex interplay between NSC intrinsic genetic programs with NSC extrinsic factors, local and systemic. In the genetic model organism, Drosophila melanogaster, NSCs, known as neuroblasts (NBs), switch from quiescence to proliferation during the embryonic to larval transition. During this time, larvae emerge from their eggshells and begin crawling, seeking out dietary nutrients. In response to animal feeding, the fat body, an endocrine organ with lipid storage capacity, produces a signal, which is released systemically into the circulating hemolymph. In response to the fat body-derived signal (FBDS), Drosophila insulin-like peptides (Dilps) are produced and released from brain neurosecretory neurons and glia, leading to downstream activation of PI3-kinase growth signaling in NBs and their glial and tracheal niche. Although this is the current model for how NBs switch from quiescence to proliferation, the nature of the FBDS extrinsic cue remains elusive. To better understand how NB extrinsic systemic cues regulate exit from quiescence, a method was developed to culture early larval brains in vitro before animal feeding. With this method, exogenous factors can be supplied to the culture media and NB exit from quiescence assayed. We found that exogenous insulin is sufficient to reactivate NBs from quiescence in whole-brain explants. Because this method is well-suited for large-scale screens, we aim to identify additional extrinsic cues that regulate NB quiescence versus proliferation decisions. Because the genes and pathways that regulate NSC proliferation decisions are evolutionarily conserved, results from this assay could provide insight into improving regenerative therapies in the clinic.

Freshly hatched OregonR wild-type brains were dissected and cultured for 24 h in supplemented Schneider&#39;s media (SSM) with insulin. Tissues were fixed and stained according to the protocol. Primary antibodies generated against Deadpan (Dpn) to detect NBs and Scribble to label cell membranes were used. The thymidine analog 5-Ethynyl-2&#8242;-deoxyuridine (Edu) was added to detect S-phase entry and NB reactivation. We found large sized Edu positive and Dpn positive NBs after 24 h in culture (Figure...

Watch this Scientific Journal Video about Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants at JoVE.com

Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

A method to reactivate quiescent neural stem cells in cultured Drosophila brain explants has been established. Using this method, the role of systemic signals can be uncoupled from tissue-intrinsic signals in the regulation of neural stem cell quiescence, entry and exit.

Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

Neural stem cells (NSCs) have the ability to proliferate, differentiate, undergo apoptosis, and even enter and exit quiescence. Many of these processes ...

neural-stem-cell-reactivation-in-cultured-drosophila-brain-explants

Research

JoVE Journal

Neuroscience

3.0K Views.  University of Virginia. A method to reactivate quiescent neural stem cells in cultured Drosophila brain explants has been established. Using this method, the role of systemic signals can be uncoupled from tissue-intrinsic signals in the regulation of neural stem cell quiescence, entry and exit. 

Video: Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

Primary Neuronal Cultures from the Brains of Late Stage Drosophila Pupae

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the removal of brains from animals at 70-78 hrs after puparium formation. The isolated brains are shown after brief incubation in papain followed by several washes in serum-free growth medium. The process of mechanical dissociation of each brain in a 5 ul drop of media on a coverslip is illustrated. The axons and dendrites of the post-mitotic neurons are sheered off near the soma during dissociation but the neurons begin to regenerate processes within a few hours of plating. Images show live cultures at 2 days. Neurons continue to elaborate processes during the first week in culture. Specific neuronal populations can be identified in culture using GAL4 lines to drive tissue specific expression of fluorescent markers such as  GFP or RFP. Whole cell recordings have demonstrated the cultured neurons form functional, spontaneously active cholinergic and GABAergic synapses. A short video segment illustrates calcium dynamics in the cultured neurons using Fura-2 as a calcium indicator dye to monitor spontaneous calcium transients and nicotine evoked calcium responses in a dish of cultured neurons. These pupal brain cultures are a useful model system in which genetic and pharmacological tools can be used to identify intrinsic and extrinsic factors that influence formation and function of central synapses.

This video demonstrates the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. Views of live cultures show neurite outgrowth and imaging of calcium levels using Fura-2.

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the ...

Biology

Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos and their dechorionation using bleach are demonstrated. Using a glass pipet attached to a mouth suction tube, we illustrate the removal of all cells from single embryos. The method for dispersing cells from each embyro into a small (5 l) drop of medium on an uncoated glass coverslip is demonstrated. A view through the microscope at 1 hour after plating illustrates the preferred cell density. Most of the cells that survive when grown in defined medium are neuroblasts that divide one or more times in culture before extending neuritic processes by 12-24 hours. A view through the microscope illustrates the level of neurite outgrowth and branching expected in a healthy culture at 2 days in vitro. The cultures are grown in a simple bicarbonate based defined medium, in a 5% CO2 incubator at 22-24&deg;C. Neuritic processes continue to elaborate over the first week in culture and when they make contact with neurites from neighboring cells they often form functional synaptic connections. Neurons in these cultures express voltage-gated sodium, calcium, and potassium channels and are electrically excitable. This culture system is useful for studying molecular genetic and environmental factors that regulate neuronal differentiation, excitability, and synapse formation/function.

This video demonstrates the preparation of primary neuronal cultures from midgastrula stage Drosophila embryos. Views of live cultures show cells 1 hour after plating and differentiated neurons after 2 days of growth in a bicarbonate-based defined medium. The neurons are electrically excitable and form synaptic connections.

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos ...

Ex vivo Culturing of Whole, Developing Drosophila Brains

We describe a method for ex vivo culturing of whole Drosophila brains. This can be used as a counterpoint to chronic genetic manipulations for investigating the cell biology and development of central brain structures by allowing acute pharmacological interventions and live imaging of cellular processes. As an example of the technique, prior work from our lab1 has shown that a previously unrecognized subcellular compartment lies between the axonal and somatodendritic compartments of axons of the Drosophila central brain. The development of this compartment, referred to as the axon initial segment (AIS)2, was shown genetically to depend on the neuron-specific cyclin-dependent kinase, Cdk5. We show here that ex vivo treatment of wild-type Drosophila larval brains with the Cdk5-specific pharmacological inhibitors roscovitine and olomoucine3 causes acute changes in actin organization, and in localization of the cell-surface protein Fasciclin 2, that mimic the changes seen in mutants that lack Cdk5 activity genetically.
A second example of the ex vivo culture technique is provided for remodeling of the connections of embryonic mushroom body (MB) gamma neurons during metamorphosis from larva to adult. The mushroom body is the center of olfactory learning and memory in the fly4, and these gamma neurons prune their axonal and dendritic branches during pupal development and then re-extend branches at a later timepoint to establish the adult innervation pattern5. Pruning of these neurons of the MB has been shown to occur via local degeneration of neurite branches6, by a mechanism that is triggered by ecdysone, a steroid hormone, acting at the ecdysone receptor B17, and that is dependent on the activity of the ubiquitin-proteasome system6. Our method of ex vivo culturing can be used to interrogate further the mechanism of developmental remodeling. We found that in the ex vivo culture setting, gamma neurons of the MB recapitulated the process of developmental pruning with a time course similar to that in vivo. It was essential, however, to wait until 1.5 hours after puparium formation before explanting the tissue in order for the cells to commit irreversibly to metamorphosis; dissection of animals at the onset of pupariation led to little or no metamorphosis in culture. Thus, with appropriate modification, the ex vivo culture approach can be applied to study dynamic as well as steady state aspects of central brain biology.

Ex vivo Culturing of Whole, Developing Drosophila Brains

This article describes a method by which one can mimic in vivo development of the Drosophila mushroom body in an ex vivo culture system.

We describe a method for ex vivo culturing of whole Drosophila brains. This can be used as a counterpoint to chronic genetic manipulations for ...

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. Understanding repair and regeneration in animals is a key question in biology. The damaged human CNS does not regenerate, and understanding how to promote the regeneration is one of main goals of medical neuroscience. The powerful genetic toolkit of Drosophila can be used to tackle the problem of CNS regeneration.
A lesion to the CNS ventral nerve cord (VNC, equivalent to the vertebrate spinal cord) is applied manually with a tungsten needle. The VNC can subsequently be filmed in time-lapse using laser scanning confocal microscopy for up to 24 hr to follow the development of the lesion over time. Alternatively, it can be cultured, then fixed and stained using immunofluorescence to visualize neuron and glial cells with confocal microscopy. Using appropriate markers, changes in cell morphology and cell state as a result of injury can be visualized. With ImageJ and purposely developed plug-ins, quantitative and statistical analyses can be carried out to measure changes in wound size over time and the effects of injury in cell proliferation and cell death. These methods allow the analysis of large sample sizes. They can be combined with the powerful genetics of Drosophila to investigate the molecular mechanisms underlying CNS regeneration and repair.

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An injury paradigm using the Drosophila larval ventral nerve cord to investigate central nervous system regeneration and repair is described. Stabbing followed by laser scanning confocal microscopy in time-lapse and fixed specimens, combined with quantitative analysis with purposefully developed software and genetics, are used to investigate the molecular mechanisms of CNS regeneration and repair.

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

An Explant System for Time-Lapse Imaging Studies of Olfactory Circuit Assembly in Drosophila

~Neurons are precisely interconnected to form circuits essential for the proper function of the brain. The Drosophila olfactory system provides an excellent model to investigate this process since 50 types of olfactory receptor neurons (ORNs) from the antennae and maxillary palps project their axons to 50 identifiable glomeruli in the antennal lobe and form synaptic connections with dendrites from 50 types of second-order projection neurons (PNs). Previous studies mainly focused on identifying important molecules that regulate the precise targeting in the olfactory circuit using fixed tissues. Here, an antennae-brain explant system that recapitulates key developmental milestones of olfactory circuit assembly in culture is described. Through dissecting the external cuticle and cleaning opaque fat bodies covering the developing pupal brain, high quality images of single neurons from live brains can be collected using two-photon microscopy. This allows time-lapse imaging of single ORN axon targeting from live tissue. This approach will help reveal important cell biological contexts and functions of previously identified important genes and identify mechanisms underpinning the dynamic process of circuit assembly.

An Explant System for Time-Lapse Imaging Studies of Olfactory Circuit Assembly in Drosophila

This protocol describes the dissection procedure, culture condition, and live imaging of an antennae-brain explant system for the study of the olfactory circuit assembly.

~Neurons are precisely interconnected to form circuits essential for the proper function of the brain. The Drosophila olfactory system provides an ...

Developmental Biology

Imaging Live Brain Explants from Drosophila melanogaster Third Instar Larvae

Live-Cell Imaging of Drosophila melanogaster Third Instar Larval Brains

Drosophila neural stem cells (neuroblasts, NBs hereafter) undergo asymmetric divisions, regenerating the self-renewing neuroblast, while also forming a differentiating ganglion mother cell (GMC), which will undergo one additional division to give rise to two neurons or glia. Studies in NBs have uncovered the molecular mechanisms underlying cell polarity, spindle orientation, neural stem cell self-renewal, and differentiation. These asymmetric cell divisions are readily observable via live-cell imaging, making larval NBs ideally suited for investigating the spatiotemporal dynamics of asymmetric cell division in living tissue. When properly dissected and imaged in nutrient-supplemented medium, NBs in explant brains robustly divide for 12-20 h. Previously described methods are technically difficult and may be challenging to those new to the field. Here, a protocol is described for the preparation, dissection, mounting, and imaging of live third-instar larval brain explants using fat body supplements. Potential problems are also discussed, and examples are provided for how this technique can be used.

Author Spotlight: Investigating Asymmetric Cell Division Dynamics: A Protocol for Live-Imaging of Drosophila Larval Brain Explants 

Here, we discuss a workflow to prepare, dissect, mount, and image live explant brains from Drosophila melanogaster third instar larvae to observe the cellular and subcellular dynamics under physiological conditions.

Drosophila neural stem cells (neuroblasts, NBs hereafter) undergo asymmetric divisions, regenerating the self-renewing neuroblast, while also forming a ...

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. The molecular clock within each pacemaker neuron generates circadian rhythms in gene expression, which underlie the rhythmic neuronal functions essential to the operation of the circuit. Investigation of the properties of the individual molecular oscillators in different subclasses of pacemaker neurons and their interaction with neuronal signaling yields a better understanding of the circadian pacemaker circuit. Here, we present a time-lapse fluorescent microscopy approach developed to monitor the molecular clockwork in clock neurons of cultured Drosophila larval brain. This method allows the multi-day recording of the rhythms of genetically encoded fluorescent circadian reporters at single-cell resolution. This setup can be combined with pharmacological manipulations to closely analyze real-time response of the molecular clock to various compounds. Beyond circadian rhythms, this multipurpose method in combination with powerful Drosophila genetic techniques offers the possibility to study diverse neuronal or molecular processes in live brain tissue.

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The goal of this protocol is to establish ex vivo Drosophila larval brain culture optimized to monitor circadian molecular rhythms with long-term fluorescence time-lapse imaging. The application of this method to pharmacological assays is also discussed.

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. ...

Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

Studies performed in Drosophila melanogaster embryos and larvae provide crucial insight into developmental processes such as cell fate specification and organogenesis. Immunostaining allows for the visualization of developing tissues and organs. However, a protective cuticle that forms at the end of embryogenesis prevents permeation of antibodies into late-stage embryos and larvae. While dissection prior to immunostaining is regularly used to analyze Drosophila larval tissues, it proves inefficient for some analyses because small tissues may be difficult to locate and isolate. Sonication provides an alternative to dissection in larval Drosophila immunostaining protocols. It allows for quick, simultaneous processing of large numbers of late-stage embryos and larvae and maintains in situ morphology. After fixation in formaldehyde, a sample is sonicated. Sample is then subjected to immunostaining with antigen-specific primary antibodies and fluorescently labeled secondary antibodies to visualize target cell types and specific proteins via fluorescence microscopy. During the process of sonication, proper placement of a sonicating probe above the sample, as well as the duration and intensity of sonication, is critical. Additonal minor modifications to standard immunostaining protocols may be required for high quality stains. For antibodies with low signal to noise ratio, longer incubation times are typically necessary. As a proof of concept for this sonication-facilitated protocol, we show immunostains of three tissue types (testes, ovaries, and neural tissues) at a range of developmental stages.

Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

Immunostaining is an effective technique for visualizing specific cell types and proteins within tissues. By utilizing sonication, the protocol described here alleviates the need to dissect Drosophila melanogaster tissues from late-stage embryos and larvae before immunostaining. We provide an efficient methodology for the immunostaining of formaldehyde-fixed whole mount larvae.

Studies performed in Drosophila melanogaster embryos and larvae provide crucial insight into developmental processes such as cell fate specification and ...

Stimulating and Analyzing Adult Neurogenesis in the Drosophila Central Brain

The molecular and cellular mechanisms underlying neurogenesis in response to disease or injury are not well understood. However, understanding these mechanisms is crucial for developing neural regenerative therapies. Drosophila melanogaster is a leading model for studies of neural development but historically has not been exploited to investigate adult brain regeneration. This is primarily because the adult brain exhibits very low mitotic activity. Nonetheless, penetrating traumatic brain injury (PTBI) to the adult Drosophila central brain triggers the generation of new neurons and new glia. The powerful genetic tools available in Drosophila combined with the simple but rigorous injury protocol described here now make adult Drosophila brain a robust model for neural regeneration research. Provided here are detailed instructions for (1) penetrating injuries to the adult central brain and (2) dissection, immunohistochemistry, and imaging post-injury. These protocols yield highly reproducible results and will facilitate additional studies to dissect mechanisms underlying neural regeneration.

Stimulating and Analyzing Adult Neurogenesis in the Drosophila Central Brain

This article provides detailed protocols for inflicting Penetrating Traumatic Brain Injury (PTBI) to adult Drosophila and examining the resulting neurogenesis.

The molecular and cellular mechanisms underlying neurogenesis in response to disease or injury are not well understood. However, understanding these ...

Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

Summary

Explore More Videos

Chapters in this video

Primary Neuronal Cultures from the Brains of Late Stage Drosophila Pupae

Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

Ex vivo Culturing of Whole, Developing Drosophila Brains

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

Live Imaging of Drosophila Larval Neuroblasts

An Explant System for Time-Lapse Imaging Studies of Olfactory Circuit Assembly in Drosophila

Live-Cell Imaging of Drosophila melanogaster Third Instar Larval Brains

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

Stimulating and Analyzing Adult Neurogenesis in the Drosophila Central Brain