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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#160;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&#160;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&#160;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 Drosophila</td>
			<td>Bloomington Drosophila Stock Center at Indiana University</td>
			<td></td>
			<td>https://bdsc.indiana.edu</td>
		</tr>
		<tr>
			<td>Wild popultions of Drosophila</td>
			<td>UC San Diego Drosophila 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 ...

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12.6K Views.  Colorado State University-Pueblo. 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. 

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

Testing Nicotine Tolerance in Aphids Using an Artificial Diet Experiment

Plants may upregulate the production of many different seconday metabolites in response to insect feeding. One of these metabolites, nicotine, is well know to have insecticidal properties. One response of tobacco plants to herbivory, or being gnawed upon by insects, is to increase the production of this neurotoxic alkaloid. Here, we will demonstrate how to set up an experiment to address this question of whether a tobacco-adapted strain of the green peach aphid, Myzus persicae, can tolerate higher levels of nicotine than the a strain of this insect that does not infest tobacco in the field.

In this video and assay is demonstrated that tests the tolerance of nicotine to two types of aphids one that infests tobacco plants in the field and one that does not.

Plants may upregulate the production of many different seconday metabolites in response to insect feeding. One of these metabolites, nicotine, is well ...

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

Our research activities target the use of biological methods for the evaluation of environmental quality, with particular reference to saltwater/brackish water and sediment. The choice of biological indicators must be based on reliable scientific knowledge and, possibly, on the availability of standardized procedures. In this article, we present a standardized protocol that used the marine crustacean Artemia to evaluate the toxicity of chemicals and/or of marine environmental matrices. Scientists propose that the brine shrimp (Artemia) is a suitable candidate for the development of a standard bioassay for worldwide utilization. A number of papers have been published on the toxic effects of various chemicals and toxicants on brine shrimp (Artemia). The major advantage of this crustacean for toxicity studies is the overall availability of the dry cysts; these can be immediately used in testing and difficult cultivation is not demanded1,2. Cyst-based toxicity assays are cheap, continuously available, simple and reliable and are thus an important answer to routine needs of toxicity screening, for industrial monitoring requirements or for regulatory purposes3. The proposed method involves the mortality as an endpoint. The numbers of survivors were counted and percentage of deaths were calculated. Larvae were considered dead if they did not exhibit any internal or external movement during several seconds of observation4. This procedure was standardized testing a reference substance (Sodium Dodecyl Sulfate); some results are reported in this work. This article accompanies a video that describes the performance of procedural toxicity testing, showing all the steps related to the protocol.

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

This study concerns the development and standardization of a valuable methodological protocol to determine long-term (14 days) lethal toxicity exerted by chemical substances, industrial wastewater or sewage and liquid environmental samples on the saltwater crustacean, Artemia franciscana.

Our research activities target the use of biological methods for the evaluation of environmental quality, with particular reference to saltwater/brackish ...

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the function and regulation of protein folding was studied in vitro, in isolated tissue culture cells and in unicellular organisms. Recent studies have uncovered links between protein homeostasis (proteostasis), metabolism, development, aging, and temperature-sensing. These findings have led to the development of new tools for monitoring protein folding in the model metazoan organism Caenorhabditis elegans. In our laboratory, we combine behavioral assays, imaging and biochemical approaches using temperature-sensitive or naturally occurring metastable proteins as sensors of the folding environment to monitor protein misfolding. Behavioral assays that are associated with the misfolding of a specific protein provide a simple and powerful readout for protein folding, allowing for the fast screening of genes and conditions that modulate folding. Likewise, such misfolding can be associated with protein mislocalization in the cell. Monitoring protein localization can, therefore, highlight changes in cellular folding capacity occurring in different tissues, at various stages of development and in the face of changing conditions. Finally, using biochemical tools ex vivo, we can directly monitor protein stability and conformation. Thus, by combining behavioral assays, imaging and biochemical techniques, we are able to monitor protein misfolding at the resolution of the organism, the cell, and the protein, respectively.

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

To study the relationship between protein homeostasis, stress and aging, we monitored changes in protein folding by following protein dysfunction, protein localization in the cell and protein stability at the organismal, cellular and protein levels, using the genetically tractable metazoan Caenorhabditis elegans as a model system.

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the ...

A Protocol for Using Gene Set Enrichment Analysis to Identify the Appropriate Animal Model for Translational Research

Recent studies that compared transcriptomic datasets of human diseases with datasets from mouse models using traditional gene-to-gene comparison techniques resulted in contradictory conclusions regarding the relevance of animal models for translational research. A major reason for the discrepancies between different gene expression analyses is the arbitrary filtering of differentially expressed genes. Furthermore, the comparison of single genes between different species and platforms often is limited by technical variance, leading to misinterpretation of the con/discordance between data from human and animal models. Thus, standardized approaches for systematic data analysis are needed. To overcome subjective gene filtering and ineffective gene-to-gene comparisons, we recently demonstrated that gene set enrichment analysis (GSEA) has the potential to avoid these problems. Therefore, we developed a standardized protocol for the use of GSEA to distinguish between appropriate and inappropriate animal models for translational research. This protocol is not suitable to predict how to design new model systems a-priori, as it requires existing experimental omics data. However, the protocol describes how to interpret existing data in a standardized manner in order to select the most suitable animal model, thus avoiding unnecessary animal experiments and misleading translational studies.

We provide a standardized protocol for the use of gene set enrichment analysis of transcriptomic data to identify an ideal mouse model for translational research. 
This protocol can be used with DNA microarray and RNA sequencing data and can further be extended to other omics data if data are available.

Recent studies that compared transcriptomic datasets of human diseases with datasets from mouse models using traditional gene-to-gene comparison ...

Laboratory Protocol for Genetic Gut Content Analyses of Aquatic Macroinvertebrates Using Group-specific rDNA Primers

Analyzing food webs is essential for a better understanding of ecosystems. For example, food web interactions can undergo severe changes caused by the invasion of non-indigenous species. However, an exact identification of field predator-prey interactions is difficult in many cases. These analyses are often based on a visual evaluation of gut content or the analysis of stable isotope ratios (&#948;15N and &#948;13C). Such methods require comprehensive knowledge about, respectively, morphologic diversity or isotopic signature from individual prey organisms, leading to obstacles in the exact identification of prey organisms. Visual gut content analyses especially underestimate soft bodied prey organisms, because maceration, ingestion and digestion of prey organisms make identification of specific species difficult. Hence, polymerase chain reaction (PCR) based strategies, for example the use of group-specific primer sets, provide a powerful tool for the investigation of food web interactions. Here, we describe detailed protocols to investigate the gut contents of macroinvertebrate consumers from the field using group-specific primer sets for nuclear ribosomal deoxyribonucleic acid (rDNA). DNA can be extracted either from whole specimens (in the case of small taxa) or out of gut contents of specimens collected in the field. Presence and functional efficiency of the DNA templates need to be confirmed directly from the tested individual using universal primer sets targeting the respective subunit of DNA. We also demonstrate that consumed prey can be determined further down to species level via PCR with unmodified group-specific primers combined with subsequent single strand conformation polymorphism (SSCP) analyses using polyacrylamide gels. Furthermore, we show that the use of different fluorescent dyes as labels enables parallel screening for DNA fragments of different prey groups from multiple gut content samples via automated fragment analysis.

Most common gut content analyses of macroinvertebrates are visual. Requiring intense knowledge about morphological diversity of prey organisms, they miss soft bodied prey and, due to strong comminution of prey, are nearly impossible for some organisms, including amphipods. We provide detailed, novel genetic approaches for macroinvertebrate prey identification in the diet of amphipods.

Analyzing food webs is essential for a better understanding of ecosystems. For example, food web interactions can undergo severe changes caused by the ...

Establishment of an Extracellular Acidic pH Culture System

Conditions of the tumor microenvironment, such as hypoxia or nutrient starvation, play critical roles in cancer progression and malignancy. However, the role of acidic extracellular pH in tumor aggressiveness and its underlying mechanism has not been extensively studied compared to hypoxic or nutrient starvation conditions. In addition, a well-defined culture method to mimic the acidic extracellular tumor microenvironment has not been fully reported.
Here we present a simple in vitro culture method to maintain acidic extracellular pH using reduced bicarbonate and increased lactate or HCl concentrations in the culture medium. The medium pH was sustained for at least 24 h and gradually decreased by 72 h following culture of PANC-1 and AsPC-1 pancreatic cancer cells. Three distinct acidic media conditions in this study highly upregulated pH-responsive genes such as MSMO1, INSIG1, and IDI1 compared to hypoxia or nutrient starvation. The upregulation of these genes can be used as a marker of acidic pH. These simple techniques are beneficial to elucidate underlying mechanisms of tumor malignancy under acidic tumor microenvironment. Therefore, our extracellular acidic pH culture system enables discovery of cellular acidic pH responses not only in cancer cells but also in primary cells, such as renal tubular cells, in relation to the other acidic disorders including, diabetic ketoacidosis, lactic acidosis, renal tubular acidosis, and respiratory acidosis.

The acidic tumor microenvironment plays critical roles in tumor progression. To assess the effects of acidic extracellular pH on cancer cells in vitro, we established simple acidic culture systems.

Conditions of the tumor microenvironment, such as hypoxia or nutrient starvation, play critical roles in cancer progression and malignancy. However, the ...

A Protocol for Laboratory Housing of Turquoise Killifish (Nothobranchius furzeri)

The development of husbandry practices in non-model laboratory fish used for experimental purposes has greatly benefited from the establishment of reference fish model systems, such as zebrafish and medaka. In recent years, an emerging fish &#8211; the turquoise killifish (Nothobranchius furzeri) &#8211; has been adopted by a growing number of research groups in the fields of biology of aging and ecology. With a captive life span of 4 - 8 months, this species is the shortest-lived vertebrate raised in captivity and allows the scientific community to test &#8211; in a short time &#8211; experimental interventions that can lead to alterations of the aging rate and life expectancy. Given the unique biology of this species, characterized by embryonic diapause, explosive sexual maturation, marked morphological and behavioral sexual dimorphism - and their relatively short adult life span -&#160;ad hoc husbandry practices are in urgent demand. This protocol reports a set of key husbandry measures that allow optimal turquoise killifish laboratory care, enabling the scientific community to adopt this species as a powerful laboratory animal model.

A Protocol for Laboratory Housing of Turquoise Killifish Nothobranchius furzeri

Laboratory housing of turquoise killifish can be scaled up&#160;to house and efficiently raise thousands of individual fish in a centralized water filtration system, employing the same infrastructure used for standard zebrafish facilities. Here we detail a list of standardized procedures that allow efficient killifish maintenance.

The development of husbandry practices in non-model laboratory fish used for experimental purposes has greatly benefited from the establishment of ...

Protocols for Testing the Toxicity of Novel Insecticidal Chemistries to Mosquitoes

New classes of insecticides with novel modes of action are needed to control insecticide resistant populations of mosquitoes that transmit diseases such as Zika, dengue and malaria. Assays for rapid, high-throughput analyses of unformulated novel chemistries against mosquito larvae and adults are presented. We describe protocols for single point-dose and dose response assays to evaluate the toxicity of small molecule chemistries to the Aedes aegypti vector of Zika, dengue and yellow fever,&#160;the malaria vector, Anopheles gambiae and the northern house mosquito, Culex quinquefasciatus, on contact and via ingestion. As an example, we evaluated the toxicity of amitriptyline, a small molecule antagonist of G protein-coupled receptors, via larval, adult topical and adult blood-feeding assay. The protocols provide a starting point to investigate insecticide potential. Results are discussed in the context of additional experiments to explore product applications and mechanisms for delivery.

Protocols are described for assessing the toxicity of chemistries to immature and adult mosquitoes for development as new classes of larvicides, adulticides and endectocides. The protocols enable high-throughput testing of multiple chemistries at single-point dose and subsequent evaluation via dose response assay to determine toxicity on contact or via ingestion.

New classes of insecticides with novel modes of action are needed to control insecticide resistant populations of mosquitoes that transmit diseases such ...

An Air-liquid Interface Bronchial Epithelial Model for Realistic, Repeated Inhalation Exposure to Airborne Particles for Toxicity Testing

For toxicity testing of airborne particles, air-liquid interface (ALI) exposure systems have been developed for in vitro tests in order to mimic realistic exposure conditions. This puts specific demands on the cell culture models. Many cell types are negatively affected by exposure to air (e.g., drying out) and only remain viable for a few days. This limits the exposure conditions that can be used in these models: usually relatively high concentrations are applied as a cloud (i.e., droplets containing particles, which settle down rapidly) within a short period of time. Such experimental conditions do not reflect realistic long-term exposure to low concentrations of particles. To overcome these limitations the use of a human bronchial epithelial cell line, Calu-3 was investigated. These cells can be cultured at ALI conditions for several weeks while retaining a healthy morphology and a stable monolayer with tight junctions. In addition, this bronchial model is suitable for testing the effects of repeated exposures to low, realistic concentrations of airborne particles using an ALI exposure system. This system uses a continuous airflow in contrast to other ALI exposure systems that use a single nebulization producing a cloud. Therefore, the continuous flow system is suitable for repeated and prolonged exposure to airborne particles while continuously monitoring the particle characteristics, exposure concentration, and delivered dose. Taken together, this bronchial model, in combination with the continuous flow exposure system, is able to mimic realistic, repeated inhalation exposure conditions that can be used for toxicity testing.

Described is the cell culture and exposure method of an in vitro bronchial model for realistic, repeated inhalation exposure to particles for toxicity testing.

For toxicity testing of airborne particles, air-liquid interface (ALI) exposure systems have been developed for in vitro tests in order to mimic realistic ...