This study was supported by the National Institute of Neurological Disorders and Stroke Grant R01NS116463 to J.K., and the Cellular and Molecular Imaging Core facility at the University of Nevada, Reno, which was supported by National Institutes of Health Grant P20GM103650 and used for research reported in this study.

1. Rearing and selecting Drosophila larvae
<ol>
	<li>Maintain/culture the fly strain (w1118, wildtype) on standard fly food medium at 25 &#730;C in a humidified incubator.</li>
	<li>Select 10 wandering 3rd instar larvae 72 h after egg laying.</li>
	<li>Place the larvae in a 35 mm empty Petri dish and gently wash them by transferring the larvae to the new dish containing tap water using forceps. Do this 2x to remove any remaining food.</li>
</ol>
2. Brain isolation from Drosophila larvae (Figure 1)
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66116/66116fig01.jpg" /> 
Figure 1: Dissection of Drosophila larval brain from 3rd instar wandering stage. (A) Schematic drawings of Drosophila larva. (B-G) Larva dissection. <a href="https://www.jove.com/files/ftp_upload/66116/66116fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Place 10 larvae on a dissection dish containing ice-cold PBS.</li>
	<li>Place the larva dorsal side up (identified by tracheal tubes running along its length) and pin each end to the bottom of the dish followed by making a small incision on the body wall at the posterior end.</li>
	<li>Cut the body wall along the dorsal midline towards the anterior end using microdissection scissors.</li>
	<li>Briefly flush the interior of the larva using a Pasteur pipette 3x with PBS in the dish to expose the brain.</li>
	<li>Locate and lift the brain using the forceps and carefully isolate it using microdissection scissors.</li>
	<li>Transfer the dissected brains to a 1.5 mL microcentrifuge tube filled with ice-cold PBS on ice; collect all larva brains. Proceed with RNA extraction using an RNA microprep kit as described in section 4. 
	NOTE: Do a dissection of 10 larvae within 15 min to prevent tissue damage and RNA degradation.</li>
</ol>
3. Drosophila S2 Schneider cells
<ol>
	<li>Grow Drosophila S2 cells in Drosophila Schneider&#39;s medium supplemented with 10% fetal bovine serum (FBS) at 25 &#176;C in a humidified incubator to a density of 8 &#215; 106 to 10 &#215; 106 cells/mL with minimum 90% viability.</li>
	<li>In a 50 mL sterile conical tube, dilute cells to 2.5 &#215; 106 cells/mL with Schneider&#39;s Drosophila medium supplemented with 10% FBS that has been prewarmed to 25 &#176;C.</li>
	<li>Transfer 8 mL of the cell suspension (20 &#215; 106 cells) into a 100 mm culture plate and add 4 mL of medium to make it to 12 mL (day 1).</li>
	<li>Incubate the cultured cells at 25 &#176;C in a humidified incubator. 
	NOTE: Cells loosely adhere to the plate after 12-16 h (day 2).</li>
	<li>Transfect the cells with appropriate DNA plasmids24.</li>
	<li>Incubate for 48 h in a humidified incubator.</li>
	<li>After incubation, collect cells by adding 5 mL of ice-cold PBS by gentle pipetting (day 4).</li>
	<li>Transfer the cells to a 15 mL tube.</li>
	<li>Pellet the cells by centrifuging at 1,000 &#215; g for 5 min at 4 &#176;C.</li>
	<li>Rinse the cells 2x with ice-cold PBS by gentle pipetting and collect the cells by centrifuging them at 1,000 &#215; g for 5 min at 4 &#176;C.</li>
	<li>Perform RNA extraction using an RNA miniprep kit. 
	NOTE: Perform the following steps inside a sterile laminar flow hood.</li>
</ol>
4. Total RNA extraction from Drosophila larvae brain and S2 cells
<ol>
	<li>Larval brain: Remove PBS by brief centrifugation (8 s short spin at 5,000 &#215; g).</li>
	<li>Add 600 &#181;L of RNA lysis buffer and homogenize 10x with a plastic pestle. Visually inspect the tube under a stereo microscope to ensure complete lysis.</li>
	<li>Centrifuge at 1,000 &#215; g for 5 min at 4 &#176;C to remove tissue debris. Transfer the cleared supernatant into a nuclease-free microcentrifuge tube.</li>
	<li>Isolate RNA using an RNA microprep kit according to the manufacturer&#39;s instructions. 
	NOTE: The use of an RNA microprep kit is essential for the larval brain samples because of the small amount of RNA present in the samples.</li>
	<li>S2 cells: Remove PBS and isolate RNA according to the manufacturer&#39;s instructions.</li>
	<li>Measure RNA yield and quality by spectrophotometry and agarose gel electrophoresis.
	<ol>
		<li>Determine the purity and quantity of extracted RNA by measuring the optical density of the extracted RNA at A260 nm and A280 nm, respectively. Make sure the A260 nm/A280 nm ratio is &#8805;2.0 and the RNA concentration is &#62;350 ng/&#181;L for downstream applications. 
		NOTE: A typical RNA yield from 10 Drosophila larva brains is ~500-800 ng/&#181;L or 2.5-4 &#181;g in 5 &#181;L. For S2 cells, the yield is ~2-3 &#181;g/&#181;L (15-30 &#181;g in 15 &#181;L). Isolated RNA can be stored at -80 &#176;C for long-term storage.</li>
	</ol>
	</li>
</ol>
5. Preparation of RNA gel and electrophoresis
<ol>
	<li>1.5% Denaturing RNA gel (100 mL) 
	NOTE: Formaldehyde is toxic through skin contact and inhalation of vapors; handle it in a chemical fume hood.
	<ol>
		<li>Dissolve three agarose tablets (1.5 g) in 82 mL of MOPS buffer (Supplemental File 1) until tablets completely break up to form fine particles.</li>
		<li>Heat the agarose slurry in a microwave until the solution is clear and all particles are completely dissolved.</li>
		<li>Cool the solution to ~60 &#176;C.</li>
		<li>Add 18 mL of 37% formaldehyde, then mix by gentle swirling. Pour the solution into the casting tray and let it solidify in a fume hood.</li>
	</ol>
	</li>
	<li>RNA sample preparation and electrophoresis
	<ol>
		<li>Dilute the RNA sample to 200 ng (in 5 &#181;L) and add 5 &#181;L of 2x RNA loading dye.</li>
		<li>Heat the samples at 70 &#730;C for 5 min in a dry bath.</li>
		<li>Load 2 &#181;L of RNA ladder in the first lane and 10 &#181;L of samples in the adjacent lanes.</li>
		<li>Perform electrophoresis in MOPS buffer at 100 V for 60 min for a 5 x 6 cm gel. 
		NOTE: Adjust the electrophoresis conditions depending on the amplicon sizes.</li>
		<li>Visualize the gel on a UV transilluminator. 
		NOTE: The presence of a single band ~600 nucleotide size indicates intact RNA preparation (see Figure 2A).</li>
	</ol>
	</li>
</ol>
6. Poly(A) tail length measurement
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66116/66116fig02v2.jpg" /> 
Figure 2: RNA sample preparation and the poly(A)-tail assay. (A) The RNA gel images show total RNA from the Drosophila larva brain (left) and S2 cells (right) on a 1.5% formaldehyde agarose gel. Single-stranded RNA ladder sizes are shown in nucleotides on lane M. Note a major RNA banding at ~600 nt, which is from rRNA. (B) Schematics of poly(A)-tail assay. Abbreviation: G/I = guanosine/inosine. <a href="https://www.jove.com/files/ftp_upload/66116/66116fig02largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>GI tailing (Figure 2B)
	<ol>
		<li>Keeping the reagents on ice, prepare the following mixture (20 &#181;L): up to 14 &#181;L of total RNA sample (1 &#181;g), 4 &#181;L of 5x Tail buffer mix, and 2 &#181;L of 10x Tail enzyme mix.</li>
		<li>Incubate at 37 &#176;C for 60 min in a thermocycler.</li>
		<li>Add 1.5 &#181;L of the tail stop solution; keep on ice for 2 min. 
		NOTE: Proceed to reverse transcription or store the GI-tailed RNA samples at -80 &#176;C until ready to proceed to reverse transcription.</li>
	</ol>
	</li>
	<li>Reverse transcription and PCR amplification
	<ol>
		<li>Synthesize cDNA by preparing the mix and incubating it under the conditions described in Supplemental File 1.</li>
		<li>Dilute the cDNA samples and perform PCR to amplify DNA as indicated in Supplemental File 1.</li>
	</ol>
	</li>
</ol>
7. PCR product analysis by agarose gel electrophoresis
<ol>
	<li>Analyze a small portion (2-5 &#181;L) of the PCR products from step 6.2.2 on a 2.5% agarose gel by electrophoresis at 100 V for 45 min for quality control.</li>
	<li>Verify the specificity of PCR for gene-specific and tail-specific reactions by sequencing the gel-extracted PCR bands.</li>
</ol>
8. Capillary electrophoresis
<ol>
	<li>Perform high-resolution gel electrophoresis on 1 &#181;L of PCR products (0.5-5 ng/&#181;L) from gene-specific and poly(A)-specific PCR using the bioanalyzer with a high-sensitivity DNA kit. Look for well-resolved peaks indicating a successful run.</li>
</ol>
9. Data analysis: poly(A) tail length measurement (Figure 3)
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66116/66116fig03v2.jpg" /> 
Figure 3: Poly(A) tail length and peak value measurement. <a href="https://www.jove.com/files/ftp_upload/66116/66116fig03largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Accessing data
	<ol>
		<li>To access the data, open the xad file in the software.</li>
		<li>Select the sample name or the ladder in the tree view panel. 
		NOTE: This will show the result as electropherograms or gel-like images for the selected samples. The lower marker 35 base pair (bp) and upper marker 10,380 bp are the internal standards used to align the ladder data (50-7,000 bp) with data from the sample wells.</li>
		<li>Zoom in and out of the electropherograms and gel-like images to display the details.</li>
	</ol>
	</li>
	<li>Performing peak and smear analysis
	<ol>
		<li>To obtain the peak sizes, open the electropherogram of a selected sample.</li>
		<li>Right-click on the electropherogram and select manual integration to manually select peaks by dragging the horizontal line.</li>
		<li>Observe the peak values in the Peak table. Identify the peak with the largest Peak Height. This is a peak of poly(A) tail length for an individual sample. In the example shown, it is 346 bp.</li>
		<li>Enable the Show/Hide Setpoints icon on the top menu and wait for a new panel to pop up on the right side.</li>
		<li>Select Advanced, scroll down to find Perform Smear Analysis, and select the check box. This will add the Region table to the electropherogram tab.</li>
		<li>Select an electropherogram from a sample and go to the Region table, which shows the From [bp] and To [bp] menu. To set the starting and ending [bp], right-click on the electropherogram and select Region to add Add Region.</li>
		<li>Right-click on any cells in the Region table and select Modify Regions to make a small new window pop up where custom regions can be set. 
		NOTE: For example, we used a region of 300 bp to 550 bp for GAPDH. The gene-specific GAPDH PCR yielded a peak at 265 bp. The universal primer (Table 1) extends the length of poly(A) PCR by 35 bp via annealing to G/I-tailed RNAs. Thus, the first adenine nucleotide on GAPDH RNA starts at 300 bp (265 + 35). We arbitrarily limited the maximum poly(A) tail length to 250 (300 + 250 = 550). From the region table, the program returns the average size within the region as 387 bp.</li>
		<li>Use equation (1) to calculate the poly(A) tail length on the mRNA of interest: 
		Poly(A) tail length = (A - B - 35) (1) 
		Where A is the average bp of poly(A)-specific PCR product from electropherogram (i.e., 387 bp for GAPDH), B is the peak bp of gene-specific PCR product from electropherogram (i.e., 265 bp for GAPDH), and &#34;35&#34; is the length of universal reverse primer tag. 
		NOTE: From the calculation above, the average poly(A) tail length of GAPDH is 387 - 265 - 35= 87 bp.</li>
	</ol>
	</li>
</ol>
10. Visualizing poly(A) tail length distribution
<ol>
	<li>Export data as a csv file under File | Export | Sample Data to retrieve sample data. 
	NOTE: The exported csv file displays run time rather than bp on the X-axis.</li>
	<li>Go to an electropherogram of a sample and check on Show Sizes on the Electropherogram tab to automatically convert the region table on Region Table from bp to run time. In the example, 70.39 s to 86.28 s correspond to 300 bp to 550 bp.</li>
	<li>Open the csv file and select the values from 70.39 s to 86.28 s of run time to generate a graph. To visualize bp sizes to the X-axis on the graph, export the electropherogram with bp sizes as an image file and overlay it on the graph generated in the spreadsheet. This will appropriately match the bp sizes on poly(A) tail distribution.</li>
</ol>

Quantifying the Poly(A) Length of Genes from the Drosophila Nervous System

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The authors have no conflicts of interest to disclose.

In this protocol, we describe the technique to dissect the Drosophila larval brain from wandering 3rd instar stage as well as the sample preparation from Drosophila S2 cells. Due to the labile nature of mRNAs, sample collection requires extra caution. For larval brain dissection, brains should not be damaged during isolation and should not be kept in solution for a prolonged duration. Keeping dissection time to 8-10 min for a round of dissection is essential. It may also be beneficial to supp...

Most eukaryotic mRNAs are posttranscriptionally polyadenylated at their 3&#8242; terminus in the nucleus by the addition of non-templated adenosines by canonical poly(A) polymerases. An intact poly(A) tail is pivotal throughout the lifecycle of mRNA, as it is essential for mRNA nuclear export1, facilitates interaction with poly(A)-binding proteins to enhance translational efficiency2, and imparts resistance against degradation3. In certain cases, the poly(A) tail can also undergo extension in the cytoplasm, facilitated by noncanonical poly(A) polymerases4. In the cytoplasm, poly (A) tail length dynamically changes and influences the life span of the mRNA molecule. Numerous polymerases and deadenylases are known for modulating tail length5,6,7. For example, the shortening of poly(A) tails correlates with translational repression, whereas the lengthening of poly(A) tails enhances translation8,9.
Accumulating genomic studies have demonstrated the fundamental significance of the poly(A) tail length across various facets of eukaryotic biology. This includes roles in germ-cell development, early embryonic development, neuronal synaptic plasticity for learning and memory, and the inflammatory response10. There have been numerous methods and assays developed for measuring poly(A) tail lengths. For example, the RNase H/oligo(dT) assay takes advantage of RNase H in the presence or absence of oligo(dT) to study poly(A) tail length11,12. Other methods to study poly(A) tail include the PCR amplification of 3&#39; ends such as rapid amplification of cDNA ends poly(A) test (RACE-PAT)12,13 and the ligase-mediated poly(A) test (LM-PAT)14. Further modifications of the PAT assay include ePAT15 and sPAT16. Enzymatic G-tailing17,18 or G/I-tailing&#160;of the 3&#39; end are other variations of the PAT assay. Further modification of these techniques includes the use of fluorescently labeled primers along with capillary gel electrophoresis for high-resolution analysis, referred to as the high-resolution poly(A) test (Hire-PAT)19. These PCR-driven assays allow fast and high-sensitivity poly(A) length quantitation.
With the development of next-generation sequencing, a high-throughput sequencing method, such as PAL-seq20 and TAIL-seq21, allows polyadenylation analyses at a transcriptome-wide scale. However, these methods provide only short sequencing reads of 36-51 nucleotides. Therefore, FLAM-Seq22 was developed for global tail length profiling of full-length mRNA and provides long reads. Nanopore technology23 provides PCR-independent, direct RNA, or direct cDNA sequencing for poly(A) tail length estimations. However, these high-throughput methods are not without limitations. They require large amounts of starting materials, are expensive, and time-consuming. Moreover, analyzing rare transcripts can be extremely challenging with high-throughput methods, and low-throughput PCR-based methods still provide an advantage when a small number of transcripts need to be analyzed, for pilot experiments, and validation of other methods.
We have recently demonstrated that Dscam1 mRNAs contain short poly(A) tails in Drosophila, which necessitates a non-canonical binding of the cytoplasmic poly(A)-binding protein on Dscam1 3&#39;UTR using the G/I tailing method24. Here we provide a streamlined procedure for tissue preparation and quantifying poly(A) length of mRNAs from the Drosophila nervous system and Drosophila S2 cells.

<table><tbody><tr><td>3-(N-morpholino) propanesulfonic acid (MOPS)</td><td>Research Product Internation (RPI)</td><td>M92020</td><td></td></tr><tr><td>Agilent High Sensitivity DNA Kit</td><td>Agilent Technologies</td><td>&amp;nbsp;5067-4626</td><td></td></tr><tr><td>Agilent software 2100 expert free download demo</td><td>Agilent Technologies</td><td></td><td>https://www.agilent.com/en/product/automated-electrophoresis/bioanalyzer-systems/bioanalyzer-software/2100-expert-software-228259</td></tr><tr><td>Apex 100 bp-Low DNA Ladder</td><td>Genesee Scientific</td><td>19-109</td><td></td></tr><tr><td>Bioanalyzer</td><td>Agilent 2100 Bioanalyzer G2938C</td><td></td><td></td></tr><tr><td>Diethyl pyrocarbonate (DEPC)</td><td>Research Product Internation (RPI)&amp;nbsp;</td><td>D43060</td><td></td></tr><tr><td>DNA dye (Gel Loading Dye, Purple (6x)</td><td>New England biolabs&amp;nbsp;</td><td>B7024S</td><td></td></tr><tr><td>&lt;em&gt;Drosophila &lt;/em&gt;S2 cell line</td><td>Drosophila Genomics Resource Center stock #181</td><td></td><td></td></tr><tr><td>Drosophila Schneider&amp;rsquo;s Medium</td><td>Thermo Fisher Scientific</td><td>21720024</td><td></td></tr><tr><td>Ehidium bromide</td><td>Genesee scientific&amp;nbsp;</td><td>20-276</td><td></td></tr><tr><td>Fetal bovine serum (FBS)</td><td>Sigma-Aldrich</td><td>F4135</td><td></td></tr><tr><td>Forceps Dumont 5&amp;nbsp;&amp;nbsp;</td><td>Fine Science tools&amp;nbsp;</td><td>11254-20</td><td></td></tr><tr><td>Nuclease free water</td><td>Thermo Fisher Scientific&amp;nbsp;</td><td>AM9932</td><td></td></tr><tr><td>PBS 10x</td><td>Research Product Internation (RPI)&amp;nbsp;</td><td>P32200</td><td></td></tr><tr><td>Poly(A) Tail-Length Assay Kit</td><td>Thermo Fisher Scientific&amp;nbsp;</td><td>764551KT</td><td></td></tr><tr><td>RiboRuler Low Range RNA Ladder</td><td>Thermo Fisher Scientific&amp;nbsp;</td><td>SM1833</td><td></td></tr><tr><td>RNA Gel Loading Dye (2x)</td><td>Thermo Fisher Scientific&amp;nbsp;</td><td>R0641</td><td></td></tr><tr><td>RNA microprep kit</td><td>Zymoresearch&amp;nbsp;</td><td>R1050&amp;nbsp;</td><td></td></tr><tr><td>RNA miniprep kit</td><td>Zymoresearch&amp;nbsp;</td><td>R1055</td><td></td></tr><tr><td>Scissors-Vannas Spring Scissors - 2.5 mm Cutting Edge</td><td>Fine Science tools&amp;nbsp;</td><td>15000-08</td><td></td></tr><tr><td>TopVision Agarose Tablets</td><td>Thermo Fisher Scientific</td><td>R2802</td><td></td></tr><tr><td>Tris-Acetate-EDTA (TAE)</td><td>Thermo Fisher Scientific</td><td>B49</td><td></td></tr></tbody></table>

measurement of poly a tail length from drosophila larva brain and cell line

Polyadenylation is a crucial posttranscriptional modification that adds poly(A) tails to the 3' end of mRNA molecules. The length of the poly(A) tail is tightly regulated by cellular processes. Dysregulation of mRNA polyadenylation has been associated with abnormal gene expression and various diseases, including cancer, neurological disorders, and developmental abnormalities. Therefore, comprehending the dynamics of polyadenylation is vital for unraveling the complexities of mRNA processing and posttranscriptional gene regulation. 
This paper presents a method for measuring poly(A) tail lengths in RNA samples isolated from Drosophila larval brains and Drosophila Schneider S2 cells. We employed the guanosine/inosine (G/I) tailing approach, which involves the enzymatic addition of G/I residues at the 3' end of mRNA using yeast poly(A) polymerase. This modification protects the RNA's 3' end from enzymatic degradation. The protected full-length poly(A) tails are then reverse-transcribed using a universal antisense primer. Subsequently, PCR amplification is performed using a gene-specific oligo that targets the gene of interest, along with a universal sequence oligo used for reverse transcription. 
This generates PCR products encompassing the poly(A) tails of the gene of interest. Since polyadenylation is not a uniform modification and results in tails of varying lengths, the PCR products display a range of sizes, leading to a smear pattern on agarose gel. Finally, the PCR products are subjected to high-resolution capillary gel electrophoresis, followed by quantification using the sizes of the poly(A) PCR products and the gene-specific PCR product. This technique offers a straightforward and reliable tool for analyzing poly(A) tail lengths, enabling us to gain deeper insights into the intricate mechanisms governing mRNA regulation.

Here, we analyzed the poly(A) tail length of Dscam1 and GAPDH from Drosophila larval brains (Figure 4). Isolated RNAs were visualized on an agarose gel for quality control. A single RNA band at around 600 nucleotide size indicates intact RNA preparation (Figure 2A). RNAs were subjected to the G/I tailing and high-resolution capillary electrophoresis using an Agilent 2100 bioanalyzer. The gel images were exported using the Agilent 2100 ...

Watch this Scientific Journal Video about Measurement of Poly A Tail Length from Drosophila Larva Brain and Cell Line at JoVE.com

Author Spotlight: Exploring the Frontier of mRNA Research with Poly A Tail Analysis Techniques

The protocol describes an efficient and reliable method for quantifying the poly(A) length of the gene of interest from the Drosophila nervous system, which can be easily adapted to tissues or cell types from other species.

Measurement of Poly A Tail Length from Drosophila Larva Brain and Cell Line

Polyadenylation is a crucial posttranscriptional modification that adds poly(A) tails to the 3' end of mRNA molecules. The length of the poly(A) tail is ...

measurement-poly-tail-length-from-drosophila-larva-brain-cell

Research

JoVE Journal

Neuroscience

840 Views.  University of Nevada, Reno. The protocol describes an efficient and reliable method for quantifying the poly(A) length of the gene of interest from the Drosophila nervous system, which can be easily adapted to tissues or cell types from other species. 

Video: Author Spotlight: Exploring the Frontier of mRNA Research with Poly A Tail Analysis Techniques

Laser Capture Microdissection of Drosophila Peripheral Neurons

The dendritic arborization (da) neurons of the Drosophila peripheral nervous system (PNS) provide an excellent model system in which to investigate the molecular mechanisms underlying class-specific dendrite morphogenesis1,2. To facilitate molecular analyses of class-specific da neuron development, it is vital to obtain these cells in a pure population. Although a range of different cell, and tissue-specific RNA isolation techniques exist for Drosophila cells, including magnetic bead based cell purification3,4, Fluorescent Activated Cell Sorting (FACS)5-8, and RNA binding protein based strategies9, none of these methods can be readily utilized for isolating single or multiple class-specific Drosophila da neurons with a high degree of spatial precision. Laser Capture Microdissection (LCM) has emerged as an extremely powerful tool that can be used to isolate specific cell types from tissue sections with a high degree of spatial resolution and accuracy. RNA obtained from isolated cells can then be used for analyses including qRT-PCR and microarray expression profiling within a given cell type10-16. To date, LCM has not been widely applied in the analysis of Drosophila tissues and cells17,18, including da neurons at the third instar larval stage of development. 
Here we present our optimized protocol for isolation of Drosophila da neurons using the infrared (IR) class of LCM. This method allows for the capture of single, class-specific or multiple da neurons with high specificity and spatial resolution. Age-matched third instar larvae expressing a UAS-mCD8::GFP19 transgene under the control of either the class IV da neuron specific ppk-GAL420 driver or the pan-da neuron specific 21-7-GAL421 driver were used for these experiments. RNA obtained from the isolated da neurons is of very high quality and can be directly used for downstream applications, including qRT-PCR or microarray analyses. Furthermore, this LCM protocol can be readily adapted to capture other Drosophila cell types a various stages of development dependent upon the cell type specific, GAL4-driven expression pattern of GFP.

Laser Capture Microdissection of Drosophila Peripheral Neurons

In this video-article we present a method for isolating single or multiple Drosophila da neurons from third instar larvae using the infrared capture (IR) class of Laser Capture Microdissection (LCM). RNA obtained from the isolated neurons can be readily used for downstream applications including qRT-PCR or microarray analyses.

The dendritic arborization (da) neurons of the Drosophila peripheral nervous system (PNS) provide an excellent model system in which to investigate the ...

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 expression is rarely tissue-specific enough to allow for precise correlation of the circuit with behavioral readouts. To overcome this major hurdle, we recently developed the FINGR method to achieve a more restrictive Gal4 expression in the tissue of interest. The FINGR method has three components: 1) the traditional Gal4/UAS system; 2) a set of FLP/FRT-mediated Gal80 converting tools; and 3) enhancer-trap FLP (ET-FLP). Gal4 is used to define the primary neural circuitry of interest. Paring the Gal4 with a UAS-effector, such as UAS-MJD78Q or UAS-Shits, regulates the neuronal activity, which is in turn manifested by alterations in the fly behavior. With an additional UAS-reporter such as UAS-GFP, the neural circuit involved in the specific behavior can be simultaneously mapped for morphological analysis. For Gal4 lines with broad expression, Gal4 expression can be restricted by using two complementary Gal80-converting tools: tubP&gt;Gal80&gt; ('flip out') and tubP&gt;stop&gt;Gal80 ('flip in'). Finally, investigators can turn Gal80 on or off, respectively, with the help of tissue-specific ET-FLP. In the flip-in mode, Gal80 will repress Gal4 expression wherever Gal4 and ET-FLP intersect. In the flip-out mode, Gal80 will relieve Gal4 repression in cells in which Gal4 and FLP overlap. Both approaches enable the restriction of the number of cells in the Gal4-defined circuitry, but in an inverse pattern. The FINGR method is compatible with the vast collection of Gal4 lines in the fly community and highly versatile for traditional clonal analysis and for neural circuit mapping. In this protocol, we demonstrate the mapping of FLP expression patterns in select ET-FLPx2 lines and the effectiveness of the FINGR method in photoreceptor cells. The principle of the FINGR method should also be applicable to other genetic model organisms in which Gal4/UAS, Gal80, and FLP/FRT are used.

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

We describe a Flippase-induced intersectional Gal80/Gal4 repression (FINGR) method, allowing tissue-specific FLP to determine Gal80 expression patterns. Wherever Gal4 and FLP overlap, Gal4 expression is turned on (Gal80 flipped out) or off (Gal80 flipped in). The FINGR method is versatile for clonal analysis and neural circuit mapping.

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 ...

Imaging Live Brain Explants from Drosophila melanogaster Third Instar Larvae

Live-Cell Imaging of Drosophila melanogaster Third Instar Larval Brains

Drosophila neural stem cells (neuroblasts, NBs hereafter) undergo asymmetric divisions, regenerating the self-renewing neuroblast, while also forming a differentiating ganglion mother cell (GMC), which will undergo one additional division to give rise to two neurons or glia. Studies in NBs have uncovered the molecular mechanisms underlying cell polarity, spindle orientation, neural stem cell self-renewal, and differentiation. These asymmetric cell divisions are readily observable via live-cell imaging, making larval NBs ideally suited for investigating the spatiotemporal dynamics of asymmetric cell division in living tissue. When properly dissected and imaged in nutrient-supplemented medium, NBs in explant brains robustly divide for 12-20 h. Previously described methods are technically difficult and may be challenging to those new to the field. Here, a protocol is described for the preparation, dissection, mounting, and imaging of live third-instar larval brain explants using fat body supplements. Potential problems are also discussed, and examples are provided for how this technique can be used.

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

Here, we discuss a workflow to prepare, dissect, mount, and image live explant brains from Drosophila melanogaster third instar larvae to observe the cellular and subcellular dynamics under physiological conditions.

Drosophila neural stem cells (neuroblasts, NBs hereafter) undergo asymmetric divisions, regenerating the self-renewing neuroblast, while also forming a ...

Developmental Biology

Monitoring Cell-to-cell Transmission of Prion-like Protein Aggregates in Drosophila Melanogaster

Protein aggregation is a central feature of most neurodegenerative diseases, including Alzheimer&#39;s disease (AD), Parkinson&#39;s disease (PD), Huntington&#39;s disease (HD), and amyotrophic lateral sclerosis (ALS). Protein aggregates are closely associated with neuropathology in these diseases, although the exact mechanism by which aberrant protein aggregation disrupts normal cellular homeostasis is not known. Emerging data provide strong support for the hypothesis that pathogenic aggregates in AD, PD, HD, and ALS have many similarities to prions, which are protein-only infectious agents responsible for the transmissible spongiform encephalopathies. Prions self-replicate by templating the conversion of natively-folded versions of the same protein, causing spread of the aggregation phenotype. How prions and prion-like proteins in AD, PD, HD, and ALS move from one cell to another is currently an area of intense investigation. Here, a Drosophila melanogaster model that permits monitoring of prion-like, cell-to-cell transmission of mutant huntingtin (Htt) aggregates associated with HD is described. This model takes advantage of powerful tools for manipulating transgene expression in many different Drosophila tissues and utilizes a fluorescently-tagged cytoplasmic protein to directly report prion-like transfer of mutant Htt aggregates. Importantly, the approach we describe here can be used to identify novel genes and pathways that mediate spreading of protein aggregates between diverse cell types in vivo. Information gained from these studies will expand the limited understanding of the pathogenic mechanisms that underlie neurodegenerative diseases and reveal new opportunities for therapeutic intervention.

Monitoring Cell-to-cell Transmission of Prion-like Protein Aggregates in Drosophila Melanogaster

Accumulating evidence supports the idea that pathogenic protein aggregates associated with neurodegenerative diseases spread between cells with prion-like properties. Here, we describe a method that enables visualization of cell-to-cell spreading of prion-like aggregates in the model organism, Drosophila melanogaster.

Protein aggregation is a central feature of most neurodegenerative diseases, including Alzheimer&#39;s disease (AD), Parkinson&#39;s disease (PD), ...

Quantitative Analysis of Neuronal Dendritic Arborization Complexity in Drosophila

Dendrites are the branched projections of a neuron, and dendritic morphology reflects synaptic organization during the development of the nervous system. Drosophila larval neuronal dendritic arborization (da) is an ideal model for studying morphogenesis of neural dendrites and gene function in the development of nervous system. There are four classes of da neurons. Class IV is the most complex with a branching pattern that covers almost the entire area of the larval body wall. We have previously characterized the effect of silencing the Drosophila ortholog of SOX5 on class IV neuronal dendritic arborization complexity (NDAC) using four parameters: the length of dendrites, the surface area of dendrite coverage, the total number of branches, and the branching structure. This protocol presents the workflow of NDAC quantitative analysis, consisting of larval dissection, confocal microscopy, and image analysis procedures using ImageJ software. Further insight into da neuronal development and its underlying mechanisms will improve the understanding of neuronal function and provide clues about the fundamental causes of neurological and neurodevelopmental disorders.

Quantitative Analysis of Neuronal Dendritic Arborization Complexity in Drosophila

This protocol focuses on quantitative analysis of neuronal dendritic arborization complexity (NDAC) in Drosophila, which can be used for studies of dendritic morphogenesis.

Dendrites are the branched projections of a neuron, and dendritic morphology reflects synaptic organization during the development of the nervous system. ...

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

With the rising prevalence of neurodegenerative diseases, it is increasingly important to understand the underlying pathophysiology that leads to neuronal dysfunction and loss. Fluorescence-based imaging tools and technologies enable unprecedented analysis of subcellular neurobiological processes, yet there is still a need for unbiased, reproducible, and accessible approaches for extracting quantifiable data from imaging studies. We have developed a simple and adaptable workflow to extract quantitative data from fluorescence-based imaging studies using Drosophila models of neurodegeneration. Specifically, we describe an easy-to-follow, semi-automated approach using Fiji/ImageJ to analyze two cellular processes: first, we quantify protein aggregate content and profile in the Drosophila optic lobe using fluorescent-tagged mutant huntingtin proteins; and second, we assess autophagy-lysosome flux in the Drosophila visual system with ratiometric-based quantification of a tandem fluorescent reporter of autophagy. Importantly, the protocol outlined here includes a semi-automated segmentation step to ensure all fluorescent structures are analyzed to minimize selection bias and to increase resolution of subtle comparisons. This approach can be extended for the analysis of other cell biological structures and processes implicated in neurodegeneration, such as proteinaceous puncta (stress granules and synaptic complexes), as well as membrane-bound compartments (mitochondria and membrane trafficking vesicles). This method provides a standardized, yet adaptable reference point for image analysis and quantification, and could facilitate reliability and reproducibility across the field, and ultimately enhance mechanistic understanding of neurodegeneration.

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

We have developed a simple and adaptable workflow to extract quantitative data from fluorescence-imaging-based cell biological studies of protein aggregation and autophagic flux in the central nervous system of Drosophila models of neurodegeneration.

With the rising prevalence of neurodegenerative diseases, it is increasingly important to understand the underlying pathophysiology that leads to neuronal ...

Purification of Transcripts and Metabolites from Drosophila Heads

For the last decade, we have tried to understand the molecular and cellular mechanisms of neuronal degeneration using Drosophila as a model organism. Although fruit flies provide obvious experimental advantages, research on neurodegenerative diseases has mostly relied on traditional techniques, including genetic interaction, histology, immunofluorescence, and protein biochemistry. These techniques are effective for mechanistic, hypothesis-driven studies, which lead to a detailed understanding of the role of single genes in well-defined biological problems. However, neurodegenerative diseases are highly complex and affect multiple cellular organelles and processes over time. The advent of new technologies and the omics age provides a unique opportunity to understand the global cellular perturbations underlying complex diseases. Flexible model organisms such as Drosophila are ideal for adapting these new technologies because of their strong annotation and high tractability. One challenge with these small animals, though, is the purification of enough informational molecules (DNA, mRNA, protein, metabolites) from highly relevant tissues such as fly brains. Other challenges consist of collecting large numbers of flies for experimental replicates (critical for statistical robustness) and developing consistent procedures for the purification of high-quality biological material. Here, we describe the procedures for collecting thousands of fly heads and the extraction of transcripts and metabolites to understand how global changes in gene expression and metabolism contribute to neurodegenerative diseases. These procedures are easily scalable and can be applied to the study of proteomic and epigenomic contributions to disease.

Purification of Transcripts and Metabolites from Drosophila Heads

We describe here the procedures for the extraction and purification of mRNA and metabolites from Drosophila heads. We are applying these techniques to better understand the cellular perturbations underlying neuronal degeneration. These methodologies can be easily scaled and adapted for other "omic" projects.

For the last decade, we have tried to understand the molecular and cellular mechanisms of neuronal degeneration using Drosophila as a model organism. ...

Biology

3D Particle Tracking for Noninvasive In Vivo Analysis of Synaptic Microtubule Dynamics in Dendrites and Neuromuscular Junctions of Drosophila

Microtubules (MTs) play critical roles in neuronal development, but many questions remain about the molecular mechanisms of their regulation and function. Furthermore, despite progress in understanding postsynaptic MTs, much less is known about the contributions of presynaptic MTs to neuronal morphogenesis. In particular, studies of in vivo MT dynamics in Drosophila sensory dendrites yielded significant insights into polymer-level behavior. However, the technical and analytical challenges associated with live imaging of the fly neuromuscular junction (NMJ) have limited comparable studies of presynaptic MT dynamics. Moreover, while there are many highly effective software strategies for automated analysis of MT dynamics in vitro and ex vivo, in vivo data often necessitate significant operator input or entirely manual analysis due to inherently inferior signal-to-noise ratio in images and complex cellular morphology. &#160;To address this, this study optimized a new software platform for automated and unbiased in vivo particle detection. Multiparametric analysis of live time-lapse confocal images of EB1-GFP labeled MTs was performed in both dendrites and the NMJ of Drosophila larvae and found striking differences in MT behaviors. MT dynamics were furthermore analyzed following knockdown of the MT-associated protein (MAP) dTACC, a key regulator of Drosophila synapse development, and identified statistically significant changes in MT dynamics compared to wild type. These results demonstrate that this novel strategy for the automated multiparametric analysis of both pre- and postsynaptic MT dynamics at the polymer-level significantly reduces human-in-the-loop criteria. The study furthermore shows the utility of this method in detecting distinct MT behaviors upon dTACC-knockdown, indicating a possible future application for functional screens of factors that regulate MT dynamics in vivo. Future applications of this method may also focus on elucidating cell type and/or compartment-specific MT behaviors, and multicolor correlative imaging of EB1-GFP with other cellular and subcellular markers of interest.&#160;

This study presents a noninvasive intravital neuronal imaging strategy combined with a new software strategy to achieve automated, unbiased tracking and analysis of in vivo microtubule (MT) plus-end dynamics in the sensory dendrites and the neuromuscular junctions of Drosophila.

Microtubules (MTs) play critical roles in neuronal development, but many questions remain about the molecular mechanisms of their regulation and function. ...

Measuring Glucose Uptake in Drosophila Models of TDP-43 Proteinopathy

Amyotrophic lateral sclerosis is a neurodegenerative disorder causing progressive muscle weakness and death within 2-5 years following diagnosis. Clinical manifestations include weight loss, dyslipidemia, and hypermetabolism; however, it remains unclear how these relate to motor neuron degeneration. Using a Drosophila model of TDP-43 proteinopathy that recapitulates several features of ALS including cytoplasmic inclusions, locomotor dysfunction, and reduced lifespan, we recently identified broad ranging metabolic deficits. Among these, glycolysis was found to be upregulated and genetic interaction experiments provided evidence for a compensatory neuroprotective mechanism. Indeed, despite upregulation of phosphofructokinase, the rate limiting enzyme in glycolysis, an increase in glycolysis using dietary and genetic manipulations was shown to mitigate locomotor dysfunction and increased lifespan in fly models of TDP-43 proteinopathy. To further investigate the effect on TDP-43 proteinopathy on glycolytic flux in motor neurons, a previously reported genetically encoded, FRET-based sensor, FLII12Pglu-700&#181;&#948;6, was used. This sensor is comprised of a bacterial glucose-sensing domain and cyan and yellow fluorescent proteins as the FRET pair. Upon glucose binding, the sensor undergoes a conformational change allowing FRET to occur. Using FLII12Pglu-700&#181;&#948;6, glucose uptake was found to be significantly increased in motor neurons expressing TDP-43G298S, an ALS causing variant. Here, we show how to measure glucose uptake, ex vivo, in larval ventral nerve cord preparations expressing the glucose sensor FLII12Pglu-700&#181;&#948;6 in the context of TDP-43 proteinopathy. This approach can be used to measure glucose uptake and assess glycolytic flux in different cell types or in the context of various mutations causing ALS and related neurodegenerative disorders.

Measuring Glucose Uptake in Drosophila Models of TDP-43 Proteinopathy

Glucose uptake is increased in Drosophila motor neurons affected by TAR DNA binding protein (TDP-43) proteinopathy, as indicated by a FRET-based, genetically encoded glucose sensor.

Amyotrophic lateral sclerosis is a neurodegenerative disorder causing progressive muscle weakness and death within 2-5 years following diagnosis. Clinical ...

TRAP-rc, Translating Ribosome Affinity Purification from Rare Cell Populations of Drosophila Embryos

Measuring levels of mRNAs in the process of translation in individual cells provides information on the proteins involved in cellular functions at a given point in time. The protocol dubbed Translating Ribosome Affinity Purification (TRAP) is able to capture this mRNA translation process in a cell-type-specific manner. Based on the affinity purification of polysomes carrying a tagged ribosomal subunit, TRAP can be applied to translatome analyses in individual cells, making it possible to compare cell types during the course of developmental processes or to track disease development progress and the impact of potential therapies at molecular level. Here we report an optimized version of the TRAP protocol, called TRAP-rc (rare cells), dedicated to identifying engaged-in-translation RNAs from rare cell populations. TRAP-rc was validated using the Gal4/UAS targeting system in a restricted population of muscle cells in Drosophila embryos. This novel protocol allows the recovery of cell-type-specific RNA in sufficient quantities for global gene expression analytics such as microarrays or RNA-seq. The robustness of the protocol and the large collections of Gal4 drivers make TRAP-rc a highly versatile approach with potential applications in cell-specific genome-wide studies.

TRAP-rc, Translating Ribosome Affinity Purification from Rare Cell Populations of Drosophila Embryos

Translating Ribosome Affinity Purification (TRAP) is able to capture cell-type-specific translation of mRNA. Here we report the first TRAP protocol dedicated to isolation of mRNA in rare cell populations of Drosophila embryos.

Measuring levels of mRNAs in the process of translation in individual cells provides information on the proteins involved in cellular functions at a given ...

Measurement of Poly A Tail Length from Drosophila Larva Brain and Cell Line

Summary

Explore More Videos

Chapters in this video

Laser Capture Microdissection of Drosophila Peripheral Neurons

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

Live-Cell Imaging of Drosophila melanogaster Third Instar Larval Brains

Monitoring Cell-to-cell Transmission of Prion-like Protein Aggregates in Drosophila Melanogaster

Quantitative Analysis of Neuronal Dendritic Arborization Complexity in Drosophila

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

Purification of Transcripts and Metabolites from Drosophila Heads

3D Particle Tracking for Noninvasive In Vivo Analysis of Synaptic Microtubule Dynamics in Dendrites and Neuromuscular Junctions of Drosophila

Measuring Glucose Uptake in Drosophila Models of TDP-43 Proteinopathy

TRAP-rc, Translating Ribosome Affinity Purification from Rare Cell Populations of Drosophila Embryos