Name: rearing the fruit fly drosophila melanogaster under axenic and gnotobiotic conditions
Uploaded: 2016-07-30T14:00:00.000+00:00
Duration: 9 min 34 s
Description: Watch this Scientific Journal Video about Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions at JoVE.com

Some details of this protocol were optimized with the assistance of Dr. Adam Dobson, who also provided helpful comments on the manuscript. This work was supported by the Foundation for the National Institutes of Health (FNIH) grant number R01GM095372 (JMC, A(CN)W, AJD, and AED). FNIH grant number 1F32GM099374-01 (PDN), and Brigham Young University startup funds (JMC, MLK, MV). Publication costs were supported by the Brigham Young University College of Life Sciences and Department of Plant and Wildlife Sciences.

1. Culture Bacteria (Start ~1 Week before Picking Eggs)
<ol>
	<li>Prepare modified MRS20 (mMRS) plates and broth tubes (Table 1). Pour 20 ml mMRS agar into each 100 mm Petri plate and allow to cool/dry overnight, or 5 ml mMRS broth into 18 mm test tubes.</li>
	<li>Streak Acetobacter pomorum, A. tropicalis, Lactobacillus brevis, and L. plantarum on mMRS agar plates. Incubate Acetobacter overnight at 30 &#176;C. Incubate Lactobacillus anaerobically by placing the plates in an airtight container and flooding with carbon dioxide before sealing. Incubate at 30 &#176;C overnight. 
	Note: L. brevis colonies may not be visible until 24-48 hr. Plates can be stored at 4 &#176;C and colonies can be used for up to 3 weeks.</li>
	<li>Two to three days before transferring dechorionated eggs to sterile diet (section 5), pick a single colony from the mMRS plate into a test tube containing mMRS broth. Grow Acetobacter with shaking and grow Lactobacillus statically for 24 hr or until turbid, both at 30 &#176;C.</li>
</ol>
2. Prepare Sterile Diet
<ol>
	<li>Prepare sterile diet (Table 2) in a 2 L Erlenmeyer flask. Microwave the diet until it has boiled 3 sequential times; mix in between each boil.</li>
	<li>Place the flask on a stir plate to maintain stirring while transferring diet to conical centrifuge tubes. Transfer 7.5 ml of diet to 50 ml centrifuge tubes. Loosely cap the tubes, and place in a covered, autoclavable polypropylene rack.</li>
	<li>Sterilize fly diet using an autoclave at 121 &#176;C and 15 psi for 25 min. Remove racks from autoclave, and immediately shake each rack horizontally to ensure diet does not separate during cooling. Be careful to not shake the racks vertically to prevent transfer of diet to the lid or rims of the tubes. Allow the diet to cool on a shaker for exactly 45 min and again shake the diet horizontally by hand. 
	Note: Diet can be stored at 4-15 &#176;C for up to a week.</li>
	<li>As an alternative to using covered autoclavable racks (steps 2.1-2.3), or to raise flies on diets containing acid preservative, perform the following steps:
	<ol>
		<li>Prepare 1 L liquid diet in a 2 L flask with a stir bar in the flask. Autoclave the diet.</li>
		<li>After autoclaving, add 10 ml preservative and stir continuously on a heated stir plate. When the agar has cooled to 50-60 &#176;C, move the flask to a heated stir plate in a biosafety cabinet and maintain flask temperature by heating at 50-60 &#176;C.</li>
		<li>In the biosafety cabinet, pipet ~7.5 ml individually into conical tubes.</li>
	</ol>
	</li>
</ol>
3. Prepare Egg-laying Cages
<ol>
	<li>Make grape-juice agar plates by microwaving 100 ml water, 10 g brewer&#39;s yeast, 10 g glucose, and 1 g of agar. Bring to a boil 3 times in step 1.1 and add 10 g of frozen grape juice concentrate to increase visibility of eggs on the agar plate.</li>
	<li>When agar has cooled to 55 &#176;C, pour 20 ml into 100 mm Petri dishes and allow to solidify.</li>
	<li>Cover the surface of the agar plates with a yeast paste by mixing 1 g brewer&#39;s yeast with 15 g water. Pour yeast paste onto the agar plate making sure the surface is covered, then pour off the excess, leaving a thin yeast residue behind. If multiple plates are used, the paste can be sequentially poured to multiple plates.</li>
	<li>Transfer agar plates into the bottom of a cage. 
	Note: A 32 oz deli container is a good substitute for acrylic fly cages.
	<ol>
		<li>Make lids for 32 oz deli containers by cutting a hole in the top and gluing a breathable mesh over the hole with non-toxic glue. If the glue is water soluble prevent water exposure to the lid.</li>
	</ol>
	</li>
	<li>Transfer 200-300 flies into the container and cover with lid. Cover the mesh-protected hole with an empty Petri dish lid to prevent evaporation from the agar surface and add a moist tissue paper inside the lid if desired. Incubate flies at 25 &#176;C overnight for 16-20 hr.</li>
</ol>
4. Collect Eggs
<ol>
	<li>Prepare a sieve for egg collection by placing nylon mesh into a plastic bushing.</li>
	<li>To retrieve the plate with eggs on it, remove flies from the cage by transferring to an empty container. If same flies will be used the next day, transfer immediately to a new cage containing a freshly yeasted grape-juice agar plate. Flies can be used for 3-5 sequential days</li>
	<li>Remove dead flies from the agar with a clean paintbrush, being careful to not break up the agar.</li>
	<li>Collect eggs by rinsing the agar plate with distilled water, gently brushing eggs from the agar surface, and pouring the slurry over the mesh. Repeat 3-4 times until all or most of the eggs have been removed from the agar plate.</li>
</ol>
5. Dechorionate Eggs and Transfer to Sterile Diet
<ol>
	<li>Prepare the biosafety cabinet by spraying the inside (including sides) with 70% ethanol. Wipe the bottom with a lab tissue, and sterilize the hood with UV light for ~15 min. Sterilize all non-biological supplies (specimen cups, paintbrush, forceps, waste container, 400 ml sterilized water, and 100 ml of 0.6% sodium hypochlorite) by spraying with ethanol and immediately placing in the biosafety cabinet. Sterilize with UV light for 15 min.</li>
	<li>Start the first of 2 sodium hypochlorite washes by placing the bushing with the eggs into a 120 ml specimen cup or other sterile container. Slowly pour ~90 ml of 0.6% sodium hypochlorite solution into the bushing until just below the rim.</li>
	<li>Rinse eggs for 2.5 min. Periodically re-suspend the eggs by using forceps to move the bushing up and down in the hypochlorite solution.</li>
	<li>Transfer the bushing directly into a second specimen cup, pre-filled with 90 ml bleach, inside the biosafety cabinet.</li>
	<li>Repeat step 5.3 inside the biosafety cabinet. At the end of the second bleach treatment, the eggs should begin to adhere to the sides of the bushing.</li>
	<li>Carry out steps 5.7-5.8 in the biosafety cabinet.</li>
	<li>Discard the bleach and wash the bushing with sterile water 3 times. Re-suspend the eggs several times during each washing by moving the bushing with forceps. By the end of the third washing most eggs should be attached to the side of the bushing.</li>
	<li>Using a paintbrush sterilized in ethanol, transfer eggs from the side of the bushing to the sterile diet. Transfer eggs individually or in small batches. Aim for 30-50 eggs per vial. Leave the caps loose to allow oxygen to enter the tube. If vials are to remain axenic, transfer to an insect incubator; otherwise, add bacteria as below.</li>
</ol>
6. Make Gnotobiotic Flies Using 4 Bacterial Species
<ol>
	<li>Prepare Bacteria
	<ol>
		<li>Prepare a sterilized biosafety cabinet with necessary supplies (pipettes, pipette tip boxes, sterilized centrifuge tubes, MRS broth, and test tube racks) as in step 4.1. Wipe the outside of test tubes with an ethanol-soaked laboratory wipe before placing in biosafety cabinet.</li>
		<li>Pellet the bacteria by first transferring 500 &#181;l of overnight growth to a sterile microfuge tube. If bacterial density is low, add up to 1.5 ml to each tube or sequentially remove supernatant and add extra culture to the same tube. Remove samples from the biosafety cabinet and centrifuge for 10 min at 10,000 x g. Use filter tips to avoid contamination between samples.</li>
		<li>Determine the density of each culture by measuring OD600. If using a multi-well plate reader, transfer 200 &#181;l of each culture to a 96-well plate in 1-, 2-, and 4- fold dilutions.</li>
		<li>Determine the amount of mMRS in which to dilute each cell pellet (5.2.2) using a plate reading spectrophotometer and the following equations. Plan to add enough broth to inoculate 50 &#956;l to each fly vial.
		<ol>
			<li>Collect OD600 readings for a 1:1, 1:2, and 1:4 dilution of each bacterial culture on a plate-reading spectrophotometer. Select the dilution for each bacterial strain that produces an OD600 value between 0.1 and 0.2 and use this value and its corresponding dilution factor as &#39;O&#39; and &#39;D&#39; in the formulas given in 6.1.4.2 or 6.1.4.3.</li>
			<li>If using the 4 species described here, normalize cells to equivalent colony forming unit (CFU)/ml densities (OD600 to CFU conversion determined previously20) using this equation: 
			E = ((O-B) x V x D)/C 
			where E = volume to resuspend pellet in (&#956;l), O = OD600 bacteria, B = OD600 blank media, D = fold-dilution, V = &#181;l bacterial culture prior to centrifugation, C = OD600 of predetermined constant. See Supplemental Code File for examples of calculations using these equations. For spectrophotometers that automatically blank, use &#34;O&#34; in place of &#34;O-B&#34;. 
			Note: The predetermined constants (units OD600, normalized to 107 CFU ml-1, constants derived in20) are as follows: A. tropicalis (0.053), A. pomorum (0.038), L. brevis (0.077), L. plantarum (0.056).</li>
			<li>If using other bacterial species (no CFU/OD600 constant is available), normalize density to OD600 = 0.1 using this equation: 
			E = ((O-B) x V x D)/0.1 OD600 
			Note: Units are the same as in step 6.1.4.2. See Supplemental Code File for examples of calculations using these equations.</li>
		</ol>
		</li>
		<li>In the biosafety cabinet, remove supernatant with a pipet tip and resuspend the pellet in fresh mMRS or PBS as calculated in step 6.1.4.2.</li>
	</ol>
	</li>
	<li>Inoculate Bacteria
	<ol>
		<li>Transfer 50 &#181;l of the bacteria to the conical tubes with sterile diet and dechorionated eggs in biosafety cabinet. Add bacteria after egg transfer to prevent contamination between vials.</li>
		<li>Place inoculated tubes in an incubator at 25&#176; C.</li>
	</ol>
	</li>
</ol>
7. Measure CFU Load/Test for Sterility
<ol>
	<li>To measure the CFU load in whole body fly homogenates, transfer 5 flies (5-7 days post eclosion) to a 1.7 ml microfuge tube containing 125 &#181;l of ceramic beads and 125 &#181;l of mMRS broth. Homogenize flies using a tissue homogenizer for 30 sec at 4.0 M/sec.
	<ol>
		<li>Alternatively, omit beads and hand homogenize in microcentrifuge tubes with plastic pestles for 1 min.</li>
		<li>If quantifying the gut microbiota, surface sterilize the flies to remove exogenous microbes22. Transfer flies to a microcentrifuge tube containing 100 &#181;l 70% ethanol for 1 min, aspirate ethanol, and transfer to a new microcentrifuge tube for homogenizations. If the DNA content of the gut will be measured, rinse for 1 min with 0.6% sodium hypochlorite before the ethanol wash.</li>
	</ol>
	</li>
	<li>Dilute the homogenate with 875 &#956;l mMRS, vortex for 5 sec, and pipet 120 &#181;l of homogenate into the first well of a microtiter plate.</li>
	<li>Perform two sequential 1:8 dilutions using 10 &#181;l homogenate and 70 &#181;l MRS in the next two wells.
	<ol>
		<li>Remove 10 &#181;l from the first well and add it to the second well containing 70 &#181;l MRS. Mix the contents of the second well thoroughly, transfer 10 &#181;l from the second well to the third well containing 70 &#181;l MRS, and mix thoroughly. This leads to 3 total concentrations of the original 1,000 &#956;l homogenate: undiluted, 1:8, and 1:64.&#160;</li>
	</ol>
	</li>
	<li>Transfer 10 &#181;l of each dilution to a mMRS plate (using a multi-channel pipet if desired). Slightly incline the dish to spread the dilution several millimeters down the agar surface and allow liquid to dry before moving the plate. The liquid dries on the plate quickly if the plates are 2 days old, reducing mixing of two neighboring droplets.</li>
	<li>Incubate at 30 &#176;C for 1-2 days. Remove plates from the incubator once distinct, individual colonies are visible, and count from a dilution with 10-100 isolated colonies.</li>
	<li>Calculate CFU per fly using the equation E&#160; = C x D/P x V/F, where E =CFU per fly, C = number of colonies counted, D = dilution, P = &#181;l plated, V = volume of fly homogenate, and F = number of flies homogenized.</li>
</ol>

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E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phylogeny+of+the+defined+murine+microbiota:+altered+Schaedler+flora.">Phylogeny of the defined murine microbiota: altered Schaedler flora.</a> Appl Environ Microbiol. 65 (8), 3287-3292 (1999).</li><li>Min, K. T., Benzer, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Wolbachia,+normally+a+symbiont+of+Drosophila,+can+be+virulent,+causing+degeneration+and+early+death.">Wolbachia, normally a symbiont of Drosophila, can be virulent, causing degeneration and early death.</a> Proc Natl Acad Sci U S A. 94 (20), 10792-10796 (1997).</li></ol>

The authors have nothing to disclose.

The method described here is one of several approaches for embryo dechorionation8,11,18,25,26,27, together with alternative methods of rearing axenic flies, including serial transfer of axenic adults18,27 or antibiotic treatment13,18. Other dechorionation methods include ethanol washes and reduce11,25,26 or extend8 hypochlorite treatment. Different wash steps may aid rearing different fly genotypes: in a previous study most of ~100 Drosophila genotypes were ...

An erratum was issued for: <a href="https://www.jove.com/t/54219/rearing-fruit-fly-drosophila-melanogaster-under-axenic-gnotobiotic">Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions</a>. The Protocol was updated.
Step 6.1.4.2 of the Protocol was updated from:
If using the 4 species described here, normalize cells to equivalent colony forming unit (CFU)/ml densities (OD600 to CFU conversion determined previously20) using this equation: 
E = ((O-B) x V x D)/C 
where E = volume to resuspend pellet in (&#956;l), O = OD600 bacteria, B = OD600 blank media, D = fold-dilution, V = &#181;l bacterial culture prior to centrifugation, C = OD600 of predetermined constant. See Supplemental Code File for examples of calculations using these equations. For spectrophotometers that automatically blank, use &#34;O&#34; in place of &#34;O-B&#34;. 
Note: The predetermined constants (units OD600, normalized to 107 CFU ml-1, constants derived in20) are as follows: A. tropicalis (0.052), A. pomorum (0.038), L. brevis (0.056), L. plantarum (0.077).
to:
If using the 4 species described here, normalize cells to equivalent colony forming unit (CFU)/ml densities (OD600 to CFU conversion determined previously20) using this equation: 
E = ((O-B) x V x D)/C 
where E = volume to resuspend pellet in (&#956;l), O = OD600 bacteria, B = OD600 blank media, D = fold-dilution, V = &#181;l bacterial culture prior to centrifugation, C = OD600 of predetermined constant. See Supplemental Code File for examples of calculations using these equations. For spectrophotometers that automatically blank, use &#34;O&#34; in place of &#34;O-B&#34;. 
Note: The predetermined constants (units OD600, normalized to 107 CFU ml-1, constants derived in20) are as follows: A. tropicalis (0.053), A. pomorum (0.038), L. brevis (0.077), L. plantarum (0.056).

An erratum was issued for: <a href="https://www.jove.com/t/54219/rearing-fruit-fly-drosophila-melanogaster-under-axenic-gnotobiotic">Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions</a>. The Protocol was updated.

Erratum: Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

Most animals are intimately associated with bacteria (&#39;microbiota&#39;) from birth to death1. Comparisons of microorganism-free (&#39;axenic&#39;) and microorganism-associated (&#39;conventional&#39;) animals have shown microbes influence diverse aspects of animal health, including metabolic, nutritional, vascular, hepatic, respiratory, immunological, endocrine, and neurological function2. The fruit fly Drosophila melanogaster is a key model for understanding many of these processes in the presence of microbes3,4 and for studying microbiota influence on animal health5,6. No bacterial species is present in every individual (&#39;core&#39;), but Acetobacter and Lactobacillus species numerically dominate the microbiota of both laboratory-reared and wild-caught D. melanogaster. Other Acetobacteraceae (including Komagataeibacter and Gluconobacter), Firmicutes (such as Enterococcus and Leuconostoc), and Enterobacteriaceae are either frequently present in Drosophila individuals at low abundance, or irregularly present at high abundance7-12.
The microbiota of Drosophila and mammals is inconstant within and across generations14,19. Microbiota inconstancy can lead to phenotypic noise when measuring microbiota-dependent traits. For example, the Acetobacteraceae influence lipid (triglyceride) storage in Drosophila15-18. If Acetobacteraceae are more abundant in flies of one vial than in another19, isogenic flies can have different phenotypes20. A solution for the problem of microbiota inconstancy in mice14 has been in practice since the 1960&#39;s, by introducing a defined community of 8 dominant microbial species to mouse pups each new generation (altered Schaedler flora), ensuring that each pup is exposed to the same key members of the mouse microbiota. This practice controls for microbiota composition even when the microbiota is not the primary target of study32, and sets precedent to ensure the presence of key microbes in a variety of experimental conditions.
To define the influence of microbes on Drosophila nutrition, several protocols for deriving axenic fly lines have been developed, including hypochlorite dechorionation of embryos (either derived de novo each generation or maintained generationally by transfer to sterile diets) and antibiotic treatment13. There are benefits to different approaches, such as ease and rapidity for both of antibiotics treatment and serial transfer, versus greater control of confounding variables with de novo dechorionation (e.g., egg density, residual contaminating microbes, off-target antibiotic effects). Regardless of the method of preparation, introduction of specified microbial species to axenic embryos permits culture of Drosophila with defined (&#39;gnotobiotic&#39;) communities. Alternatively, mimicking the use of Schaedler flora, this community could be inoculated to conventionally-laid eggs (following steps 6-7 only) to ensure the presence of trait-influencing microbes in each vial and avoid complications of microbiota inconstancy. Here we describe the protocol for raising axenic and gnotobiotic Drosophila by de novo dechorionation of embryos, and for confirming the presence of introduced or contaminating microbial taxa.

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rearing the fruit fly drosophila melanogaster under axenic and gnotobiotic conditions

The influence of microbes on myriad animal traits and behaviors has been increasingly recognized in recent years. The fruit fly Drosophila melanogaster is a model for understanding microbial interactions with animal hosts, facilitated by approaches to rear large sample sizes of Drosophila under microorganism-free (axenic) conditions, or with defined microbial communities (gnotobiotic). This work outlines a method for collection of Drosophila embryos, hypochlorite dechorionation and sterilization, and transfer to sterile diet. Sterilized embryos are transferred to sterile diet in 50 ml centrifuge tubes, and developing larvae and adults remain free of any exogenous microbes until the vials are opened. Alternatively, flies with a defined microbiota can be reared by inoculating sterile diet and embryos with microbial species of interest. We describe the introduction of 4 bacterial species to establish a representative gnotobiotic microbiota in Drosophila. Finally, we describe approaches for confirming bacterial community composition, including testing if axenic Drosophila remain bacteria-free into adulthood.

Successful rearing of axenic flies is confirmed by isolation of no CFUs from whole-body homogenizations of D. melanogaster adults (Figure 1). Alternatively, if the plated homogenate yields colonies, the vials are contaminated and should be discarded. For gnotobiotic flies, each of the four bacterial isolates were isolated from pools of 5 adult males, demonstrating differences in total viable CFUs associated with adult flies (Figure 1). Each bacte...

Watch this Scientific Journal Video about Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions at JoVE.com

Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

A method for rearing Drosophila melanogaster under axenic and gnotobiotic conditions is presented. Fly embryos are dechorionated in sodium hypochlorite, transferred aseptically to sterile diet, and reared in closed containers. Inoculating diet and embryos with bacteria leads to gnotobiotic associations, and bacterial presence is confirmed by plating whole-body Drosophila homogenates.

Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

The influence of microbes on myriad animal traits and behaviors has been increasingly recognized in recent years. The fruit fly Drosophila melanogaster is ...

rearing-fruit-fly-drosophila-melanogaster-under-axenic-gnotobiotic

Research

JoVE Journal

Developmental Biology

21.0K Views.  Brigham Young University. A method for rearing Drosophila melanogaster under axenic and gnotobiotic conditions is presented. Fly embryos are dechorionated in sodium hypochlorite, transferred aseptically to sterile diet, and reared in closed containers. Inoculating diet and embryos with bacteria leads to gnotobiotic associations, and bacterial presence is confirmed by plating whole-body Drosophila homogenates.

Video: Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

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

Drosophila melanogaster Larva Injection Protocol

The use of unconventional models to study innate immunity and pathogen virulence provides a valuable alternative to mammalian models, which can be costly and raise ethical issues. Unconventional models are notoriously cheap, easy to handle and culture, and do not take much space. They are genetically amenable and possess complete genome sequences, and their use presents no ethical considerations. The fruit fly Drosophila melanogaster, for instance, has provided great insights into a variety of behavior, development, metabolism, and immunity research. More specifically, D. melanogaster adult flies and larvae possess several innate defense reactions that are shared with vertebrate animals. The mechanisms regulating immune responses have been mostly revealed through genetic and molecular studies in the D. melanogaster model. Here a novel larval injection technique is provided, which will further promote investigations of innate immune processes in D. melanogaster larvae and explore the pathogenesis of a wide range of microbial infections.

Drosophila melanogaster Larva Injection Protocol

Drosophila melanogaster adult flies have been extensively utilized as model organisms to investigate the molecular mechanisms underlying host antimicrobial innate immune responses and microbial infection strategies. To promote the D. melanogaster larva stage as an additional or alternative model system, a larval injection technique is described.

The use of unconventional models to study innate immunity and pathogen virulence provides a valuable alternative to mammalian models, which can be costly ...

Immunology and Infection

Quantification of Macronutrients Intake in a Thermogenetic Neuronal Screen using Drosophila Larvae

Foraging and feeding behaviors allow animals to access sources of energy and nutrients essential for their development, health, and fitness. Investigating the neuronal regulation of these behaviors is essential for the understanding of the physiological and molecular mechanisms underlying nutritional homeostasis. The use of genetically tractable animal models such as worms, flies, and fish greatly facilitates these types of studies. In the last decade, the fruit fly Drosophila melanogaster has been used as a powerful animal model by neurobiologists investigating the neuronal control of feeding and foraging behaviors. While undoubtedly valuable, most studies examine adult flies. Here, we describe a protocol that takes advantage of the simpler larval nervous system to investigate neuronal substrates controlling feeding behaviors when larvae are exposed to diets differing in their protein and carbohydrates content. Our methods are based on a quantitative colorimetric no-choice feeding assay, performed in the context of a neuronal thermogenetic-activation screen. As a read-out, the amount of food eaten by larvae over a 1 h interval was used when exposed to one of the three dye-labeled diets that differ in their protein to carbohydrates (P:C) ratios. The efficacy of this protocol is demonstrated in the context of a neurogenetic screen in larval Drosophila, by identifying candidate neuronal populations regulating the amount of food eaten in diets of different macronutrient quality. We were also able to classify and group the genotypes tested into phenotypic classes. Besides a brief review of the currently available methods in the literature, the advantages and limitations of these methods are discussed and, also, some suggestions are provided about how this protocol might be adapted to other specific experiments.

Quantification of Macronutrients Intake in a Thermogenetic Neuronal Screen using Drosophila Larvae

Described here is a protocol that enables the colorimetric quantification of the amount of food eaten within a defined interval of time by Drosophila melanogaster larvae exposed to diets of different macronutrient quality. These assays are conducted in the context of a neuronal thermogenetic screen.

Foraging and feeding behaviors allow animals to access sources of energy and nutrients essential for their development, health, and fitness. Investigating ...

Neuroscience

Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

Neural stem cells (NSCs) have the ability to proliferate, differentiate, undergo apoptosis, and even enter and exit quiescence. Many of these processes are controlled by the complex interplay between NSC intrinsic genetic programs with NSC extrinsic factors, local and systemic. In the genetic model organism, Drosophila melanogaster, NSCs, known as neuroblasts (NBs), switch from quiescence to proliferation during the embryonic to larval transition. During this time, larvae emerge from their eggshells and begin crawling, seeking out dietary nutrients. In response to animal feeding, the fat body, an endocrine organ with lipid storage capacity, produces a signal, which is released systemically into the circulating hemolymph. In response to the fat body-derived signal (FBDS), Drosophila insulin-like peptides (Dilps) are produced and released from brain neurosecretory neurons and glia, leading to downstream activation of PI3-kinase growth signaling in NBs and their glial and tracheal niche. Although this is the current model for how NBs switch from quiescence to proliferation, the nature of the FBDS extrinsic cue remains elusive. To better understand how NB extrinsic systemic cues regulate exit from quiescence, a method was developed to culture early larval brains in vitro before animal feeding. With this method, exogenous factors can be supplied to the culture media and NB exit from quiescence assayed. We found that exogenous insulin is sufficient to reactivate NBs from quiescence in whole-brain explants. Because this method is well-suited for large-scale screens, we aim to identify additional extrinsic cues that regulate NB quiescence versus proliferation decisions. Because the genes and pathways that regulate NSC proliferation decisions are evolutionarily conserved, results from this assay could provide insight into improving regenerative therapies in the clinic.

Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

A method to reactivate quiescent neural stem cells in cultured Drosophila brain explants has been established. Using this method, the role of systemic signals can be uncoupled from tissue-intrinsic signals in the regulation of neural stem cell quiescence, entry and exit.

Neural stem cells (NSCs) have the ability to proliferate, differentiate, undergo apoptosis, and even enter and exit quiescence. Many of these processes ...

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.

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.

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

Biology

Oral Bacterial Infection and Shedding in Drosophila melanogaster

The fruit fly Drosophila melanogaster is one of the best developed model systems of infection and innate immunity. While most work has focused on systemic infections, there has been a recent increase of interest in the mechanisms of gut immunocompetence to pathogens, which require methods to orally infect flies. Here we present a protocol to orally expose individual flies to an opportunistic bacterial pathogen (Pseudomonas aeruginosa) and a natural bacterial pathogen of D. melanogaster (Pseudomonas entomophila). The goal of this protocol is to provide a robust method to expose male and female flies to these pathogens. We provide representative results showing survival phenotypes, microbe loads, and bacterial shedding, which is relevant for the study of heterogeneity in pathogen transmission. Finally, we confirm that Dcy mutants (lacking the protective peritrophic matrix in the gut epithelium) and Relish mutants (lacking a functional immune deficiency (IMD) pathway), show increased susceptibility to bacterial oral infection. This protocol, therefore, describes a robust method to infect flies using the oral route of infection, which can be extended to the study of a variety genetic and environmental sources of variation in gut infection outcomes and bacterial transmission.

Oral Bacterial Infection and Shedding in Drosophila melanogaster

This protocol describes methods to orally expose and infect the fruit fly Drosophila melanogaster with bacterial pathogens, and to measure the number of infectious bacteria shed following gut infection. We further describe the effect of immune mutants on fly survival following oral bacterial infection.

The fruit fly Drosophila melanogaster is one of the best developed model systems of infection and innate immunity. While most work has focused on systemic ...

Establishment and Maintenance of Gnotobiotic American Cockroaches (Periplaneta americana)

Gnotobiotic animals are a powerful tool for the study of controls on microbiome structure and function. Presented here is a protocol for the establishment and maintenance of gnotobiotic American cockroaches (Periplaneta americana). This approach includes built-in sterility checks for ongoing quality control. Gnotobiotic insects are defined here as cockroaches that still contain their vertically transmitted endosymbiont (Blattabacterium) but lack other microbes that normally reside on their surface and in their digestive tract. For this protocol, egg cases (oothecae) are removed from a (nonsterile) stock colony and surface sterilized. Once collected and sterilized, the oothecae are incubated at 30 &#176;C for approximately 4&#8722;6 weeks on brain-heart infusion (BHI) agar until they hatch or are removed due to contamination. Hatched nymphs are transferred to an Erlenmeyer flask containing a BHI floor, sterile water, and sterile rat food. To ensure that the nymphs are not housing microbes that are unable to grow on BHI in the given conditions, an additional quality control measure uses restriction fragment-length polymorphism (RFLP) to test for nonendosymbiotic microbes. Gnotobiotic nymphs generated using this approach can be inoculated with simple or complex microbial communities and used as a tool in gut microbiome studies.

Establishment and Maintenance of Gnotobiotic American Cockroaches Periplaneta americana

This protocol is used in establishing and maintaining gnotobiotic American cockroaches (Periplaneta americana) by surface sterilizing the egg cases (oothecae) prior to hatching. These gnotobiotic insects contain their vertically transmitted Blattabacterium endosymbionts but have axenic guts.

Gnotobiotic animals are a powerful tool for the study of controls on microbiome structure and function. Presented here is a protocol for the establishment ...

Drosophila Maintenance

Drosophila melanogaster, commonly known as fruit flies, are a frequently used model organism for life science research. Although starting a collection of these critters may seem as easy as leaving a banana on your kitchen counter for too long, a productive fly colony in the lab requires careful husbandry and maintenance.
This video demonstrates the necessary steps for maintaining a healthy fly stock. The overview begins with the preparation and storage of the yeast and sugar-containing media on which flies feed. Next, the vessels most commonly used for housing Drosophila are shown, as well as how and when to move flies between these containers. Finally, the presentation also includes examples of the ways in which housing and feeding conditions are manipulated for biological experiments.

Maintenance and Husbandry of Drosophila melanogaster

Drosophila melanogaster, commonly known as fruit flies, are a frequently used model organism for life science research. Although starting a collection of ...

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Biology I: yeast, Drosophila and C. elegans

Drosophila melanogaster Embryo and Larva Harvesting and Preparation

Drosophila melanogaster embryos and larvae are easy to manipulate and develop rapidly by mechanisms that are analogous to other organisms, including mammals. For these reasons, many researchers utilize fly embryos and larvae to answer questions in diverse fields ranging from behavioral to developmental biology. Prior to experimentation, however, the embryos and larvae must first be collected.
This video will first demonstrate how "egg-laying cups" are used to collect Drosophila embryos on agar plates. The harvest and dechorionation of embryos will then be described. Next, the video will demonstrate how to identify and manipulate Drosophila in one of the three larval stages that follow the embryo stage. Finally, examples of some of the ways in which fly embryos and larvae are used in biological research are provided.

Drosophila melanogaster Embryo and Larva Harvesting and Preparation

Drosophila melanogaster embryos and larvae are easy to manipulate and develop rapidly by mechanisms that are analogous to other organisms, including ...

Drosophila Egg Collection and Dechorionation: A Method to Remove the Outermost Egg Layer

Source: Koyle, M. L., et al. <a href="https://www.jove.com/video/54219/" target="_blank">Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions</a>. J. Vis. Exp. (2016).
The Drosophila embryo is surrounded by protective outer membranes. To gain access to the embryo, the chorion&#8211;the outermost of the membranes&#8211;is often removed. This video describes a commonly used method of dechorionation using sodium hypochlorite. The example protocol demonstrates the procedure while employing sterile techniques necessary for axenic or gnotobiotic animal rearing.

Source: Koyle, M. L., et al. Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions. J. Vis. Exp. (2016).
The Drosophila ...

Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

Summary

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Establishment and Maintenance of Gnotobiotic American Cockroaches (Periplaneta americana)

Drosophila Maintenance

Drosophila melanogaster Embryo and Larva Harvesting and Preparation

Drosophila Egg Collection and Dechorionation: A Method to Remove the Outermost Egg Layer