This work was supported by National Institutes of Health Grant R01 GM54408 awarded to L.C. Griffith. We would like to thank Fred Wolf for all his help in designing our square chamber and setting up our assay, Frank Mello at the Brandeis University machine shop for building our chamber, and Dan Valente and Tim Lebestky for helpful conversations regarding analysis issues.

Part 1: Feeding and Management of flies
<ol>
	<li>Flies should be grown in bottles containing yeast-free standard media. You will need large numbers of flies, so crosses need to be set up accordingly. Flies should be grown in a 12-hour light:dark cycle at 25&#186;C.</li>
	<li>Flies should be collected soon after eclosion (1-3 days). Flies can be worked with at this stage using a carbon dioxide diffuser, and should be sorted into test tubes containing autoclaved or yeast-free media. We use males only, stored 10 to a test tube.</li>
	<li>Allow at least a day or two after use of carbon dioxide before running the behavioral experiments. We run flies in our behavioral assay 3-5 days following ECLOSION.</li>
	<li>Assays should always be done within the same 2-3 hour time window each day to avoid circadian rhythm issues.</li>
</ol>
Part 2: Setting up the tracking system in a controlled environment
<ol>
	<li>All experiments are done in an environment room which maintains a constant temperature of 25&#186;C with 70% humidity.</li>
	<li>Suspend a digital video camera over the area to be recorded (camera facing downwards). Since our tracking software is contrast-based, we suspend the camera over a light box. The camera records videos directly onto a Dell computer via a firewire connection. We use a Sharp digital video camera, hooked up to a Dell computer with Windows Moviemaker (Vista version) for video acquisition.</li>
	<li>Air pulses will be administered in this protocol, so a source for air flow must be present in the room. Using rubber tubing, connect the air source to a carbon filter for air filtration.</li>
	<li>Using more rubber tubing, connect the other side of the carbon filter to an Erlenmeyer flask via a hollow rubber stopper, and fill the flask with about a half inch of water. This will humidify the air.</li>
	<li>The flask should also have a sidearm, which will connect to a Y-shaped valve via rubber tubing.</li>
	<li>One branch of the Y-valve should be connected to a flowmeter. The other branch should deliver air to the square locomotion chamber.</li>
</ol>
Part 3: Delivering flies to the locomotion chamber
<ol>
	<li>The square locomotion chamber is based on the design from Wolf et al. 20021, with the addition of small drop trays to prevent the flies from flying within the chamber. The chamber should be taken apart, and all components should be thoroughly wiped down with 70% ethanol and dried before use.</li>
	<li>Anesthesia immediately before conducting a behavioral assay can potentially compromise performance. To avoid this, flies are gently knocked into the chamber using a funnel with a small outlet. To make an outlet of the right size, attach a blue pipette tip (for P1000) to the end of a funnel. Use scissors to cut off the very end of the pipette tip to make the opening big enough for a fly to pass through.</li>
	<li>The top of our chamber is a small piece of plexiglass secured by screws around the edges. When the screws are removed, the holes can be used to deliver flies into the chamber. Position the plexiglass top so that the screw hole sits directly over the inner chamber instead of where a screw normally sits. Take a test tube containing the flies and place it upside down over the funnel, which should then be placed into the screw hole.</li>
	<li>Do NOT move the funnel, as this could hurt or damage the flies. Instead, gently bang the whole chamber, funnel and all, until all flies are inside the chamber. This knocking should be done on a mouse pad to absorb the shock of this banging. When flies are inside, remove the funnel, and re-position the plexiglass top over the screw beds. Then re-screw the plexiglass top into place.</li>
</ol>
Part 4: Running the Locomotor Assay
<ol>
	<li>Once the flies are properly loaded, let them acclimate in the chamber for 30 minutes. This acclimation period should take place in the controlled environment (25&#186;C, 70% humidity), and the chamber should be placed on top of the light box (which should be turned on). Turn other room lights off.</li>
	<li>Following the acclimation period, switch the Y-valve in the air flow system such that the air will only flow to the flowmeter. Turn on the air, and ramp the air up to the desired speed (4.0-6.0 L/min). We generally use 5.0-5.5 L/min, but any speed in that range will work.</li>
	<li>Once the desired airspeed is attained, switch the Y-valve so that the air flows to the locomotion chamber. Time this air pulse for 15 seconds, and then abruptly turn the air off. As you turn the air off, switch the camera to record mode. (This part takes practice to do all at once).</li>
	<li>Stop the recording after the desired time and SAVE. We record 30 second trials at 10 frames per second (at least 8 trials per genotype).</li>
</ol>
Part 5: Analysis of Videos
<ol>
	<li>We use the Dynamic Image Analysis System (DIAS) software 3.2 for motion tracking2. In order to use this software, we first convert our videos to an AVI file format with Quicktime 7.5.5.</li>
	<li>To track the flies based on contrast, we use the &#8220;Autotrace by Threshold&#8221; function. We then use the &#8220;Make Path from Trace&#8221; function to digitize these traces.</li>
	<li>To output the instantaneous speeds of each fly, we use the &#8220;Compute Parameters&#8221; function to produce a database file (DIAS-specific).&#160; Using this function, we also smooth the data using a &#8220;5,15,60,15,5&#8221; Tukey Smoothing Window.</li>
	<li>Once this information is outputted as a database file, it can be opened in Microsoft Excel. We use a Matlab script to compile the instantaneous speeds within excel and calculate additional parameters necessary for understanding movement dynamics and bout structure.</li>
</ol>
Representative Results:
Figure 1 shows representative traces from wild type Canton S organisms (Figure 1, left panel) and flies null for the dCASK gene which were generated by crossing two large overlapping deletions (Df(3R)x307 and Df(3R)x313) (Figure 1, right panel). dCASK null flies have previously been shown to have locomotor problems using Buridan&#8217;s Paradigm3, and in our paradigm, compared to Canton S, they show greatly decreased locomotion. We always run wild type organisms first to make sure that conditions and behavior are within the normal range on a given day. We have observed deviations from standard responses in less than 5% of testing days. These differences can usually be attributed to problems with parameters of our controlled environment.
<img alt="Figure 1" src="/files/ftp_upload/old/1096/1096fig1.jpg" />
Figure 1. DIAS-generated traces of both Canton S wild type flies (above, left) and Locomotor-deficient dCASK Null flies generated from overlapping deletions (above,right). The traces represent what is seen in the recorded videos. Both pictures contrain traces from 8-10 flies run for 30 seconds in the video tracking assay following an air pulse.
<img alt="Figure 2" src="/files/ftp_upload/old/1096/1096fig2.jpg" />
Figure 2. Wild type flies were run in the locomotor assay (above) following an air pulse (purple bars) and without an air pulse (blue). In all parameters calculated by the analysis program, there were no significant differences between the two conditions (determined with two-tailed student t-test). This demonstrates that following an air pulse, flies locomote normally in our setup.

<ol>
	<li>Wolf, F. W., Rodan, A. R., Tsai, L. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-Resolution+Analysis+of+Ethanol-Induced+Locomotor+Stimulation+in+Drosophila.">High-Resolution Analysis of Ethanol-Induced Locomotor Stimulation in Drosophila.</a> J Neurosci. 22 (24), 11035-11044 (2002).</li><li>Soll, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+of+computers+in+understanding+how+animal+cells+crawl.">The use of computers in understanding how animal cells crawl.</a> International review of cytology. 163, 43-104 (1995).</li><li>Martin, J. R., Ollo, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+Drosophila+Ca2+/calmodulin-dependent+protein+kinase+(Caki)+is+localized+in+the+central+nervous+system+and+implicated+in+walking+speed.">A new Drosophila Ca2+/calmodulin-dependent protein kinase (Caki) is localized in the central nervous system and implicated in walking speed.</a> The EMBO journal. 15 (8), 1865-1876 (1996).</li><li>Levine, J. D., Funes, P., Dowse, H. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Resetting+the+Circadian+Clock+by+Social+Experience+in+Drosophila+melanogaster.">Resetting the Circadian Clock by Social Experience in Drosophila melanogaster.</a> Science. 298, 2010-2012 (2002).</li></ol>

In our setup, a moderately strong air pulse transiently stops fly movement. When the air pulse ends, the flies are released from this stationary state and locomote normally, as shown in figure 2. Because this air pulse effectively synchronizes locomotion of the population, we use it to begin the trial so that we can also study the onset of movement following a momentum-breaking stimulus. The assay can, however, be done without an air pulse since locomotor parameters following initiation are unaffected by the air pulse...

<!DOCTYPE html PUBLIC "-//W3C//DTD XHTML 1.0 Transitional//EN" "http://www.w3.org/TR/xhtml1/DTD/xhtml1-transitional.dtd">
<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Square Chamber</td>
<td>Tool</td>
<td>Machine Shop</td>
<td>N/A</td>
<td>Design from Wolf et al. 20021</td>
</tr>
<tr>
<td>Digital Camera</td>
<td>Camera</td>
<td>Sharp</td>
<td>ViewcamZ VL-23</td>
<td>&nbsp;</td>
<tr>
<td>Flowmeter</td>
<td>Tool</td>
<td>Cole Parmer</td>
<td>SY-32003-12</td>
<td>&nbsp;</td>
<tr>
<td>Light Box</td>
<td>Tool</td>
<td>DNASTAR, Inc.</td>
<td>Seq-Easy</td>
<td>&nbsp;</td>
<tr>
<td>Charcoal Filter</td>
<td>Tool</td>
<td>Fisher Scientific</td>
<td>09-744-37</td>
<td>&nbsp;</td>
<tr>
<td>DIAS 3.2</td>
<td>Software</td>
<td>Soll technologies</td>
<td>N/A</td>
<td>www.solltechnologies.com</td>
</tr>		</tbody>
		</table>
		</div>

high-resolution video tracking of locomotion in adult drosophila melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field. Studying locomotor behavior in Drosophila melanogaster relies on the ability to be able to quantify changes in motion during or in response to a given task. For this reason, a high-resolution video tracking system, such as the one we describe in this paper, is a valuable tool for measuring locomotion in real-time. Our protocol involves the use of an initial air pulse to break the flies momentum, followed by a thirty second filming period in a square chamber. A tracking program is then used to calculate the instantaneous speed of each fly within the chamber in 10 msec increments. Analysis software then compiles this data, and outputs a variety of parameters such as average speed, max speed, time spent in motion, acceleration, etc. This protocol will discuss proper feeding and management of flies for behavioral tasks, handling flies without anesthetization or immobilization, setting up a controlled environment, and running the assay from start to finish.

Watch this Scientific Journal Video about High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster at JoVE.com

High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster

The study of complex locomotor behavior in Drosophila melanogaster is dependent upon the ability to quantify changes in a given fly's movement. This article demonstrates how to do this using a high-resolution tracking system.

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field.  Studying ...

high-resolution-video-tracking-locomotion-adult-drosophila

Research

JoVE Journal

Biology

13.4K Views.  Brandeis. The study of complex locomotor behavior in Drosophila melanogaster is dependent upon the ability to quantify changes in a given fly's movement. This article demonstrates how to do this using a high-resolution tracking system.

Video: High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster

Dissection of Larval CNS in Drosophila Melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to examine the expression of various novel and marker proteins.  Staining of whole larvae is impractical because the tough cuticle prevents antibodies from penetrating inside the body cavity.  In order to stain these tissues it is necessary to dissect the animal prior to fixing and staining.  In this article we demonstrate how to dissect Drosophila larvae without damaging the CNS.  Begin by tearing the larva in half with a pair of fine forceps, and then turn the cuticle "inside-out" to expose the CNS.  If the dissection is performed carefully the CNS will remain attached to the cuticle.  We usually keep the CNS attached to the cuticle throughout the fixation and staining steps, and only completely remove the CNS from the cuticle just prior to mounting the samples on glass slides.  We also show some representative images of a larval CNS stained with Eve, a transcription factor expressed in a subset of neurons in the CNS.  The article concludes with a discussion of some of the practical uses of this technique and the potential difficulties that may arise.

In this article we demonstrate how to dissect the central nervous system from third instar Drosophila larvae.

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to ...

High-resolution Measurement of Odor-Driven Behavior in Drosophila Larvae

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral paradigm, the experimenter usually assumes that the rapid diffusion of odorant molecules from the source leads to the creation of a stable gradient in the dish. To establish a quantitative correlation between sensory inputs and behavioral responses, it is necessary to achieve a more thorough characterization of the odorant stimulus conditions. In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral ...

Visually Mediated Odor Tracking During Flight in Drosophila

Flying insects use visual cues to stabilize their heading in a wind stream. Many animals additionally track odors carried in the wind. As such, visual stabilization of upwind tracking directly aids in odor tracking. But do olfactory signals directly influence visual tracking behavior independently from wind cues? Additionally, recent advances in olfactory molecular genetics and neurophysiology have motivated novel quantitative behavioral analyses to assess the behavioral influence of (e.g.) genetically inactivating specific olfactory activation circuits. We modified a magnetic tether system originally devised for vision experiments by equipping the arena with narrow laminar flow odor plumes. Here we focus on experiments that can be performed after a fly is tethered and is able to navigate in the magnetic arena. We show how to acquire video images optimized for measuring body angle, how to judge stable odor tracking, and we illustrate two experiments to examine the influence of visual cues on odor tracking.

Here we describe how to optimize the acquired video image for an olfactory magnetic-tether (OMT) apparatus. We also describe two sample experimental protocols for studying visuo-olfactory fusion.

Flying insects use visual cues to stabilize their heading in a wind stream. Many animals additionally track odors carried in the wind. As such, visual ...

Dissection of Oenocytes from Adult Drosophila melanogaster

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides protection against desiccation and other environmental challenges. Several of these cuticular hydrocarbon (CHC) compounds also function as semiochemical signals, and as such mediate pheromonal communications between members of the same species, or in some instances between different species, and influence behavior. Specialized cells referred to as oenocytes are regarded as the primary site for CHC synthesis. However, relatively little is known regarding the involvement of the oenocytes in the regulation of the biosynthetic, transport, and deposition pathways contributing to CHC output. Given the significant role that CHCs play in several aspects of insect biology, including chemical communication, desiccation resistance, and immunity, it is important to gain a greater understanding of the molecular and genetic regulation of CHC production within these specialized cells. The adult oenocytes of D. melanogaster are located within the abdominal integument, and are metamerically arrayed in ribbon-like clusters radiating along the inner cuticular surface of each abdominal segment. In this video article we demonstrate a dissection technique used for the preparation of oenocytes from adult D. melanogaster. Specifically, we provide a detailed step-by-step demonstration of (1) how to fillet prepare an adult Drosophila abdomen, (2) how to identify the oenocytes and discern them from other tissues, and (3) how to remove intact oenocyte clusters from the abdominal integument. A brief experimental illustration of how this preparation can be used to examine the expression of genes involved in hydrocarbon synthesis is included. The dissected preparation demonstrated herein will allow for the detailed molecular and genetic analysis of oenocyte function in the adult fruit fly.

Dissection of Oenocytes from Adult Drosophila melanogaster

In insects, the oenocytes produce cuticular hydrocarbon compounds. These compounds protect against desiccation and facilitate chemical communication. Here we demonstrate a dissection technique used to isolate the oenocytes from adult Drosophila melanogaster, and illustrate how this preparation can be utilized to study genes involved in hydrocarbon synthesis.

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides ...

Isolation of Drosophila melanogaster Testes

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline interactions. Testes are typically isolated from adult males 0-3 days after eclosion from the pupal case. The testes of wild-type flies are easily distinguished from other tissues because they are yellow, but the testes of white mutant flies, a common genetic background for laboratory experiments are similar in both shape and color to the fly gut. Performing dissection on a glass microscope slide with a black background makes identifying the testes considerably easier. Testes are removed from the flies using dissecting needles. Compared to protocols that use forceps for testes dissection, our method is far quicker, allowing a well-practiced individual to dissect testes from 200-300 wild-type flies per hour, yielding 400-600 testes. Testes from white flies or from mutants that reduce testes size are harder to dissect and typically yield 200-400 testes per hour.

Isolation of Drosophila melanogaster Testes

Drosophila melanogaster testes can be rapidly and efficiently isolated from adult males using dissecting needles. With practice, one can readily isolate in one or two days an amount of testes sufficient for the analysis of DNA or RNA by high throughput sequencing or more traditional molecular biology methods or of protein for antibody- or enzyme-based assays.

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline ...

Insulin Injection and Hemolymph Extraction to Measure Insulin Sensitivity in Adult Drosophila melanogaster

Conserved nutrient sensing mechanisms exist between mammal and fruit fly where peptides resembling mammalian insulin and glucagon, respectively function to maintain glucose homeostasis during developmental larval stages 1,2. Studies on largely post-mitotic adult flies have revealed perturbation of glucose homeostasis as the result of genetic ablation of insulin-like peptide (ILP) producing cells (IPCs) 3. Thus, adult fruit flies hold great promise as a suitable genetic model system for metabolic disorders including type II diabetes. To further develop the fruit fly system, comparable physiological assays used to measure glucose tolerance and insulin sensitivity in mammals must be established. To this end, we have recently described a novel procedure for measuring oral glucose tolerance response in the adult fly and demonstrated the importance of adult IPCs in maintaining glucose homeostasis 4,5. Here, we have modified a previously described procedure for insulin injection 6 and combined it with a novel hemolymph extraction method to measure peripheral insulin sensitivity in the adult fly. Uniquely, our protocol allows direct physiological measurements of the adult fly's ability to dispose of a peripheral glucose load upon insulin injection, a methodology that makes it feasible to characterize insulin signaling mutants and potential interventions affecting glucose tolerance and insulin sensitivity in the adult fly.

Insulin Injection and Hemolymph Extraction to Measure Insulin Sensitivity in Adult Drosophila melanogaster

Conserved insulin signaling pathways found in the fruit fly Drosophila melanogaster make this organism a potential tool for modeling metabolic disorders including type II diabetes. To this end, it is critical to establish physiological assays to effectively measure systemic insulin action in peripheral glucose disposal in the adult fly.

Conserved nutrient sensing mechanisms exist between mammal and fruit fly where peptides resembling mammalian insulin and glucagon, respectively function ...

Measurement of Lifespan in Drosophila melanogaster

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced physical performance and increased risk of disease. Individual aging is manifest at the population level as an increase in age-dependent mortality, which is often measured in the laboratory by observing lifespan in large cohorts of age-matched individuals. Experiments that seek to quantify the extent to which genetic or environmental manipulations impact lifespan in simple model organisms have been remarkably successful for understanding the aspects of aging that are conserved across taxa and for inspiring new strategies for extending lifespan and preventing age-associated disease in mammals.
The vinegar fly, Drosophila melanogaster, is an attractive model organism for studying the mechanisms of aging due to its relatively short lifespan, convenient husbandry, and facile genetics. However, demographic measures of aging, including age-specific survival and mortality, are extraordinarily susceptible to even minor variations in experimental design and environment, and the maintenance of strict laboratory practices for the duration of aging experiments is required. These considerations, together with the need to practice careful control of genetic background, are essential for generating robust measurements. Indeed, there are many notable controversies surrounding inference from longevity experiments in yeast, worms, flies and mice that have been traced to environmental or genetic artifacts1-4. In this protocol, we describe a set of procedures that have been optimized over many years of measuring longevity in Drosophila using laboratory vials. We also describe the use of the dLife software, which was developed by our laboratory and is available for download (<a href="http://sitemaker.umich.edu/pletcherlab/software">http://sitemaker.umich.edu/pletcherlab/software</a>). dLife accelerates throughput and promotes good practices by incorporating optimal experimental design, simplifying fly handling and data collection, and standardizing data analysis. We will also discuss the many potential pitfalls in the design, collection, and interpretation of lifespan data, and we provide steps to avoid these dangers.

Measurement of Lifespan in Drosophila melanogaster

Drosophila melanogaster is a powerful model organism for exploring the molecular basis of longevity regulation. This protocol will discuss the steps involved in generating a reproducible, population-based measurement of longevity as well as potential pitfalls and how to avoid them.

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Maintenance of a Drosophila melanogaster Population Cage

Large quantities of DNA, RNA, proteins and other cellular components are often required for biochemistry and molecular biology experiments. The short life cycle of Drosophila enables collection of large quantities of material from embryos, larvae, pupae and adult flies, in a synchronized way, at a low economic cost. A major strategy for propagating large numbers of flies is the use of a fly population cage. This useful and common tool in the Drososphila community is an efficient way to regularly produce milligrams to tens of grams of embryos, depending on uniformity of developmental stage desired. While a population cage can be time consuming to set up, maintaining a cage over months takes much less time and enables rapid collection of biological material in a short period. This paper describes a detailed and flexible protocol for the maintenance of a Drosophila melanogaster population cage, starting with 1.5 g of harvested material from the previous cycle.

Maintenance of a Drosophila melanogaster Population Cage

This manuscript reports a detailed protocol for culturing, on a regular basis, a population of Drosophila melanogaster using a fly population cage.

Large quantities of DNA, RNA, proteins and other cellular components are often required for biochemistry and molecular biology experiments. The short life ...

Optical Cross-Sectional Muscle Area Determination of Drosophila Melanogaster Adult Indirect Flight Muscles

Muscle mass wasting, known as muscle atrophy, is a common phenotype in Drosophila models of neuromuscular diseases. We have used the indirect flight muscles (IFMs) of flies, specifically the dorso-longitudinal muscles (DLM), as the experimental subject to measure the atrophic phenotype brought about by different genetic causes. In this protocol, we describe how to embed fly thorax muscles for semi thin sectioning, how to obtain a good contrast between muscle and the surrounding tissue, and how to process optical microscope images for semiautomatic acquisition of quantifiable data and analysis. We describe three specific applications of the methodological pipeline. First, we show how the method can be applied to quantify muscle degeneration in a myotonic dystrophy fly model; second, measurement of muscle cross-sectional area can help to identify genes that either promote or prevent muscle atrophy and/or muscle degeneration; third, this protocol can be applied to determine whether a candidate compound is able to significantly modify a given atrophic phenotype induced by a disease-causing mutation or by an environmental trigger.

Optical Cross-Sectional Muscle Area Determination of Drosophila Melanogaster Adult Indirect Flight Muscles

We report a method to quantify muscle area, which is an indirect method to determine muscle mass in Drosophila adults. We demonstrate the application of our methodology by analyzing the indirect flight muscles in a Drosophila model of Myotonic Dystrophy disease.

Muscle mass wasting, known as muscle atrophy, is a common phenotype in Drosophila models of neuromuscular diseases. We have used the indirect flight ...

High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster

Summary

Explore More Videos

Chapters in this video

Dissection of Larval CNS in Drosophila Melanogaster

High-resolution Measurement of Odor-Driven Behavior in Drosophila Larvae

Visually Mediated Odor Tracking During Flight in Drosophila

Dissection of Oenocytes from Adult Drosophila melanogaster

Isolation of Drosophila melanogaster Testes

Insulin Injection and Hemolymph Extraction to Measure Insulin Sensitivity in Adult Drosophila melanogaster

Measurement of Lifespan in Drosophila melanogaster

Imaging Through the Pupal Case of Drosophila melanogaster

Maintenance of a Drosophila melanogaster Population Cage

Optical Cross-Sectional Muscle Area Determination of Drosophila Melanogaster Adult Indirect Flight Muscles