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.

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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 ...

Polyadenylation is a post-transcription modification. This add poly-A tails to the three-prime end of mRNA molecules. Dysregulation of polyadenylation has been associated with abnormal gene expression and various diseases, including neurological disorders.Our method provides a cost-effective, high-resolution analysis for measuring poly-A lengths. High-throughput methods for poly-A tail analysis, like next generation sequencing, and a high-throughput sequencing provides high-content information. However, analyzing rare transcripts can be extremely challenging with these methods.PCR-based methods are usually low-throughput. However, they can be easily used to analyze a small number of transcripts. The GI-tailing method is one of them.With this method, we discovered that Dscam mRNAs contain short poly-A tails. Dscam is a neuronal cell adhesion molecule. It is involved in many important neurodevelopmental processes.Because Dscam mRNA contains short poly-A tails unlike most mRNAs, Dscam mRNA translation is under unique gene regulation, which is one of the questions we are trying to answer.

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803 Views.  University of Nevada, Reno. Polyadenylation is a post-transcription modification. This add poly-A tails to the three-prime end of mRNA molecules. Dysregulation of polyadenylation has been associated with abnormal gene expression and various diseases, including neurological disorders.Our method provides a cost-effective, high-resolution analysis for measuring poly-A lengths. High-throughput methods for poly-A tail analysis, like next generation sequencing, and a high-throughput sequencing provides high-content inf...

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

Immunohistological Labeling of Microtubules in Sensory Neuron Dendrites, Tracheae, and Muscles in the Drosophila Larva Body Wall

To understand how differences in complex cell shapes are achieved, it is important to accurately follow microtubule organization. The Drosophila larval body wall contains several cell types that are models to study cell and tissue morphogenesis. For example tracheae are used to examine tube morphogenesis1, and the dendritic arborization (DA) sensory neurons of the Drosophila larva have become a primary system for the elucidation of general and neuron-class-specific mechanisms of dendritic differentiation2-5 and degeneration6.
The shape of dendrite branches can vary significantly between neuron classes, and even among different branches of a single neuron7,8. Genetic studies in DA neurons suggest that differential cytoskeletal organization can underlie morphological differences in dendritic branch shape4,9-11. We provide a robust immunological labeling method to assay in vivo microtubule organization in DA sensory neuron dendrite arbor (Figures 1, 2, Movie 1). This protocol illustrates the dissection and immunostaining of first instar larva, a stage when active sensory neuron dendrite outgrowth and branching organization is occurring 12,13.
In addition to staining sensory neurons, this method achieves robust labeling of microtubule organization in muscles (Movies 2, 3), trachea (Figure 3, Movie 3), and other body wall tissues. It is valuable for investigators wishing to analyze microtubule organization in situ in the body wall when investigating mechanisms that control tissue and cell shape.

Immunohistological Labeling of Microtubules in Sensory Neuron Dendrites, Tracheae, and Muscles in the Drosophila Larva Body Wall

To understand how complex cell shapes, such as neuronal dendrites, are achieved during development, it is important to be able to accurately assay microtubule organization. Here we describe a robust immunohistological labeling method to examine microtubule organization of dendritic arborization neuron sensory dendrites, trachea, muscle, and other Drosophila larva body wall tissues.

To understand how differences in complex cell shapes are achieved, it is important to accurately follow microtubule organization. The Drosophila larval ...

A Method to Inflict Closed Head Traumatic Brain Injury in Drosophila

Traumatic brain injury (TBI) affects millions of people each year, causing impairment of physical, cognitive, and behavioral functions and death. Studies using Drosophila have contributed important breakthroughs in understanding neurological processes. Thus, with the goal of understanding the cellular and molecular basis of TBI pathologies in humans, we developed the High Impact Trauma (HIT) device to inflict closed head TBI in flies. Flies subjected to the HIT device display phenotypes consistent with human TBI such as temporary incapacitation and progressive neurodegeneration. The HIT device uses a spring-based mechanism to propel flies against the wall of a vial, causing mechanical damage to the fly brain. The device is inexpensive and easy to construct, its operation is simple and rapid, and it produces reproducible results. Consequently, the HIT device can be combined with existing experimental tools and techniques for flies to address fundamental questions about TBI that can lead to the development of diagnostics and treatments for TBI. In particular, the HIT device can be used to perform large-scale genetic screens to understand the genetic basis of TBI pathologies.

A Method to Inflict Closed Head Traumatic Brain Injury in Drosophila

Here we describe a method to inflict closed head traumatic brain injury (TBI) in Drosophila. This method provides a gateway to investigate the cellular and molecular mechanisms that underlie TBI pathologies using the vast array of experimental tools and techniques available for flies.

Traumatic brain injury (TBI) affects millions of people each year, causing impairment of physical, cognitive, and behavioral functions and death. Studies ...

Foraging Path-length Protocol for Drosophila melanogaster Larvae

The Drosophila melanogaster larval path-length phenotype is an established measure used to study the genetic and environmental contributions to behavioral variation. The larval path-length assay was developed to measure individual differences in foraging behavior that were later linked to the foraging gene. Larval path-length is an easily scored trait that facilitates the collection of large sample sizes, at minimal cost, for genetic screens. Here we provide a detailed description of the current protocol for the larval path-length assay first used by Sokolowski. The protocol details how to reproducibly handle test animals, perform the behavioral assay and analyze the data. An example of how the assay can be used to measure behavioral plasticity in response to environmental change, by manipulating feeding environment prior to performing the assay, is also provided. Finally, appropriate test design as well as environmental factors that can modify larval path-length such as food quality, developmental age and day effects are discussed.

Foraging Path-length Protocol for Drosophila melanogaster Larvae

We provide a detailed protocol for a Drosophila melanogaster foraging path-length assay. We discuss the preparation and handling of test animals, how to perform the assay and analyze the data.

The Drosophila melanogaster larval path-length phenotype is an established measure used to study the genetic and environmental contributions to behavioral ...

Electrophysiological Method for Whole-cell Voltage Clamp Recordings from Drosophila Photoreceptors

Whole-cell voltage clamp recordings from Drosophila melanogaster photoreceptors have revolutionized the field of invertebrate visual transduction, enabling the use of D. melanogaster molecular genetics to study inositol-lipid signaling and Transient Receptor Potential (TRP) channels at the single-molecule level. A handful of labs have mastered this powerful technique, which enables the analysis of the physiological responses to light under highly controlled conditions. This technique allows control over the intracellular and extracellular media; the membrane voltage; and the fast application of pharmacological compounds, such as a variety of ionic or pH indicators, to the intra- and extracellular media. With an exceptionally high signal-to-noise ratio, this method enables the measurement of dark spontaneous and light-induced unitary currents (i.e.&#160;spontaneous and quantum bumps) and macroscopic Light-induced Currents (LIC) from single D. melanogaster photoreceptors. This protocol outlines, in great detail, all the key steps necessary to perform this technique, which includes both electrophysiological and optical recordings. The fly retina dissection procedure for the attainment of intact and viable ex vivo isolated ommatidia in the bath chamber is described. The equipment needed to perform whole-cell and fluorescence imaging measurements are also detailed. Finally, the pitfalls in using this delicate preparation during extended experiments are explained.

Electrophysiological Method for Whole-cell Voltage Clamp Recordings from Drosophila Photoreceptors

Whole-cell recordings from Drosophila melanogaster photoreceptors enable the measurement of spontaneous dark bumps, quantum bumps, macroscopic responses to light, and current-voltage relationships under various conditions. In combination with D. melanogaster genetic manipulation tools, this method enables the study of the ubiquitous inositol-lipid signaling pathway and its target, the TRP channel.

Whole-cell voltage clamp recordings from Drosophila melanogaster photoreceptors have revolutionized the field of invertebrate visual transduction, ...

Live-cell Measurement of Odorant Receptor Activation Using a Real-time cAMP Assay

The enormous sizes of the mammalian odorant receptor (OR) families present difficulties to find their cognate ligands among numerous volatile chemicals. To efficiently and accurately deorphanize ORs, we combine the use of a heterologous cell line to express mammalian ORs and a genetically modified biosensor plasmid to measure cAMP production downstream of OR activation in real time. This assay can be used to screen odorants against ORs and vice versa. Positive odorant-receptor interactions from the screens can be subsequently confirmed by testing against various odor concentrations, generating concentration-response curves. Here we used this method to perform a high-throughput screening of an odorous compound against a human OR library expressed in Hana3A cells and confirmed that the positively-responding receptor is the cognate receptor for the compound of interest. We found this high-throughput detection method to be efficient and reliable in assessing OR activation and our data provide an example of its potential use in OR functional studies.

Characterizing the function of odorant receptors serves an indispensable part in the deorphanization process. We describe a method to measure the activation of odorant receptors in real time using a cAMP assay.

The enormous sizes of the mammalian odorant receptor (OR) families present difficulties to find their cognate ligands among numerous volatile chemicals. ...

Simple Generation of a High Yield Culture of Induced Neurons from Human Adult Skin Fibroblasts

Induced neurons (iNs), the product of somatic cells directly converted to neurons, are a way to obtain patient-derived neurons from tissue that is easily accessible. Through this route, mature neurons can be obtained in a matter of a few weeks. Here, we describe a straightforward and rapid one-step protocol to obtain iNs from dermal fibroblasts obtained through biopsy samples from adult human donors. We explain each step of the process, including the maintenance of the dermal fibroblasts, the freezing procedure to build a stock of the cell line, seeding of the cells for reprogramming, as well as the culture conditions during the conversion process. In addition, we describe the preparation of glass coverslips for electrophysiological recordings, long-term coating conditions, and fluorescence activated cell sorting (FACS). We also illustrate examples of the results to be expected. The protocol described here is easy to perform and can be applied to human fibroblasts derived from human skin biopsies from patients with various different diagnoses and ages. This protocol generates a sufficient amount of iNs which can be used for a wide array of biomedical applications, including disease modeling, drug screening, and target validation.

Direct neuronal reprogramming generates neurons that maintain the age of the starting somatic cell. Here, we describe a single vector-based method to generate induced neurons from dermal fibroblasts obtained from adult human donors.

Induced neurons (iNs), the product of somatic cells directly converted to neurons, are a way to obtain patient-derived neurons from tissue that is easily ...

Cell-Type Specific Protein Purification and Identification from Complex Tissues Using a Mutant Methionine tRNA Synthetase Mouse Line

Understanding protein homeostasis in vivo is key to knowing how the cells work in both physiological and disease conditions. The present protocol describes in vivo labeling and subsequent purification of newly synthesized proteins using an engineered mouse line to direct protein labeling to specific cellular populations. It is an inducible line by Cre recombinase expression of L274G-Methionine tRNA synthetase (MetRS*), enabling azidonorleucine (ANL) incorporation to the proteins, which otherwise will not occur. Using the method described here, it is possible to purify cell-type-specific proteomes labeled in vivo and detect subtle changes in protein content due to sample complexity reduction.

This protocol describes how to perform cell-type-specific protein labeling with azidonorleucine (ANL) using a mouse line expressing a mutant L274G-Methionine tRNA synthetase (MetRS*) and the necessary steps for labeled cell-type-specific proteins isolation. We outline two possible ANL administration routes in live mice by (1) drinking water and (2) intraperitoneal injections.

Understanding protein homeostasis in vivo is key to knowing how the cells work in both physiological and disease conditions. The present protocol ...

Monitoring Brain Metabolite Transport in Drosophila Using FRET Sensors

Visualizing Monocarboxylates and Other Relevant Metabolites in the Ex Vivo Drosophila Larval Brain Using Genetically Encoded Sensors

The high energy requirements of brains due to electrical activity are one of their most distinguishing features. These requirements are met by the production of ATP from glucose and its metabolites, such as the monocarboxylates lactate and pyruvate. It is still unclear how this process is regulated or who the key players are, particularly in Drosophila.
Using genetically encoded F&#246;rster resonance energy transfer-based sensors, we present a simple method for measuring the transport of monocarboxylates and glucose in glial cells and neurons in an ex-vivo Drosophila larval brain preparation. The protocol describes how to dissect and adhere a larval brain expressing one of the sensors to a glass coverslip. 
We present the results of an entire experiment in which lactate transport was measured in larval brains by knocking down previously identified monocarboxylate transporters in glial cells. Furthermore, we demonstrate how to rapidly increase neuronal activity and track metabolite changes in the active brain. The described method, which provides all necessary information, can be used to analyze other Drosophila living tissues.

Author Spotlight: Advances in Brain Energy Metabolism Research Using the Drosophila Model

Here we present a protocol to visualize the transport of monocarboxylates, glucose, and ATP in glial cells and neurons using genetically encoded F&#246;rster resonance energy transfer-based sensors in an ex-vivo Drosophila larval brain preparation.

The high energy requirements of brains due to electrical activity are one of their most distinguishing features. These requirements are met by the ...

Single-Neuron Ablation in Drosophila for Studying Nervous System Adaptation

Laser Cell Ablation in Intact Drosophila Larvae Reveals Synaptic Competition

The protocol describes single-neuron ablation with a 2-photon laser system in the central nervous system (CNS) of intact Drosophila melanogaster larvae. Using this non-invasive method, the developing nervous system can be manipulated in a cell-specific manner. Disrupting the development of individual neurons in a network can be used to study how the nervous system can compensate for the loss of synaptic input. Individual neurons were specifically ablated in the giant fiber system of Drosophila, with a focus on two neurons: the presynaptic giant fiber (GF) and the postsynaptic tergotrochanteral motor neuron (TTMn). The GF synapses with the ipsilateral TTMn, which is crucial to the escape response. Ablating one of the GFs in the 3rd instar brain, just after the GF starts axonal growth, permanently removes the cell during the development of the CNS. The remaining GF reacts to the absent neighbor and forms an ectopic synaptic terminal to the contralateral TTMn. This atypical, bilaterally symmetric terminal innervates both TTMns, as demonstrated by dye coupling, and drives both motor neurons, as demonstrated by electrophysiological assays. In summary, the ablation of a single interneuron demonstrates synaptic competition between a bilateral pair of neurons that can compensate for the loss of one neuron and restore normal responses to the escape circuit.

Author Spotlight: Investigating the Mechanisms of Neural Circuit Assembly and Synapse Formation in Drosophila

This protocol demonstrates the laser cell ablation of individual neurons in intact Drosophila larvae. The method enables the study of the effect of reducing competition between neurons in the developing nervous system.

The protocol describes single-neuron ablation with a 2-photon laser system in the central nervous system (CNS) of intact Drosophila melanogaster larvae. ...