This work was supported by funding from the Ellison Medical Foundation (SDP, <a href="http://www.ellisonfoundation.org/index.jsp" target="_blank">http://www.ellisonfoundation.org/index.jsp</a>), NIH K01AG031917 (NJL, <a href="http://www.nih.gov" target="_blank">http://www.nih.gov/</a>), NIH 5T32GM007315-35 (JR) and NIH R01AG030593 (SDP). This work utilized the resources of the Drosophila Aging Core (DAC) of the Nathan Shock Center of Excellence in the Biology of Aging funded by the National Institute of Aging P30-AG-013283 (<a href="http://www.nih.gov" target="_blank">http://www.nih.gov/</a>). The authors would like to thank the Pletcher Laboratory for helpful discussions and particularly Brian Chung for critical reading of the manuscript. We would like to acknowledge Nick Asher and Kathryn Borowicz for assistance with data collection.

We recommend storing experimental foods, yeast paste, and grape agar plates that appear in the protocol at 4 &deg;C and using them within 1-2 months as long as mold and dryness have not set in. Standard environmental conditions for both the larval and adult stage involve maintenance of flies in an incubator at 25 &deg;C with a 12:12 hr light dark cycle and 60% relative humidity.
1. Preparation of Experimental Food
<ol>
	<li>For larval growth, we use a modified Caltech Medium5, which is abbreviated in this protocol as CT.</li>
	<li>We recommend a diet for adult Drosophila (SY) that consists of sugar (sucrose) and yeast (lyophilized whole brewer's yeast) in a 2% agar base, that has been boiled, supplemented with antibiotics and anti-fungal agents, and distributed (10 ml per vial)6. Food should be allowed to solidify and evaporate for 12-24 hr prior to storage. Because the nutrient environment can substantially impact longevity, consistency in cooking processes is essential both within an experiment and for comparison between experiments.</li>
	<li>If a pharmacological agent is to be added to the adult food, this drug can be mixed into a small batch of food and layered (2 ml) on the food surface, with control vials receiving layers containing vehicle alone.</li>
</ol>
2. Preparation of a Live Yeast Paste
Combine 5-6 ml of water with 3 g of active dry yeast and mix well. The consistency of the yeast paste should be that of a smooth peanut butter.
3. Preparation of a Grape Agar Plate
<ol>
	<li>Add a packet of a grape agar premix in 500 ml of distilled water in a 1,000 ml flask and follow the instructions on the packet to dissolve the grape agar mix.</li>
	<li>Carefully pour a thick layer of the mixture into 100 mm Petri dishes while avoiding bubble formation. One packet of premix produces approximately 14 grape agar plates.</li>
	<li>Let the media cool and solidify in room temperature with lids on for 15 min. Plates can be kept at 4 &deg;C, wrapped in plastic wrap, or used immediately.</li>
</ol>
4. Collection of Synchronized Eggs
All media used in this protocol, i.e. yeast, grape agar plates, CT and 10% SY food, should be at room temperature.
<ol>
	<li>Spread a 2-3 cm diameter layer of yeast paste on a grape agar plate and set aside.</li>
	<li>Place a large egg collection cage on a CO2 pad with the mesh-side down to anesthetize the flies. Nitrogen-based anesthesia is an alternative to CO2, used by some laboratories, that can produce high quality results7.</li>
	<li>Using a funnel, transfer 150-200 pairs of flies into the egg collection cage.</li>
	<li>Place the grape agar plate to cover the open end of the cage and secure with an end cap.</li>
	<li>Lay the cage on its side until flies wake up. Then, keep the cage, grape agar plate side down, in the incubator overnight.</li>
	<li>On the next day, swap the grape plate with a newly yeasted grape plate. Only put about 1 cm diameter of yeast paste onto the surface of the plate. Discard the 1st day grape plate.</li>
	<li>Allow embryos to collect on the surface of the grape agar plate for 16-22 hr. Once done, collect the grape agar plate and discard the parent flies.</li>
	<li>Wash the surface of the grape agar plates with 1x Phosphate Buffered Saline (PBS). Eggs can be mobilized by gently scraping with a cotton swab. Take care not to scratch, damage, or scrape thin pieces of agar off the surface of the plate. With the aid of a funnel, pour the washed eggs into a 15 ml conical tube.</li>
	<li>Let the eggs settle to the bottom of the tube and pour off the supernatant, being careful not to lose any eggs. The remaining volume should be around 2-3 ml.</li>
	<li>Add 8-10 ml of PBS to the tube and repeat the above step 2-3 more times to thoroughly wash the eggs until the supernatant is clear. Getting rid of residual yeast is the key in this step.</li>
	<li>After the wash, drain all supernatant until remaining volume is 2 ml. Aliquot 32 &mu;l of eggs into CT bottles using a wide-bore pipette tip. Eggs in the pipette tip should be compact, with little to no liquid aspirated. This can be achieved by inserting the pipette tip deep into the egg settlement and quickly releasing the plunger to aspirate.</li>
	<li>Place seeded CT bottles back in the incubator throughout fly development.</li>
</ol>
5. Collection of Age-matched Adult Flies
<ol>
	<li>Adults will typically eclose from day 9 onward. Discard the flies that emerged on the first day and place bottles back in the incubator overnight. This practice will avoid inadvertent selection for early emergents and allow for the collection of a maximum number of synchronized flies.</li>
	<li>16-22 hr later, transfer the day-old adult flies into 10% SY food bottles. If needed, another batch can be collected the next day.</li>
	<li>Return flies back into the incubator and allow flies to reach sexual maturity and mate for two days. Record the day of transfer to 10% SY bottles as the first day of adulthood.</li>
</ol>
6. Sorting Flies and Setting up Longevity Experiment
<ol>
	<li>Anesthetize small groups of flies on the anesthetic pad, then sort males and females into two groups using a paintbrush. If using CO2, it is critical to minimize exposure to prevent possible long-lasting health issues that can compromise the integrity of the longevity experiment.</li>
	<li>Place 30 flies of the same gender into individual vials. Assuming no balancer chromosomes in the population and healthy progeny, each bottle should produce around 3-4 vials of each sex. Aliquoting flies into individual vials should take no more than 3-4 min, so that, in total, flies are only exposed to anesthesia for a maximum of 9-10 min.</li>
	<li>Repeat steps 6.1-6.2 until there are 8-10 vial replicates for each gender and experimental treatment.</li>
</ol>
7. Setting Up the Excel spreadsheet to Track the Longevity Experiment
<ol>
	<li>We recommend randomizing vial position rather than grouping vials by experimental condition in order to avoid bias associated with the vial location in the incubator and to obscure the vial identity to the experimenter. To do this, first assign a randomized numerical ID to each vial in a spreadsheet program, then arrange the vials in trays by ID number. If you are using the dLife experiment management software, follow the tutorial on experiment setup to generate the ID number for each vial.</li>
	<li>(Optional) If you are using an RFID reader or barcode reader in association with dLife, attach an RFID or barcode tag to each vial and associate the tag with the vial's numerical ID in dLife. The program will recognize each vial when scanned with a reader and guide one to record the data in the correct location in a spreadsheet. Usage of a tag reader in conjunction with dLife significantly reduces data collection time and recording error.</li>
</ol>
8. Maintaining the Longevity Experiment
The vials containing fresh food should be at room temperature for each transfer.
<ol>
	<li>During the experimental period, transfer flies onto new vials containing fresh food every 2 days (young females), or 3 times a week (males or females &gt;3 weeks of age). This step will ensure that the feeding environment for young females is not disrupted by the presence of larvae. This transfer should be completed without anesthesia, which can induce acute mortality, particularly in older flies (Pletcher, personal observations).</li>
	<li>During each vial transfer, record the age, count the dead flies in the old vial, and the dead flies that are carried to the new vial. Record this information separately in two columns in a spreadsheet (either dLife or your own spreadsheet). This will ensure that the carried flies are not double-counted. The total number of deaths (dead + carried) should at least equal the number of carried flies from the previous transfer. Subtract the number of previously carried flies from the total number of deaths to determine the number of new deaths.</li>
	<li>A fly is considered right-censored if it left the experiment prior to natural death through escape or accidental death. Animals exiting the experiment in this way should be entered into a separate column on the day that the fly exited the experiment. Censored flies are not recorded as dead (see below).</li>
	<li>Continue to repeat steps 8.1-8.2 until the last survivor is dead. Be aware that as the flies age, some flies may lie on their back and appear dead due to their inactiveness. Therefore when counting carried (dead) flies, tap on the side of the vials to determine if there are leg movements. If so, these flies are still alive. In the case where flies remain stuck to the food in the old vial but alive, they should not be counted as dead and should be rescued by further tapping of the vial to dislodge the fly. Censoring such flies should be used with caution as it may result in experimental bias.</li>
</ol>
9. Data Analysis
<ol>
	<li>The survivorship curve displays the probability that an individual survives to a given age and is typically calculated using a Kaplan-Meier approach (Figure 1)8. In the absence of right-censored data, the formula can be simplified such that age-specific survivorship at age x (Sx) is determined by dividing the number of individuals alive at the start of a census time at age x (Nx) by the total number of flies in the experiment (N0); Sx=Nx/N0. 9.2) Survivorship curves are the most common form of data presentation, and they can be tested for equality between groups using a log-rank test. Reasonable inference can be drawn from as few as 50-100 individuals in a cohort. Survivorship is a cumulative measure, however, and therefore deaths that are not aging-related, such as those early in life, will depress survivorship throughout the lifespan making it difficult to ascertain effects specific to aging.</li>
	<li>A second data visualization method is the age-specific mortality function, which displays the risk of dying through each age interval and presents a more nuanced description of survival data9,10. Mortality measures are independent from one age to another, and the shape of the mortality curve is useful for inference about the dynamics of aging, particularly when treatments are adjusted during adult life. Estimates of age-specific mortality, however, lack both precision and accuracy with small sample sizes, and often require several-to-many hundreds of individuals per cohort for a reliable estimate11.</li>
	<li>Other methods for estimating longevity differences include parametric (e.g. Gompertz) and semi-parametric (e.g. Cox regression) models. These models can be powerful but should be applied with caution because of the assumptions made about the shape of the mortality curves and the nature of treatment effects, which could lead to incorrect inference11,12.</li>
</ol>

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No conflicts of interest declared.

The protocol presented here describes a method for producing reproducible measurements of adult longevity in Drosophila that is adaptable for assessment of genetic, pharmacological, and environmental interventions. Crucial aspects of the protocol include carefully controlling the larval development environment, minimizing adult stress, and minimizing bias across experimental groups and controls. We also present the use of the dLife lifespan experiment management software. By simply attaching a bar code or RFID t...

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>Active Dry Yeast</td>
 <td>Fleishmann's Yeast</td>
 <td>2192</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Grape Agar Powder Premix</td>
 <td>Genesee Scientific</td>
 <td>47-102</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Large Embryo Collection Cages</td>
 <td>Genesee Scientific</td>
 <td>59-101</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Large Replacement End Caps</td>
 <td>Genesee Scientific</td>
 <td>59-103</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>6 oz Square Bottom Bottles, polypropylene</td>
 <td>Genesee Scientific</td>
 <td>32-130</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Flugs Closures for Stock Bottles</td>
 <td>Genesee Scientific</td>
 <td>49-100</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Drosophila Vials, Wide, Polystrene</td>
 <td>Genesee Scientific</td>
 <td>32-117</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Flugs Closures for Wide Vials</td>
 <td>Genesee Scientific</td>
 <td>49-101</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Wide Orifice Aardvark Pipet Tips, 200 ul</td>
 <td>Denville Scientific</td>
 <td>P1105-CP</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Flystuff Flypad, Standard Size</td>
 <td>Genesee Scientific</td>
 <td>59-114</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>BD Falcon 15 ml Conical Centrifuge Tubes</td>
 <td>Fisher Scientific</td>
 <td>14-959-70C</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Fisherbrand Petri Dishes with Clear Lids, Raised Ridge; 100 O.D. x 15 mm H;</td>
 <td>Fisher Scientific</td>
 <td>08-757-12</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Kimax* Colorware Flasks 1,000 ml yellow</td>
 <td>Fisher Scientific</td>
 <td>10-200-47</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>PBS pH 7.4 10x</td>
 <td>Invitrogen</td>
 <td>70011044</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Gelidium Agar</td>
 <td>Mooragar</td>
 <td>n/a</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Brewer's Yeast</td>
 <td>MP Biomedicals</td>
 <td>0290331280</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Granulated Sugar</td>
 <td>Kroger</td>
 <td>n/a</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Tegosept</td>
 <td>Genesee Scientific</td>
 <td>20-266</td>
 <td>Fly Food Preservative</td>
 </tr>
 <tr>
 <td>Propionic Acid, 99%</td>
 <td>Acros Organics</td>
 <td>149300025</td>
 <td>Fly Food Preservative</td>
 </tr>
 <tr>
 <td>Kanamycin Sulfate</td>
 <td>ISC BioExpress</td>
 <td>0408-10G</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Tetracycline HCl</td>
 <td>VWR</td>
 <td>80058-724</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>

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.

A simplified scheme of the protocol is presented in Figure 1, where key steps are outlined. The synchronization part of the protocol can be used for various assays that require age-matched adult flies.
Typical survivorship curves of wild-type flies are shown in Figure 2a, using the dLife experiment management software (Figure 2b,c). Adult males usually live shorter, with both populations achieving a mean and median longevity of &gt;50 days on ...

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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.

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 ...

measurement-of-lifespan-in-drosophila-melanogaster

Research

JoVE Journal

Biology

33.9K Views.  University of Michigan. 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.

Video: Measurement of Lifespan in 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 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.

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 ...

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained and restrained whole larvae. For direct control of the environment around the heart another approach utilizes the dissected larvae and removal of the internal organs in order to bathe the heart in desired compounds. The exposed heart also allows membrane potentials to be monitored which can give insight of the ionic currents generated by the myocytes and for electrical conduction along the heart tube. These approaches have various advantages and disadvantages for future experiments that are discussed. The larval heart preparation provides an additional model besides the Drosophila skeletal NMJ to investigate the role of intracellular calcium regulation on cellular function. Learning more about the underlying ionic currents that shape the action potentials in myocytes in various species, one can hope to get a handle on the known ionic dysfunctions associated to specific genes responsible for various diseases in mammals.

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various ways to monitor heart function in the larva of Drosophila for assessing questions dealing with the function of gap junctions, ion channel mutations, modulation of pacemaker activity and pharmacological studies.

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained  and  ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

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 ...

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 ...

Enhanced Northern Blot Detection of Small RNA Species in Drosophila Melanogaster

The last decades have witnessed the explosion of scientific interest around gene expression control mechanisms at the RNA level. This branch of molecular biology has been greatly fueled by the discovery of noncoding RNAs as major players in post-transcriptional regulation. Such a revolutionary perspective has been accompanied and triggered by the development of powerful technologies for profiling short RNAs expression, both at the high-throughput level (genome-wide identification) or as single-candidate analysis (steady state accumulation of specific species). Although several state-of-art strategies are currently available for dosing or visualizing such fleeing molecules, Northern Blot assay remains the eligible approach in molecular biology for immediate and accurate evaluation of RNA expression. It represents a first step toward the application of more sophisticated, costly technologies and, in many cases, remains a preferential method to easily gain insights into RNA biology. Here we overview an efficient protocol (Enhanced Northern Blot) for detecting weakly expressed microRNAs (or other small regulatory RNA species) from Drosophila melanogaster whole embryos, manually dissected larval/adult tissues or in vitro cultured cells. A very limited amount of RNA is required and the use of material from flow cytometry-isolated cells can be also envisaged.

Enhanced Northern Blot Detection of Small RNA Species in Drosophila Melanogaster

The aim of this publication is to visualize and discuss the operative steps of an Enhanced Northern Blot protocol on RNA extracted from Drosophila melanogaster embryos, cells, and tissues. This protocol is particularly useful for the efficient detection of small RNA species.

The last decades have witnessed the explosion of scientific interest around gene expression control mechanisms at the RNA level. This branch of molecular ...

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 ...

Measurement of Carbon Dioxide Production from Radiolabeled Substrates in Drosophila melanogaster

The power of Drosophila genetics is increasingly being applied to questions of hormone signaling and metabolism and to the development of models of human disease in this organism. Sensitive methods for measurements of parameters such as metabolic rates are needed to drive the understanding of physiology and disease in small animals such as the fruit fly. The method described here assesses fuel oxidation in small numbers of adult flies fed food containing trace amounts of 14C-labeled substrates such as glucose or fatty acid. After the feeding period and any additional experimental manipulations, flies are transferred to short tubes capped with mesh, which are then placed in glass vials containing KOH-saturated filter paper that traps exhaled, radiolabeled CO2 generated from oxidation of radiolabeled substrates as potassium bicarbonate, KHCO3. This radiolabeled bicarbonate is measured by scintillation counting. This is a quantitative, reproducible, and simple approach for the study of fuel oxidation. The use of radiolabeled glucose, fatty acids, or amino acids allows determination of the contribution of these different fuel sources to energy metabolism under different conditions such as feeding and fasting and in different genetic backgrounds. This complements other approaches used to measure in vivo energy metabolism and should further the understanding of metabolic regulation.

Measurement of Carbon Dioxide Production from Radiolabeled Substrates in Drosophila melanogaster

This paper describes a method for the measurement of fuel oxidation in Drosophila melanogaster in which trace amounts of specific radiolabeled metabolic substrates are fed to flies. The exhaled radiolabeled CO2 that is a produced from fuel oxidation is collected and measured.

The power of Drosophila genetics is increasingly being applied to questions of hormone signaling and metabolism and to the development of models of human ...