This research was funded by the NINDS of the National Institutes of Health under award number K01NS094545, andgrants from the Lefler Center for the Study of Neurodegenerative Disorders. We acknowledge Liming Tan and Jason McEwan for valuable conversations.

1. Planning Before the Experiment
<ol>
	<li>Before starting the experiment, perform a small-scale pilot experiment to confirm if the genetics work as expected to specifically label the desired cells.
	<ol>
		<li>As a first pass, use immunohistochemistry and light microscopy to determine if unwanted cells are labeled. Ideally, genetic markers will be specific within the retina and optic lobe (Figure 3). Only optic lobes are used for cell dissociation and so labeling in the central brain is not an issue. At the same time, determine how many cells of interest are labeled per brain.</li>
		<li>If the genetic label(s) have the desired specificity, perform a pilot FACS experiment (Figure 4) to determine if a pure population of cells can be reproducibly isolated, and also to assess how many cells of interest can be isolated from each brain. If needed, assess the purity of isolated cells through quantitative PCR to determine the enrichment or de-enrichment of specific genes.</li>
	</ol>
	</li>
	<li>Determine the number of crosses needed to obtain the desired amount of material for the experiment. 
	NOTE: Based on experimental experience, ~17&#8211;25% of fluorescently labeled cells are consistently isolated with this protocol, and high-quality RNA-sequencing data are obtained from as few as 1,000 FACS purified cells (fewer cells may be sufficient).
	<ol>
		<li>To obtain 1,000 cells for RNA-seq analyses, ensure that there are at least 40 cells of interest fluorescently labeled per brain (20 cells per optic lobe). Around ~10 cells of interest should be purified per brain, so dissect 100 brains.</li>
		<li>To obtain ~100 flies at the appropriate stage of development to dissect in each experiment, use ~100 crosses if selecting against balancers (this will vary depending on the genotypes of the parents). If flies are homozygous for the alleles/transgenes of interest fewer crosses can be set.</li>
		<li>Limit the dissection window to 1 h to maximize cellular health and minimize non-specific changes to gene expression and genome organization that can occur after brains are dissected, but this can be adjusted. Determine the number of dissectors required to dissect 100 brains in 1 h, which depends on the skill of the dissectors.</li>
	</ol>
	</li>
</ol>
2. Fly Work
NOTE: Flies are raised at 25 &#176;C with ~50% humidity unless otherwise noted. Below the genotype of females is indicated as Genotype F, and that of males as Genotype M.
<ol>
	<li>Expanding stocks
	<ol>
		<li>Obtain ~100 vials of young flies (no more than a week old) with Genotype F and ~30 vials of Genotype M.</li>
		<li>On Thursday of Week 1, flip 100 vials of Genotype F onto fresh food. Starting on Friday, flip the vials each day through Tuesday of Week 2. These 6 flips will be used to collect virgins. Also on Wednesday of Week 2, flip flies of Genotype M onto fresh food and clear out the parents on Friday of Week 2. These vials will be used to collect males for the crosses.</li>
	</ol>
	</li>
	<li>Collecting virgins
	<ol>
		<li>From Monday of Week 3, collect virgins from the Genotype F vials. Collect 2-3 times a day and store the virgins at 18 &#176;C with ~50% humidity. Repeat this each day through Tuesday of Week 4. It is highly recommended to collect as many virgins as possible, since a lot of them die&#160;before the crosses are set. Typically, ~2,500 virgins should be collected to set 100 crosses.</li>
	</ol>
	</li>
	<li>Setting crosses
	<ol>
		<li>On Friday of Week 4, set the desired numbers of crosses (typically 100&#8211;200 crosses per experiment) with 15 virgins of Genotype F and 7 males of Genotype M for each cross. It is very important to use young and healthy males with intact wings.</li>
		<li>Flip the crosses every day from Sunday through Saturday of Week 5. Supplement the crosses with fresh virgins or males if dead flies are found. It is essential to prevent the food from drying out, so water the vials as needed. Dry food will significantly decrease the yield.</li>
	</ol>
	</li>
	<li>Negative control for FACS sorting
	<ol>
		<li>Include a negative control without the desired cells being labeled for the FACS sorting. Ideally, the negative control should have the reporter (e.g. UAS-GFP) but not the driver (e.g. GAL4), so that basal expression of the reporter (UAS-constructs can be &#34;leaky&#34;) can be determined to set appropriate gating for the FACS sorting. Flip the flies for the negative control at the same time with the experimental flies, and stage them in the same way.</li>
	</ol>
	</li>
	<li>Staging pupae
	<ol>
		<li>To collect cells at a specific stage of pupal development (e.g. 40 h after puparium formation [h APF]), stage the pupae in a 1 h time window to match with the 1 h period allotted for dissections (discussed above). FACS should be performed directly after cell dissociation, which occurs after dissections and takes ~40 min. Thus, the exact time of staging and dissections is determined by FACS availability.</li>
		<li>To obtain pupae at 40 h APF, stage on Monday of Week 6 at a specific time point (e.g., 5 pm) to dissect pupae 40 h later (e.g., 9 am on Wednesday). Use a wet paintbrush to gently pick white pre-pupae and place them on the wall in pre-warmed vials. Only pick the pupae with desired genotype.</li>
	</ol>
	</li>
	<li>Biological replicates
	<ol>
		<li>Do at least 3 biological replicates for each experiment. As a simple way, repeat the experiment three times in the same week (Week 6) using the same crosses. For example, stage pupae for the second and third replicates on Tuesday and Wednesday, respectively. Dissect these pupae on Thursday and Friday, respectively.</li>
	</ol>
	</li>
</ol>
3. Sample Preparation
NOTE: All the reagents used in this protocol are listed in the Table of Materials.
<ol>
	<li>Prepare solutions for dissections and tissue dissociation
	<ol>
		<li>Prepare the proteolytic enzyme blend stock solution (see Table of Materials) by reconstituting the lyophilized enzyme with ddH2O to a concentration of 26 W&#252;nsch units (Wu)/mL. For a vial containing 260 Wu (large vial), add 10 mL of ddH2O to the vial, and make single-use aliquots (4.2 &#181;L are required per dissociation). Store stock solution at -20 &#176;C. Each experiment requires dissociation of two samples (negative control and experimental tissue).</li>
		<li>Prepare 10x Rinaldini&#39;s Solution (RS) and Complete Schneider&#39;s Media (CSM) the day before the dissection (Monday of Week 6). Prepare 5 mL of CSM for each dissector, and about 5 mL of 1x RS and 5 mL of CSM for dissociation of 2 samples (negative control and experimental). Store the 10x RS at 4 &#176;C for up to a month, and CSM at 4 &#176;C for up to a week.
		<ol>
			<li>To prepare 100 mL of 10x RS, dissolve 8 g of NaCl, 200 mg of KCl, 50 mg of NaH2PO4, 1 g of NaHCO3 and 1 g of glucose in 100 mL of ddH2O in a 250 mL beaker. Filter sterilize with a 0.22 mm filter.</li>
			<li>Add 20 mg of L-Glutathione to 1 mL of ddH2O in a microtube.</li>
			<li>To prepare 50 mL of CSM, add 5 mL of heat inactivated bovine serum, 0.1 mL of 10 mg/mL insulin solution, 5 mL of 200 mM L-Glutamine solution, 12.5 &#956;L of 20 mg/mL L-Glutathione, and 1 mL of Penicillin-Streptomycin solution (5,000 units of penicillin and 5 mg of streptomycin/mL) to 38.9 mL of Schneider&#39;s media. Filter sterilize with a 0.22 mm filter.</li>
			<li>Dilute 10x RS with ddH2O to make 1x RS working solution. Prepare 500 &#956;L of 1x RS in non-adhesive microtubes for each sample.</li>
		</ol>
		</li>
		<li>Turn on a water bath and set the temperature at 37 &#176;C. Make sure the temperature is accurate by using a thermometer. The water bath must be at the correct temperature prior to papain activation. It is recommended to set the temperature the evening before a morning dissection.</li>
	</ol>
	</li>
	<li>Prepare papain and proteolytic enzyme blend 
	NOTE: Make papain solution fresh before every dissociation.
	<ol>
		<li>On Tuesday of Week 6 around 45 min before starting the dissections, retrieve 1 vial of papain powder from 4 &#176;C and incubate at room temperature. Also set aside CSM in an Eppendorf tube at room temperature to later add to the papain powder.</li>
		<li>min prior to the start of dissections use a syringe to add the room temperature CSM to the vial of papain (directly through the cap) so that the concentration is 100 units/mL (the amount of powder in each vial is slightly different; for a vial containing 123 units add 1.23 mL of CSM). To mix, first invert the vial to capture any powder stuck on the inside of the cap, and then pipette up and down.</li>
		<li>Add the vial to a beaker containing 37 &#176;C water in the water bath. Make sure the vial of papain solution is not floating. Activate the papain for 30 min at 37 &#176;C.</li>
		<li>After adding the papain to a beaker in the water bath, retrieve an aliquot of proteolytic enzyme blend (26 Wu/mL) from -20 &#176;C and incubate it at room temperature. After 30 min of activation, incubate the papain at room temperature.</li>
	</ol>
	</li>
	<li>Dissections
	<ol>
		<li>On Tuesday of Week 6, dissect the staged pupal brains in cold CSM with clean forceps, and cut off the optic lobes by pinching at the junction of the optic lobe and the central brain.</li>
		<li>Add the lobes to the bottom of a microtube containing 500 &#956;L of 1x RS (one microtube for the experimental tissue and one for the negative control tissue). Always keep the tubes on ice.</li>
		<li>Dissect as many lobes as possible in 1 h. For the negative control tissue, 10 lobes are enough. Pool dissected optic lobes into a single microtube to eliminate tissue loss during transfer between microtubes.</li>
	</ol>
	</li>
	<li>Cell dissociation
	<ol>
		<li>Carefully remove the 1x RS from the microtube with dissected optic lobes using a P200 pipette, leaving enough to cover the lobes. Carefully add 500 &#956;L of 1x RS to the side of the tube, so as to disturb the brains as little as possible. Always keep the tube on ice.</li>
		<li>Let the lobes settle to the bottom. Just once, take up 200 &#956;L and pipette out to mix (lobes should move around). Let the lobes settle again.</li>
		<li>For two samples (experimental and negative control), aliquot 600 &#956;L of activated room temperature papain into a microtube. Add 4.2 &#956;L of room temperature proteolytic enzyme blend into 600 &#956;L papain solution to obtain a final concentration of 0.18 Wu/mL. Mix by pipetting up and down. This is the dissociation solution.</li>
		<li>Carefully remove the 1x RS from each sample and add 300 &#956;L of dissociation solution to the lobes. Incubate the samples at 25 &#176;C, 1,000 rpm for 15 min in a microtube thermomixer.</li>
		<li>At the 5 and 10 min time points, stop the thermomixer and let the lobes sink to the bottom of the microtube. Take up 200 &#956;L of solution and pipet out to mix.</li>
		<li>After 15 min, let the lobes settle to the bottom and carefully remove the dissociation solution from the lobes (try not to disturb the lobes).</li>
		<li>Wash twice with 1x RS. To do this, carefully add 500 &#956;L 1x RS (try not to disturb the lobes). To mix, gently pipet up 200 &#956;L, and then pipet out into the microtube. Let the lobes sink to the bottom and repeat.</li>
		<li>Wash each sample twice with 500 &#956;L of CSM (same as previous step).</li>
		<li>Carefully remove the CSM and add another 200 &#956;L of CSM to the tube. Using a P200 pipette, disrupt the tissue by pipetting up and down without foaming, until the solution is homogenous (i.e., can no longer see any remnants of tissue).</li>
		<li>Filter the cell suspension through a 30 &#956;m mesh cap into a 5 mL FACS tube on ice (use a P200 and make sure the entire suspension goes through). Add another 500 &#956;L of CSM to rinse the dissociation microtube, and filter the solution into the FACS tube.</li>
		<li>Carefully take off the cap of the FACS tube, collect the solution underneath the mesh filter with a P200 and add it to the rest of the suspension in the FACS tube. The samples are ready for FACS.</li>
		<li>Be sure to have microtubes for collecting FACS purified cells ready in advance. For RNA-seq, sort cells from each sample directly into microtubes containing 300 &#956;L of RLT buffer (lysis buffer) from the RNA purification kit (add 1:100 2-Mercaptoethanol before use).</li>
	</ol>
	</li>
	<li>FACS sorting 
	NOTE: After dissociation, immediately sort the cells by FACS (1 h max). Make sure to book FACS sessions accordingly when planning for the experiment. A cell sorter with 100 &#956;m nozzle size is typically used.
	<ol>
		<li>Compare the cell distribution in the negative control and the experimental sample to set up the gating for the sorting. Ideally, establish the gating in the pilot FACS experiment, and the negative control sample will provide confirmation or fine tuning of the pre-established gating. The cells of interest should be well separated from the background cells that are present in both the negative control and the experimental sample (Figure 4) (test this in the pilot experiments).</li>
		<li>Collect the sorted cells in the prepared collection tubes.</li>
	</ol>
	</li>
	<li>RNA purification
	<ol>
		<li>After sorting the cells directly into 300 &#956;L of RLT buffer, pipet up and down to lyse the cells.</li>
		<li>Purify RNA following the standard protocol of Purification of Total RNA from Animal and Human Cells using the purification kit (see Table of Materials) with on-column DNA digestion7. Elute the RNA in 20 &#956;L of RNase-free water.</li>
		<li>Freeze the RNA samples with liquid nitrogen and store at -80 &#176;C.</li>
		<li>For each experiment perform at least 3 replicates for control and experimental conditions. Prepare the cDNA libraries for all samples at the same time and run all the samples on the same flow cell to control for technical variability. Once all the RNA samples are prepared, use a speed vac to reduce the volume to 2&#8211;5 &#956;L (each sample).</li>
	</ol>
	</li>
</ol>
4. Preparing cDNA Libraries and Sequencing by Smart-seq2
NOTE: To minimize technical variability, make all the cDNA libraries at the same time, and sequence them on the same flow cell.
<ol>
	<li>Use an input of 1.5 &#956;L of concentrated RNA to make cDNA libraries by following the standard protocol for Smart-seq217 except for the following modifications: use a different high-fidelity PCR master mix (see Table of Materials) for PCR amplification; use an on-column PCR purification kit (see Table of Materials) instead of magnetic beads to purify PCR products.</li>
	<li>The Smart-seq2 protocol includes a PCR pre-amplification step after reverse transcription as well as a final enrichment PCR step after tagmentation. The number of cycles in the PCR amplification depends on the abundance of the input material. With 1,000 cells as input, typically use 15 cycles in the PCR pre-amplification step and 12 cycles for the final enrichment PCR step.</li>
	<li>Pool the libraries and sequence them on a single flow cell (e.g., the NextSeq platform).</li>
</ol>

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The authors declare that they have no competing financial interests.

This protocol is simple and not technically difficult to execute, but there are several key steps that if overlooked will cause a considerable reduction in cellular yield. (Step 2.3.2.) It is crucial that crosses are healthy, and that the food does not dry out. Regular watering of crosses is essential to maximize the number of flies available for dissection that are of the right genotype and at the correct stage of development. How often crosses need to be watered will vary depending on the food used and the housing cond...

The Drosophila visual system is an outstanding model for studying the genetic basis of development and behavior. It comprises a stereotyped cellular architecture1 and an advanced genetic toolkit for manipulating specific cell types2,3. A major strength of this system is the ability to autonomously interrogate gene function in cell types of interest with single cell resolution, using genetic mosaic methods4,5. We sought to combine these genetic tools with recent advances in next generation sequencing to perform cell type-specific transcriptomic and genomic analyses in single cell clones in genetic mosaic experiments.
To accomplish this, it is essential to develop a robust and efficient method to selectively isolate low abundant cell populations in the brain. Previously, we developed a protocol for isolating specific cell types in the visual system during pupal development through FACS, and determining their transcriptomes using RNA-seq6. In these experiments, the vast majority of cells of a particular type (e.g. R7 photoreceptors) were fluorescently labeled. Using this method, in genetic mosaic experiments, wherein only a subset of cells of the same type are labeled, we failed to isolate enough cells to obtain quality sequencing data. To address this, we sought to increase the cellular yield by improving the cell dissociation protocol.
Our approach was to decrease the total length of the protocol to maximize the health of dissociated cells, improve cellular health by altering dissection and dissociation buffers, and reduce the amount of mechanical disruption to the dissected tissue. We tested the improved protocol7 using mosaic analysis with a repressible cell marker5 (MARCM), which allows generation of single fluorescently labeled clones of particular cell types, that are wild type or mutant for a gene of interest in an otherwise heterozygous fly. Where, under identical conditions, our earlier protocol failed to generate enough material for RNA-seq, the improved protocol was successful. We reproducibly achieve a high cellular yield (~25% of labeled cells) and obtain high quality RNA-seq data from as little as 1,000 cells7.
A number of protocols have been previously described to isolate particular cell types in Drosophila6,8,9,10,11,12,13,14,15,16. These protocols are mostly intended for isolating cells that are abundant within the brain. Our protocol is optimized for isolating low abundant cell populations (fewer than 100 cells per brain) in the visual system using FACS for subsequent transcriptomic and genomic analyses. With this protocol, we aim to provide a way to reproducibly isolate low abundant cells from the fly visual system by FACS and obtaining high quality transcriptome data by RNA-seq.

<table><tbody><tr><td>Liberase TM</td><td>Roche</td><td>5401127001</td><td>Proteolytic enzyme blend</td></tr><tr><td>NaCl</td><td>Sigma-Aldrich</td><td>S3014</td><td></td></tr><tr><td>KCl</td><td>Sigma-Aldrich</td><td>P9541</td><td></td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>S9638</td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>&amp;nbsp;S5761</td><td></td></tr><tr><td>Glucose</td><td>Sigma-Aldrich</td><td>G0350500</td><td></td></tr><tr><td>L-Glutathione</td><td>Sigma-Aldrich</td><td>G6013</td><td></td></tr><tr><td>Heat Inactivated Bovine Serum</td><td>Sigma-Aldrich</td><td>F4135</td><td></td></tr><tr><td>Insulin Solution</td><td>Sigma-Aldrich</td><td>I0516</td><td></td></tr><tr><td>L-Glutamine</td><td>Sigma-Aldrich</td><td>G7513</td><td></td></tr><tr><td>Penicillin-Streptomycin Solution</td><td>Sigma-Aldrich</td><td>P4458</td><td></td></tr><tr><td>Schneider&#039;s Culture Medium</td><td>Gibco</td><td>21720024</td><td></td></tr><tr><td>Papain</td><td>Worthington</td><td>LK003178</td><td></td></tr><tr><td>2-Mercaptoethanol&amp;nbsp;</td><td>Sigma-Aldrich</td><td>M6250-100ML</td><td></td></tr><tr><td>RNeasy Micro Kit</td><td>Qiagen</td><td>74004</td><td>RNA purification kit</td></tr><tr><td>RNase-free DNase</td><td>Qiagen</td><td>79254</td><td></td></tr><tr><td>SuperScript II Reverse Transcriptase</td><td>Life Technologies</td><td>18064-014</td><td></td></tr><tr><td>dNTP Mix</td><td>Life Technologies</td><td>R0191</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt; Solution</td><td>Sigma-Aldrich</td><td>M1028-10X1ML</td><td></td></tr><tr><td>Betaine Solution</td><td>Sigma-Aldrich</td><td>B0300-1VL</td><td></td></tr><tr><td>RNaseOUT</td><td>Life Technologies</td><td>10777-019</td><td></td></tr><tr><td>Q5 High-Fidelity 2x Master Mix</td><td>New England Biolabs</td><td>M0492S</td><td></td></tr><tr><td>MinElute PCR Purification Kit</td><td>Qiagen</td><td>28004</td><td></td></tr><tr><td>Nextera XT DNA Library Prepration Kit</td><td>Illumina</td><td>FC-131-1024</td><td></td></tr><tr><td>Nextera XT Index Kit</td><td>Illumina</td><td>FC-131-1001</td><td></td></tr><tr><td>Test Tube with Cell Strainer Snap Cap</td><td>Falcon</td><td>352235</td><td></td></tr><tr><td>Bottle-Top Vacuum Filter Systems</td><td>Corning</td><td>CLS431153</td><td></td></tr><tr><td>ThermoMixer F1.5</td><td>Eppendorf</td><td>5384000020</td><td></td></tr><tr><td>FACSAria Flow Cytometer</td><td>BD Biosciences</td><td>656700</td><td></td></tr><tr><td>HiSeq 2500 Sequencing System</td><td>Illumina</td><td>SY&amp;ndash;401&amp;ndash;2501</td><td></td></tr></tbody></table>

purification of low-abundant cells in the drosophila visual system

Recent improvements in the sensitivity of next generation sequencing have facilitated the application of transcriptomic and genomic analyses to small numbers of cells. Utilizing this technology to study development in the Drosophila visual system, which boasts a wealth of cell type-specific genetic tools, provides a powerful approach for addressing the molecular basis of development with precise cellular resolution. For such an approach to be feasible, it is crucial to have the capacity to reliably and efficiently purify cells present at low abundance within the brain. Here, we present a method that allows efficient purification of single cell clones in genetic mosaic experiments. With this protocol, we consistently achieve a high cellular yield after purification using fluorescence activated cell sorting (FACS) (~25% of all labeled cells), and successfully performed transcriptomics analyses on single cell clones generated through mosaic analysis with a repressible cell marker (MARCM). This protocol is ideal for applying transcriptomic and genomic analyses to specific cell types in the visual system, across different stages of development and in the context of different genetic manipulations.

General scheme 
A general scheme of the protocol is shown in Figure 1. The protocol is divided into three major parts: fly work, sample preparation, sequencing and data analysis. The &#34;Planning before the experiment&#34; session of the protocol is not included in the general scheme for simplicity.
Timeline 
A calendar of the major parts of the protocol...

Watch this Scientific Journal Video about Purification of Low-abundant Cells in the Drosophila Visual System at JoVE.com

Purification of Low-abundant Cells in the Drosophila Visual System

Here, we present a cell dissociation protocol for efficiently isolating cells present at low abundance within the Drosophila visual system through fluorescence activated cell sorting (FACS).

Recent improvements in the sensitivity of next generation sequencing have facilitated the application of transcriptomic and genomic analyses to small ...

purification-of-low-abundant-cells-in-the-drosophila-visual-system

Research

JoVE Journal

Genetics

6.2K Views.  Harvard Medical School. Here, we present a cell dissociation protocol for efficiently isolating cells present at low abundance within the Drosophila visual system through fluorescence activated cell sorting (FACS).

Video: Purification of Low-abundant Cells in the Drosophila Visual System

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 ...

Developmental Biology

Single Drosophila Ommatidium Dissection and Imaging

The fruit fly Drosophila melanogaster has made invaluable contributions to neuroscience research and has been used widely as a model for neurodegenerative diseases because of its powerful genetics1. The fly eye in particular has been the organ of choice for neurodegeneration research, being the most accessible and life-dispensable part of the Drosophila nervous system. However the major caveat of intact eyes is the difficulty, because of the intense autofluorescence of the pigment, in imaging intracellular events, such as autophagy dynamics2, which are paramount to understanding of neurodegeneration.
We have recently used the dissection and culture of single ommatidia3 that has been essential for our understanding of autophagic dysfunctions in a fly model of Dentatorubro-Pallidoluysian Atrophy (DRPLA)3, 4.
We now report a comprehensive description of this technique (Fig. 1), adapted from electrophysiological studies5, which is likely to expand dramatically the possibility of fly models for neurodegeneration. This method can be adapted to image live subcellular events and to monitor effective drug administration onto photoreceptor cells (Fig. 2). If used in combination with mosaic techniques6-8, the responses of genetically different cells can be assayed in parallel (Fig. 2).

Single Drosophila Ommatidium Dissection and Imaging

The limiting factor in the use of the adult Drosophila eye to study neurodegeneration and cell biology is the difficult imaging of intracellular processes. We describe the dissection of single ommatidia to generate a bona-fide primary neuronal cell culture, which can be subject to drug treatment and advanced imaging.

The fruit fly Drosophila melanogaster has made invaluable contributions to neuroscience research and has been used widely as a model for neurodegenerative ...

Neuroscience

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including lens-secreting cone cells, pigment cells, a bristle cell and eight photoreceptors (PRs) R1-R8 2. The PRs have specialized microvillar structures, the rhabdomeres, which contain light-sensitive pigments, the Rhodopsins (Rhs). The rhabdomeres of six PRs (R1-R6) form a trapezoid and contain Rh1 3 4. The rhabdomeres of R7 and R8 are positioned in tandem in the center of the trapezoid and share the same path of light. R7 and R8 PRs stochastically express different combinations of Rhs in two main subtypes5: In the 'p' subtype, Rh3 in pR7s is coupled with Rh5 in pR8s, whereas in the 'y' subtype, Rh4 in yR7s is associated with Rh6 in yR8s 6 7 8.
Early specification of PRs and development of ommatidia begins in the larval eye-antennal imaginal disc, a monolayer of epithelial cells. A wave of differentiation sweeps across the disc9 and initiates the assembly of undifferentiated cells into ommatidia10-11. The 'founder cell' R8 is specified first and recruits R1-6 and then R7 12-14. Subsequently, during pupal development, PR differentiation leads to extensive morphological changes 15, including rhabdomere formation, synaptogenesis and eventually rh expression.
In this protocol, we describe methods for retinal dissections and immunohistochemistry at three defined periods of retina development, which can be applied to address a variety of questions concerning retinal formation and developmental pathways. Here, we use these methods to visualize the stepwise PR differentiation at the single-cell level in whole mount larval, midpupal and adult retinas (Figure 1).

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The Drosophila retina is a crystal-like lattice composed of a small number of cell types that are generated in a stereotyped manner 1. Its amenability to sophisticated genetic analysis allows the study of complex developmental programs. This protocol describes dissections and immunohistochemistry of retinas at three discrete developmental stages, with a focus on photoreceptor differentiation.

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including ...

Dissection of the Drosophila Pupal Retina for Immunohistochemistry, Western Analysis, and RNA Isolation

The Drosophila pupal retina provides an excellent model system for the study of morphogenetic processes during development. In this paper, we present a reliable protocol for the dissection of the delicate Drosophila pupal retina. Our surgical approach utilizes readily-available microdissection tools to open pupae and precisely extract eye-brain complexes. These can be fixed, subjected to immunohistochemistry, and retinas then mounted onto microscope slides and imaged if the goal is to detect cellular or subcellular structures. Alternatively, unfixed retinas can be isolated from brain tissue, lysed in appropriate buffers and utilized for protein gel electrophoresis or mRNA extraction (to assess protein or gene expression, respectively). Significant practice and patience may be required to master the microdissection protocol described, but once mastered, the protocol enables relatively quick isolation of mainly undamaged retinas.

Dissection of the Drosophila Pupal Retina for Immunohistochemistry, Western Analysis, and RNA Isolation

This paper presents a surgical method for dissecting Drosophila pupal retinas along with protocols for the processing of tissue for immunohistochemistry, western analysis, and RNA-extraction.&#160;

The Drosophila pupal retina provides an excellent model system for the study of morphogenetic processes during development. In this paper, we present a ...

Isolation and Purification of Drosophila Peripheral Neurons by Magnetic Bead Sorting

The Drosophila peripheral nervous system (PNS) is a powerful model for investigating the complex processes of neuronal development and dendrite morphogenesis at the functional and molecular levels. To aid in these analyses, we have developed a strategy for the isolation of a subclass of PNS neurons called dendritic arborization (da) neurons that have been widely used for studying dendrite morphogenesis1,2. These neurons are very difficult to isolate as a pure population, due in part to their extremely low occurrence and their difficult-to-reach location below the tough chitinous larval cuticle. Our newly developed method overcomes these challenges, and is based on a fast and specific cell enrichment using antibody-coated magnetic beads. For our magnetic bead sorting studies, we have used age-matched third instar larvae expressing a mouse CD8 tagged GFP fusion protein (UAS-mCD8-GFP)3 under the control of either the class IV dendritic arborization (da) neuron-specific pickpocket (ppk)-GAL4 driver4 or the control of the pan-da neuron-specific GAL421-7 driver5. Although this protocol has been optimized for isolating PNS cells which are attached to the inner wall of the larval cuticle, by varying a few parameters, the same protocol could be used to isolate many different cell types attached to the cuticle at larval or pupal stages of development (e.g. epithelia, muscle, oenocytes etc.), or other cell types from larval organs depending upon the GAL4-specific driver expression pattern. The RNA isolated by this method is of high quality and can be readily used for downstream genomic analyses such as microarray gene expression profiling studies. This approach offers a powerful new tool to perform studies on isolated Drosophila dendritic arborization (da) neurons thereby providing novel insights into the molecular mechanisms underlying dendrite morphogenesis.

Isolation and Purification of Drosophila Peripheral Neurons by Magnetic Bead Sorting

In this video-article we present a method for the isolation and purification of Drosophila peripheral neurons using a fast magnetic bead assisted cell sorting strategy. RNA obtained from the isolated cells can be readily used for downstream applications including microarray analyses.

The Drosophila peripheral nervous system (PNS) is a powerful model for investigating the complex processes of neuronal development and dendrite ...

Biology

Purification of Endogenous Drosophila Transient Receptor Potential Channels

Drosophila phototransduction is one of the fastest known G protein-coupled signaling pathways. To ensure the specificity and efficiency of this cascade, the calcium (Ca2+)-permeable cation channel, transient receptor potential (TRP), binds tightly to the scaffold protein, inactivation-no-after-potential D (INAD), and forms a large signaling protein complex with eye-specific protein kinase C (ePKC) and phospholipase C&#946;/No receptor potential A (PLC&#946;/NORPA). However, the biochemical properties of the Drosophila TRP channel remain unclear. Based on the assembling mechanism of INAD protein complex, a modified affinity purification plus competition strategy was developed to purify the endogenous TRP channel. First, the purified histidine (His)-tagged NORPA 863-1095 fragment was bound to Ni-beads and used as bait to pull down the endogenous INAD protein complex from Drosophila head homogenates. Then, excessive purified glutathione S-transferase (GST)-tagged TRP 1261-1275 fragment was added to the Ni-beads to compete with the TRP channel. Finally, the TRP channel in the supernatant was separated from the excessive TRP 1261-1275 peptide by size-exclusion chromatography. This method makes it possible to study the gating mechanism of the Drosophila TRP channel from both biochemical and structural angles. The electrophysiology properties of purified Drosophila TRP channels can also be measured in the future.

Purification of Endogenous Drosophila Transient Receptor Potential Channels

Based on the assembling mechanism of the INAD protein complex, in this protocol, a modified affinity purification plus competition strategy was developed to purify the endogenous Drosophila TRP channel.

Drosophila phototransduction is one of the fastest known G protein-coupled signaling pathways. To ensure the specificity and efficiency of this cascade, ...

Biochemistry

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 ...

Enhancing Protein Visualization with Antibody-Mediated Live Imaging

Visualizing Low-Abundance Proteins and Post-Translational Modifications in Living Drosophila Embryos via Fluorescent Antibody Injection

Visualization of proteins in living cells using GFP (Green Fluorescent Protein) and other fluorescent tags has greatly improved understanding of protein localization, dynamics, and function. Compared to immunofluorescence, live imaging more accurately reflects protein localization without potential artifacts arising from tissue fixation. Importantly, live imaging enables quantitative and temporal characterization of protein levels and localization, crucial for understanding dynamic biological processes such as cell movement or division. However, a major limitation of fluorescent tagging approaches is the need for sufficiently high protein expression levels to achieve successful visualization. Consequently, many endogenously tagged fluorescent proteins with relatively low expression levels cannot be detected. On the other hand, ectopic expression using viral promoters can sometimes lead to protein mislocalization or functional alterations in physiological contexts. To address these limitations, an approach is presented that utilizes highly sensitive antibody-mediated protein detection in living embryos, essentially performing immunofluorescence without the need for tissue fixation. As proof of principle, endogenously GFP-tagged Notch receptor that is barely detectable&#160;in living embryos can be successfully visualized after antibody injection. Furthermore, this approach was adapted to visualize post-translational modifications (PTMs) in living embryos, allowing the detection of temporal changes in tyrosine phosphorylation patterns during early embryogenesis and revealing a novel subpopulation of phosphotyrosine (p-Tyr) underneath apical membranes. This approach can be modified to accommodate other protein-specific, tag-specific, or PTM-specific antibodies and should be compatible with other injection-amenable model organisms or cell lines. This protocol opens new possibilities for live imaging of low-abundance proteins or PTMs that were previously challenging to detect using traditional fluorescent tagging methods.

Author Spotlight: Unveiling the Dynamics of Mechanical and Biochemical Signals in Animal Morphogenesis Research 

This protocol describes the customized antibody-based fluorescence labeling and injection into early Drosophila embryos to enable live imaging of low-abundance proteins or post-translational modifications that are challenging to detect using traditional GFP/mCherry-tag approaches.

Visualization of proteins in living cells using GFP (Green Fluorescent Protein) and other fluorescent tags has greatly improved understanding of protein ...

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a plethora of mutant strains and technical tricks have been developed to allow sophisticated questions to be asked and answered in a reasonable amount of time. Fundamental insight into the interplay of components of all known major signaling pathways has been obtained in forward and reverse genetic Drosophila studies. The fly eye has proven to be exceptionally well suited for mutational analysis, since, under laboratory conditions, flies can survive without functional eyes. Furthermore, the surface of the insect eye is composed of some 800 individual unit eyes (facets or ommatidia) that form a regular, smooth surface when looked at under a dissecting microscope. Thus, it is easy to see whether a mutation might affect eye development or growth by externally looking for the loss of the smooth surface ('rough eye' phenotype; Fig. 1) or overall eye size, respectively (for examples of screens based on external eye morphology see e.g.1). Subsequent detailed analyses of eye phenotypes require fixation, plastic embedding and thin-sectioning of adult eyes.
The Drosophila eye develops from the so-called eye imaginal disc, a bag of epithelial cells that proliferate and differentiate during larval and pupal stages (for review see e.g. 2). Each ommatidium consists of 20 cells, including eight photoreceptors (PR or R-cells; Fig. 2), four lens-secreting cone cells, pigment cells ('hexagon' around R-cell cluster) and a bristle. The photoreceptors of each ommatidium, most easily identified by their light sensitive organelles, the rhabdomeres, are organized in a trapezoid made up of the six "outer" (R1-6) and two "inner" photoreceptors (R7/8; R8 [Fig. 2] is underneath R7 and thus only seen in sections from deeper areas of the eye). The trapezoid of each facet is precisely aligned with those of its neighbors and the overall anteroposterior and dorsoventral axes of the eye (Fig. 3A). In particular, the ommatidia of the dorsal and ventral (black and red arrows, respectively) halves of the eye are mirror images of each other and correspond to two chiral forms established during planar cell polarity signaling (for review see e.g. 3).
The method to generate semi-thin eye sections (such as those presented in Fig. 3) described here is slightly modified from the one originally described by Tomlinson and Ready4. It allows the morphological analysis of all cells except for the transparent cone cells. In addition, the pigment of R-cells (blue arrowheads in Fig. 2 and 3) can be used as a cell-autonomous marker for the genotype of a R-cell, thus genetic requirements of genes in a subset of R-cells can readily be determined5,6.

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

A standard approach to prepare adult Drosophila eyes for semi-thin sectioning and light microscopic analysis is presented here. The protocol can be used for gross morphological analysis of eye defects, or with the indicated adjustments can be used to determine genetic requirements of genes in specific cell types of the eye (e.g. clonal analysis of photoreceptors) or for electron microscopic analysis.

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a ...

The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells

The Drosophila eye is widely used as a model for studies of development and neuronal degeneration. With the powerful mitotic recombination technique, elegant genetic screens based on clonal analysis have led to the identification of signaling pathways involved in eye development and photoreceptor (PR) differentiation at larval stages. We describe here the Tomato/GFP-FLP/FRT method, which can be used for rapid clonal analysis in the eye of living adult Drosophila. Fluorescent photoreceptor cells are imaged with the cornea neutralization technique, on retinas with mosaic clones generated by flipase-mediated recombination. This method has several major advantages over classical histological sectioning of the retina: it can be used for high-throughput screening and has proved an effective method for identifying the factors regulating PR survival and function. It can be used for kinetic analyses of PR degeneration in the same living animal over several weeks, to demonstrate the requirement for specific genes for PR survival or function in the adult fly. This method is also useful for addressing cell autonomy issues in developmental mutants, such as those in which the establishment of planar cell polarity is affected.

The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells

The Tomato/GFP-FLP/FRT method involves visualizing mosaic photoreceptor cells in living Drosophila. It can be used to follow individual photoreceptor cell fates in the retina for days or weeks. This method is ideal for studies of retinal degeneration and neurodegenerative diseases or photoreceptor cell development.

The Drosophila eye is widely used as a model  for studies of development and neuronal degeneration. With the powerful mitotic  recombination technique, ...

Purification of Low-abundant Cells in the Drosophila Visual System

Summary

Explore More Videos

Chapters in this video

Live-imaging of the Drosophila Pupal Eye

Single Drosophila Ommatidium Dissection and Imaging

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

Dissection of the Drosophila Pupal Retina for Immunohistochemistry, Western Analysis, and RNA Isolation

Isolation and Purification of Drosophila Peripheral Neurons by Magnetic Bead Sorting

Purification of Endogenous Drosophila Transient Receptor Potential Channels

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

Visualizing Low-Abundance Proteins and Post-Translational Modifications in Living Drosophila Embryos via Fluorescent Antibody Injection

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells