We thank Mark G. Perkins, Pearce Laboratory, Department of Anesthesiology, University of Wisconsin-Madison, for the construction of the SAA prototype. The work is supported by the National Institute of General Medical Sciences (NIGMS) with R01GM134107 and by the R&#38;D fund of the Department of Anesthesiology, University of Wisconsin-Madison.

NOTE: See the Table of Materials for details about all the materials used in the protocol.
1. Construction of the SAA
<ol>
	<li>Make the frame by cutting wood and assembling the frame using the dimensions in Figure 1A.</li>
	<li>Modify 50 mL conical tube caps.
	<ol>
		<li>Drill two holes in each cap with a 9/32 in drill bit. Sand the holes to clean up the ragged plastic. Sand the top of the cap to roughen the surface (this helps with glue adhesion).</li>
		<li>Cut 5 mL serological pipettes to size (3 in for inflow and 1.5 in for outflow) by scoring the plastic and then breaking it clean at the scored line. Sand the ends of the cut/broken pipettes.</li>
		<li>Glue netting to the tubes (allow proper drying time for the adhesive). Cut the netting to the size of the tube after the adhesive dries.</li>
		<li>Insert the tubes into the holes of the conical caps with both tubes extending (3/4 in) above the cap; ensure that the inflow tube extends longer into the tube than the outflow (Figure 1B).</li>
		<li>Apply glue to the tops of the caps around the tubes to secure the parts together (allow proper drying time for the adhesive before continuing).</li>
	</ol>
	</li>
	<li>Attach the caps to the frame, and route the tubing (Figure 1C).
	<ol>
		<li>Attach adhesive cable tie-downs to the frame (3.25 in apart, center to center).</li>
		<li>Affix the caps to the frame using zip ties; cut the zip tie tag ends short.</li>
		<li>Cut and connect lengths (9 in) of Tygon tubing to the inflow/outflow tubes on each modified cap (Figure 1D). Starting at the upstream end, attach first to the inflow, and then subsequently attach tubing from the outflow to the inflow of the next position.</li>
		<li>Add a flow indicator to the most downstream &#34;inflow&#34; (position 10, Figure 1E).</li>
		<li>Put a 50 mL conical tube on the first position, and fill it with water to just below the inflow tube (Figure 1F).</li>
	</ol>
	</li>
	<li>Prepare the interfacing for the vaporizer. Remove the plungers, cut notches out of two 10 mL dispensing syringes (1/2 in deep x 1/4 in wide, Figure 1G), and insert them into the vaporizer inflow and outflow with the notches facing directly toward the front of the vaporizer to align with the holes (Figure 1H). Optional: Glue the modified syringes into place. If affordable, use a commercial manifold (see the Table of Materials for one option).</li>
	<li>Hook up the entire system. Use Tygon tubing to attach the components together in the following order: carrier gas tank with regulator &#62; gas specific flowmeter &#62; vaporizer &#62; SAA (Figure 1C).</li>
	<li>Fill empty positions on the array with empty 50 mL conical tubes. Turn on the gas tank, open the flowmeter to ~2 L/min, and turn on the vaporizer to 0%. Confirm gas flow through the system by checking the flowmeter upstream of the vaporizer and the flow indicator downstream of the last chamber of the SAA for flow. Alternatively, insert the downstream tubing end into water, and look for bubbles. 
	&#8203;NOTE: As the system is not pressurized, a water column higher than a couple of centimeters will stop the flow. If there is no flow at the downstream end of the array, check the following: the vaporizer needs to be on to allow flow; check that the tank regulator and flow meters are allowing flow; check the array positions to make sure the tubes are screwed on tightly; and check for leaks around the adhesive on the modified caps.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65144/65144fig01.jpg" /> 
Figure 1: Construction of the SAA. (A) Schematic, with measurements, of the wooden frame that supports the SAA. (B) Schematized cross-section, with measurements, of a modified cap with inflow and outflow tubes made of 5 mL serological pipettes. (C) Assembled SAA (reproduced from Olufs et al.5) (D) Details of a modified 50 mL conical cap showing inflow and outflow tubes. (E) Downstream (position 10) outflow with the flow indicator. (F) Upstream (position 1) water-filled tube to humidify the carrier gas. The red arrow indicates the water level. (G) Modified 10 mL dispensing syringe for the makeshift manifold. The red circle highlights the cut-out notch located between the 8 mL and 10 mL marks (or 1/2 in x 1/4 in). (H) Rear view of the Tec7 vaporizer showing the insertion and orientation of the modified syringes. Only one syringe is in place in this view to show, on the left, the hole (red arrow) that needs to be aligned with the notch of the modified syringe. Note: Misalignment of this cut-out notch and the outflow opening will disrupt anesthetic administration. This part is a potential weak spot in this custom-made system. If funds are available, a commercial manifold should be used. Abbreviation: SAA = serial anesthesia array. <a href="https://www.jove.com/files/ftp_upload/65144/65144fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Prior to anesthetic exposure
<ol>
	<li>Twenty-four hours or more prior to the anesthetic exposure, sort the fly cohorts as needed for the experiment using the preferred method (e.g., CO2 or ether).</li>
</ol>
3. Operation of the SAA
<ol>
	<li>Transfer flies from food vials into empty 50 mL conical tubes (without CO2).
	<ol>
		<li>Count and record any dead flies prior to exposure.</li>
	</ol>
	</li>
	<li>Uncap and screw 50 mL conical tubes with flies onto the SAA.</li>
	<li>Turn on the carrier gas, and set to the desired flow rate. 
	NOTE: We typically use 1-2 L/min.</li>
	<li>Set the anesthetic vaporizer to the desired concentration. 
	NOTE: We typically use 2% for isoflurane and 3.5% for sevoflurane, which are equipotent doses in mammals10.</li>
	<li>Expose the flies for the desired duration (min: 15 min). 
	NOTE: A minimum exposure time of 15 min is recommended to avoid possible variability in equilibration across the positions of the SAA. In this system, it takes 2-3 min for the anesthetics to equilibrate across all the positions.</li>
	<li>At the end of the exposure, flush the system with fresh gas flow (vaporizer set to 0%) at 1.5 L/min for 5 min corresponding to approximately 10x volumes of the total SAA volume.</li>
</ol>
4. Checklist prior to starting an experiment
<ol>
	<li>Open the high-pressure regulator (on top of the air tank) completely, and then close it half a turn to ensure carrier gas flow.</li>
	<li>Follow tubing for each line to the i) flowmeters and ii) vaporizer (make sure the inflow/outflow are connected correctly), and iii) check the anesthetic level in the vaporizers.</li>
	<li>After loading the chambers with subjects, check that the air/gas is flowing with the bubble test or the flow indicator. 
	NOTE: Some vaporizers do not allow air flow when the dial is in the off position.</li>
	<li>When gas is flowing, confirm that both the flowmeter and the downstream flow indicator indicate flow.</li>
	<li>At the end of the experiment, allow for 4-5 min of airflow to wash out the anesthetic.</li>
</ol>

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J., 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=A+method+to+inflict+closed+head+traumatic+brain+injury+in+Drosophila.">A method to inflict closed head traumatic brain injury in Drosophila.</a> Journal of Visualized Experiments. (100), e52905 (2015).</li><li>Loewen, C. A., Ganetzky, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mito-nuclear+interactions+affecting+lifespan+and+neurodegeneration+in+a+Drosophila+model+of+Leigh+syndrome.">Mito-nuclear interactions affecting lifespan and neurodegeneration in a Drosophila model of Leigh syndrome.</a> Genetics. 208 (4), 1535-1552 (2018).</li><li>Johnson-Schlitz, D., 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=Anesthetic+preconditioning+of+traumatic+brain+injury+is+ineffective+in+a+Drosophila+model+of+obesity.">Anesthetic preconditioning of traumatic brain injury is ineffective in a Drosophila model of obesity.</a> Journal of Pharmacology and Experimental Therapeutics. 381 (3), 229-235 (2022).</li><li>Li, H., 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=Isoflurane+postconditioning+reduces+ischemia-induced+nuclear+factor-kappaB+activation+and+interleukin+1beta+production+to+provide+neuroprotection+in+rats+and+mice.">Isoflurane postconditioning reduces ischemia-induced nuclear factor-kappaB activation and interleukin 1beta production to provide neuroprotection in rats and mice.</a> Neurobiology of Disease. 54, 216-224 (2013).</li><li>Leibovitch, B. 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The authors have no conflicts of interest to declare.

Critical steps in the construction of the SAA include ensuring tight fittings to avoid leakage of the anesthetic mixture of gases. The SAA must be housed in a fume hood to avoid contamination of the laboratory space. All the elements from the carrier gas cylinders to the flow indicator downstream of the SAA should be checked as outlined in the checklist.
Other methods of administering VGAs to flies are complicated to operate (the inebriometer)21, have low throughput22, do not allow the simultaneous exposure of multiple populations23, do not allow precise control of the anesthetic concentration21, or have a readout that is difficult to translate into clinically accepted terms24.
The current version of the SAA relies on a commercial vaporizer, and hence, toxicologic studies are limited to volatile anesthetics. If used with other volatile substances, a vaporizer could be used &#34;off label&#34; after calibrating the output. Alternatively, a different method of vaporizing the volatile substances could be applied, which would require dedicated measurements to titrate the drug concentrations, as described previously25.
Apart from the flow indicators, there are no alarms (i.e., if the tanks empty, the flow through the SAA will be interrupted). Depending on the intensity of the use, the SAA may need cleaning, tightening, and possibly replacement of the Tygon tubing. We have performed &#34;maintenance&#34; on our original SAA twice in 7 years of use.
This method for anesthetizing fruit flies allows the use of the genetic toolbox available to Drosophila researchers in a high-throughput system. Multiple cohorts of flies of different populations (e.g., genotype, age, sex) can be simultaneously exposed to identical anesthetic concentrations and the desired combination of carrier gas (air, O2, N2O, noble gases) suitable to the research question at hand.
We show here that the SAA has been useful for revealing unexpected changes in resilience to isoflurane toxicity in the ND2360114 fly line and that standard laboratory fly lines differ in their responsiveness to AP.&#160;Identifying these findings was possible because of the tight control of the experimental conditions and the high throughput of the SAA.
The SAA can be adapted to study the effects of other volatile organic compounds (VOCs) on insects (e.g., honeybees). For VOCs with vapor pressures close to those of volatile anesthetics (isoflurane: 240 mmHg at 20 &#176;C), conventional vaporizers could be used, but the output would have to be calibrated. The commercial vaporizer for desflurane is heated, potentially offering additional flexibility.

The existence of collateral anesthetic effects (i.e., effects that are not immediately observable but may have delayed behavioral consequences) is generally accepted, but the understanding of their mechanisms and risk factors remains rudimentary1,2. Their delayed manifestation and subtleness limit the number of potentially important variables that can be investigated in mammalian models within reasonable time frames and at an acceptable cost. The fruit fly (Drosophila melanogaster) offers unique advantages in the context of neurodegenerative disease3 and for toxicologic screening4 that have, to date, not been applied to the study of anesthetic collateral effects. 
We developed the serial anesthesia array (SAA) to facilitate the use of fruit flies in the study of anesthetic pharmacodynamics and pharmacogenetics. A key advantage of the SAA is the simultaneous exposure to identical experimental conditions of multiple cohorts. When paired with the experimental flexibility of fruit flies, the high throughput of the SAA allows the exploration of biological and environmental variables on a scale impossible in mammalian models. 
In principle, the SAA is simply a series of connected anesthetizing locations (chambers made of 50 mL vials) through which a carrier gas delivers volatile agents. The system's first chamber contains distilled water through which the carrier gas is humidified (flies are sensitive to dehydration), and it terminates with a simple flow indicator that indicates the gas flow through the system. Fine nets placed on the openings of the connecting tubing separate the chambers to prevent the migration of flies between the chambers. The number of locations "in series" is limited by the resistance to the unpressurized gas flow (tubing, nets). 
We characterized the kinetics of this SAA prototype in a previous publication5. Although the exact pharmacokinetic properties will vary between SAAs, the relevant basics that have been tested experimentally are as follows: (i) an initial flow of 1.5-2 L/min equilibrates all the chambers (total volume of &plusmn;550 mL) with the desired concentration of anesthetic within 2 min; (ii) the concentration of anesthetic vapor delivered to the chambers does not change appreciably between the first and the last location because the amount of anesthetic contained in the volume of gas in an individual chamber (50 mL) far exceeds the amount taken up by any number of flies; and (iii) once the chambers have equilibrated, the carrier gas flow can be reduced (50-100 mL/min or less) to avoid waste and contamination of the environment (volatile anesthetics have greenhouse gas properties). The minimal flow necessary to maintain a steady-state concentration of vapor depends primarily on the leakiness of the SAA, as the uptake of vapor by the flies is negligible. Under these standard conditions (2% isoflurane and 1.5 L/min carrier gas flow), flies are anesthetized (i.e., immobile) in all positions of the array within 3-4 min, with unnoticeable differences between positions. VGAs can be administered for minutes to hours, and our typical exposure paradigms are in the range of 15 min to 2 h. To flush the system, the vaporizer is turned off, and flow is maintained to exchange approximately 10x volumes of the array (1.5 L/min for 5 min). The speed of anesthetic elimination will vary with the set rate of flow.
Volatile anesthetic agents interact with numerous still unidentified targets, including the immune-inflammatory system6. The contribution of individual molecular targets to primary versus collateral outcomes (the "anesthetic state" vs. long- and short-term "side effects") is poorly understood. Therefore, a sensitive, high-throughput fly system is valuable to inform experiments in higher animals, despite the obvious differences between flies and mammals7. Some differences can, in fact, be advantageous; for example, the fly's immune system differs from that of higher animals in that it lacks the adaptive arm of the response8. While this may seem like a limitation for understanding disease in humans, it offers a unique opportunity to study the interaction of VGAs with the innate immune-inflammatory response in isolation from the adaptive response9. This allows studies of the pharmacologic effects of VGA on inflammation and their modulation by the varied genetic backgrounds present in a population.

<table><tbody><tr><td>&lt;strong&gt;Serial Anesthesia Array:&amp;nbsp;&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>5 mL Serological Pipettes</td><td>Fisher Scientific</td><td>13-676-10C</td><td>Polystyrene, 5mL serological pipette</td></tr><tr><td>50 mL Conical Tubes</td><td>Fisher Scientific</td><td>1495949A</td><td>Polypropylene, 50 mL</td></tr><tr><td>Cable Tie Mounting Pad</td><td>Grainger</td><td>6EEE6</td><td>1.25 inch L x 1 inch W x 0.28 inch H</td></tr><tr><td>Dispensing Syringe</td><td>Grainger</td><td>5FVE0</td><td>10 mL with Luer-Lock Connection</td></tr><tr><td>Fabric Mesh Netting</td><td></td><td></td><td>1 mm mesh</td></tr><tr><td>Flow Indicator</td><td>Grainger</td><td>8RH52</td><td>5/16 to 1/2 inch connection size, paddle wheel style</td></tr><tr><td>Tygon Tubing</td><td>Tygon</td><td>E-3603</td><td>ID: 5/16, OD: 7/16, wall: 1/16</td></tr><tr><td>Wood Frame</td><td></td><td></td><td>10 feet of 2 inch x 3/4 inch</td></tr><tr><td>Zip Tie</td><td></td><td></td><td>&gt;5inch</td></tr><tr><td>&lt;strong&gt;Vaporizer Interface (Budget Alternative to Manifold):&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Dispensing Syringe</td><td>Grainger</td><td>5FVE0</td><td>10 mL with Luer-Lock Connection</td></tr><tr><td>&lt;strong&gt;Commercial Manifold and Vaporizers:&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>1/4 inch Equal Barbed Y Connector</td><td>Somni Scientific</td><td>BF-9000</td><td></td></tr><tr><td>1/8 inch NPT to 1/4 inch Barbed Elbow (Plastic)</td><td>Somni Scientific</td><td>BF-9004</td><td></td></tr><tr><td>AIR 0-4 LPM Flowmeter w/ black knob</td><td>Somni Scientific</td><td>FP-4002</td><td></td></tr><tr><td>Flowmeter auxiliary mounting bracket</td><td>Somni Scientific</td><td>NonInvPart</td><td></td></tr><tr><td>Medical Air, 1/8 inch NPT Male x DISS Male</td><td>Somni Scientific</td><td>GF-11012</td><td></td></tr><tr><td>TT-2 Table Top Anesthesia System, built in dual diverter valve system. Includes 6&#039; color coded tubing X2. (Vaporizer not Included)</td><td>Somni Scientific</td><td>TT-17000</td><td></td></tr><tr><td>Tec 7 Isoflurane Vaporizer</td><td>GE Datex-Ohmeda</td><td>1175-9101-000</td><td>Agent-specific vaporizer (Isoflurane)</td></tr><tr><td>Tec 7 Sevoflurane Vaporizer</td><td>GE Datex-Ohmeda</td><td>1175-9301-000</td><td>Agent-specific vaporizer (Sevoflurane)</td></tr></tbody></table>

the serial anesthesia array for the high-throughput investigation of volatile agents using drosophila melanogaster

Volatile general anesthetics (VGAs) are used worldwide on millions of people of all ages and medical conditions. High concentrations of VGAs (hundreds of micromolar to low millimolar) are necessary to achieve a profound and unphysiological suppression of brain function presenting as "anesthesia" to the observer. The full spectrum of the collateral effects triggered by such high concentrations of lipophilic agents is not known, but interactions with the immune-inflammatory system have been noted, although their biological significance is not understood.
To investigate the biological effects of VGAs in animals, we developed a system termed the serial anesthesia array (SAA) to exploit the experimental advantages offered by the fruit fly (Drosophila melanogaster). The SAA consists of eight chambers arranged in series and connected to a common inflow. Some parts are available in the lab, and others can be easily fabricated or purchased. A vaporizer, which is necessary for the calibrated administration of VGAs, is the only commercially manufactured component. VGAs constitute only a small percentage of the atmosphere flowing through the SAA during operation, as the bulk (typically over 95%) is carrier gas; the default carrier is air. However, oxygen and any other gases can be investigated. 
The SAA's principal advantage over prior systems is that it allows the simultaneous exposure of multiple cohorts of flies to exactly titrable doses of VGAs. Identical concentrations of VGAs are achieved within minutes in all the chambers, thus providing indistinguishable experimental conditions. Each chamber can contain from a single fly to hundreds of flies. For example, the SAA can simultaneously examine eight different genotypes or four genotypes with different biological variables (e.g., male vs. female, old vs. young). We have used the SAA to investigate the pharmacodynamics of VGAs and their pharmacogenetic interactions in two experimental fly models associated with neuroinflammation-mitochondrial mutants and traumatic brain injury (TBI).

An SAA video link is provided here: Perouansky Research Methods - Department of Anesthesiology - UW-Madison (wisc.edu) (https://anesthesia.wisc.edu/research/researchers/perouansky-laboratory/perouansky-research-methods/)&#160;Our lab has used the SAA to (i) study the effect of genotype on behavioral sensitivity to anesthetics5; (ii) screen mitochondrial mutants for the collateral effects of anesthetics11; and (iii) investigate the pharmacodynamics of isoflurane and sevoflurane on outcomes in traumatic brain injury (TBI)12,13,14,15,16,17. The published results clearly demonstrate that the genetic background influences the pharmacodynamics of clinically used VGAs with respect to both the conventional phenotype of anesthesia and the collateral effects of anesthetic toxicity, as well as tissue protection5,11,13,14,15.
Representative example 1 (Figure 2):Genetic drift in resilience to isoflurane toxicity detected by reliably reproducible experimental conditions 
The discovery of a gradual quantitative change in VGA-induced mortality among separately cultured ND2360114 flies is an example of the usefulness of reliable comparisons of anesthetic pharmacodynamics across experimental groups using the SAA. ND23 is a gene encoding a subunit in the core of Complex I of the mETC (analogous to Ndufs8 in mammals)18. Mutations in this subunit are a cause of Leigh syndrome, a lethal mitochondrial disease. We observed a gradual weakening of the isoflurane-induced mortality phenotype over time in various homozygous ND2360114 stocks cultured simultaneously under standard laboratory conditions (i.e., without exposure to VGAs). This evolutionary adaptation to isoflurane toxicity occurred in the absence of any exposure to VGAs and is probably a collateral effect of &#34;survival of the fittest&#34; within the mutant stocks. This gradual change in isoflurane sensitivity would have remained unrecognized without our confidence that the experimental conditions were identical across the assays and over time. We conclude that selection favors modifiers of the effects of ND2360114, with coincidental increased resilience to isoflurane toxicity. As inflammation in the central nervous system plays an important role in the pathogenesis of Leigh syndrome, the witnessed evolution of resistance may be due to adaptive changes in the innate immune-inflammatory response, with resistance to isoflurane toxicity being an accidental byproduct.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65144/65144fig02.jpg" /> 
Figure 2: Variation in isoflurane toxicity-induced mortality as a result of evolutionary pressure in ND2360114 flies. Seven lines (A-G) isolated from a single population through single-pair matings, expanded, and tested for 24 h mortality (PM24) following a 2 h exposure to 2% isoflurane (at 10-13 days old) show variability in the phenotype arising from a single population. Data shown as box and whisker plots. The boxes represent the second and third quartiles of the data, with the whiskers extending to the minimum and maximum data points. The mean and median are indicated by &#34;+&#34; and horizontal lines, respectively. The percent mortalities of the individual replicates (N) are shown as circles. N = 3-4 vials of 20-50 flies/vial. P-value for an ordinary one-way ANOVA; p = 0.012 indicates a significant difference among the means. <a href="https://www.jove.com/files/ftp_upload/65144/65144fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Representative example 2 (Figure 3): Illustration of a high-throughput application of the SAA to reveal genetic background effects on isoflurane pharmacodynamics 
As an example of the high throughput of the system, Figure 3 illustrates the effects of identical exposures to isoflurane (15 min of 2% isoflurane) prior to traumatic brain injury (TBI)16, a protocol testing anesthetic preconditioning (AP) in this fly model13,15,19. The readout is mortality 24 h after TBI corrected for natural attrition (MI24). In this model, all the flies regained mobility (i.e., were alive) within 30 min after TBI, and the mortality recorded in the MI24 was a result of secondary brain injury (sBI). In the four fly lines, AP with isoflurane reduced the MI24 to various degrees, indicating that responsiveness to AP is a quantitative trait. As the inflammatory response is an important factor in morbidity from sBI, AP may involve modulation of the immune system20.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65144/65144fig03.jpg" /> 
Figure 3: Influence of genetic background on the suppression of mortality (MI24) by preconditioning with isoflurane. Preconditioning flies with 15 min of 2% isoflurane (purple) reduced the mortality index at 24 h (MI24) in w1118 and y1w1118 strains (p &#60; 0.0001 and p = 0.036, respectively). The MI24 was not significantly lower in the preconditioned Oregon R (OR) and Canton S (CS) lines (p = 0.16 and p = 0.27, respectively). Data shown as box and whisker plots. The boxes represent the second and third quartiles of the data, with the whiskers extending to the minimum and maximum data points. The mean and median are indicated by &#34;+&#34; and horizontal lines, respectively. The MI24 values of the individual replicates (N) are shown as circles. N = 15-33 vials of 30-40 flies/vial for TBI-treated flies. N = 2-15 vials of 30-40 flies/vial for untreated controls. P-values from an unpaired, two-tailed Student&#39;s t-test. <a href="https://www.jove.com/files/ftp_upload/65144/65144fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about The Serial Anesthesia Array for the High-Throughput Investigation of Volatile Agents Using Drosophila melanogaster at JoVE.com

The Serial Anesthesia Array for the High-Throughput Investigation of Volatile Agents Using Drosophila melanogaster

The fruit fly (Drosophila melanogaster) is widely used for biological and toxicological research. To expand the utility of flies, we developed an instrument, the serial anesthesia array, that simultaneously exposes multiple fly samples to volatile general anesthetics (VGAs), making it possible to investigate the collateral effects (toxic and protective) of VGAs.

The Serial Anesthesia Array for the High-Throughput Investigation of Volatile Agents Using Drosophila melanogaster

Volatile general anesthetics (VGAs) are used worldwide on millions of people of all ages and medical conditions. High concentrations of VGAs (hundreds of ...

the-serial-anesthesia-array-for-high-throughput-investigation

Research

JoVE Journal

Genetics

695 Views.  University of Wisconsin-Madison. De SAA maakt verwerking met een hoge doorvoer mogelijk voor het onderzoeken van de farmacologie van vluchtige stoffen in ongewervelde modelorganismen die zijn begiftigd met een geavanceerd brein en een uitgebreid repertoire aan gedragingen. In combinatie met de genetische gereedschapskist die beschikbaar is in Drosophila melanogaster, maakt deze techniek het mogelijk om de farmacogenetica van vluchtige stoffen snel en betaalbaar te bestuderen. Veel mensen worden blootgesteld aan vluchtige sto...

Video: The Serial Anesthesia Array for the High-Throughput Investigation of Volatile Agents Using Drosophila melanogaster

High-resolution Measurement of Odor-Driven Behavior in Drosophila Larvae

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral paradigm, the experimenter usually assumes that the rapid diffusion of odorant molecules from the source leads to the creation of a stable gradient in the dish. To establish a quantitative correlation between sensory inputs and behavioral responses, it is necessary to achieve a more thorough characterization of the odorant stimulus conditions. In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral ...

Biology

High-Throughput Method for Measuring Alcohol Sedation Time of Individual Drosophila melanogaster

Drosophila melanogaster provides an excellent model to study the genetic underpinnings of alcohol sensitivity. In contrast to studies in human populations, the Drosophila model allows strict control over genetic background, and virtually unlimited numbers of individuals of the same genotype can be reared rapidly under well-controlled environmental conditions without regulatory restrictions and at relatively low cost. Flies exposed to ethanol undergo physiological and behavioral changes that resemble human alcohol intoxication, including loss of postural control, sedation, and development of tolerance. Here, we describe a simple, low-cost, high-throughput assay for assessing alcohol sedation sensitivity in large numbers of single flies. The assay is based on video recording of single flies introduced without anesthesia in 24-well cell culture plates in a set-up that enables synchronous initiation of alcohol exposure. The system enables a single person to collect individual ethanol sedation data on as many as 2,000 flies within an 8 h work period. The assay can, in principle, be extended to assess the effects of exposure to any volatile substance and applied to measure effects of acute toxicity of volatiles on other insects, including other fly species.

High-Throughput Method for Measuring Alcohol Sedation Time of Individual Drosophila melanogaster

Current methods to measure alcohol sensitivity in Drosophila are designed to test groups of flies. We present a simple, low-cost, high-throughput assay for assessing alcohol sedation sensitivity in large numbers of single flies. The method does not require specialized tools and can be performed in any laboratory using common materials.

Drosophila melanogaster provides an excellent model to study the genetic underpinnings of alcohol sensitivity. In contrast to studies in human ...

Behavior

Straightforward Assay for Quantification of Social Avoidance in Drosophila melanogaster

Drosophila melanogaster is an emerging model to study different aspects of social interactions. For example, flies avoid areas previously occupied by stressed conspecifics due to an odorant released during stress known as the Drosophila stress odorant (dSO). Through the use of the T-maze apparatus, one can quantify the avoidance of the dSO by responder flies in a very affordable and robust assay. Conditions necessary to obtain a strong performance are presented here. A stressful experience is necessary for the flies to emit dSO, as well as enough emitter flies to cause a robust avoidance response to the presence of dSO. Genetic background, but not their group size, strongly altered the avoidance of the dSO by the responder flies. Canton-S and Elwood display a higher performance in avoiding the dSO than Oregon and Samarkand strains. This behavioral assay will allow identification of mechanisms underlying this social behavior, and the assessment of the influence of genes and environmental conditions on both emission and avoidance of the dSO. Such an assay can be included in batteries of simple diagnostic tests used to identify social deficiencies of mutants or environmental conditions of interest.

Straightforward Assay for Quantification of Social Avoidance in Drosophila melanogaster

Here, we present a protocol to quantify the avoidance of stressed individuals. This paradigm is powerful yet user-friendly and can be used to assess the influence of genes and environment on one kind of social interaction in Drosophila melanogaster.

Drosophila melanogaster is an emerging model to study different aspects of social interactions. For example, flies avoid areas previously occupied by ...

Neuroscience

An Inexpensive, Scalable Behavioral Assay for Measuring Ethanol Sedation Sensitivity and Rapid Tolerance in Drosophila

Alcohol use disorder (AUD) is a serious health challenge. Despite a large hereditary component to AUD, few genes have been unambiguously implicated in their etiology. The fruit fly, Drosophila melanogaster, is a powerful model for exploring molecular-genetic mechanisms underlying alcohol-related behaviors and therefore holds great promise for identifying and understanding the function of genes that influence AUD. The use of the Drosophila model for these types of studies depends on the availability of assays that reliably measure behavioral responses to ethanol. This report describes an assay suitable for assessing ethanol sensitivity and rapid tolerance in flies. Ethanol sensitivity measured in this assay is influenced by the volume and concentration of ethanol used, a variety of previously reported genetic manipulations, and also the length of time the flies are housed without food immediately prior to testing. In contrast, ethanol sensitivity measured in this assay is not affected by the vigor of fly handling, sex of the flies, and supplementation of growth medium with antibiotics or live yeast. Three different methods for quantitating ethanol sensitivity are described, all leading to essentially indistinguishable ethanol sensitivity results. The scalable nature of this assay, combined with its overall simplicity to set-up and relatively low expense, make it suitable for small and large scale genetic analysis of ethanol sensitivity and rapid tolerance in Drosophila.

An Inexpensive, Scalable Behavioral Assay for Measuring Ethanol Sedation Sensitivity and Rapid Tolerance in Drosophila

Straightforward assays for measuring ethanol sensitivity and rapid tolerance in Drosophila facilitate the use of this model organism for investigating these important ethanol-related behaviors. Here, a relatively simple, scalable assay for measuring ethanol sensitivity and rapid tolerance in flies is described.

Alcohol use disorder (AUD) is a serious health challenge. Despite a large hereditary component to AUD, few genes have been unambiguously implicated in ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

High-resolution Quantification of Odor-guided Behavior in Drosophila melanogaster Using the Flywalk Paradigm

In their natural environment, insects such as the vinegar fly Drosophila melanogaster are bombarded with a huge amount of chemically distinct odorants. To complicate matters even further, the odors detected by the insect nervous system usually are not single compounds but mixtures whose composition and concentration ratios vary. This leads to an almost infinite amount of different olfactory stimuli which have to be evaluated by the nervous system.
To understand which aspects of an odor stimulus determine its evaluation by the fly, it is therefore desirable to efficiently examine odor-guided behavior towards many odorants and odor mixtures. To directly correlate behavior to neuronal activity, behavior should be quantified in a comparable time frame and under identical stimulus conditions as in neurophysiological experiments. However, many currently used olfactory bioassays in Drosophila neuroethology are rather specialized either towards efficiency or towards resolution.
Flywalk, an automated odor delivery and tracking system, bridges the gap between efficiency and resolution. It allows the determination of exactly when an odor packet stimulated a freely walking fly, and to determine the animal&#180;s dynamic behavioral reaction.

High-resolution Quantification of Odor-guided Behavior in Drosophila melanogaster Using the Flywalk Paradigm

The automated tracking system Flywalk is used for high-resolution quantification of odor-guided behavior in Drosophila melanogaster.

In their natural environment, insects such as the vinegar fly Drosophila melanogaster are bombarded with a huge amount of chemically distinct odorants. To ...

Olfactory Behaviors Assayed by Computer Tracking Of Drosophila in a Four-quadrant Olfactometer

A key challenge in neurobiology is to understand how neural circuits function to guide appropriate animal behaviors. Drosophila melanogaster is an excellent model system for such investigations due to its complex behaviors, powerful genetic techniques, and compact nervous system. Laboratory behavioral assays have long been used with Drosophila to simulate properties of the natural environment and study the neural mechanisms underlying the corresponding behaviors (e.g. phototaxis, chemotaxis, sensory learning and memory)1-3. With the recent availability of large collections of transgenic Drosophila lines that label specific neural subsets, behavioral assays have taken on a prominent role to link neurons with behaviors4-11. Versatile and reproducible paradigms, together with the underlying computational routines for data analysis, are indispensable for rapid tests of candidate fly lines with various genotypes. Particularly useful are setups that are flexible in the number of animals tested, duration of experiments and nature of presented stimuli. The assay of choice should also generate reproducible data that is easy to acquire and analyze. Here, we present a detailed description of a system and protocol for assaying behavioral responses of Drosophila flies in a large four-field arena. The setup is used here to assay responses of flies to a single olfactory stimulus; however, the same setup may be modified to test multiple olfactory, visual or optogenetic stimuli, or a combination of these. The olfactometer setup records the activity of fly populations responding to odors, and computational analytical methods are applied to quantify fly behaviors. The collected data are analyzed to get a quick read-out of an experimental run, which is essential for efficient data collection and the optimization of experimental conditions.

Olfactory Behaviors Assayed by Computer Tracking Of Drosophila in a Four-quadrant Olfactometer

We describe here a behavioral setup and data analysis method for assaying olfactory responses of up to 100 vinegar flies (Drosophila melanogaster). This system may be used with single or multiple olfactory stimuli, and adaptable for optogenetic activation or silencing of neuronal subsets.

A key challenge in neurobiology is to understand how neural circuits function to guide appropriate animal behaviors. Drosophila melanogaster is an ...

Electrophysiological Recording from Drosophila Trichoid Sensilla in Response to Odorants of Low Volatility

Insects rely on their sense of smell to guide a wide range of behaviors that are critical for their survival, such as food-seeking, predator avoidance, oviposition, and mating. Myriad chemicals of varying volatilities have been identified as natural odorants that activate insect Olfactory Receptor Neurons (ORNs). However, studying the olfactory responses to low-volatility odorants has been hampered by an inability to effectively present such stimuli using conventional odor-delivery methods. Here, we describe a procedure that permits the effective presentation of low-volatility odorants for in vivo Single-Sensillum Recording (SSR). By minimizing the distance between the odor source and the target tissue, this method allows for the application of biologically salient but hitherto inaccessible odorants, including palmitoleic acid, a stimulatory pheromone with a demonstrated effect on ORNs involved in courtship and mating behavior1. Our procedure thus affords a new avenue to assay a host of low-volatility odorants for the study of insect olfaction and pheromone communication.

Electrophysiological Recording from Drosophila Trichoid Sensilla in Response to Odorants of Low Volatility

The overall goal of this protocol is to demonstrate how to present odorants of low volatility for single-sensillum recording from Drosophila olfactory receptor neurons that respond to long-chain cuticular pheromones.

Insects rely on their sense of smell to guide a wide range of behaviors that are critical for their survival, such as food-seeking, predator avoidance, ...

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 ...

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Decades of research in many model organisms have led to the current concept of synaptic plasticity underlying learning and memory formation. Learning-induced changes in synaptic transmission are often distributed across many neurons and levels of processing in the brain. Therefore, methods to visualize learning-dependent synaptic plasticity across neurons are needed. The fruit fly Drosophila melanogaster represents a particularly favorable model organism to study neuronal circuits underlying learning. The protocol presented here demonstrates a way in which the processes underlying the formation of associative olfactory memories, i.e., synaptic activity and their changes, can be monitored in vivo. Using the broad array of genetic tools available in Drosophila, it is possible to specifically express genetically encoded calcium indicators in determined cell populations and even single cells. By fixing a fly in place, and opening the head capsule, it is possible to visualize calcium dynamics in these cells whilst delivering olfactory stimuli. Additionally, we demonstrate a set-up in which the fly can be subjected, simultaneously, to electric shocks to the body. This provides a system in which flies can undergo classical olfactory conditioning - whereby a previously na&#239;ve odor is learned to be associated with electric shock punishment - at the same time as the representation of this odor (and other untrained odors) is observed in the brain via two-photon microscopy. Our lab has previously reported the generation of synaptically localized calcium sensors, which enables one to confine the fluorescent calcium signals to pre- or postsynaptic compartments. Two-photon microscopy provides a way to spatially resolve fine structures. We exemplify this by focusing on neurons integrating information from the mushroom body, a higher-order center of the insect brain. Overall, this protocol provides a method to examine the synaptic connections between neurons whose activity is modulated as a result of olfactory learning.

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Here we present a protocol with which pre- and/or postsynaptic calcium can be visualized in the context of Drosophila learning and memory. In vivo calcium imaging using synaptically localized calcium sensors is combined with a classical olfactory conditioning paradigm such that the synaptic plasticity underlying this type of associative learning may be determined.

The Serial Anesthesia Array for the High-Throughput Investigation of Volatile Agents Using Drosophila melanogaster

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