This publication was supported by a grant from the Department of Education (PR Award #P031C160025-17, Project title: 84.031C) to the Colorado State University-Pueblo (CSU-Pueblo) Communities to Build Active STEM Engagement (C-BASE). We thank Current Zoology and Elsevier for providing the rights to use the representative results published in previous papers, as well as the editors of JoVE for providing us with the opportunity to publish this protocol. We would also like to thank the C-BASE Program, Dr. Brian Vanden Heuvel (C-BASE and Department of Biology, CSU-Pueblo), CSU-Pueblo Biology department, Thomas Graziano, Dr. Bernard Possidente (Department of Biology, Skidmore College), and Dr. Claire Varian Ramos (Department of Biology, Colorado State University-Pueblo) for their support and assistance.

The following protocol is an experimental protocol used to rear species in the Drosophila genus on contaminated medium when oral ingestion of a toxin is appropriate; other forms of exposure are possible using the Drosophila model11,15,16,26. The methods described in this protocol have been previously described by Hirsch et al.19 and Peterson et al.23,24,25.
1. Set Up Stock Populations of Drosophila in the Research Laboratory
<ol>
	<li>Set up an environmentally-controlled incubator (or small room) to house stock populations of Drosophila by setting the incubators for a constant temperature, light:dark cycle, and humidity, depending upon the preferences of the test species. 
	NOTE: Preferred environmental conditions will vary depending upon the native ecology of the species chosen for the study. For example, D. melanogaster is native to sub-Saharan Africa27 and is typically maintained at 25 &#176;C, 12:12 light:dark cycle, and approximately 60% humidity16,18,19,20,21,22,23,24,25,28,29,30. On the other hand, D. montana range extends throughout most of Canada and the midwest USA, a much colder region; therefore, D. montana is typically maintained at 19&#8211;20 &#176;C and sometimes a 24 h light regime to simulate conditions during the mating season31. For a more detailed description of the geographic ranges of various species of Drosophila, see the Drosophila Speciation Patterns website32.</li>
	<li>Obtain a preferred Drosophila species and/or inbred line from either a stock center (see Table of Materials), another research laboratory upon request, or collect wild, genetically variable populations from the field. 
	NOTE: The following steps explain the methods to collect wild, genetically variable populations of Drosophila to maintain in the research laboratory. These methods have been modified from Markow and O&#39;Grady33 and Werner and Jaenike34 to collect the widest diversity of species at once, rather than target particular species with one bait source.
	<ol>
		<li>Freeze half a dozen ripe bananas in the freezer overnight and defrost before setting bait traps.</li>
		<li>Prepare multiple 1&#8211;2 L plastic bottles by cutting a u-shaped slit in the front of the bottle to allow flies to be captured in the bait bottle and not escape. Cap the plastic bottles with their bottle caps so the flies do not escape via the lids.</li>
		<li>Add the defrosted banana to the bottom of the bottles so that the bottom of the bottles contains approximately one-inch of banana. Place a slice of ripe tomato in the bottle. Add a yeast slurry (the leftover yeast from the beer making process) to the banana at the bottom of the bottle so that the banana gets to soak in the yeast slurry.</li>
		<li>Add wooden sticks (in an upright vertical position) to the bottle so the flies have a clean substrate to walk on away from the yeast slurry and banana. 
		Figure 1 illustrates the final product of these methods.</li>
		<li>Hang bait bottles in trees overnight and check every 24 h. Mouth aspirate flies out of bottles and individually place females in vials with medium to create iso-female lines. 
		NOTE: Multi-female lines can be created, however, only if the females of each species can be clearly identified. In addition, flies within the genus Drosophila occupy different ecological niches and will have different dietary requirements depending upon those niches (Werner and Jaenike34); see Werner and Jaenike34 for diet recommendations and food recipes.</li>
		<li>Examine the adult F1 offspring under the dissection microscope to identify the species of the collected Drosophila (see Markow and O&#39;Grady33 and Werner and Jaenike34 for assistance in identifying various species).</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57450/57450fig1.jpg" /> 
Figure 1: Pictorial representation of traps and bait used to collect wild populations of Drosophila in the field. (A) Fly traps set at a local field site in Colorado. (B) A closer view of the fly traps set at this field site. <a href="/files/ftp_upload/57450/57450fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="3">
	<li>Maintain the iso-female or the multi-female lines in an environmentally-controlled incubator or room with constant temperature, light:dark cycle, and humidity. To do this, house flies in vials or bottles in preferred medium and allow the gravid females to lay eggs in the medium. Monitor the vials for the presence of larvae and pupae. 
	NOTE: Flies within the genus Drosophila occupy different ecological niches and will have different dietary requirements and environmental abiotic preferences depending upon those niches33,34. Environmental preferences and dietary recommendations (and further instruction on fly husbandry) can be found in Elgin and Miller28, Shaffer et al.29, Stocker and Gallant30, Markow and O&#39;Grady33, and Werner and Jaenike34. If using wild-caught species, local environmental conditions can be simulated in the incubators until the species can be identified.</li>
	<li>Transfer stocks frequently to fresh medium, discarding old vials, to maintain healthy lines and avoid infection from mites. 
	NOTE: The frequency of transfer will depend on the life cycle of the species. For example, transfer Drosophila melanogaster every 2 weeks to fresh medium.For further information on maintaining lines in the laboratory, see Rand et al.16, Elgin and Miller28, Shaffer et al.29, Stocker and Gallant30, Greenspan35, and Science Education Database36.</li>
</ol>
2. Rear Drosophila in the Contaminated Medium
NOTE: If testing Drosophila in the laboratory for the first time or with a new contaminant(s), identify the lethal dose (see Castaneda et al.37 and Massie et al.38 for methods) and the LD50 (see Castaneda et al.37 and Akins et al.39 for methods) first. Then, run a dose-response curve to identify biologically-relevant concentrations for the desired phenotypic output; see Hirsch et al.19 and Zhou et al.40 for methods.
<ol>
	<li>Prepare stock solutions of the contaminated medium at the desired concentration(s), depending upon the chemistry of the contaminant. 
	NOTE: For example, to prepare stock solutions of PbAc: Prepare stock solutions of lead acetate (PbAc) medium by adding contaminant to distilled water (dH20) until medium made with contaminant water reaches desired concentration. For example, a stock solution of 1,000 &#181;M PbAc, can be prepared by adding PbAc to dH20 until it reaches 1,000 &#181;M PbAc. Further, dilute the stock solution (e.g. the 1,000 &#181;M PbAc solution) to the desired concentration (such as 500 &#181;M PbAc) and maintain these solutions as stock as well.</li>
	<li>Prepare medium, following manufacturer&#39;s guidelines to serve as the control medium. Prepare additional medium, following manufacturer&#39;s guidelines; however, supplement prepared contaminant solution for dH20. 
	NOTE: For example, if using an instant Drosophila medium, add approximately one teaspoon instant medium to a plastic vial. Add approximately 5&#8211;5.5 mL dH20 to the medium. Sprinkle a few grains of live baker&#39;s yeast to prepare control medium. To prepare experimental medium, supplement the stock solution (such as 500 &#181;M PbAc) for dH20.</li>
	<li>Transfer reproductively viable mature males and females from stock populations into the control and the experimental medium. 
	NOTE: The time post-eclosion to reproductive maturity is different between the Drosophila species41.
	<ol>
		<li>Gently tap the vial of stock flies down with the dominant hand. Ensure that the flies automatically move to the bottom of the vial. With the other hand, remove the cap of the vial while tapping the vial and place a fresh vial of control or contaminated medium on top of the vial with the flies. Hold the vials together and flip them over, gently tapping, so that the flies automatically are transferred to the fresh vial of control or contaminated medium. While still tapping the vial with the flies, cap the vial.</li>
		<li>Repeat with more vials, making sure to standardize the number of flies in each vial. 
		NOTE: The total number of adults transferred via single transfer or anesthesia will depend on the size of the vials used to avoid overcrowding.</li>
		<li>Incubate adults in a standard environmental condition (i.e. an incubator) and allow the adults to mate and lay eggs in the medium for 24&#8211;96 h.</li>
		<li>After 24&#8211;96 h, discard adults in a morgue (a flask filled with mineral oil and capped with a tight-fitting funnel) leaving behind fertilized eggs (which will later become the experimental subjects) to mature for testing. Place the vials in the incubator to allow the eggs to develop.</li>
		<li>Monitor the vials for wandering-instar larvae by looking for larvae that are emerging from the medium.</li>
	</ol>
	</li>
</ol>
3. Collect Experimental Subjects at Various Developmental Stages
NOTE: Experimental subjects can be collected at any developmental stage, placed in the blind coded 15-mL conical tubes, and tested for accumulation. Methods for testing the accumulation of contaminants will depend on the contaminant being studied. For example, accumulation of PbAc can be tested using Inductively-Coupled Plasma Mass Spectrometry (ICP-MS)42. In addition, experimental subjects can be collected at any developmental stage to be tested for a variety of phenotypic effects of contaminants. Figure 2 illustrates the Drosophila life cycle43. Figure 3 illustrates the experimental protocol for exposure and the different developmental stages for collection.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57450/57450fig2.jpg" /> 
Figure 2: Conceptual overview of the life cycle of D. melanogaster (the most commonly used Drosophila model system). The stages of Drosophila life cycle are: 1) egg, 2) first-instar larva, 3) second-instar larva, 4) third-instar larva, 5) wandering third-instar larva, 6) white-eye pupa, 7) red-eye pupa, 8) newly-eclosed adult, and 9) mature adult. <a href="/files/ftp_upload/57450/57450fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57450/57450fig3.jpg" /> 
Figure 3: Conceptual overview of the methods for orally exposing Drosophila to contaminated medium in both the parental (F0) and subsequent generations (F1 and onward). (A) Methods for oral exposure during development in the exposed generation. (B) Methods to test the transfer of contaminants to offspring (F1 to the desired generation). This figure has been modified from Peterson et al.24 <a href="/files/ftp_upload/57450/57450fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Collect wandering-third instar larvae
	<ol>
		<li>Start monitoring vials when lights turn on in the incubator, as larvae will emerge from the medium and move upwards on the side of the vial within an h after lights turn on in the incubator. Within this h, remove the wandering-third instar larvae from the sides of the vial carefully using a wooden stick or tweezers. 
		NOTE: The number of larvae available for collection will depend on the number of eggs laid in &#34;2.3.4&#34;.</li>
		<li>To remove excess medium from the larvae, place the larvae in a small beaker with dH2O. Pour the dH2O out of the beaker and place the larvae on a delicate task wiper. Using a delicate task wiper, gently remove the excess dH2O from the larvae.</li>
		<li>Maintain experimental populations in an environmentally-controlled incubator.</li>
	</ol>
	</li>
	<li>Collect newly-eclosed adults
	<ol>
		<li>Monitor vials for eclosion by observing the coloration of the pupae along the sides of the vials. 
		NOTE: Pupae will darken during development. Developmental time, particularly pre-eclosion, depends on the species tested.</li>
		<li>When the first adults begin to eclose, dump and discard these adults into a morgue containing mineral oil.</li>
		<li>When the lights turn on in the incubator the following morning, dump and discard any adults of unknown age (or virginity) that may have eclosed overnight or during the morningbefore lights on.</li>
		<li>Approximately 4 h later, anesthetize any adults that emerged as newly-eclosed adults with a CO2 gun in the vials. Place adults on a CO2 plate under a dissection microscope. Sex adults by looking for sex combs on the forelimbs of males and ovipositors in females. 
		NOTE: D. melanogaster must be collected within 6 h of eclosion to avoid mating but other species may have longer developmental times (and therefore, do not need to be collected within this time frame).</li>
		<li>Separate adults on the CO2 plate using a wooden stick. Gently transfer adults in sex-specific groups using a wooden stick to the medium matching pre-existing history.</li>
	</ol>
	</li>
	<li>Collect mature adults post-eclosion
	<ol>
		<li>Allow adults to remain on the medium matching pre-eclosion exposure from egg stage to the desired age post-eclosion in an environmentally-controlled incubator.</li>
		<li>Singly transfer adults to the control medium for 24 h prior to testing to allow adults to groom excess contaminated medium off their bodies.</li>
	</ol>
	</li>
</ol>
4. Rear Experimental Subjects to Test the Effects of Multigenerational or Transgenerational Exposure.
<ol>
	<li>To rear the parental generation (a.k.a the P0 or F0 generations), transfer adults from stock populations to control and the experimental medium following the steps in &#34;2.1&#34; to &#34;2.3&#34; and &#34;3.1&#34; to &#34;3.3&#34;.</li>
	<li>When the adults are reproductively mature (see Pitnick et al.41), singly transfer (as stated in 2.3.1) one vial of males to a fresh vial of control or experimental medium. Singly transfer one vial of females to the fresh vial that now contains males. Allow adults to mate and lay eggs in the medium for 24-96 h. Dump and discard adults into a morgue containing mineral oil and re-incubate vials to allow offspring to develop.</li>
	<li>Repeat steps 4.1 through 4.2 depending on the desired number of generations.</li>
</ol>

The authors have nothing to disclose.

Drosophila melanogaster has been established as a powerful model for a range of biological processes due to the extensive conservation of genes and pathways between D. melanogaster and humans13,14. For the same reasons that it&#39;s a powerful model for medical science, Drosophila has emerged as a suitable model system to study the impact of anthropogenic pollution on a range of toxicological endpoints. Several laboratories are successf...

Every year, approximately 1,000 new chemicals are introduced by the chemical industry1,2; however, the environmental impacts of only a small percentage of these chemicals are tested before distribution2,3. Although large-scale catastrophes are uncommon, sublethal and chronic exposure to a large variety of pollutants are widespread in both humans and wildlife4,5. The historical focus of ecotoxicology and environmental toxicology was to test lethality, single chemical exposure, acute exposure, and the physiological effects of exposure, as a means of measuring the impact of pollutants on survival6,7,8,9,10. Although there is a shift towards ethical and non-invasive approaches to animal testing, current approaches are limiting because of the role that development, emergent properties, and external factors (such as population-level and ecosystem-level interactions) play in mediating ecologically-important endpoints8. Therefore, there is a need for methods that incorporate a more holistic approach without sacrificing wildlife and/or vertebrates in the laboratory.
Invertebrate model systems, such as Drosophila melanogaster, are an attractive alternative to address the need for a more holistic approach to toxicity testing. D. melanogaster, was originally developed as an invertebrate model system for human-related genetic research about a century ago11.D. melanogaster is now prominently used as a vertebrate model alternative for several reasons: 1) the conservation of genes and pathways between D. melanogaster and humans; 2) short generation time compared to vertebrate models; 3) inexpensive cost of maintenance; 4) ease in generating large sample sizes; and 5) plethora of phenotypic- and ecologically-relevant endpoints available for testing11,12,13,14,15,16,17.
Several laboratories11,15,16,17,18,19,20,21,22,23,24,25 are now using D. melanogaster as a vertebrate model alternative for toxicity testing to understand the impacts of pollution on humans. Local wild species of Drosophila can be utilized, as well, as toxicity models for wildlife (and humans) to answer ecologically-, behaviorally-, and evolutionarily-relevant questions at multiple biological levels of organization. Using species within the Drosophila genus as a model, several measurable endpoints are possible11,15,16,18,19,20,21,22,23,24,25. In addition, using the Drosophila model, toxicologists can: 1) ethically link effects at multiple biological levels of the organization; 2) incorporate the role of emergent factors and development; 3) study ecologically-important endpoints (in addition to medically-important endpoints); 4) test multiple stressors simultaneously; 5) and test long-term multigenerational (e.g. evolutionary and transgenerational) implications of stressors. Therefore, using Drosophila as a model system enables a multitude of approaches, not limited to studying mechanistic approaches with inbred strains of D. melanogaster in the laboratory.
In this paper, we present the methods for rearing and collecting Drosophila to answer various toxicological questions. More specifically, we describe the methodology for 1) rearing Drosophila in medium laced with one or more pollutants; 2) collecting Drosophila throughout development (e.g. wandering third-instar larvae, pupal cases, newly-eclosed adults, and mature adults); and 3) rearing Drosophila in the contaminated medium to test intergenerational and transgenerational transmission, as well as evolutionary implications of long-term toxicant exposure. Using this protocol, previous authors18,19,20,21,22,23,24,25 have reported different physiological, genetic, and behavioral effects of developmental lead (Pb2+) exposure. This protocol enables toxicologists to use a more holistic toxicological approach, which is essential to understanding how pollutants are risk factors for both humans and wildlife in an ever increasingly polluted environment.

<table><tbody><tr><td>Carolina Biological Instant Drosophila Medium Formula 4-24</td><td>Carolina Biological</td><td>173204</td><td></td></tr><tr><td>Drosophila vials, Narrow (PS), Polystyrene, Superbulk, 1000 vials/unit</td><td>Genessee Scientific</td><td>32-116SB</td><td>Used to store flies</td></tr><tr><td>Flugs Closures for vials and bottles, Narrow plastic vials</td><td>Genessee Scientific</td><td>49-102</td><td>Used to store flies</td></tr><tr><td>Cardboard trays, trays only, narrow</td><td>Genessee Scientific</td><td>32-124</td><td>Used to organize populations of flies</td></tr><tr><td>Cardboard trays, dividers only, narrow</td><td>Genessee Scientific</td><td>32-126</td><td>Used to organize populations of flies</td></tr><tr><td>Thermo Scientific&amp;nbsp;Nalgene Square Wide-Mouth HDPE Bottles with Closure</td><td>Fischer Scientific</td><td>03-312D</td><td>Useful for storage of contaminants</td></tr><tr><td>Thermo Scientific&amp;nbsp;Nalgene Color-Coded LDPE Wash Bottles</td><td>Fischer Scientific</td><td>03-409-17C</td><td>Useful for storage of contaminants</td></tr><tr><td>Eppendorf&amp;nbsp;Repeater M4 Manual Handheld Pipette Dispenser</td><td>Fischer Scientific</td><td>14-287-150</td><td>Used to prepare medium</td></tr><tr><td>Combitips Advanced Pipetter Tips - Standard, Eppendorf Quality Tips</td><td>Fischer Scientific</td><td>13-683-708</td><td>Used to prepare medium</td></tr><tr><td>Flypad, Standard Size (8.1 X 11.6cm)</td><td>Genessee Scientific</td><td>59-114</td><td>Used to anesthetize flies</td></tr><tr><td>Flystuff foot valve</td><td>Genessee Scientific</td><td>59-121</td><td>Used to anesthetize flies</td></tr><tr><td>Tubing, green (1 continguous foot/unit)</td><td>Genessee Scientific</td><td>59-124G</td><td>Used to anesthetize flies</td></tr><tr><td>Mineral Oil, Light, White, High Purity Grade, 500 mL HDPE Bottle</td><td>VWR</td><td>97064-130</td><td>Used to make a morgue</td></tr><tr><td>Glass Erlenmeyer Flask Set - 3 Sizes - 50, 150 and 250ml, Karter Scientific 214U2</td><td>Walmart</td><td>Not applicable</td><td>Used to make a morgue</td></tr><tr><td>BGSET5 Glass Beaker Set Of 5</td><td>Walmart</td><td></td><td></td></tr><tr><td>Inbred or wildtype line of &lt;em&gt;Drosophila&lt;/em&gt;</td><td>Bloomington &lt;em&gt;Drosophila&lt;/em&gt; Stock Center at Indiana University</td><td></td><td>https://bdsc.indiana.edu</td></tr><tr><td>Wild popultions of &lt;em&gt;Drosophila&lt;/em&gt;</td><td>UC San Diego &lt;em&gt;Drosophila&lt;/em&gt; Stock Center</td><td></td><td>https://stockcenter.ucsd.edu/info/welcome.php</td></tr></tbody></table>

experimental protocol for using drosophila as an invertebrate model system for toxicity testing in the laboratory

Emergent properties and external factors (population-level and ecosystem-level interactions, in particular) play important roles in mediating ecologically-important endpoints, though they are rarely considered in toxicological studies. D. melanogaster is emerging as a toxicology model for the behavioral, neurological, and genetic impacts of toxicants, to name a few. More importantly, species in the genus Drosophila can be utilized as a model system for an integrative framework approach to incorporate emergent properties and answer ecologically-relevant questions in&#160;toxicology research. The aim of this paper is to provide a protocol for exposing species in the genus Drosophila to pollutants to be used as a model system for a range of phenotypic outputs and ecologically-relevant questions. More specifically, this protocol can be used to 1) link multiple biological levels of organization and understand the impact of toxicants on both individual- and population-level fitness; 2) test the impact of toxicants at different stages of developmental exposure; 3) test multigenerational and evolutionary implications of pollutants; and 4) test multiple contaminants and stressors simultaneously.

By orally exposing Drosophila to a contaminant(s) throughout development, various toxicological questions can be tested by exposing Drosophila at different levels of biological organization. This section presents representative results obtained using this protocol in previously published papers23,24. In particular, this protocol was previously used to evaluate the accumulation, elimination and sequestration of le...

Watch this Scientific Journal Video about Experimental Protocol for Using Drosophila As an Invertebrate Model System for Toxicity Testing in the Laboratory at JoVE.com

Experimental Protocol for Using Drosophila As an Invertebrate Model System for Toxicity Testing in the Laboratory

In this paper, we provide a detailed protocol for exposing species in the genus Drosophila to pollutants with the goal of studying the impact of exposure on a range of phenotypic outputs at different developmental stages and for more than one generation.

Experimental Protocol for Using Drosophila As an Invertebrate Model System for Toxicity Testing in the Laboratory

Emergent properties and external factors (population-level and ecosystem-level interactions, in particular) play important roles in mediating ...

experimental-protocol-for-using-drosophila-as-an-invertebrate-model

Research

JoVE Journal

Biology

12.6K Views.  Colorado State University-Pueblo. In deze paper bieden wij een gedetailleerd protocol voor het blootstellen van de soort van het geslacht Drosophila aan verontreinigende stoffen, met het doel van het bestuderen van de impact van blootstelling over een scala van fenotypische resultaten op verschillende ontwikkelingsstadia en voor meer dan één generatie.

Video: Experimental Protocol for Using Drosophila As an Invertebrate Model System for Toxicity Testing in the Laboratory

Drosophila Adult Olfactory Shock Learning

Drosophila have been used in classical conditioning experiments for over 40 years, thus greatly facilitating our understanding of memory, including the elucidation of the molecular mechanisms involved in cognitive diseases1-7. Learning and memory can be assayed in larvae to study the effect of neurodevelopmental genes8-10 and in flies to measure the contribution of adult plasticity genes1-7. Furthermore, the short lifespan of Drosophila facilitates the analysis of genes mediating age-related memory impairment5,11-13. The availability of many inducible promoters that subdivide the Drosophila nervous system makes it possible to determine when and where a gene of interest is required for normal memory as well as relay of different aspects of the reinforcement signal3,4,14,16.
Studying memory in adult Drosophila allows for a detailed analysis of the behavior and circuitry involved and a measurement of long-term memory15-17. The length of the adult stage accommodates longer-term genetic, behavioral, dietary and pharmacological manipulations of memory, in addition to determining the effect of aging and neurodegenerative disease on memory3-6,11-13,15-21.
Classical conditioning is induced by the simultaneous presentation of a neutral odor cue (conditioned stimulus, CS+) and a reinforcement stimulus, e.g., an electric shock or sucrose, (unconditioned stimulus, US), that become associated with one another by the animal1,16. A second conditioned stimulus (CS-) is subsequently presented without the US. During the testing phase, Drosophila are simultaneously presented with CS+ and CS- odors. After the Drosophila are provided time to choose between the odors, the distribution of the animals is recorded. This procedure allows associative aversive or appetitive conditioning to be reliably measured without a bias introduced by the innate preference for either of the conditioned stimuli. Various control experiments are also performed to test whether all genotypes respond normally to odor and reinforcement alone.

Drosophila Adult Olfactory Shock Learning

The method to measure adult Drosophila associative memory is described. The assay is based on the ability of the fly to associate an odor presented with a negative reinforcer (electric shock) and then recall this information at a later time, allowing memory to be measured.

Drosophila have been used in classical conditioning experiments for over 40 years, thus greatly facilitating our understanding of memory, including the ...

Neuroscience

Methods to Test Endocrine Disruption in Drosophila melanogaster

In recent years there has been growing evidence that all organisms and the environment are exposed to hormone-like chemicals, known as endocrine disruptor chemicals (EDCs). These chemicals may alter the normal balance of endocrine systems and lead to adverse effects, as well as an increasing number of hormonal disorders in the human population or disturbed growth and reduced reproduction in the wildlife species. For some EDCs, there are documented health effects and restrictions on their use. However, for most of them, there is still no scientific evidence in this sense. In order to verify potential endocrine effects of a chemical in the full organism, we need to test it in appropriate model systems, as well as in the fruit fly, Drosophila melanogaster. Here we report detailed in vivo protocols to study endocrine disruption in Drosophila, addressing EDC effects on the fecundity/fertility, developmental timing, and lifespan of the fly. In the last few years, we used these Drosophila life traits to investigate the effects of exposure to 17-&#945;-ethinylestradiol (EE2), bisphenol A (BPA), and bisphenol AF (BPA F). Altogether, these assays covered all Drosophila life stages and made it possible to evaluate endocrine disruption in all hormone-mediated processes. Fecundity/fertility and developmental timing assays were useful to measure the EDC impact on the fly reproductive performance and on developmental stages, respectively. Finally, the lifespan assay involved chronic EDC exposures to adults and measured their survivorship. However, these life traits can also be influenced by several experimental factors that had to be carefully controlled. So, in this work, we suggest a series of procedures we have optimized for the right outcome of these assays. These methods allow scientists to establish endocrine disruption for any EDC or for a mixture of different EDCs in Drosophila, although to identify the endocrine mechanism responsible for the effect, further essays could be needed.

Methods to Test Endocrine Disruption in Drosophila melanogaster

Endocrine disruptor chemicals (EDCs) represent a serious problem for organisms and for natural environments. Drosophila melanogaster represents an ideal model to study EDC effects in vivo. Here, we present methods to investigate endocrine disruption in Drosophila, addressing EDC effects on fecundity, fertility, developmental timing, and lifespan of the fly.

In recent years there has been growing evidence that all organisms and the environment are exposed to hormone-like chemicals, known as endocrine disruptor ...

Developmental Biology

Functional Analysis of the Larval Feeding Circuit in Drosophila

The serotonergic feeding circuit in Drosophila melanogaster larvae can be used to investigate neuronal substrates of critical importance during the development of the circuit. Using the functional output of the circuit, feeding, changes in the neuronal architecture of the stomatogastric system can be visualized. Feeding behavior can be recorded by observing the rate of retraction of the mouth hooks, which receive innervation from the brain. Locomotor behavior is used as a physiological control for feeding, since larvae use their mouth hooks to traverse across an agar substrate. Changes in feeding behavior can be correlated with the axonal architecture of the neurites innervating the gut. Using immunohistochemistry it is possible to visualize and quantitate these changes. Improper handling of the larvae during behavior paradigms can alter data as they are very sensitive to manipulations. Proper imaging of the neurite architecture innervating the gut is critical for precise quantitation of number and size of varicosities as well as the extent of branch nodes. Analysis of most circuits allow only for visualization of neurite architecture or behavioral effects; however, this model allows one to correlate the functional output of the circuit with the impairments in neuronal architecture.

Functional Analysis of the Larval Feeding Circuit in Drosophila

The feeding circuit in Drosophila melanogaster larvae serves a simple yet powerful model that allows changes in feeding rate to be correlated with alterations in the stomatogastric neural circuitry. This circuit is composed of central serotonergic neurons that send projections to the mouth hooks as well as the foregut.

The serotonergic  feeding circuit in Drosophila  melanogaster larvae can be used to investigate neuronal substrates of  critical importance during the ...

An Affordable and Efficient "Homemade" Platform for Drosophila Behavioral Studies, and an Accompanying Protocol for Larval Mitochondrial Respirometry

The usefulness of Drosophila as a model organism for the study of human diseases, behaviors and basic biology is unquestionable. Although practical, Drosophila research lacks popularity in developing countries, possibly due to the misinformed idea that establishing a lab and performing relevant experiments with such tiny insects is difficult and requires expensive, specialized apparatuses. Here, we describe how to build an affordable flylab to quantitatively analyze a myriad of behavioral parameters in D. melanogaster, by 3D-printing many of the necessary pieces of equipment. We provide protocols to build in-house vial racks, courtship arenas, apparatuses for locomotor assays, etc., to be used for general fly maintenance and to perform behavioral experiments using adult flies and larvae. We also provide protocols on how to use more sophisticated systems, such as a high resolution oxygraph, to measure mitochondrial oxygen consumption in larval samples, and show its association with behavioral changes in the larvae upon the xenotopic expression of the mitochondrial alternative oxidase (AOX). AOX increases larval activity and mitochondrial leak respiration, and accelerates development at low temperatures, which is consistent with a thermogenic role for the enzyme. We hope these protocols will inspire researchers, especially from developing countries, to use Drosophila to easily combine behavior and mitochondrial metabolism data, which may lead to information on genes and/or environmental conditions that may also regulate human physiology and disease states.

An Affordable and Efficient "Homemade" Platform for Drosophila Behavioral Studies, and an Accompanying Protocol for Larval Mitochondrial Respirometry

We provide protocols for anyone with a "maker culture" mind to start building a flylab for quantitative analysis of a myriad of behavioral parameters in Drosophila melanogaster, by 3D-printing many of the necessary pieces of equipment. We also describe a high resolution respirometry protocol using larvae to combine behavioral and mitochondrial metabolism data.

The usefulness of Drosophila as a model organism for the study of human diseases, behaviors and basic biology is unquestionable. Although practical, ...

Behavior

Taste Preference Assay for Adult Drosophila

Olfactory and gustatory perception of the environment is vital for animal survival. The most obvious application of these chemosenses is to be able to distinguish good food sources from potentially dangerous food sources. Gustation requires physical contact with a chemical compound which is able to signal through taste receptors that are expressed on the surface of neurons. In insects, these gustatory neurons can be located across the animal's body allowing taste to play an important role in many different behaviors. Insects typically prefer compounds containing sugars, while compounds that are considered bitter tasting are avoided. Given the basic biological importance of taste, there is intense interest in understanding the molecular mechanisms underlying this sensory modality. We describe an adult Drosophila taste assay which reflects the preference of the animals for a given tastant compound. This assay may be applied to animals of any genetic background to examine the taste preference for a desired soluble compound.

Taste Preference Assay for Adult Drosophila

Taste is an important sensory process which facilitates attraction to beneficial substances and avoidance of toxic substances. This protocol describes a simple ingestion assay for determining Drosophila gustatory preference for a given chemical compound.

Olfactory and gustatory perception of the environment is vital for animal survival.  The most obvious application of these chemosenses is to be able to ...

Toxicity Study of Zinc Oxide Nanoparticles in Cell Culture and in Drosophila melanogaster

Zinc oxide nanoparticles (ZnO NPs) have a wide range of applications, but the number of reports on ZnO NP-associated toxicity has grown rapidly in recent years. However, studies that elucidate the underlying mechanisms for ZnO NP-induced toxicity are scanty. We determined the toxicity profiles of ZnO NPs using both in vitro and in vivo experimental models. A significant decrease in cell viability was observed in ZnO NP-exposed MRC5 lung fibroblasts, showing that ZnO NPs exert cytotoxic effects. Similarly, interestingly, gut exposed to ZnO NPs exhibited a dramatic increase in reactive oxygen species levels (ROS) in the fruit fly Drosophila. More in-depth studies are required to establish a risk assessment for the increased usage of ZnO NPs by consumers.

Toxicity Study of Zinc Oxide Nanoparticles in Cell Culture and in Drosophila melanogaster

We describe a detailed protocol for evaluating the toxicological profiles of zinc oxide nanoparticles (ZnO NPs) in particular, the type of cell death in human MRC5 lung fibroblasts and ROS formation in the fruit fly Drosophila.

Zinc oxide nanoparticles (ZnO NPs) have a wide range of applications, but the number of reports on ZnO NP-associated toxicity has grown rapidly in recent ...

Immunology and Infection

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar Drosophila Melanogaster

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield deleterious consequences, yet the current methods for recording nervous system activity of an individual animal is costly and laborious. This suction electrode preparation of the third-instar larval central nervous system of Drosophila melanogaster, is a tractable system for testing the physiological effects of neuroactive agents, determining the physiological role of various neural pathways to CNS activity, as well as the influence of genetic mutations to neural function. This ex vivo preparation requires only moderate dissecting skill and electrophysiological expertise to generate reproducible recordings of insect neuronal activity. A wide variety of chemical modulators, including peptides, can then be applied directly to the nervous system in solution with the physiological saline to measure the influence on the CNS activity. Further, genetic technologies, such as the GAL4/UAS system, can be applied independently or in tandem with pharmacological agents to determine the role of specific ion channels, transporters, or receptors to arthropod CNS function. In this context, the assays described herein are of significant interest to insecticide toxicologists, insect physiologists, and developmental biologists for which D. melanogaster is an established model organism. The goal of this protocol is to describe an electrophysiological method to enable the measurement of electrogenesis of the central nervous system in the model insect, Drosophila melanogaster, which is useful for testing a diversity of scientific hypotheses.

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar  Drosophila Melanogaster

This protocol describes a method to record the descending electrical activity of the Drosophila melanogaster central nervous system to enable the cost-efficient and convenient testing of pharmacological agents, genetic mutations of neural proteins, and/or the role of unexplored physiological pathways.

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield ...

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

The regrowth capacity of damaged neurons governs neuroregeneration and functional recovery after nervous system trauma. Over the past few decades, various intrinsic and extrinsic inhibitory factors involved in the restriction of axon regeneration have been identified. However, simply removing these inhibitory cues is insufficient for successful regeneration, indicating the existence of additional regulatory machinery. Drosophila melanogaster, the fruit fly, shares evolutionarily conserved genes and signaling pathways with vertebrates, including humans. Combining the powerful genetic toolbox of flies with two-photon laser axotomy/dendriotomy, we describe here the Drosophila sensory neuron &#8211; dendritic arborization (da) neuron injury model as a platform for systematically screening for novel regeneration regulators. Briefly, this paradigm includes a) the preparation of larvae, b) lesion induction to dendrite(s) or axon(s) using a two-photon laser, c) live confocal imaging post-injury and d) data analysis. Our model enables highly reproducible injury of single labeled neurons, axons, and dendrites of well-defined neuronal subtypes, in both the peripheral and central nervous system.

A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System

Here, we present a protocol using the Drosophila sensory neuron - dendritic arborization (da) neuron injury model, which combines in vivo live imaging, two-photon laser axotomy/dendriotomy, and the powerful fly genetic toolbox, as a platform for screening potential promoters and inhibitors of neuroregeneration.

The regrowth capacity of damaged neurons governs neuroregeneration and functional recovery after nervous system trauma. Over the past few decades, various ...

Medicine

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.

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.

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

Lifespan Protocol for Drosophila melanogaster: Generating Survival Curves to Identify Differences in Fly Longevity

Source: Bovier, T. F., et al. <a href="https://www.jove.com/video/59535/" target="_blank">Methods to Test Endocrine Disruption in Drosophila melanogaster</a>. J. Vis. Exp. (2019).
Scientists use the Drosophila lifespan protocol to measure how different experimental conditions affect flies&#39; longevity and generate survival curves in the process to quantify the experimental outcomes. The sample protocol features the assay being applied to study the effects on longevity from exposure to varying concentrations of endocrine disruptor chemicals, or EDCs, such as 17-&#945;-ethinylestradiol (EE2).

Source: Bovier, T. F., et al. Methods to Test Endocrine Disruption in Drosophila melanogaster. J. Vis. Exp. (2019).
Scientists use the Drosophila lifespan ...

Experimental Protocol for Using Drosophila As an Invertebrate Model System for Toxicity Testing in the Laboratory

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