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have been shown to divide for over 10 h13,32, while samples imaged with a multi-well slide usually divide less often (unpublished observations).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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<li>Sunchu, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Asymmetric+chromatin+retention+and+nuclear+envelopes+separate+chromosomes+in+fused+cells+in+vivo%5BTitle%5D)+AND+%22Communications+Biology%22%5BJournal%5D)">Asymmetric chromatin retention and nuclear envelopes separate chromosomes in fused cells in vivo.</a> Communications Biology. 5 (1), 953(2022).</li>
<li>Oliveira, A. C., Rebelo, A. R., Homem, C. C. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Integrating+animal+development%3A+How+hormones+and+metabolism+regulate+developmental+transitions+and+brain+formation%5BTitle%5D)+AND+%22Developmental+Biology%22%5BJournal%5D)">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/pubmed?term=((Environmental+control+of+the+cell+cycle+in+Drosophila%3A+nutrition+activates+mitotic+and+endoreplicative+cells+by+distinct+mechanisms%5BTitle%5D)+AND+%22Development%22%5BJournal%5D)">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/pubmed?term=((Drosophila+Aurora-A+kinase+inhibits+neuroblast+self-renewal+by+regulating+aPKC%2FNumb+cortical+polarity+and+spindle+orientation%5BTitle%5D)+AND+%22Genes+%26+Development%22%5BJournal%5D)">Drosophila Aurora-A kinase inhibits neuroblast self-renewal by regulating aPKC/Numb cortical polarity and spindle orientation.</a> Genes & 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/pubmed?term=((Long-term+live+cell+imaging+and+automated+4D+analysis+of+Drosophila+neuroblast+lineages%5BTitle%5D)+AND+%22PLoS+ONE%22%5BJournal%5D)">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/pubmed?term=((Live+imaging+of+neuroblast+lineages+within+intact+larval+brains+in+Drosophila%5BTitle%5D)+AND+%22Cold+Spring+Harbor+Protocols%22%5BJournal%5D)">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/pubmed?term=((Jupiter%2C+a+new+Drosophila+protein+associated+with+microtubules%5BTitle%5D)+AND+%22Cell+Motility+and+the+Cytoskeleton%22%5BJournal%5D)">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/pubmed?term=((The+last-born+daughter+cell+contributes+to+division+orientation+of+Drosophila+larval+neuroblasts%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">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/pubmed?term=((An+immobilization+technique+for+long-term+time-lapse+imaging+of+explanted+Drosophila+tissues%5BTitle%5D)+AND+%22Frontiers+in+Cell+and+Developmental+Biology%22%5BJournal%5D)">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><tbody><tr><td>0.22 &amp;micro;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&#039;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 &amp;micro;&amp;micro;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

JoVE Journal

Developmental Biology

3.3K Views.  University of Washington. Ons onderzoek richt zich op het bestuderen van de impact van asymmetrische celdeling of ACD, op stamcelproliferatie en differentiatie. We gebruiken Drosophila neurale stamcellen of neuroblasten als een model om de mechanismen, voorschriften en effecten van asymmetrische celdeling op ontwikkeling en neurogenese te begrijpen. De recente ontwikkelingen in kunstmatige intelligentie hebben grote vooruitgang geboekt in beeldkwantificering en -analyse, wat het gebied van beeldvorming van levenscelle...

Video: Auteur in de schijnwerpers: onderzoek naar de dynamiek van asymmetrische celdeling: een protocol voor live-beeldvorming van explantaten van de hersenen van Drosophila-larven 

Live-imaging of the Drosophila Pupal Eye

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track during live cell imaging. These processes include: retinal rotation, cell growth and organismal movement. Additionally, irregularities in the topology of the epithelium, including subtle bumps and folds often introduced as the pupa is prepared for imaging, make it challenging to acquire in-focus images of more than a few ommatidia in a single focal plane. The workflow outlined here remedies these issues, allowing easy analysis of cellular processes during Drosophila pupal eye development. Appropriately-staged pupae are arranged in an imaging rig that can be easily assembled in most laboratories. Ubiquitin-DE-Cadherin:GFP and GMR-GAL4&#8211;driven UAS-&#945;-catenin:GFP are used to visualize cell boundaries in the eye epithelium 1-3. After deconvolution is applied to fluorescent images captured at multiple focal planes, maximum projection images are generated for each time point and enhanced using image editing software. Alignment algorithms are used to quickly stabilize superfluous motion, making individual cell behavior easier to track.

Live-imaging of the Drosophila Pupal Eye

This protocol presents an efficient method for imaging the live Drosophila pupal eye neuroepithelium. This method compensates for tissue movement and uneven topology, enhances visualization of cell boundaries through the use of multiple GFP-tagged junction proteins, and uses an easily-assembled imaging rig.

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track ...

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of its convenient genetics and fully sequenced genome. Additionally, the larval nervous system is an ideal model system to study mechanisms of axonal transport as the larval segmental nerves contain bundles of axons with their cell bodies located within the brain and their nerve terminals ending along the length of the body. Here we describe the procedure for visualization of synaptic vesicle proteins within larval segmental nerves. If done correctly, all components of the nervous system, along with associated tissues such as muscles and NMJs, remain intact, undamaged, and ready to be visualized. 3rd instar larvae carrying various mutations are dissected, fixed, incubated with synaptic vesicle antibodies, visualized and compared to wild type larvae. This procedure can be adapted for several different synaptic or neuronal antibodies and changes in the distribution of a variety of proteins can be easily observed within larval segmental nerves.

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster larvae provide an ideal model system to investigate the mechanisms of axonal transport within larval segmental nerves. Using this procedure, 3rd instar larvae carrying various mutations can be compared to wild type larvae.

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of ...

Neuroscience

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 Imaging of Glial Cell Migration in the Drosophila Eye Imaginal Disc

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons. While recent progress has been made to describe the live migration of glial cells in the developing pupal wing (1), studies of Drosophila glial cell migration have typically involved the examination of fixed tissue. Live microscopic analysis of motile cells offers the ability to examine cellular behavior throughout the migratory process, including determining the rate of and changes in direction of growth. Paired with use of genetic tools, live imaging can be used to determine more precise roles for specific genes in the process of development. Previous work by Silies et al. (2) has described the migration of glia originating from the optic stalk, a structure that connects the developing eye and brain, into the eye imaginal disc in fixed tissue. Here we outline a protocol for examining the live migration of glial cells into the Drosophila eye imaginal disc. We take advantage of a Drosophila line that expresses GFP in developing glia to follow glial cell progression in wild type and in mutant animals.

Here we describe a protocol to examine the migration of glial cells into the developing Drosophila eye using live microscopic analysis paired with GFP tagged glial cells.

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons.  While ...

Biology

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Live Imaging of Drosophila Larval Neuroblasts

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

Live Imaging of Drosophila Larval Neuroblasts

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

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

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three preparations of the brain and visual system that cover the range from the developing eye disc-brain complex in the developing pupae to individual eye and brain dissection from adult flies. All protocols are optimized for the live culture of the preparations. However, we also present the conditions for fixed tissue immunohistochemistry where applicable. Finally, we show live imaging conditions for these preparations using conventional and resonant 4D confocal live imaging in a perfusion chamber. Together, these protocols provide a basis for live imaging on different time scales ranging from functional intracellular assays on the scale of minutes to developmental or degenerative processes on the scale of many hours.

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

This protocol describes three Drosophila preparations: 1) adult brain dissection, 2) adult retina dissection and 3) developing eye disc- brain complexes dissection. Emphasis is laid on special preparation techniques and conditions for live imaging, although all preparations can be used for fixed tissue immunohistochemistry.

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

Real-Time Imaging of Germline Stem Cells in Drosophila Ex Vivo Germaria Culture

Extended Live Imaging of Female Drosophila melanogaster Germline Stem Cell Niches

Live imaging methods allow the analysis of dynamic cellular processes in detail and in real-time. The Drosophila ovary represents an excellent model to explore the dynamics of a myriad of developmental processes, such as cell division, stemness, differentiation, migration, apoptosis, autophagy, cellular adhesion, etc., over time. Recently, we have implemented an extended ex vivo culture and live imaging of the female Drosophila GSC niche. Using a Drosophila line harboring a GFP::Par-1 transgene as an example, this method allows the visualization of the GSCs&#39; asymmetric division within their niche and the description of the changes in the spectrosome morphology along the cell cycle. Here, we present a detailed protocol for the ex vivo culture of Drosophila germaria, enabling prolonged visualization of the female GSC niche. Importantly, this protocol is broadly applicable to live imaging GSCs with multiple fluorescently tagged proteins of interest that are available in stock centers and/or in the Drosophila research community.

Here, we provide a detailed protocol to perform live imaging of the asymmetric division of germline stem cells (GSCs) in the Drosophila ovarian niche. We use a transgenic line that ubiquitously expresses a green fluorescent protein (GFP) fusion of the spectrosome protein Par-1.

Live imaging methods allow the analysis of dynamic cellular processes in detail and in real-time. The Drosophila ovary represents an excellent model to ...

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

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

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

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

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

Live-Cell Imaging of Drosophila melanogaster Third Instar Larval Brains

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