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>NaH2PO4</td>
			<td>Sigma-Aldrich</td>
			<td>S9638</td>
			<td></td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>Sigma-Aldrich</td>
			<td>&#160;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&#39;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&#160;</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>MgCl2 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&#8211;401&#8211;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

Universidad de Cartagena UDEC

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

High Resolution Fluorescent In Situ Hybridization in Drosophila Embryos and Tissues Using Tyramide Signal Amplification

In our efforts to determine the patterns of expression and subcellular localization of Drosophila RNAs on a genome-wide basis, and in a variety of tissues, we have developed numerous modifications and improvements to our original fluorescent in situ hybridization (FISH) protocol. To facilitate throughput and cost effectiveness, all steps, from probe generation to signal detection, are performed using exon 96-well microtiter plates. Digoxygenin (DIG)-labelled antisense RNA probes are produced using either cDNA clones or genomic DNA as templates. After tissue fixation and permeabilization, probes are hybridized to transcripts of interest and then detected using a succession of anti-DIG antibody conjugated to biotin, streptavidin conjugated to horseradish peroxidase (HRP) and fluorescently conjugated tyramide, which in the presence of HRP, produces a highly reactive intermediate that binds to electron dense regions of immediately adjacent proteins. These amplification and localization steps produce a robust and highly localized signal that facilitates both cellular and subcellular transcript localization. The protocols provided have been optimized to produce highly specific signals in a variety of tissues and developmental stages. References are also provided for additional variations that allow the simultaneous detection of multiple transcripts, or transcripts and proteins, at the same time.

High Resolution Fluorescent In Situ Hybridization in Drosophila Embryos and Tissues Using Tyramide Signal Amplification

The described RNA in situ hybridization protocol allows the detection of RNA in whole Drosophila embryos or dissected tissues. Using 96-well microtiter plates and tyramide signal amplification, transcripts can be detected at high resolution, sensitivity, and throughput, and at a relatively low cost.

In our efforts to determine the patterns of expression and subcellular localization of Drosophila RNAs on a genome-wide basis, and in a variety of ...

Production and Purification of Baculovirus for Gene Therapy Application

Baculovirus has traditionally been used for the production of recombinant protein and vaccine. However, more recently, baculovirus is emerging as a promising vector for gene therapy application. Here, baculovirus is produced by transient transfection of the baculovirus plasmid DNA (bacmid) in an adherent culture of Sf9 cells. Baculovirus is subsequently expanded in Sf9 cells in a serum-free suspension culture until the desired volume is obtained. It is then purified from the culture supernatant using heparin affinity chromatography. Virus supernatant is loaded onto the heparin column which binds baculovirus particles in the supernatant due to the affinity of heparin for baculovirus envelop glycoprotein. The column is washed with a buffer to remove contaminants and baculovirus is eluted from the column with a high-salt buffer. The eluate is diluted to an isotonic salt concentration and baculovirus particles are further concentrated using ultracentrifugation. Using this method, baculovirus can be concentrated up to 500-fold with a 25% recovery of infectious particles. Although the protocol described here demonstrates the production and purification of the baculovirus from cultures up to 1 L, the method can be scaled-up in a closed-system suspension culture to produce a clinical-grade vector for gene therapy application.

In this protocol, baculovirus is produced by transient transfection of baculovirus plasmid into Sf9 cells and amplified in a serum-free suspension culture. The supernatant is purified by heparin affinity chromatography and further concentrated by ultracentrifugation. This protocol is useful for the production and purification of baculovirus for gene therapy application.

Baculovirus has traditionally been used for the production of recombinant protein and vaccine. However, more recently, baculovirus is emerging as a ...

CRISPR-Mediated Reorganization of Chromatin Loop Structure

Recent studies have clearly shown that long-range, three-dimensional chromatin looping interactions play a significant role in the regulation of gene expression, but whether looping is responsible for or a result of alterations in gene expression is still unknown. Until recently, how chromatin looping affects the regulation of gene activity and cellular function has been relatively ambiguous, and limitations in existing methods to manipulate these structures prevented in-depth exploration of these interactions. To resolve this uncertainty, we engineered a method for selective and reversible chromatin loop re-organization using CRISPR-dCas9 (CLOuD9). The dynamism of the CLOuD9 system has been demonstrated by successful localization of CLOuD9 constructs to target genomic loci to modulate local chromatin conformation. Importantly, the ability to reverse the induced contact and restore the endogenous chromatin conformation has also been confirmed. Modulation of gene expression with this method establishes the capacity to regulate cellular gene expression and underscores the great potential for applications of this technology in creating stable de novo chromatin loops that markedly affect gene expression in the contexts of cancer and development.

Chromatin looping plays a significant role in gene regulation; however, there have been no technological advances that allow for selective and reversible modification of chromatin loops. Here we describe a powerful system for chromatin loop re-organization using CRISPR-dCas9 (CLOuD9), demonstrated to selectively and reversibly modulate gene expression at targeted loci.

Recent studies have clearly shown that long-range, three-dimensional chromatin looping interactions play a significant role in the regulation of gene ...

Measuring Exercise Levels in Drosophila melanogaster Using the Rotating Exercise Quantification System (REQS)

Drosophila melanogaster is a new model organism for studies in exercise biology. To date, two main exercise systems, the Power Tower and the Treadwheel have been described. However, a method to measure the amount of additional animal activity induced through the exercise treatment has been lacking. The Rotating Exercise Quantification System (REQS) fills this need, providing a measure of animal activity for animals experiencing rotational exercise. This protocol details how to use the REQS to assess animal activity during rotational exercise and illustrates the type of data that can be generated. Here, we demonstrate how the REQS is used to measure sex- and strain-specific differences in exercise induced activity. The REQS can also be used to evaluate the impact of various other experimental parameters such as age, diet, or population size on exercise induced activity. In addition, it can be used to compare the efficacy of different exercise training protocols. Importantly, it provides an opportunity to standardize exercise treatments between strains, allowing the researcher to achieve equal amounts of activity between groups if needed. Thus, the REQS is a notable new resource for exercise biologists working with the Drosophila model system and complements existing exercise systems.

Measuring Exercise Levels in Drosophila melanogaster Using the Rotating Exercise Quantification System REQS

The Rotating Exercise Quantification System (REQS) can induce exercise in Drosophila melanogaster through rotation while simultaneously measuring the amount of activity performed by the animals. Here, we present a point-by-point protocol detailing how to measure activity levels of animals experiencing rotational exercise treatments using the REQS.

Drosophila melanogaster is a new model organism for studies in exercise biology. To date, two main exercise systems, the Power Tower and the Treadwheel ...

Characterizing Histone Post-translational Modification Alterations in Yeast Neurodegenerative Proteinopathy Models

Neurodegenerative diseases, such as amyotrophic lateral sclerosis (ALS) and Parkinson&#39;s disease (PD), cause the loss of hundreds of thousands of lives each year. Effective treatment options able to halt disease progression are lacking. Despite the extensive sequencing efforts in large patient populations, the majority of ALS and PD cases remain unexplained by genetic mutations alone. Epigenetics mechanisms, such as the post-translational modification of histone proteins, may be involved in neurodegenerative disease etiology and progression and lead to new targets for pharmaceutical intervention. Mammalian in vivo and in vitro models of ALS and PD are costly and often require prolonged and laborious experimental protocols. Here, we outline a practical, fast, and cost-effective approach to determining genome-wide alterations in histone modification levels using Saccharomyces cerevisiae as a model system. This protocol allows for comprehensive investigations into epigenetic changes connected to neurodegenerative proteinopathies that corroborate previous findings in different model systems while significantly expanding our knowledge of the neurodegenerative disease epigenome.

This protocol outlines experimental procedures to characterize genome-wide changes in the levels of histone post-translational modifications (PTM) occurring in connection with the overexpression of proteins associated with ALS and Parkinson&#39;s disease in Saccharomyces cerevisiae models. After SDS-PAGE separation, individual histone PTM levels are detected with modification-specific antibodies via Western blotting.

Neurodegenerative diseases, such as amyotrophic lateral sclerosis (ALS) and Parkinson&#39;s disease (PD), cause the loss of hundreds of thousands of lives ...

In vivo Application of the REMOTE-control System for the Manipulation of Endogenous Gene Expression

Here we describe a protocol for implementing the REMOTE-control system (Reversible Manipulation of Transcription at Endogenous loci), which allows for reversible and tunable expression control of an endogenous gene of interest in living model systems. The REMOTE-control system employs enhanced lac repression and tet activation systems to achieve down- or upregulation of a target gene within a single biological system. Tight repression can be achieved from repressor binding sites flexibly located far downstream of a transcription start site by inhibiting transcription elongation. Robust upregulation can be attained by enhancing the transcription of an endogenous gene by targeting tet transcriptional activators to the cognate promoter. This reversible and tunable expression control can be applied and withdrawn repeatedly in organisms. The potency and versatility of the system, as demonstrated for endogenous Dnmt1 here, will allow more precise in vivo functional analyses by enabling investigation of gene function at various expression levels and by testing the reversibility of a phenotype.

This protocol outlines the steps needed to generate a model system in which the transcription of an endogenous gene of interest can be conditionally controlled in live animals or cells using enhanced lac repressor and/or tet activator systems.

Here we describe a protocol for implementing the REMOTE-control system (Reversible Manipulation of Transcription at Endogenous loci), which allows for ...

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Drosophila S2 cells are an important tool in studying mitosis in tissue culture, providing molecular insights into this fundamental cellular process in a rapid and high-throughput manner. S2 cells have proven amenable to both fixed- and live-cell imaging applications. Notably, live-cell imaging can yield valuable information about how loss or knockdown of a gene can affect the kinetics and dynamics of key events during cell division, including mitotic spindle assembly, chromosome congression, and segregation, as well as overall cell cycle timing. Here we utilize S2 cells stably transfected with fluorescently tagged mCherry:&#945;-tubulin to mark the mitotic spindle and GFP:CENP-A (referred to as &#8216;CID&#8217; gene in Drosophila) to mark the centromere to analyze the effects of key mitotic genes on the timing of cell divisions, from prophase (specifically at Nuclear Envelope Breakdown; NEBD) to the onset of anaphase. This imaging protocol also allows for the visualization of the spindle microtubule and chromosome dynamics throughout mitosis. Herein, we aim to provide a simple yet comprehensive protocol that will allow readers to easily adapt S2 cells for live imaging experiments. Results obtained from such experiments should expand our understanding of genes involved in the cell division by defining their role in several simultaneous and dynamic events. Observations made in this cell culture system can be validated and further investigated in vivo using the impressive toolkit of genetic approaches in flies.

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Cell divisions can be visualized in real time using fluorescently tagged proteins and time-lapse microscopy. Using the protocol presented here, users can analyze cell division timing dynamics, mitotic spindle assembly, and chromosome congression and segregation. Defects in these events following RNA interference (RNAi)-mediated gene knockdown can be assessed and quantified.

Drosophila S2 cells are an important tool in studying mitosis in tissue culture, providing molecular insights into this fundamental cellular process in a ...

Investigation of the Transcriptional Role of a RUNX1 Intronic Silencer by CRISPR/Cas9 Ribonucleoprotein in Acute Myeloid Leukemia Cells

The bulk of the human genome (~98%) is comprised of non-coding sequences. Cis-regulatory elements (CREs) are non-coding DNA sequences that contain binding sites for transcriptional regulators to modulate gene expression. Alterations of CREs have been implicated in various diseases including cancer. While promoters and enhancers have been the primary CREs for studying gene regulation, very little is known about the role of silencer, which is another type of CRE that mediates gene repression. Originally identified as an adaptive immunity system in prokaryotes, CRISPR/Cas9 has been exploited to be a powerful tool for eukaryotic genome editing. Here, we present the use of this technique to delete an intronic silencer in the human RUNX1 gene and investigate the impacts on gene expression in OCI-AML3 leukemic cells. Our approach relies on electroporation-mediated delivery of two preassembled Cas9/guide RNA (gRNA) ribonucleoprotein (RNP) complexes to create two double-strand breaks (DSBs) that flank the silencer. Deletions can be readily screened by fragment analysis. Expression analyses of different mRNAs transcribed from alternative promoters help evaluate promoter-dependent effects. This strategy can be used to study other CREs and is particularly suitable for hematopoietic cells, which are often difficult to transfect with plasmid-based methods. The use of a plasmid- and virus-free strategy allows simple and fast assessments of gene regulatory functions.

Investigation of the Transcriptional Role of a RUNX1 Intronic Silencer by CRISPR/Cas9 Ribonucleoprotein in Acute Myeloid Leukemia Cells

Direct delivery of preassembled Cas9/guide RNA ribonucleoprotein complexes is a fast and efficient means for genome editing in hematopoietic cells. Here, we utilize this approach to delete a RUNX1 intronic silencer and examine the transcriptional responses in OCI-AML3 leukemic cells.

The bulk of the human genome (~98%) is comprised of non-coding sequences. Cis-regulatory elements (CREs) are non-coding DNA sequences that contain binding ...

A FACS-based Protocol to Isolate RNA from the Secondary Cells of Drosophila Male Accessory Glands

To understand the function of an organ, it is often useful to understand the role of its constituent cell populations. Unfortunately, the rarity of individual cell populations often makes it difficult to obtain enough material for molecular studies. For example, the accessory gland of the Drosophila male reproductive system contains two distinct secretory cell types. The main cells make up 96% of the secretory cells of the gland, while the secondary cells (SC) make up the remaining 4% of cells (about 80 cells per male). Although both cell types produce important components of the seminal fluid, only a few genes are known to be specific to the SCs. The rarity of SCs has, thus far, hindered transcriptomic analysis study of this important cell type. Here, a method is presented that allows for the purification of SCs for RNA extraction and sequencing. The protocol consists in first dissecting glands from flies expressing a SC-specific GFP reporter and then subjecting these glands to protease digestion and mechanical dissociation to obtain individual cells. Following these steps, individual, living, GFP-marked cells are sorted using a fluorescent activated cell sorter (FACS) for RNA purification. This procedure yields SC-specific RNAs from ~40 males per condition for downstream RT-qPCR and/or RNA sequencing in the course of one day. The rapidity and simplicity of the procedure allows for the transcriptomes of many different flies, from different genotypes or environmental conditions, to be determined in a short period of time.

A FACS-based Protocol to Isolate RNA from the Secondary Cells of Drosophila Male Accessory Glands

Here, we present a protocol to dissociate and sort a specific cell population from the Drosophila male accessory glands (secondary cells) for RNA sequencing and RT-qPCR. Cell isolation is accomplished through FACS purification of GFP-expressing secondary cells after a multistep-dissociation process requiring dissection, proteases digestion and mechanical dispersion.

To understand the function of an organ, it is often useful to understand the role of its constituent cell populations. Unfortunately, the rarity of ...

Efficient Transcriptionally Controlled Plasmid Expression System for Investigation of the Stability of mRNA Transcripts in Primary Alveolar Epithelial Cells

Studying posttranscriptional regulation is fundamental to understanding the modulation of a given messenger RNA (mRNA) and its impact on cell homeostasis and metabolism. Indeed, fluctuations in transcript expression could modify the translation efficiency and ultimately the cellular activity of a transcript. Several experimental approaches have been developed to investigate the half-life of mRNA although some of these methods have limitations that prevent the proper study of posttranscriptional modulation. A promoter induction system can express a gene of interest under the control of a synthetic tetracycline-regulated promoter. This method allows the half-life estimation of a given mRNA under any experimental condition without disturbing cell homeostasis. One major drawback of this method is the necessity to transfect cells, which limits the use of this technique in isolated primary cells that are highly resistant to conventional transfection techniques. Alveolar epithelial cells in primary culture have been used extensively to study the cellular and molecular biology of the alveolar epithelium. The unique characteristics and phenotype of primary alveolar cells make it essential to study the posttranscriptional modulations of genes of interest in these cells. Therefore, our aim was to develop a novel tool to investigate the posttranscriptional modulations of mRNAs of interest in alveolar epithelial cells in primary culture. We designed a fast and efficient transient transfection protocol to insert a transcriptionally controlled plasmid expression system into primary alveolar epithelial cells. This cloning strategy, using a viral epitope to tag the construct, allows for the easy discrimination of construct expression from that of endogenous mRNAs. Using a modified &#916;&#916; quantification cycle (Cq) method, the expression of the transcript can then be quantified at different time intervals to measure its half-life. Our data demonstrate the efficiency of this novel approach in studying posttranscriptional regulation in various pathophysiological conditions in primary alveolar epithelial cells.

Here, we present a tool that can be used to study the posttranscriptional modulation of a transcript in primary alveolar epithelial cells by using an inducible expression system coupled to a pipette electroporation technique.

Studying posttranscriptional regulation is fundamental to understanding the modulation of a given messenger RNA (mRNA) and its impact on cell homeostasis ...

Purification of Low-abundant Cells in the Drosophila Visual System

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Rozdziały w tym wideo

High Resolution Fluorescent In Situ Hybridization in Drosophila Embryos and Tissues Using Tyramide Signal Amplification

Production and Purification of Baculovirus for Gene Therapy Application

CRISPR-Mediated Reorganization of Chromatin Loop Structure

Measuring Exercise Levels in Drosophila melanogaster Using the Rotating Exercise Quantification System (REQS)

Characterizing Histone Post-translational Modification Alterations in Yeast Neurodegenerative Proteinopathy Models

In vivo Application of the REMOTE-control System for the Manipulation of Endogenous Gene Expression

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Investigation of the Transcriptional Role of a RUNX1 Intronic Silencer by CRISPR/Cas9 Ribonucleoprotein in Acute Myeloid Leukemia Cells

A FACS-based Protocol to Isolate RNA from the Secondary Cells of Drosophila Male Accessory Glands

Efficient Transcriptionally Controlled Plasmid Expression System for Investigation of the Stability of mRNA Transcripts in Primary Alveolar Epithelial Cells