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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>10 &#181;L Pipette tips</td>
			<td>Denville Sci</td>
			<td>P2102</td>
			<td></td>
		</tr>
		<tr>
			<td>1000 &#181;L Pipette tips</td>
			<td>Denville Sci</td>
			<td>P2103-N</td>
			<td></td>
		</tr>
		<tr>
			<td>1000 &#181;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 &#181;L Pipette</td>
			<td>Gilson</td>
			<td>P20</td>
			<td></td>
		</tr>
		<tr>
			<td>200 &#181;L Pipette tips</td>
			<td>Gilson</td>
			<td>P200</td>
			<td></td>
		</tr>
		<tr>
			<td>200 &#181;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 &#176;C refrigerator</td>
			<td>Fisher Scientific</td>
			<td></td>
			<td>Provides an ideal temperature for &#62;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:&#160; 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&#39;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

Localizing Protein in 3D Neural Stem Cell Culture: a Hybrid Visualization Methodology

The importance of 3-dimensional (3D) topography in influencing neural stem and progenitor cell (NPC) phenotype is widely acknowledged yet challenging to study. When dissociated from embryonic or post-natal brain, single NPCs will proliferate in suspension to form neurospheres. Daughter cells within these cultures spontaneously adopt distinct developmental lineages (neurons, oligodendrocytes, and astrocytes) over the course of expansion despite being exposed to the same extracellular milieu. This progression recapitulates many of the stages observed over the course of neurogenesis and gliogenesis in post-natal brain and is often used to study basic NPC biology within a controlled environment. Assessing the full impact of 3D topography and cellular positioning within these cultures on NPC fate is, however, difficult. To localize target proteins and identify NPC lineages by immunocytochemistry, free-floating neurospheres must be plated on a substrate or serially sectioned. This processing is required to ensure equivalent cell permeabilization and antibody access throughout the sphere. As a result, 2D epifluorescent images of cryosections or confocal reconstructions of 3D Z-stacks can only provide spatial information about cell position within discrete physical or digital 3D slices and do not visualize cellular position in the intact sphere. Here, to reiterate the topography of the neurosphere culture and permit spatial analysis of protein expression throughout the entire culture, we present a protocol for isolation, expansion, and serial sectioning of post-natal hippocampal neurospheres suitable for epifluorescent or confocal immunodetection of target proteins. Connexin29 (Cx29) is analyzed as an example. Next, using a hybrid of graphic editing and 3D modelling softwares rigorously applied to maintain biological detail, we describe how to re-assemble the 3D structural positioning of these images and digitally map labelled cells within the complete neurosphere. This methodology enables visualization and analysis of the cellular position of target proteins and cells throughout the entire 3D culture topography and will facilitate a more detailed analysis of the spatial relationships between cells over the course of neurogenesis and gliogenesis in vitro.
Both Imbeault and Valenzuela contributed equally and should be considered joint first authors.

Here, we describe how to produce, expand, and immunolabel postnatal hippocampal neural progenitor cells (NPCs) in three-dimensional (3D) culture. Next, using hybrid visualization technologies, we demonstrate how digital images of immunolabelled cryosections can be used to reconstruct and map the spatial position of immunopositive cells throughout the entire 3D neurosphere.

The importance of 3-dimensional (3D) topography in influencing neural stem and progenitor cell (NPC) phenotype is widely acknowledged yet challenging to ...

Isolation and Culture of Neural Crest Cells from Embryonic Murine Neural Tube

The embryonic neural crest (NC) is a multipotent progenitor population that originates at the dorsal aspect of the neural tube, undergoes an epithelial to mesenchymal transition (EMT) and migrates throughout the embryo, giving rise to diverse cell types 1-3. NC also has the unique ability to influence the differentiation and maturation of target organs4-6. When explanted in vitro, NC progenitors undergo self-renewal, migrate and differentiate into a variety of tissue types including neurons, glia, smooth muscle cells, cartilage and bone. 
NC multipotency was first described from explants of the avian neural tube7-9. In vitro isolation of NC cells facilitates the study of NC dynamics including proliferation, migration, and multipotency. Further work in the avian and rat systems demonstrated that explanted NC cells retain their NC potential when transplanted back into the embryo10-13. Because these inherent cellular properties are preserved in explanted NC progenitors, the neural tube explant assay provides an attractive option for studying the NC in vitro. 
To attain a better understanding of the mammalian NC, many methods have been employed to isolate NC populations. NC-derived progenitors can be cultured from post-migratory locations in both the embryo and adult to study the dynamics of post-migratory NC progenitors11,14-20, however isolation of NC progenitors as they emigrate from the neural tube provides optimal preservation of NC cell potential and migratory properties13,21,22. Some protocols employ fluorescence activated cell sorting (FACS) to isolate a NC population enriched for particular progenitors11,13,14,17. However, when starting with early stage embryos, cell numbers adequate for analyses are difficult to obtain with FACS, complicating the isolation of early NC populations from individual embryos. Here, we describe an approach that does not rely on FACS and results in an approximately 96% pure NC population based on a Wnt1-Cre activated lineage reporter23. 
The method presented here is adapted from protocols optimized for the culture of rat NC11,13. The advantages of this protocol compared to previous methods are that 1) the cells are not grown on a feeder layer, 2) FACS is not required to obtain a relatively pure NC population, 3) premigratory NC cells are isolated and 4) results are easily quantified. Furthermore, this protocol can be used for isolation of NC from any mutant mouse model, facilitating the study of NC characteristics with different genetic manipulations. The limitation of this approach is that the NC is removed from the context of the embryo, which is known to influence the survival, migration and differentiation of the NC2,24-28.

Isolation of embryonic neural crest from the neural tube facilitates the use of in vitro methods for studying migration, self-renewal, and multipotency of neural crest.

The embryonic neural crest (NC) is a multipotent progenitor population that originates at the dorsal aspect of the neural tube, undergoes an epithelial to ...

Design and Assembly of an Ultra-light Motorized Microdrive for Chronic Neural Recordings in Small Animals

The ability to chronically record from populations of neurons in freely behaving animals has proven an invaluable tool for dissecting the function of neural circuits underlying a variety of natural behaviors, including navigation1 , decision making 2,3, and the generation of complex motor sequences4,5,6. Advances in precision machining has allowed for the fabrication of light-weight devices suitable for chronic recordings in small animals, such as mice and songbirds. The ability to adjust the electrode position with small remotely controlled motors has further increased the recording yield in various behavioral contexts by reducing animal handling.6,7
 Here we describe a protocol to build an ultra-light motorized microdrive for long-term chronic recordings in small animals. Our design evolved from an earlier published version7, and has been adapted for ease-of use and cost-effectiveness to be more practical and accessible to a wide array of researchers. This proven design 8,9,10,11 allows for fine, remote positioning of electrodes over a range of ~ 5 mm and weighs less than 750 mg when fully assembled. We present the complete protocol for how to build and assemble these drives, including 3D CAD drawings for all custom microdrive components.

The design, fabrication and assembly of an ultra-light motorized microdrive is described. The device provides a cost-effective and easy-to-use solution for chronic recordings of single units in small behaving animals.

The ability to chronically record from populations of neurons in freely behaving animals has proven an invaluable tool for dissecting the function of ...

A Computer-assisted Multi-electrode Patch-clamp System

The patch-clamp technique is today the most well-established method for recording electrical activity from individual neurons or their subcellular compartments. Nevertheless, achieving stable recordings, even from individual cells, remains a time-consuming procedure of considerable complexity. Automation of many steps in conjunction with efficient information display can greatly assist experimentalists in performing a larger number of recordings with greater reliability and in less time. In order to achieve large-scale recordings we concluded the most efficient approach is not to fully automatize the process but to simplify the experimental steps and reduce the chances of human error while efficiently incorporating the experimenter's experience and visual feedback. With these goals in mind we developed a computer-assisted system which centralizes all the controls necessary for a multi-electrode patch-clamp experiment in a single interface, a commercially available wireless gamepad, while displaying experiment related information and guidance cues on the computer screen. Here we describe the different components of the system which allowed us to reduce the time required for achieving the recording configuration and substantially increase the chances of successfully recording large numbers of neurons simultaneously.

Multi-electrode patch-clamp recordings constitute a complex task. Here we show how, by automating of many of the experimental steps, it is possible to accelerate the process leading to qualitative improvement in performance and number of recordings.

The patch-clamp technique is today the most  well-established method for recording electrical activity from individual  neurons or their subcellular ...

Assessing Cell Cycle Progression of Neural Stem and Progenitor Cells in the Mouse Developing Brain after Genotoxic Stress

Neurons of the cerebral cortex are generated during brain development from different types of neural stem and progenitor cells (NSPC), which form a pseudostratified epithelium lining the lateral ventricles of the embryonic brain. Genotoxic stresses, such as ionizing radiation, have highly deleterious effects on the developing brain related to the high sensitivity of NSPC. Elucidation of the cellular and molecular mechanisms involved depends on the characterization of the DNA damage response of these particular types of cells, which requires an accurate method to determine NSPC progression through the cell cycle in the damaged tissue. Here is shown a method based on successive intraperitoneal injections of EdU and BrdU in pregnant mice and further detection of these two thymidine analogues in coronal sections of the embryonic brain. EdU and BrdU are both incorporated in DNA of replicating cells during S phase and are detected by two different techniques (azide or a specific antibody, respectively), which facilitate their simultaneous detection. EdU and BrdU staining are then determined for each NSPC nucleus in function of its distance from the ventricular margin in a standard region of the dorsal telencephalon. Thus this dual labeling technique allows distinguishing cells that progressed through the cell cycle from those that have activated a cell cycle checkpoint leading to cell cycle arrest in response to DNA damage.
An example of experiment is presented, in which EdU was injected before irradiation and BrdU immediately after and analyzes performed within the 4 hr&#160;following irradiation. This protocol provides an accurate analysis of the acute DNA damage response of NSPC in function of the phase of the cell cycle at which they have been irradiated. This method is easily transposable to many other systems in order to determine the impact of a particular treatment on cell cycle progression in living tissues.

Administration of two analogs of thymidine, EdU and BrdU, in pregnant mice allows the analysis of cell cycle progression in neural and progenitor cells in the embryonic mouse brain. This method is useful to determine the effects of genotoxic stress, including ionizing radiation, during brain development.

Neurons of the cerebral cortex are generated during brain development from different types of neural stem and progenitor cells (NSPC), which form a ...

Laser-guided Neuronal Tracing In Brain Explants

We present a technique which combines an in vitro tracer injection protocol, which uses a series of electrical and pressure pulses to increase dye uptake through electroporation in brain explants with targeted laser illumination and matching filter goggles during the procedure. The described technique of in vitro electroporation by itself yields relatively good visual control for targetting certain areas of the brain. By combining it with laser excitation of fluorescent genetic markers and their read-out through band-passing filter goggles, which can pick up the emissions of the genetically labeled cells/nuclei and the fluorescent tracing dye, a researcher can substantially increase the accuracy of injections by finding the area of interest and controlling for the dye-spread/uptake in the injection area much more efficiently. In addition, the laser illumination technique allows to study the functionality of a given neurocircuit by providing information about the type of neurons projecting to a certain area in cases where the GFP expression is linked to the type of transmitter expressed by a subpopulation of neurons.

We describe a technique to label neurons and their processes via anterograde or retrograde tracer injections into brain nuclei using an in vitro preparation. We modified an existing method of in vitro tracer electroporation by taking advantage of fluorescently labeled mouse mutants and basic optical equipment in order to increase labeling accuracy.

We present a technique which combines an in vitro tracer injection protocol, which uses a series of electrical and pressure pulses to increase dye uptake ...

Enumeration of Neural Stem Cells Using Clonal Assays

Neural stem cells (NSCs) have the ability to self-renew and generate the three major neural lineages &#8212; astrocytes, neurons and oligodendrocytes. NSCs and neural progenitors (NPs) are commonly cultured in vitro as neurospheres. This protocol describes in detail how to determine the NSC frequency in a given cell population under clonal conditions. The protocol begins with the seeding of the cells at a density that allows for the generation of clonal neurospheres. The neurospheres are then transferred to chambered coverslips and differentiated under clonal conditions in conditioned medium, which maximizes the differentiation potential of the neurospheres. Finally, the NSC frequency is calculated based on neurosphere formation and multipotency capabilities. Utilities of this protocol include the evaluation of candidate NSC markers, purification of NSCs, and the ability to distinguish NSCs from NPs. This method takes 13 days to perform, which is much shorter than current methods to enumerate NSC frequency.

Neural stem cells (NSCs) refer to cells&#160;which can self-renew and differentiate into the three neural lineages. Here, we describe a protocol to determine NSC frequency in a given cell population using neurosphere formation and differentiation under clonal conditions.

Neural stem cells (NSCs) have the ability to self-renew and generate the three major neural lineages &#8212; astrocytes, neurons and oligodendrocytes. ...

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat open surfaces is complex,&#160;including: pausing, turning, and meandering. This complexity in the repertoire of movement hinders detailed analysis of the events occurring during a single crawl stride cycle. To overcome this obstacle, linear agarose channels were made that constrain larval behavior to straight, sustained, rhythmic crawling. In principle, because agarose channels and the Drosophila larval body are both optically clear, the movement of larval structures labeled by genetically-encoded fluorescent probes can be monitored in intact, freely-moving larvae. In the past, larvae were placed in linear channels and crawling at the level of whole organism, segment, and muscle were analyzed1. In the future, larvae crawling in channels can be used for calcium imaging to monitor neuronal activity. Moreover, these methods can be used with larvae of any genotype and with any researcher-designed channel. Thus the protocol presented below is widely applicable for studies using the Drosophila larva as a model to understand motor control.

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

The Drosophila larva is a powerful model system to study neural control of behavior. This publication describes the use of linear agarose channels to elicit sustained bouts of linear crawling and methods to quantify the dynamics of larval structures during repetitive crawling behavior.

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat ...

Development of a Refined Protocol for Trans-scleral Subretinal Transplantation of Human Retinal Pigment Epithelial Cells into Rat Eyes

Degenerative retinal diseases such as age-related macular degeneration (AMD) are the leading cause of irreversible vision loss worldwide. AMD is characterized by the degeneration of retinal pigment epithelial (RPE) cells, which are a monolayer of cells functionally supporting and anatomically wrapping around the neural retina. Current pharmacological treatments for the non-neovascular AMD (dry AMD) only slow down the disease progression but cannot restore vision, necessitating studies aimed at identifying novel therapeutic strategies. Replacing the degenerative RPE cells with healthy cells holds promise to treat dry AMD in the future. Extensive preclinical studies of stem cell replacement therapies for AMD involve the transplantation of stem cell-derived RPE cells into the subretinal space of animal models, in which the subretinal injection technique is applied. The approach most frequently used in these preclinical animal studies is through the trans-scleral route, which is made difficult by the lack of direct visualization of the needle end and can often result in retinal damage. An alternative approach through the vitreous allows for direct observation of the needle end position, but it carries a high risk of surgical traumas as more eye tissues are disturbed. We have developed a less risky and reproducible modified trans-scleral injection method that uses defined needle angles and depths to successfully and consistently deliver RPE cells into the rat subretinal space and avoid excessive retinal damage. Cells delivered in this manner have been previously demonstrated to be efficacious in the Royal College of Surgeons (RCS) rat for at least 2 months. This technique can be used not only for cell transplantation but also for delivery of small molecules or gene therapies.

Subretinal injection has been widely applied in preclinical studies of stem cell replacement therapy for age-related macular degeneration. In this visualized article, we describe a less risky, reproducible and precisely modified subretinal injection technique via the trans-scleral approach to deliver cells into rat eyes.

Degenerative retinal diseases such as age-related macular degeneration (AMD) are the leading cause of irreversible vision loss worldwide. AMD is ...

Modelling Zika Virus Infection of the Developing Human Brain In Vitro Using Stem Cell Derived Cerebral Organoids

The recent emergence of Zika virus (ZIKV) in susceptible populations has led to an abrupt increase in microcephaly and other neurodevelopmental conditions in newborn infants. While mosquitos are the main route of viral transmission, it has also been shown to spread via sexual contact and vertical mother-to-fetus transmission. In this latter case of transmission, due to the unique viral tropism of ZIKV, the virus is believed to predominantly target the neural progenitor cells (NPCs) of the developing brain.
Here a method for modeling ZIKV infection, and the resulting microcephaly, that occur when human cerebral organoids are exposed to live ZIKV is described. The organoids display high levels of virus within their neural progenitor population, and exhibit severe cell death and microcephaly over time. This three-dimensional cerebral organoid model allows researchers to conduct species-matched experiments to observe and potentially intervene with ZIKV infection of the developing human brain. The model provides improved relevance over standard two-dimensional methods, and contains human-specific cellular architecture and protein expression that are not possible in animal models.

Modelling Zika Virus Infection of the Developing Human Brain In Vitro Using Stem Cell Derived Cerebral Organoids

This protocol describes a technique used to model Zika virus infection of the developing human brain. Using wildtype or engineered stem cell lines, researchers may use this technique to uncover the various mechanisms or treatments that may affect early brain infection and resulting microcephaly in Zika virus-infected embryos.

The recent emergence of Zika virus (ZIKV) in susceptible populations has led to an abrupt increase in microcephaly and other neurodevelopmental conditions ...