This research is supported by R35GM148160 (C. C.) and a National Institutes of Health (NIH) Training Grant T32 GM007270 (R. C. S)

NOTE: Figure 1 shows the materials required to perform this study.
1. Considerations and preparations for the experiment
<ol>
	<li>Prevent the larvae from overcrowding. 
	NOTE: The quality of explant larval brains is directly related to the health and quality of the larvae prior to dissection. Larvae that are malnourished from overcrowding will generally yield lower-quality brains30.
	<ol>
		<li>Ensure that no more than 20-30 larvae are present per meal cap dish to avoid malnutrition. Examples of these can be seen in Figure 2.</li>
	</ol>
	</li>
	<li>Filter and aliquot Schneider&#39;s medium before use.
	<ol>
		<li>For each dissection, prepare fresh imaging and dissection medium by supplementing aliquoted Schneider&#39;s insect medium with 1% bovine growth serum (BGS). A volume of 5 mL of dissection and imaging medium is usually sufficient for an imaging experiment.</li>
		<li>Warm the supplemented medium to room temperature (RT) before use.</li>
	</ol>
	</li>
	<li>Consider the length of the movie to be collected, and use that to factor the supplementation of the imaging medium, the mounting approach, and the acquisition settings of the microscope. 
	NOTE: Under optimal conditions, NBs in brains supplemented with only BGS will robustly divide for upward of 3 h.
	<ol>
		<li>Supplement the imaging medium by adding larval fat body tissues to the imaging medium to support divisions past 4 h when conducting experiments that require longer movies. 
		&#8203;NOTE: Fat bodies secrete mitogens that support NB proliferation31, and whole fat bodies from 10 larvae are sufficient to support four to five brains. Further, samples imaged with a membrane-bound slide have been shown to divide for over 10 h13,32, while samples imaged with a multi-well slide usually divide less often (unpublished observations).</li>
		<li>Alternatively, implement a more complex imaging medium for longer movies, as described previously33. Minimize photodamage by adjusting the exposure time, laser power, and sampling frequency for best results.</li>
	</ol>
	</li>
</ol>
2. Larvae staging and collection (Figure 2)
<ol>
	<li>Cross 1-5 day old female virgin flies with 1-7 day old adult male flies to produce progeny with the desired genotype. For optimal yield, cross 10-15 female virgins with 5-10 males. Deposit these flies into a fly cage with a meal cap (Figure 2A-C), and incubate at 25 &#176;C.</li>
	<li>Swap the meal cap daily. This prevents the meal caps from becoming overcrowded with larvae, which reduces the quality of the dissected brains.</li>
	<li>If the meal cap is significantly covered in larvae (i.e., &#62;30), split this meal cap in half, and replace one half with a fresh meal cap that has also been cut in half. Alternatively, swap the meal caps on a more frequent basis (i.e., every 12 h instead of every 24 h). Examples of overpopulated meal caps can be seen in Figure 2E, F.</li>
	<li>Incubate the meal cap with larvae at 25 &#176;C until the larvae reach the desired age.</li>
</ol>
3. Larval fat body dissection (Figure 3)
NOTE: This protocol describes dissections using a 3-well dissection dish.
<ol>
	<li>Pipette ~400 &#181;L of dissection and imaging medium into each well of a 3-well dissection dish.</li>
	<li>Wash ten 72-96-h old well-fed wild-type larvae by gently holding them with dissection forceps and dipping them in and out of dissection solution in the bottom-most well until all food particulates have been washed off. After rinsing, move the clean larvae to the middle well.</li>
	<li>Using one set of tweezers, hold the larva by the mouth hooks. With the other set of tweezers, rupture one side of the larva&#39;s cuticle.</li>
	<li>This rupture will cause the fat bodies to spill out of the larva. The fat bodies are off-white and semi-translucent and will have a lattice-like structure (Figure 3I). The fat bodies will also tend to stick to themselves and the dissection tweezers. Once identified, collect as much fat body from each larva as possible, and transfer it with the forceps to the top-most well with 400 &#181;L of RT dissection medium.</li>
</ol>
4. Larval brain dissection (Figure 3)
<ol>
	<li>Wash the experimental larvae in dissection and imaging medium as above to free them of food residues. For best results, avoid storing undissected larvae in the dissection solution. This will cause the larvae to &#34;drown&#34; and will negatively impact the quality of the dissected brains.</li>
	<li>Using one set of tweezers, hold the larva by the mouth hooks. Using another set of tweezers, gently cut/rip off approximately 1/3 of the larva from the posterior side (Figure 3A). This will cause elements of the digestive tract, fat bodies, connective tissue, and nervous system to &#34;burst&#34; out of the ruptured side of the larva (Figure 3B).</li>
	<li>Using one set of tweezers, hold the larva by the mouth hooks. With the other set of tweezers, gently brush the cuticle toward the mouth hooks while &#34;pushing&#34; inward with the tweezers holding the mouth hooks until the entire larva is turned inside out. This motion is similar to turning a sock &#34;inside out&#34; (Figure 3C, D).</li>
	<li>Invert the larva so that the central nervous system and other tissues face outward while still being connected to the cuticle. At this step, locate the central nervous system (CNS) to avoid accidental removal. Using tweezers, gently remove all the non-CNS tissue, leaving only the CNS and brain attached to the cuticle (Figure 3E).</li>
	<li>The brain will be attached to the cuticle via axonal connections. Using microdissection scissors, cut these axonal connections to release the brain from the cuticle. To do this, first gently cut underneath the brain lobes (Figure 3F). Repeat with the connections under the ventral nerve cord. 
	NOTE: This step may be done with tweezers if microdissection scissors are not available. Take special care when using tweezers to ensure that the brain tissue is not over-stretched during removal from the cuticle because mechanical stress will negatively affect the brain health.</li>
	<li>Transfer the dissected brain into a well with dissection and imaging medium. For imaging experiments longer than 3 h, use dissection and imaging medium supplemented with fat bodies as described above. Dissect the larvae in batches to keep the dissection time under 20 min.</li>
</ol>
5. Mounting and imaging (Figure 4)
<ol>
	<li>For imaging with a membrane-bound slide34:
	<ol>
		<li>Collect both dissected brains and isolated fat bodies in the last well of the dissection dish.</li>
		<li>Assemble half of the slide by placing a gas-permeable membrane over the back of the slide, and press the split ring into the center, holding it in place (Figure 4A-C).</li>
		<li>Using a 200 &#181;L micropipette, transfer up to 10 dissected brains and as much fat body as possible (see above) in ~130-140 &#181;L of dissection and imaging medium to the membrane. Make sure to deposit the medium with the samples in the center of the gas-permeable membrane (Figure 4D, E).</li>
		<li>Orient the brains for the population of NBs to be imaged and for the type of microscope being used (Figure 4E). Position the sample as close to the microscope&#39;s objective as possible. For example, to image NBs in the central brain lobes, orient the brains such that the brain lobes are closest to the objective (Figure 4H).</li>
		<li>Once the brains are oriented, gently place a glass coverslip on top of the solution on the membrane. This will cause the solution containing the brains and fat bodies to spread over the entirety of the membrane (Figure 4F).</li>
		<li>Blot excessive solution by holding a laboratory tissue close to the coverslip edge. The optimal amount of solution is achieved when the brains touch the coverslip without being squashed. If reorientation is required at this step, carefully move the coverslip to move the brains.</li>
		<li>Immobilize the coverslip by applying melted petroleum jelly along the edges of the coverslip with a paintbrush. Allow the jelly to solidify (Figure 4G).</li>
	</ol>
	</li>
	<li>For imaging with a multi-well imaging slide (Figure 4):
	<ol>
		<li>Add 400 &#181;L of imaging medium to a well of a multi-well slide (in the experiment performed here, a chambered 8-well micro[&#181;]-slide was used; Figure 4I). Transfer the previously dissected fat bodies to this well (see step 3.4).</li>
		<li>Deposit up to 10 brains in a cluster near the center of the well (Figure 4J).</li>
		<li>Orient the brains for the population of NBs to be imaged and for the type of microscope being used, as described in step 5.1.4 (Figure 4K). Arrange the samples so they are close to each other. This will minimize the distance the stage must move between samples, which reduces sample drift during acquisition.</li>
		<li>Once the brains have been oriented in the well, allow the brains to settle for 2-5 min. This increases their stability during transport/imaging. Prepare the microscope for acquisition during this time.</li>
		<li>Cover the &#181;-slide with the slide cover, and transfer it to the microscope. Begin acquisition with the lowest laser power and exposure time possible to minimize photobleaching.</li>
	</ol>
	</li>
</ol>
6. Data processing and management best practices
<ol>
	<li>Process the data as needed according to the available analysis software.
	<ol>
		<li>For the example shown here, save the acquired data with SlideBook software as a SlideBook Image File (.sld).</li>
		<li>To convert into Imaris&#39; proprietary file type (.ims) using the Imaris File Converter, open the Imaris File Converter in a separate window. Click on and drag the .sld files into the &#34; input&#34; section of the Imaris File Converter.</li>
		<li>Determine the desired output location for the converted files, and click on &#34;Start all.&#34;</li>
		<li>After conversion, view and annotate the data in the Imaris software. 
		NOTE: Alternatives for image analysis can be used in place of Imaris, such as Fiji (https://hpc.nih.gov/apps/Fiji.html), Aivia (https://www.aivia-software.com/), Volocity (https://www.volocity4d.com/), or others.</li>
	</ol>
	</li>
	<li>Retain as much of the original data as possible for proper record-keeping. For example, if the acquisition software is saved in one file format but is converted to a different format for analysis, retain the acquired version of the data.</li>
	<li>For data analysis, maintain a record of as many details as possible about the sample and acquisition settings. Key information to retain includes the genotype of the dissected larvae, the age of the larvae prior to dissection, the state of the meal cap they were reared in, the laser power used during imaging, the exposure time, the length of acquisition, and the temporal resolution.</li>
</ol>
7. Example quantification of cell cycle length (Figure 5)
NOTE: in this example, larvae expressing the polarity marker Pins (Pins::EGFP16) and the microtubule-binding protein Jupiter25 (cherry::Jupiter13) were imaged. The subsequent analysis was performed using Imaris software.
<ol>
	<li>Open the movie using the image analysis software of choice. Scroll through the length of the movie to identify dividing NBs, and label them for future reference. Identify the dividing NBs by their distinct mitotic spindles (Figure 5C-E).</li>
	<li>Identify a reference cell cycle stage to determine the cell cycle length. In this example, metaphase is used as a reference.</li>
	<li>Manually determine the number of frames between successive metaphases, and convert it to minutes or hours to determine the time taken to complete one cell cycle.
	<ol>
		<li>Do this by taking the temporal resolution of the movie and multiplying it by the number of frames between metaphases. For example, if the temporal resolution of the movie is one frame every 5 min, and metaphases are observed in frame 13 and frame 35, the time between these metaphases would be 110 min ([35 &#8722; 13] &#215; 5).</li>
	</ol>
	</li>
	<li>Plot the data with any appropriate software. The data shown here were plotted using PRISM software.</li>
</ol>
8. Example quantification of cell spindle alignment (Figure 5)
NOTE: In this example, the analysis is performed using Imaris software.
<ol>
	<li>Open the movie file in Imaris or another software of choice. Scroll through the length of the movie to identify dividing NBs, and label them for future reference.</li>
	<li>Determine the vector formed by the spindle poles using the apical and basal centrosomes (represented by m), as follows: 
	<img alt="Equation 1" src="/files/ftp_upload/65538/65538eq01.jpg" style="margin: auto;" /> 
	where Ax, Ay, and Az are the coordinates of the apical centrosome, and Bx, By, and Bz are the coordinates of the basal centrosome. Similarly, the axis of the division vector (represented by n) is formed by the midpoint of the apical Pins::EGFP crescent and the basal cortex: 
	<img alt="Equation 2" src="/files/ftp_upload/65538/65538eq02.jpg" style="margin: auto;" /> 
	where Ax, Ay, and Az are the coordinates of the midpoint of the Pins::EGFP crescent, and Bx, By, and Bz are the coordinates of the midpoint of the basal cortex.</li>
	<li>Determine the magnitude of the vectors m and n: 
	Magnitude of m: <img alt="Equation 3" src="/files/ftp_upload/65538/65538eq03.jpg" style="margin: auto;" /> 
	Magnitude of n: <img alt="Equation 4" src="/files/ftp_upload/65538/65538eq04.jpg" style="margin: auto;" /></li>
	<li>Determine the dot product (represented by k) of m and n: 
	<img alt="Equation 5" src="/files/ftp_upload/65538/65538eq05.jpg" style="margin: auto;" /></li>
	<li>Using the dot product k and vector magnitudes m and n, determine the angle between the vectors: 
	<img alt="Equation 6" src="/files/ftp_upload/65538/65538eq06.jpg" style="margin: auto;" /></li>
	<li>Plot the data in the software of choice. The data shown here were prepared in Microsoft Excel and visualized in PRISM.</li>
</ol>

Imaging Live Brain Explants from Drosophila melanogaster Third Instar Larvae

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R., Cabernard, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+centriolar+protein+Bld10/Cep135+is+required+to+establish+centrosome+asymmetry+in+Drosophila+neuroblasts.">The centriolar protein Bld10/Cep135 is required to establish centrosome asymmetry in Drosophila neuroblasts.</a> Current Biology. 24 (13), 1548-1555 (2014).</li><li>LaFoya, B., Prehoda, K. 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F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Integrating+animal+development:+How+hormones+and+metabolism+regulate+developmental+transitions+and+brain+formation.">Integrating animal development: How hormones and metabolism regulate developmental transitions and brain formation.</a> Developmental Biology. 475, 256-264 (2021).</li><li>Britton, J. S., Edgar, B. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Environmental+control+of+the+cell+cycle+in+Drosophila:+nutrition+activates+mitotic+and+endoreplicative+cells+by+distinct+mechanisms.">Environmental control of the cell cycle in Drosophila: nutrition activates mitotic and endoreplicative cells by distinct mechanisms.</a> Development. 125 (11), 2149-2158 (1998).</li><li>Lee, C. -. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+Aurora-A+kinase+inhibits+neuroblast+self-renewal+by+regulating+aPKC/Numb+cortical+polarity+and+spindle+orientation.">Drosophila Aurora-A kinase inhibits neuroblast self-renewal by regulating aPKC/Numb cortical polarity and spindle orientation.</a> Genes &amp; Development. 20 (24), 3464-3474 (2006).</li><li>Homem, C. C. F., Reichardt, I., Berger, C., Lendl, T., Knoblich, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+live+cell+imaging+and+automated+4D+analysis+of+Drosophila+neuroblast+lineages.">Long-term live cell imaging and automated 4D analysis of Drosophila neuroblast lineages.</a> PLoS ONE. 8 (11), e79588 (2013).</li><li>Cabernard, C., Doe, C. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+imaging+of+neuroblast+lineages+within+intact+larval+brains+in+Drosophila.">Live imaging of neuroblast lineages within intact larval brains in Drosophila.</a> Cold Spring Harbor Protocols. 2013 (10), 970-977 (2013).</li><li>Karpova, N., Bobinnec, Y., Fouix, S., Huitorel, P., Debec, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Jupiter,+a+new+Drosophila+protein+associated+with+microtubules.">Jupiter, a new Drosophila protein associated with microtubules.</a> Cell Motility and the Cytoskeleton. 63 (5), 301-312 (2006).</li><li>Loyer, N., Januschke, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+last-born+daughter+cell+contributes+to+division+orientation+of+Drosophila+larval+neuroblasts.">The last-born daughter cell contributes to division orientation of Drosophila larval neuroblasts.</a> Nature Communications. 9 (1), 3745 (2018).</li><li>Bostock, M. P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+immobilization+technique+for+long-term+time-lapse+imaging+of+explanted+Drosophila+tissues.">An immobilization technique for long-term time-lapse imaging of explanted Drosophila tissues.</a> Frontiers in Cell and Developmental Biology. 8, 590094 (2020).</li></ol>

The authors have no financial disclosures to declare.

This protocol outlines one approach for the imaging of live explant brains from Drosophila melanogaster larvae. The protocol described here allows for explant brains to be observed for 12-20 h under the right experimental conditions. Special consideration must be given to the preparation of samples and the design of the desired experiments. As mentioned above, one of the most critical factors that determines the quality of the dissected tissue is the health of the larvae. To achieve the highest quality possible,...

Asymmetric cell division (ACD) is the process by which subcellular components such as RNA, proteins, and organelles are partitioned unequally between daughter cells1,2. This process is commonly seen in stem cells, which undergo ACD to give rise to daughter cells with different developmental fates. Drosophila NBs divide asymmetrically to produce one NB, which retains its stemness, and one ganglion mother cell (GMC). The GMC undergoes further divisions to produce differentiating neurons or glia3. Asymmetrically dividing NBs are abundant in the developing brains of third-instar larvae, which are readily observed via microscopy. At the third instar larval stage, there are roughly 100 NBs present in each central brain lobe3,4,5,6.
Asymmetric cell division is a highly dynamic process. Live-cell imaging protocols have been used to measure and quantify the dynamics of cell polarity7,8,9,10, spindle orientation11,12,13, the dynamics of the actomyosin cortex14,15,16,17,18, microtubule and centrosome biology19,20,21,22,23,24,25,26,27, and membrane10,28 and chromatin dynamics29. Qualitative and quantitative descriptions of ACD rely on robust methods and protocols to image dividing NBs in intact living brains. The following protocol outlines methods to prepare, dissect, and image third instar larval brains for live-cell imaging in vivo using two different mounting approaches. These methods are best suited for researchers interested in the spatiotemporal dynamics of stem cell divisions, as well as divisions in other brain cells, as they allow for short- and long-term observations of cellular events. Additionally, these techniques are readily accessible to newcomers to the field. We demonstrate the effectiveness and adaptability of this approach with larval brains expressing fluorescently tagged microtubule and cortical fusion proteins. We additionally discuss methods of analysis and considerations for application in other studies.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.22 &#181;m polyethersulfone (PES) Membrane</td>
			<td>Genesee</td>
			<td>25-231</td>
			<td>Vacuum-driven filters</td>
		</tr>
		<tr>
			<td>Agar</td>
			<td>Genesee</td>
			<td>20-248</td>
			<td>granulated agar</td>
		</tr>
		<tr>
			<td>Analytical Computer</td>
			<td>Dell</td>
			<td>NA</td>
			<td>Intel Xeon Gold 5222 CPU with two 3.80 GHz processors running Windows 10 on a 64-bit operating system</td>
		</tr>
		<tr>
			<td>Bovine Growth Serum</td>
			<td>HyClone</td>
			<td>SH30541.02</td>
			<td></td>
		</tr>
		<tr>
			<td>Chambered Imaging Slides</td>
			<td>Ibidi</td>
			<td>80826</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal Microscope</td>
			<td>Nikon</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Custom-machined metal slide</td>
			<td>NA</td>
			<td>NA</td>
			<td>See Cabernard and Doe 2013 (Ref. 34) for specifications</td>
		</tr>
		<tr>
			<td>Dissection Dishes</td>
			<td>Fisher Scientific</td>
			<td>5024343</td>
			<td>3-well porcelain micro spot plate</td>
		</tr>
		<tr>
			<td>Dissection Forceps</td>
			<td>World Precision Instruments</td>
			<td>Dumont #5</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection Microscope</td>
			<td>Leica</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection Scissors</td>
			<td>Fine Science Tools (FST)</td>
			<td>15003-08</td>
			<td></td>
		</tr>
		<tr>
			<td>Embryo collection cage</td>
			<td>Genesee</td>
			<td>59-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Flypad with access to CO2 to anesthetize adult flies</td>
			<td>Genesee</td>
			<td>59-172</td>
			<td></td>
		</tr>
		<tr>
			<td>Gas-permeable membrane</td>
			<td>YSI</td>
			<td>98095</td>
			<td>Gas-permeable membrane</td>
		</tr>
		<tr>
			<td>Glass Cover Slides</td>
			<td>Electron Microscopy Sciences</td>
			<td>72204-03</td>
			<td># 1.5; 22 mm x 40 mm glass coverslips</td>
		</tr>
		<tr>
			<td>Imaris</td>
			<td>Oxford Instruments</td>
			<td>NA</td>
			<td>Alternatives: Fiji, Volocity, Aivia</td>
		</tr>
		<tr>
			<td>Imaris File Converter</td>
			<td>Oxford Instruments</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Instant Yeast</td>
			<td>Saf-Instant</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Molasses</td>
			<td>Genesee</td>
			<td>62-117</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dish</td>
			<td>Greiner Bio-One</td>
			<td>628161</td>
			<td>60 mm x 15 mm Petri dish</td>
		</tr>
		<tr>
			<td>Petroleum Jelly</td>
			<td>Vaseline</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Schneider&#39;s Insect Medium with L-glutamine and sodium bicarbonate liquid</td>
			<td>Millipore Sigma</td>
			<td>S0146</td>
			<td></td>
		</tr>
		<tr>
			<td>SlideBook acquisition software</td>
			<td>3i</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Vacuum-Driven Filtration Unit with a 0.22 &#181;&#181;m PES membrane filter</td>
			<td>Genesee Scientific, GenClone</td>
			<td>25-231</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Dissection and imaging of central brain lobe NBs expressing Pins::EGFP and Cherry::Jupiter 
To showcase this protocol, larvae expressing UAS-driven Cherry::Jupiter13 and endogenously tagged Pins::EGFP16 (w; worGal4, UAS-cherry::jupiter/CyO; Pins::EGFP/TM6B, Tb) were imaged for 4 h using the described protocol using multi-well imaging slides (Figure 5C,D). Additional data were taken from larvae expressing U...

Watch this Scientific Journal Video about Investigating Asymmetric Cell Division Dynamics: A Protocol for Live-Imaging of Drosophila Larval Brain Explants at JoVE.com

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.

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 ...

live-cell-imaging-drosophila-melanogaster-third-instar-larval

Research

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Developmental Biology

3.2K Views.  University of Washington. 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.

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

Upright Imaging of Drosophila Egg Chambers

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in architecture, is well-characterized, and is amenable to genetic analysis. Each egg chamber consists of germ-line cells surrounded by a single epithelial layer of somatic follicle cells. Subsets of follicle cells undergo differentiation during specific stages to become several different cell types. Standard techniques primarily allow for a lateral view of egg chambers, and therefore a limited view of follicle cell organization and identity. The upright imaging protocol describes a mounting technique that enables a novel, vertical view of egg chambers with a standard confocal microscope. Samples are first mounted between two layers of glycerin jelly in a lateral (horizontal) position on a glass microscope slide. The jelly with encased egg chambers is then cut into blocks, transferred to a coverslip, and flipped to position egg chambers upright. Mounted egg chambers can be imaged on either an upright or an inverted confocal microscope. This technique enables the study of follicle cell specification, organization, molecular markers, and egg development with new detail and from a new perspective.

Upright Imaging of Drosophila Egg Chambers

The upright imaging method described in this protocol allows for the detailed visualization of the poles of a developing Drosophila melanogaster egg. This end-on view provides a new perspective into the arrangements and morphologies of multiple cell types in the follicular epithelium.

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in ...

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

In vertebrates, hematopoiesis is regulated by inductive microenvironments (niches). Likewise, in the invertebrate model organism&#160;Drosophila melanogaster, inductive microenvironments known as larval Hematopoietic Pockets (HPs) have been identified as anatomical sites for the development and regulation of blood cells (hemocytes), in particular of the self-renewing macrophage lineage.&#160;HPs are segmentally repeated pockets between the epidermis and muscle layers of the larva, which also comprise sensory neurons of the peripheral nervous system. In the larva, resident (sessile) hemocytes are exposed to anti-apoptotic, adhesive and proliferative cues from these sensory neurons and potentially other components of the HPs, such as the lining muscle and epithelial layers. During normal development, gradual release of resident hemocytes from the HPs fuels the population of circulating hemocytes, which culminates in the release of most of the resident hemocytes at the beginning of metamorphosis. Immune assaults, physical injury or mechanical disturbance trigger the premature release of resident hemocytes into circulation. The switch of larval hemocytes between resident locations and circulation raises the need for a common standard/procedure to selectively isolate and quantify these two populations of blood cells from single Drosophila larvae. Accordingly, this protocol describes an automated method to release and quantify the resident and circulating hemocytes from single larvae. The method facilitates ex vivo approaches, and may be adapted to serve a variety of developmental stages of Drosophila and other invertebrate organisms.

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

Drosophila blood cells, or hemocytes, cycle between resident sites and circulation. In the larva, resident (sessile) hemocytes localize to inductive microenvironments, the Hematopoietic Pockets, while circulating hemocytes move freely in the hemolymph. The goal of this protocol is the standardized isolation and quantification of these two, behaviorally distinct but interchanging, hemocyte populations.

In vertebrates, hematopoiesis is regulated by inductive microenvironments (niches). Likewise, in the invertebrate model organism&#160;Drosophila ...

Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

The influence of microbes on myriad animal traits and behaviors has been increasingly recognized in recent years. The fruit fly Drosophila melanogaster is a model for understanding microbial interactions with animal hosts, facilitated by approaches to rear large sample sizes of Drosophila under microorganism-free (axenic) conditions, or with defined microbial communities (gnotobiotic). This work outlines a method for collection of Drosophila embryos, hypochlorite dechorionation and sterilization, and transfer to sterile diet. Sterilized embryos are transferred to sterile diet in 50 ml centrifuge tubes, and developing larvae and adults remain free of any exogenous microbes until the vials are opened. Alternatively, flies with a defined microbiota can be reared by inoculating sterile diet and embryos with microbial species of interest. We describe the introduction of 4 bacterial species to establish a representative gnotobiotic microbiota in Drosophila. Finally, we describe approaches for confirming bacterial community composition, including testing if axenic Drosophila remain bacteria-free into adulthood.

Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

A method for rearing Drosophila melanogaster under axenic and gnotobiotic conditions is presented. Fly embryos are dechorionated in sodium hypochlorite, transferred aseptically to sterile diet, and reared in closed containers. Inoculating diet and embryos with bacteria leads to gnotobiotic associations, and bacterial presence is confirmed by plating whole-body Drosophila homogenates.

The influence of microbes on myriad animal traits and behaviors has been increasingly recognized in recent years. The fruit fly Drosophila melanogaster is ...

Culture of Adult Transgenic Zebrafish Retinal Explants for Live-cell Imaging by Multiphoton Microscopy

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The M&#252;ller glia re-enter the cell cycle and produce neuronal progenitor cells that undergo subsequent rounds of cell divisions and differentiate into the lost neuronal cell types. Both M&#252;ller glia and neuronal progenitor cell nuclei replicate their DNA and undergo mitosis in distinct locations of the retina, i.e. they migrate between the basal Inner Nuclear Layer (INL) and the Outer Nuclear Layer (ONL), respectively, in a process described as Interkinetic Nuclear Migration (INM). INM has predominantly been studied in the developing retina. To examine the dynamics of INM in the adult regenerating zebrafish retina in detail, live-cell imaging of fluorescently-labeled M&#252;ller glia/neuronal progenitor cells is required. Here, we provide the conditions to isolate and culture dorsal retinas from Tg[gfap:nGFP]mi2004 zebrafish that were exposed to constant intense light for 35 h. We also show that these retinal cultures are viable to perform live-cell imaging experiments, continuously acquiring z-stack images throughout the thickness of the retinal explant for up to 8 h using multiphoton microscopy to monitor the migratory behavior of gfap:nGFP-positive cells. In addition, we describe the details to perform post-imaging analysis to determine the velocity of apical and basal INM. To summarize, we established conditions to study the dynamics of INM in an adult model of neuronal regeneration. This will advance our understanding of this crucial cellular process and allow us to determine the mechanisms that control INM.

Zebrafish retinal regeneration has mostly been studied using fixed retinas. However, dynamic processes such as interkinetic nuclear migration occur during the regenerative response and require live-cell imaging to investigate the underlying mechanisms. Here, we describe culture and imaging conditions to monitor Interkinetic Nuclear Migration (INM) in real-time using multiphoton microscopy.

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The ...

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the proteins involved in these pathways are evolutionarily conserved between mammals and other eukaryotes, such as the fruit fly Drosophila melanogaster, suggesting that similar organizing principles exist during the development of these organisms. Importantly, Drosophila has been used extensively to identify cellular and molecular mechanisms regulating processes that are required in mammals including neurogenesis, differentiation, axonal guidance, and synaptogenesis. Flies have also been used successfully to model a variety of human neurodevelopmental diseases. Here we describe a protocol for the step-by-step microdissection, fixation, and immunofluorescent localization of proteins within the adult Drosophila brain. This protocol focuses on two example neuronal populations, mushroom body neurons and retinal photoreceptors, and includes optional steps to trace individual mushroom body neurons using Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. Example data from both wild-type and mutant brains are shown along with a brief description of a scoring criteria for axonal guidance defects. While this protocol highlights two well-established antibodies for investigating the morphology of mushroom body and photoreceptor neurons, other Drosophila brain regions and the localization of proteins within other brain regions can also be investigated using this protocol.

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

This protocol describes the dissection and immunostaining of adult Drosophila melanogaster brain tissues. Specifically, this protocol highlights the use of Drosophila mushroom body and photoreceptor neurons as example neuronal subsets that can be accurately used to uncover general principles underlying many aspects of neuronal development.

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the ...

Live Cell Imaging of Chromosome Segregation During Mitosis

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled by multiple mechanisms that include the Spindle Assembly Checkpoint (SAC). The SAC is part of a complex feedback system that is responsible for prevention of a cell progress through mitosis unless all chromosomal kinetochores have attached to spindle microtubules. Chromosomal lagging and abnormal chromosome segregation is an indicator of dysfunctional cell cycle control checkpoints and can be used to measure the genomic stability of dividing cells. Deregulation of the SAC can result in the transformation of a normal cell into a malignant cell through the accumulation of errors during chromosomal segregation. Implementation of the SAC and the formation of the kinetochore complex are tightly regulated by interactions between kinases and phosphatase such as Protein Phosphatase 2A (PP2A). This protocol describes live cell imaging of lagging chromosomes in mouse embryonic fibroblasts isolated from mice that had a knockout of the PP2A-B56&#947; regulatory subunit. This method overcomes the shortcomings of other cell cycle control imaging techniques such as flow cytometry or immunocytochemistry that only provide a snapshot of a cell cytokinesis status, instead of a dynamic spatiotemporal visualization of chromosomes during mitosis.

This protocol describes an easy and convenient method to label and visualize live chromosomes in mitotic cells using Histone2B-GFP BacMam 2.0 label and a spinning disc confocal microscopy system.

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled ...

Preparation of Drosophila Larval Samples for Gas Chromatography-Mass Spectrometry (GC-MS)-based Metabolomics

Recent advances in the field of metabolomics have established the fruit fly Drosophila melanogaster as a powerful genetic model for studying animal metabolism. By combining the vast array of Drosophila genetic tools with the ability to survey large swaths of intermediary metabolism, a metabolomics approach can reveal complex interactions between diet, genotype, life-history events, and environmental cues. In addition, metabolomics studies can discover novel enzymatic mechanisms and uncover previously unknown connections between seemingly disparate metabolic pathways. In order to facilitate more widespread use of this technology among the Drosophila community, here we provide a detailed protocol that describes how to prepare Drosophila larval samples for gas chromatography-mass spectrometry (GC-MS)-based metabolomic analysis. Our protocol includes descriptions of larval sample collection, metabolite extraction, chemical derivatization, and GC-MS analysis. Successful completion of this protocol will allow users to measure the relative abundance of small polar metabolites, including amino acids, sugars, and organic acids involved in glycolysis and the TCA cycles.

Preparation of Drosophila Larval Samples for Gas Chromatography-Mass Spectrometry GC-MS-based Metabolomics

This protocol describes how to prepare Drosophila larvae for GC-MS-based metabolomic analysis.

Recent advances in the field of metabolomics have established the fruit fly Drosophila melanogaster as a powerful genetic model for studying animal ...

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

There are currently over 160 distinct Drosophila cell lines distributed by the Drosophila Genomics Resource Center (DGRC). With genome engineering, the number of novel cell lines is expected to increase. The DGRC aims to familiarize researchers with using Drosophila cell lines as an experimental tool to complement and drive their research agenda. Procedures for working with a variety of Drosophila cell lines with distinct characteristics are provided, including protocols for thawing, culturing, and cryopreserving cell lines. Importantly, this publication demonstrates the best practices required to work with Drosophila cell lines to minimize the risk of contaminations from adventitious microorganisms or from other cell lines. Researchers who become familiar with these procedures will be able to delve into the many applications that use Drosophila cultured cells including biochemistry, cell biology and functional genomics.

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

Drosophila cell lines are important reagents for both fundamental and biomedical research. This article provides protocols for thawing, subculturing, and the cryopreservation of commonly used Drosophila cell lines to assist researchers in incorporating the use of these reagents in their research.

There are currently over 160 distinct Drosophila cell lines distributed by the Drosophila Genomics Resource Center (DGRC). With genome engineering, the ...

Methods to Test Endocrine Disruption in Drosophila melanogaster

In recent years there has been growing evidence that all organisms and the environment are exposed to hormone-like chemicals, known as endocrine disruptor chemicals (EDCs). These chemicals may alter the normal balance of endocrine systems and lead to adverse effects, as well as an increasing number of hormonal disorders in the human population or disturbed growth and reduced reproduction in the wildlife species. For some EDCs, there are documented health effects and restrictions on their use. However, for most of them, there is still no scientific evidence in this sense. In order to verify potential endocrine effects of a chemical in the full organism, we need to test it in appropriate model systems, as well as in the fruit fly, Drosophila melanogaster. Here we report detailed in vivo protocols to study endocrine disruption in Drosophila, addressing EDC effects on the fecundity/fertility, developmental timing, and lifespan of the fly. In the last few years, we used these Drosophila life traits to investigate the effects of exposure to 17-&#945;-ethinylestradiol (EE2), bisphenol A (BPA), and bisphenol AF (BPA F). Altogether, these assays covered all Drosophila life stages and made it possible to evaluate endocrine disruption in all hormone-mediated processes. Fecundity/fertility and developmental timing assays were useful to measure the EDC impact on the fly reproductive performance and on developmental stages, respectively. Finally, the lifespan assay involved chronic EDC exposures to adults and measured their survivorship. However, these life traits can also be influenced by several experimental factors that had to be carefully controlled. So, in this work, we suggest a series of procedures we have optimized for the right outcome of these assays. These methods allow scientists to establish endocrine disruption for any EDC or for a mixture of different EDCs in Drosophila, although to identify the endocrine mechanism responsible for the effect, further essays could be needed.

Methods to Test Endocrine Disruption in Drosophila melanogaster

Endocrine disruptor chemicals (EDCs) represent a serious problem for organisms and for natural environments. Drosophila melanogaster represents an ideal model to study EDC effects in vivo. Here, we present methods to investigate endocrine disruption in Drosophila, addressing EDC effects on fecundity, fertility, developmental timing, and lifespan of the fly.

In recent years there has been growing evidence that all organisms and the environment are exposed to hormone-like chemicals, known as endocrine disruptor ...

Assay for Blood-brain Barrier Integrity in Drosophila melanogaster

Proper nervous system development includes the formation of the blood-brain barrier, the diffusion barrier that tightly regulates access to the nervous system and protects neural tissue from toxins and pathogens. Defects in the formation of this barrier have been correlated with neuropathies, and the breakdown of this barrier has been observed in many neurodegenerative diseases. Therefore, it is critical to identify the genes that regulate the formation and maintenance of the blood-brain barrier to identify potential therapeutic targets. In order to understand the exact roles these genes play in neural development, it is necessary to assay the effects of altered gene expression on the integrity of the blood-brain barrier. Many of the molecules that function in the establishment of the blood-brain barrier have been found to be conserved across eukaryotic species, including the fruit fly, Drosophila melanogaster. Fruit flies have proven to be an excellent model system for examining the molecular mechanisms regulating nervous system development and function. This protocol describes a step-by-step procedure to assay for blood-brain barrier integrity during the embryonic and larval stages of D. melanogaster development.

Assay for Blood-brain Barrier Integrity in Drosophila melanogaster

Blood-brain barrier integrity is critical for nervous system function. In Drosophila melanogaster, the blood-brain barrier is formed by glial cells during late embryogenesis. This protocol describes methods to assay for blood-brain barrier formation and maintenance in D. melanogaster embryos and third instar larvae.

Proper nervous system development includes the formation of the blood-brain barrier, the diffusion barrier that tightly regulates access to the nervous ...