We thank our staff for help with the testing and optimization of this protocol: Ameya Belamkar, Marilyn Clark, Alexander Fitt, Emma Rose Gallant, Ethan Golditch, Matthew Lowe, Morgan Marsh, Kyle McClung, Andy Puga, Darcy Rose, Cameron Stockbridge, and Noelle Zolman. We also thank our colleagues from the Precision Toxicology Group, particularly our Exposure Group counterparts, for helping to identify the goals of the protocol.
This project received funding from the European Union&#39;s Horizon 2020 Research and Innovation program under Grant Agreement No. 965406. The work presented in this publication was performed as part of the ASPIS Cluster. This output reflects only the authors&#39; views, and the European Union cannot be held responsible for any use that may be made of the information contained therein. This publication was also made possible with support from the Indiana Clinical and Translational Sciences Institute, which is funded in part by Award Number UL1TR002529 from the National Institutes of Health, National Center for Advancing Translational Sciences, Clinical and Translational Sciences Award. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. Portions of this project were supported by funds from Indiana University awarded to JRS and the PhyloTox consortium. JMH and EMP were supported by NIH award P40OD018537 to Bloomington Drosophila Stock Center.

<ol>
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No conflicts of interest to disclose.

The fruit fly Drosophila melanogaster is emerging as a powerful system for NAMs16,18,19,21. By leveraging the unparalleled genetic resources available to the fly community, combined with recent advances in genomics and metabolomics, chemical safety studies using Drosophila are capable of quickly identifying the molecular mechanisms by which individual compounds interfere with metabolism, physiology, and cell signaling (for example, see39). This inexpensive protocol is designed to rapidly define dose-response curves and subsequently generate samples for RNA-seq and metabolomics analysis. Moreover, this flexible protocol can be adapted for use with any genotype and can accommodate many classes of chemicals.
A notable aspect of this protocol is the choice of liquid food used in the chemical exposure, which is based on a previous study, but differs from the solid media used by most toxicological studies of Drosophila18,22. This specific liquid media was selected to reflect the nutritional content of the standard, solid BDSC media that the flies are also fed in this protocol, to ensure the flies receive consistent nutrition. The simplicity of liquid feeding media has many advantages. Liquid media is easier to handle than solid food, which needs to be either melted and resolidified or reconstituted from powder. Liquid media also increases the system&#39;s throughput, ensures even chemical distribution throughout the feeding media, and decreases the time spent working with hazardous compounds. Additionally, the media does not require solutions to be heated, which facilitates the testing of volatile test compounds. Finally, because of the relatively few components included in the food solution, undesirable side reactions are minimized between the test chemical and other dietary components. The yeast used in the food is also inactive, further limiting the reactivity of the feeding medium. However, please note that the method is not suitable for testing developmental or larval toxicity.
Some of the materials used in the protocol can be substituted, such as using glass fly vials rather than polypropylene. However, the materials used were selected to be both inert and disposable to avoid unwanted chemical reactions between reagents and chemical exposures that could result from cleaning glassware.
The use of liquid food necessitates a vehicle for food delivery. Cellulose acetate filter paper was selected for this purpose due to its flexibility and inert nature28. Other researchers used similar protocols but with other vehicles, such as delicate task wipes or glass fiber filter29,30. The cellulose acetate filter paper suited these needs because it is an inert vehicle which can be cut to the ideal shape to fit it into the bottom of the fly vials without large gaps between the paper and vial wall, preventing death due to flies becoming stuck in media or the vehicle itself.
An important limitation of this system is that the maximum testable concentration of a chemical is tied to the solubility of the chemical. Non-water-soluble compounds require an additional solvent, which can lead to additional or synergistic effects with the chemical of interest. This can also create situations in which it is not possible to prepare stock solutions that are concentrated enough to achieve the desired endpoint in all organisms, therefore limiting analysis of the resulting data31. To address this, chemicals with low water solubility can be tested by adding up to 0.5% dimethyl sulfoxide to the food solution. Other solvents could be used as well, but additional research is needed for each solvent of interest to determine the maximum acceptable solvent concentration within the solution to maximize solubility while minimizing solvent effects on the organism.
Extensive characterization of the olfaction response in Drosophila has described how flies avoid consuming toxic compounds40,41, leading to reduced feeding on treated media. The blue dye assay addresses this phenomenon by allowing researchers to efficiently screen the feeding behaviors of the flies fed each concentration of experimental chemical42,43,44. The presence or absence of blue in the fly&#39;s gastrointestinal tract indicates if the fly has been eating the toxicant-containing medium. Although more sophisticated methods of assessing fly feeding behaviors exist, such as the Fly Liquid-Food Interaction Counter45, this qualitative method is better suited for higher-throughput screening.
A notable aspect of this protocol is that it has been optimized for a 48 h exposure period without the need to transfer flies or add additional liquid to the exposure vial. Using a humidity chamber and placing the chambers in an incubator kept at high humidity prevented the filter paper containing the feeding media from drying out during this timeframe. The protocol can be adapted for longer exposure durations, but the method must be adjusted to ensure that the filter paper does not become dry and cause significant changes in solution concentration or lethality due to desiccation.
Finally, an important characteristic of this protocol is that it can readily accommodate genetic variants, which allows researchers to utilize the vast array of genetic tools for Drosophila to expand these preliminary studies on wild-type organisms to better understand mechanisms of chemical action in vivo. In this regard, the protocol outlined above could be easily modified to complement a previously described JoVE&#160;protocol by Peterson and Long that allows for toxicological analysis of wild-caught flies18.
Because of the wide variety of previous studies on the toxicity of sodium arsenite in Drosophila32,33,34,35,36, Oregon-R flies were treated with this compound to demonstrate the efficacy of our system. Male flies exhibited an LD50 of 0.65 mM, and females exhibited an LD50 of 0.90 mM. This aligns with previous studies of sodium arsenite-treated adult Drosophila. For example, Goldstein and Babich37 found that 50% of flies (mixed sexes) died after 7 days of exposure to 0.5 mM NaAsO2. Although this is a slightly lower dose than was presently observed, the differences between their methods and this method (including the use of solid exposure media, a longer time scale, and mixed sexes) likely account for this difference. Importantly, both methods resulted in overall similar LD50 values.
Observations from experiments using this protocol can be used to find genetic and molecular targets for subsequent behavioral or mechanistic studies. The exposure method can also be used to treat Drosophila for sampling for metabolomics and proteomics, making this protocol well suited to the growing field of precision toxicology (modeled from the precision medicine field46). In this regard, exposed flies can be collected after step 8 for subsequent genomic and metabolomics analysis. Samples collected in step 8 can then be processed, as described by Li and Tennessen47, starting with step 3.
Ultimately, the data acquired from the experiments described above, as well as any subsequent metabolomics and proteomics data, would ideally be used in cross-species comparisons. As previously noted26, such cross-species studies are powerful and capable of determining how individual chemicals interfere with conserved biological pathways. Thus, the protocol described above can be used to find evolutionary commonalities in response to individual toxicants across phyla and help inform chemical safety regulation.

Humans are constantly exposed to chemicals from a variety of sources, including air1, food2, water3,4, medications5, cleaning agents6, personal care products7, industrial chemicals7, and building materials7. Moreover, thousands of new chemicals are introduced each year8, many of which are not properly vetted for health and environmental safety. This lack of adequate chemical safety testing stems in part from an over-reliance on mammalian models, such as mice and rats. While such rodent models are informative, chemical safety testing in these systems is expensive, time-consuming, and often causes unacceptable levels of suffering to the test animal9.
The financial and ethical burdens associated with mammalian chemical safety testing, as well as the time-consuming nature of mammalian studies, are major contributing factors to the paucity of data surrounding new chemicals. To address this issue, the U.S. Environmental Protection Agency (EPA), the European Chemicals Agency (ECHA), Health Canada, and other agencies are implementing measures that incorporate new approach methodologies (NAMs) into regulatory frameworks10, thus placing North American and European policy in line with international goals to replace, reduce, and refine the use of animals (the 3Rs principal)11,12,13,14. NAMs encompass a variety of assays primarily based on in vitro and in silico models that provide a mechanistic understanding of chemical toxicity instead of observing adversity inflicted on mammalian test species, thereby increasing the rate of data generation for chemical risk assessment while still producing high fidelity outputs15. However, these methods are not yet proven to safeguard against systemic toxicity, including the disruption of vital biological processes involving interorgan communication and endocrine signaling. Further, they cannot account for the bioaccumulation of chemicals within specific tissues, the ability of individual compounds to be absorbed and secreted, and the interplay between behavior and chemical exposure.
Due to the limitations of in vitro and computational models, the successful use of NAMs to reduce or replace mammalian models should also include invertebrate in vivo models, such as the fruit fly, Drosophila melanogaster. Previous studies in the fly have demonstrated that this organism is well suited for studying the conserved genetic pathways that protect animal cells against toxic molecules16,17,18,19,20,21,22. Moreover, the fly shows remarkable genetic similarity to humans, including functional homologs to over 65% of human diseases23,24,25 and an even greater conservation of important functional pathways26. These features, combined with their relatively short life cycle, low maintenance cost, and readily observable behavioral responses, make Drosophila well-suited for use as a toxicological model27,28,29,30. Moreover, flies have much higher throughput than rodent models and capture effects on metabolism, physiology, and hormone signaling that are not readily detectable by other non-organismal NAMs9.
The protocol described here represents a framework for testing the effects of chemical exposure on adult Drosophila. The method is designed to be efficient, inexpensive, and reproducible, while also minimizing the time researchers must be in contact with the test chemical and accommodating sample collection for metabolomics and other omics approaches. The protocol is optimized for testing a single chemical per experiment, but can easily accommodate other experimental parameters, such as varied solvents or combinations of chemicals.

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			<td>15 mL Centrifuge Tubes&#160;</td>
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			<td>5 ml Tubes&#160;</td>
			<td>VWR&#160;</td>
			<td>60818-576</td>
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			<td>Benchmark Dose Software version 3.2</td>
			<td>U.S. Environmental Protection Agency</td>
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			<td>Cardboard trays</td>
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			<td>trays and dividers for narrow vials&#160;</td>
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			<td>Combitips advanced, 50 mL</td>
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			<td>Combitips&#160;advanced, Biopur, 50&#160;mL, light gray, colorless tips</td>
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			<td>Cotton balls</td>
			<td>Genesee Scientific flystuff</td>
			<td>51-101</td>
			<td>Cotton balls, large, fits narrow vials&#160;</td>
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			<td>Delicate task wipes</td>
			<td>Kimtech</td>
			<td>34155</td>
			<td>Kimtech Science Kimwipes Delicate Task Wipes, 1 Ply / 8.2&#34; x 4.39&#34;</td>
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		<tr>
			<td>Drosophila Vial Plugs, Cellulose Acetate (aka, Flugs)</td>
			<td>VWR</td>
			<td>89168-888</td>
			<td>Wide</td>
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			<td>FD&#38;C Blue No. 1</td>
			<td>Spectrum Chemical</td>
			<td>FD110</td>
			<td>CAS number 3844-45-9</td>
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			<td>Flies&#160;</td>
			<td>BDSC</td>
			<td>Stock #2057&#160;</td>
			<td>OregonR wildtype&#160;</td>
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			<td>Gloves (nitrile)</td>
			<td>Kimtech</td>
			<td>55082/55081/55083</td>
			<td>Kimtech purple nitrile exam gloves, 5.9 mil, ambidextrous 9.5&#34;&#160;</td>
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			<td>Grade 1 CHR cellulose chromatography paper</td>
			<td>Cytvia</td>
			<td>3001-917</td>
			<td>Sheet, 46 x 57 cm</td>
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			<td>Mesh for humidity chamber</td>
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			<td>Multipette / Repeater (X) stream&#160;</td>
			<td>Eppendorf&#160;</td>
			<td>022460811</td>
			<td>Repeater Xstream&#160;</td>
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			<td>Plastic grate</td>
			<td>Plaskolite</td>
			<td>18469 (from lowes)</td>
			<td>Plaskolite 24 in x 48 in 7.85 sq ft louvered ceiling light panels, cut down to fit in rubbermaid tubs</td>
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		<tr>
			<td>Plastic trays for glass vials</td>
			<td>Genesee Scientific flystuff</td>
			<td>59-207</td>
			<td>Narrow fly vial reload tray&#160;</td>
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			<td>Polypropylene Drosophila Vial</td>
			<td>VWR</td>
			<td>75813-156</td>
			<td>Wide (28.5 mm)</td>
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			<td>Rubbermaid tubs</td>
			<td>Rubbermaid</td>
			<td>3769017 (from Lowes)</td>
			<td>Rubbermaid Roughneck Tote 10 gallon 18&#34; L x 12&#34; W x 8 1/2&#34; H</td>
		</tr>
		<tr>
			<td>Sucrose ultra pure</td>
			<td>MP Biomedicals, Inc.</td>
			<td>821721</td>
			<td></td>
		</tr>
		<tr>
			<td>Tube racks for wide-mouthed tubes</td>
			<td>Thermo scientific&#160;</td>
			<td>5970-0230</td>
			<td>Nalgene Unwire Test tube racks, for 30 mm tubes&#160;</td>
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			<td>Water Purification System</td>
			<td>Millipore Milli-Q</td>
			<td>ZMQ560F01</td>
			<td>Millipore Milli-Q Biocel Water Purifier&#160;</td>
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		<tr>
			<td>Yeast extract</td>
			<td>Acros Organics</td>
			<td>451120050</td>
			<td>CAS number 84604-16-0</td>
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assessment of chemical toxicity in adult drosophila melanogaster

Human industries generate hundreds of thousands of chemicals, many of which have not been adequately studied for environmental safety or effects on human health. This deficit of chemical safety information is exacerbated by current testing methods in mammals that are expensive, labor-intensive, and time-consuming. Recently, scientists and regulators have been working to develop new approach methodologies (NAMs) for chemical safety testing that are cheaper, more rapid, and reduce animal suffering. One of the key NAMs to emerge is the use of invertebrate organisms as replacements for mammalian models to elucidate conserved chemical modes of action across distantly related species, including humans. To advance these efforts, here, we describe a method that uses the fruit fly, Drosophila melanogaster,&#160;to assess chemical safety. The protocol describes a simple, rapid, and inexpensive procedure to measure the viability and feeding behavior of exposed adult flies. In addition, the protocol can be easily adapted to generate samples for genomic and metabolomic approaches. Overall, the protocol represents an important step forward in establishing Drosophila as a standard model for use in precision toxicology.

The fly has long served as a model in studies for determining sodium arsenite (NaAsO2) toxicity35,36,37,38. To demonstrate the efficacy of the protocol, male and female flies were exposed to NaAsO2, with the goal of comparing these results with earlier studies. Using the methodology described above, adult Oregon-R (BDSC stock #2057) males and females were exposed to a range of NaAsO2 concentrations (0, 0.01, 0.02, 0.1, 0.2, 1, and 2 mM) and scored for lethality 48 h after the start of exposure (Figure 1A,B).
The purpose of this initial analysis was to identify the approximate range of concentrations that would allow a more precise characterization of NaAsO2 toxicity. In subsequent experiments, concentrations were selected (0, 0.2, 0.5, 1, 1.5, 2, 2.5, 3, 3.5, and 5 mM) that more precisely defined the NaAsO2 dose-response curve (Figure 1C,D). Note that the resulting analysis examined several concentrations that induced 100% lethality. Data were analyzed using the Environmental Protection Agency&#39;s publicly available Benchmark Dose Software version 3.2.0.125. The data were modeled as &#34;dichotomous&#34; and the Dichotomous Hill model was used for subsequent analyses. Based on this model, the final LD10, LD25, and LD50 of male flies fed NaAsO2 were 0.30 mM, 0.50 mM, and 0.65 mM, respectively. For female flies, these values were slightly higher, with an LD10 of 0.30 mM, an LD25 of 0.65 mM, and an LD50 of 0.90 mM. Overall, the values obtained using this method are similar to those previously reported for arsenic toxicity in the Drosophila melanogaster35,36,37,38, thus validating the methodology.
In addition to the six replicates used to calculate the dose-response curve, male and female flies were also fed NaAsO2 exposure solutions that contained 1% FD&#38;C blue, which is easily visible in the digestive tract using light microscopy. Based on the presence of blue dye within the intestine of NaAsO2-fed flies, both male and female flies continued to feed 24 h after the beginning of the chemical exposure, regardless of the NaAsO2 concentration present within the liquid media (Figure 2). However, the ingested food was observed to be occasionally regurgitated at doses above 0.2 mM for females and 0.5 mM for males (Figure 2). These findings suggest that regurgitation could serve a key role in the Drosophila response to arsenic poisoning.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65029/65029fig01.jpg" /> 
Figure 1: Dose-response curves for male and female Drosophila treated with NaAsO2 for 48 h. All graphs show the estimated proportions of dead flies at each NaAsO2 concentration tested based on the Dichotomous Hill model. (A,B) A broad range of NaAsO2 concentrations were tested to approximate the dose at which each sex of fly begins to die. (A) shows the male data, and (B) shows the female data. N = 4 vials, with 20 flies per vial. (C,D) A narrower range of NaAsO2 concentrations was tested to determine precise doses at which 10%, 25%, and 50% of each sex of flies died. These doses are indicated to the right of each graph. (C) shows the male data, and (D) shows the female data. N = 6 vials, with 20 flies per vial. <a href="https://www.jove.com/files/ftp_upload/65029/65029fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65029/65029fig02.jpg" /> 
Figure 2: Representative results from the blue dye assay of male and female Drosophila treated with NaAsO2 for 24 h. Micrographs show flies fed increasing concentrations of NaAsO2. Row (A) shows male flies, and row (B) shows female flies, with the concentration of NaAsO2 increasing from left to right. The abdomen shows a small amount of blue near the thorax at low concentrations, indicating that exposure media entered the gut. At higher concentrations, blue dye begins to accumulate around the mouth, suggesting that exposure media is being regurgitated. The scale bar is 1 mm. <a href="https://www.jove.com/files/ftp_upload/65029/65029fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Assessment of Chemical Toxicity in Adult Drosophila Melanogaster at JoVE.com

Assessment of Chemical Toxicity in Adult Drosophila Melanogaster

This protocol describes an efficient and inexpensive method that uses liquid media to assess the effects of chemical toxicants on the viability of adult Drosophila melanogaster.

Assessment of Chemical Toxicity in Adult Drosophila Melanogaster

Human industries generate hundreds of thousands of chemicals, many of which have not been adequately studied for environmental safety or effects on human ...

assessment-of-chemical-toxicity-in-adult-drosophila-melanogaster

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JoVE Journal

Biology

3.1K Views.  Indiana University. This protocol describes an efficient and inexpensive method that uses liquid media to assess the effects of chemical toxicants on the viability of adult Drosophila melanogaster. 

Video: Assessment of Chemical Toxicity 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.

Dissection z larw CNS na 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 ...

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 ...

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 ...

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 ...