We thank Dr. Linda Restifo at the University of Arizona for suggesting testing the O2 consumption of CASK mutants and for sending CASK mutants and their congenic controls. Publication fees were provided by the Departmental Reinvestment Fund from the Biology Department at the University of College Park. Space and some equipment were provided by the Universities at Shady Grove.

1. Fly rearing and collection
<ol>
	<li>Maintain flies at 25 &#176;C in narrow vials containing standard Drosophila food. 
	NOTE: The sample size for each genotype should comprise at least nine replicates, each consisting of a single respirometer chamber containing 15-25 flies, set up as described below.</li>
	<li>Transfer the flies every 2-3 days.</li>
	<li>Anesthetize flies with CO2, collect groups of 15-25 males of each genotype, and place each group into fresh, unyeasted food vials. 
	NOTE: Males were used to reduce variability due to reproductive status. The method applies to both sexes.</li>
	<li>Allow the flies to recover at 25 &#176;C for at least 24 h. 
	NOTE: By the time of the experiment, flies should be 1-4 days old. The frequency of collections described in step 1.3 can be set to narrow the age range of the flies.</li>
</ol>
2. Setup and assembly of respirometer chamber
<ol>
	<li>Turn on the water bath and set it to the desired temperature for the experiment. 
	NOTE: The experiments below were conducted at 25 &#176;C using 50 mL Schlenk tubes as chambers. Components are to be assembled as shown in Figures 1A, 1B, and 1C.</li>
	<li>Clean the ground glass joints of chambers and sensor plugs thoroughly by spraying 70% ethanol onto a laboratory wipe (not directly onto the joint) and wiping dust and old grease from the sensor plug (Figure 1A). Wipe off ethanol with a fresh laboratory wipe.</li>
	<li>Place 1 cm piece of cotton roll soaked in purified water into the bottom of the chamber to stabilize the humidity.
	<ol>
		<li>Add enough water (~0.5 mL) to form a small pool at the bottom of the cotton roll.</li>
	</ol>
	</li>
	<li>Wipe off any water that has spilled onto the joint of the chamber.</li>
	<li>Transfer the flies to labeled polypropylene tubes using a funnel.
	<ol>
		<li>Plug the tube with a cotton roll. 
		NOTE: Tubes consist of a 5 mL polypropylene test tube, trimmed to 5.5 cm in length and perforated with a hot knife to allow the free exchange of air with the experimental chamber. CO2 anesthesia is known to cause metabolic abnormalities, so flies are transferred without anesthesia which requires more care to avoid losing the flies.</li>
	</ol>
	</li>
	<li>Add one ventilated tube with flies into each respirometer chamber (on top of wet cotton).</li>
	<li>Fill soda lime cartridges (4-5 pellets per tube) and place them on the top of the tube containing flies inside the chamber. 
	NOTE: Soda lime cartridges consist of 800 &#181;L centrifuge tubes perforated 4-5 times with a power drill.</li>
	<li>Fill O2 generators with saturated copper sulfate (CuSO4) solution below level of vent holes 
	NOTE: O2 generators consist of screw-cap centrifuge tubes with 4 holes drilled below the threads. Platinum (Pt) and Copper (Cu) electrodes are soldered to two-pin connector, inserted into holes drilled in cap, and affixed with epoxy. Electrolysis of CuSO4 generates the O2 consumed by the experimental organism. CuSO4 is toxic to invertebrates, avoid spills or leakage and clean up immediately.</li>
	<li>Connect the filled O2 generator to two-pin connector on the sensor plug. 
	NOTE: The copper cathode must connect with the negative output of the controller and the platinum anode to the positive wire. Reversed connections will cause the failure of the experiment.</li>
	<li>Place two small dabs of clear silicone grease on opposite sides of the ground glass joint of the sensor plug.</li>
	<li>Insert the plug into the chamber and rotate the plug (or chamber) with moderate pressure to spread the grease in the joint.
	<ol>
		<li>Wipe off excess grease with a laboratory wipe.</li>
	</ol>
	</li>
	<li>Snap plastic Keck clamps onto joints to secure plugs in chambers. The assembled chamber should look like Figure 1C.</li>
	<li>Repeat the above steps for the number of chambers used for the day&#39;s experiment. 
	NOTE: The number of chambers that can be recorded is limited by the number of available chambers, controllers and USB inputs to the computer. For the present experiments, seven chambers were normally run in parallel. Experimental flies such as mutants should be matched with appropriate controls. A chamber set up identically but without flies should be included in each experiment as a control for environmental variation. Chambers containing different treatments (mutant, wildtype, no-fly) should be rotated between experiments.</li>
	<li>Place assembled chambers into a rack in the water bath with stopcocks open (Figure 1E). 
	NOTE: To avoid circadian variation, chambers were placed into the bath between 9:30 and 9:50 am for all experiments described here.</li>
	<li>Leave stopcocks open (Keep the handle parallel to the stopcock). 
	NOTE: Be careful not to allow water to enter the stopcocks.</li>
	<li>Allow the chambers to equilibrate with stopcocks open for about 30 min. 
	&#8203;NOTE: While the chamber is equilibrated, connect the electronics and set up data acquisition as described below.</li>
</ol>
3. Setting up controllers and computer
<ol>
	<li>Be sure that the switches supplying current to the O2 generators are in the OFF position (away from the connector; Figure 1D).</li>
	<li>Plug each controller box into an available Universal serial bus (USB) port. 
	NOTE: Construction and programming of controller units described elsewhere12.</li>
	<li>Connect controllers to respirometer chambers using 6-conductor cables.</li>
	<li>Check that the organic light emitting diode (OLED) displays of the controllers (Figure 1D) are displaying environmental parameters.</li>
	<li>Briefly turn on O2 generators using the switch on the controller (Figure 1D).
	<ol>
		<li>If the current value increases from zero to between 35 and 55 mA, the controller and chamber are ready for experiments.</li>
	</ol>
	</li>
	<li>Determine which COM ports are being used by the controllers, as described below.,
	<ol>
		<li>Click the Start Icon in Microsoft Windows.</li>
		<li>Click the Settings Icon.</li>
		<li>Click Bluetooth and Devices.</li>
		<li>Ensure that the controllers and their COM ports appear in the list of devices.</li>
	</ol>
	</li>
	<li>Open PuTTY on the desktop and set up a log file for each channel of the respirometer as described below. 
	NOTE: PuTTY is a free secure shell and telnet client that is used to transfer data to the computer via COM ports.
	<ol>
		<li>Select COM port for a controller by typing the number of the port in the &#34;Serial line&#34; box (Figure 2A).</li>
		<li>Click on Logging.</li>
		<li>Select Printable Output in &#34;Session logging&#34; (Figure 2B).</li>
		<li>Under Log File Name click Browse.</li>
		<li>In the folder of the choice, create a filename containing descriptive information (e.g., date, species, COM port number).</li>
		<li>Click Save.</li>
		<li>Click Open. A window will open showing comma-delimited data being logged (Figure 2C).</li>
		<li>Repeat for all other controllers in use for the experiment. Input to each COM port will appear as a separate window (Figure 2D).</li>
	</ol>
	</li>
</ol>
4. Running experiments
<ol>
	<li>Once chambers have equilibrated for 30 min, seal them by closing stopcocks.</li>
	<li>Cover the bath and chambers with a polystyrene foam box to maintain a stable environment.</li>
	<li>Allow to equilibrate for another hour.</li>
	<li>Turn on the current to the O2 generator of each chamber using the switch on the controller box.</li>
	<li>Once the O2 generators are activated, ensure that the pressure increases to pre-set OFF pressure. 
	NOTE: 1017 hPa, which is slightly above atmospheric pressure, was used as the &#34;OFF&#34; pressure in this series of experiments. Return to the ambient pressure will indicate leakage of gas from the chambers. Further, it allows the same pressure to be used across experiments regardless of ambient barometric pressure. The &#34;ON&#34; pressure was 1016 hPa, meaning that pressure only needed to drop 1 hPa before the O2 generator was activated. This provided adequate sensitivity to measure O2 consumption in Drosophila. Once a chamber is pressurized to the &#34;OFF&#34; setting, current should drop to zero.</li>
	<li>Let the experiment run for 3 or more h. 
	NOTE: Higher VO2 at&#160;elevated temperatures can allow for shorter experiment times. Monitor occasionally to ensure that equipment is functioning but avoid excessive activity near the chambers that may affect temperature stability.</li>
</ol>
5. Finishing experiment
<ol>
	<li>Turn off O2 generators on all controllers. 
	NOTE: Do first to avoid running the O2 generators while the chambers are open.</li>
	<li>Open the stopcocks to unseal the chambers.</li>
	<li>Leave the PuTTY windows open for another 5-15 min to provide a final baseline.</li>
	<li>Close the PuTTY window for each controller, ending recordings. 
	&#8203;NOTE: All experiments ended between 4:50 and 5:10 pm.</li>
	<li>Disconnect sensors from cables.</li>
	<li>Move chambers to dry rack.</li>
	<li>Remove sensor plugs one at a time from the chambers.</li>
	<li>Disconnect the O2 generators and place them in the tube rack.</li>
	<li>Wipe grease off the sensor plug and keep it in the rack.</li>
	<li>Clean grease from chamber joints and remove tubes with flies and soda lime.</li>
	<li>Anesthetize flies in each tube with CO2, tap onto a weight boat and weigh on a microbalance.
	<ol>
		<li>Log the weight and number of flies for each tube.</li>
	</ol>
	</li>
	<li>Discard flies or set them aside for additional procedures.</li>
	<li>Dump soda lime from cartridges into the waste container.</li>
	<li>Open the O2 generator and discard the CuSO4 solution into the waste container.
	<ol>
		<li>Rinse electrodes and tube with purified water.</li>
		<li>Place the tube racks for drying.</li>
	</ol>
	</li>
</ol>
6. Analysis of charge transfer data
<ol>
	<li>Import Data as comma-delimited text into a spreadsheet, with each record comprising a separate worksheet.</li>
	<li>Record the current and time data for each pulse of the O2 generator. Starting with the first pulse after the chamber was pressurized, record the start time and end time (as row numbers) of each current pulse. That is the row number when the current goes above zero (usually to about 45-50 mA) to the last row that is above zero.</li>
	<li>Make a table on the worksheet to record the following data:
	<ol>
		<li>The average current amplitude during the pulse: = AVERAGE([first row of pulse]:[last row of pulse]) for each pulse (from the current column).</li>
		<li>Pulse duration: ([Last row of pulse] - [first row of pulse[-one row]])/1000 for each pulse (from the time in milliseconds column).</li>
		<li>Total experiment time: [time at start of last pulse] - [time at end of first pulse after chamber pressurized] (from the time in minutes column).</li>
	</ol>
	</li>
	<li>Then calculate charge transfer (Q) for each pulse (average current X duration)</li>
	<li>Sum the charge from all pulses to calculate Total Charge (Qtot).</li>
</ol>
7. Analysis of O2 consumption
<ol>
	<li>Set up a new spreadsheet for all data and enter or calculate the following for each chamber:
	<ol>
		<li>Qtot (total charge)</li>
		<li>Moles (= Q &#247; 96485 &#215; 4)</li>
		<li>mL O2 (= moles &#215; 22413 mL/mol)</li>
		<li>Total time (from the data analysis above)</li>
		<li>mL min-1 (= ml O2 &#247; total time)</li>
		<li>Weight in grams (flies anesthetized and weighed measured after the experiment)</li>
		<li>mL min-1 g-1 (= mL min-1 &#247; weight in grams)</li>
		<li>mL/h/g (the above &#215; 60)</li>
		<li>mg/fly (= weight of flies &#247; number of flies)</li>
		<li>&#956;L fly-1 h-1 (= (mL min-1 &#215; 3600) &#247; number of flies).</li>
	</ol>
	</li>
	<li>Tabulate data for each treatment (genotype, e.g.)</li>
	<li>Compare treatments using ANOVA, t-test, or Mann-Whitney u-test 13.</li>
</ol>

Adapting Microrespirometry for Drosophila - Insights into CASK Mutant Metabolism

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W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lack+of+correlation+between+body+mass+and+metabolic+rate+in+Drosophila+melanogaster.">Lack of correlation between body mass and metabolic rate in Drosophila melanogaster.</a> Journal of Insect Physiology. 50 (5), 445-453 (2004).</li><li>Norton, F. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Permeation+of+gases+through+solids.">Permeation of gases through solids.</a> Journal of Applied Physics. 28 (1), 34-39 (1957).</li><li>Hoegh-Guldberg, O., Manahan, D. 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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+overexpression+of+copper-+and+zinc-containing+superoxide+dismutase+and+catalase+retards+age-related+oxidative+damage+and+increases+metabolic+potential+in+Drosophila+melanogaster.">Simultaneous overexpression of copper- and zinc-containing superoxide dismutase and catalase retards age-related oxidative damage and increases metabolic potential in Drosophila melanogaster.</a> Journal of Biological Chemistry. 270 (26), 15671-15674 (1995).</li><li>Van Voorhies, W. A., Melvin, R. G., Ballard, J. W. O., Williams, J. 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The authors declare no conflicts of interest.

The above procedure demonstrates measurement of O2 consumption in D. Melanogaster using an electronic coulometric microrespirometer. The resulting data for O2 consumption in wild-type D. melanogaster were within the ranges described in most previous publications using diverse methods (Table 1) although somewhat lower than that reported by others3,6.
Critical steps addressed the t...

The ability to measure metabolic rate is crucial for a complete understanding of an organism in its environmental context. For example, it is necessary to measure metabolic rate in order to understand its role in lifespan1, the role of diet in metabolism2, or the threshold for hypoxic stress3.
There are two general approaches to measuring the metabolic rate4. Direct calorimetry measures energy expenditure directly by measuring heat production. Indirect calorimetry measures energy production through other means, often via respirometric measurement of O2 consumption (VO2), CO2 production or both. Although direct calorimetry has been applied to small ectotherms, including Drosophila melanogaster5, respirometry is technically simpler and more commonly used.
Several forms of respirometry have been used successfully to measure metabolic rate in wildtype and mutant D. melanogaster and have provided insight into the metabolic effects of temperature6, social environment3, diet3,7,and neurodevelopmental disorders8. These fall into two classes, which vary considerably in cost and complexity. Manometry is the simplest and least expensive9,10, in which flies are placed into a sealed chamber that contains a CO2 absorbent and which is connected via a thin capillary to a fluid reservoir. As O2 is consumed and CO2 absorbed, pressure in the chamber decreases and fluid is drawn into the capillary. The fluid-filled volume of the capillary is therefore proportional to VO2. More elaborate versions, which compensate for the force exerted by the fluid in the capillary, have also been used on D. melanogaster1. Manometry has the advantages of being simple and inexpensive, but, because it is sensitive to pressure, requires constant environmental conditions. Further, because consumed O2 is not replaced, the partial pressure of O2 (PO2) gradually decreases inside the chambers.
Respirometry using gas analysis is also regularly used for D. melanogaster. In this case, gases are sampled at regular intervals from sealed chambers containing flies and sent to an infrared analyzer2,6,11. This type of apparatus has the advantages that it is available commercially, is less sensitive to environmental conditions, and gases are refreshed during sampling so that PO2 remains stable. However, the equipment can be expensive and complex to operate.
A recently developed coulometric microrespirometer12 provides an inexpensive, sensitive, and stable alternative to existing systems. In practice, an organism is placed into an airtight chamber where it consumes O2 and the exhaled CO2 is removed by an absorbent material, resulting in a net decrease in chamber pressure. When the internal pressure decreases to a pre-set threshold (ON-threshold), current is passed through an electrolytic O2 generator, returning pressure to a second threshold (OFF-threshold) stopping electrolysis. Charge transfer across the O2 generator is directly proportional to the amount of O2 required to re-pressurize the chamber and can therefore be used to measure the O2 consumed by the organism4. The method is highly sensitive, measures VO2 precisely, and the regular replacement of O2 can maintain PO2 at a nearly constant level for hours or days.
The coulometric microrespirometer used in this study employs a multi-modal (pressure, temperature, and humidity) electronic sensor. The sensor is operated by a microcontroller that detects small changes in pressure and activates O2 generation when a low pressure threshold is reached12. This apparatus is assembled from off the shelf parts, can be used with a wide variety of chambers and experimental environments, and has been employed successfully to examine the effects of body mass and temperature on the beetle Tenebrio molitor. In the present study, the microrespirometer has been adapted to measure O2 consumption in D. melanogaster, which has approximately 1% of the mass of T. molitor. Sensitivity of the apparatus has been increased by reducing the threshold for activating O2 generation, and environmental stability has been enhanced by conducting experiments in a temperature-controlled water bath and by maintaining humidity inside the chambers at or near 100%.
The CASK (Calmodulin-dependent Serine Protein Kinase) protein, part of the family of membrane-associated guanylate kinases (MAGUK), is a molecular scaffold in different multi-protein complexes, and mutations in CASK are associated with neurodevelopmental disorders in humans and in D. melanogaster13,14. A viable D. melanogaster mutant, CASK&#916;18, disrupts activity of dopaminergic neurons15 and reduces activity levels by more than 50% compared to congenic controls14,16. Because of the reduced activity levels of CASK mutants and the role of catecholamines in regulating metabolism17 we hypothesized that their standard metabolic rate, and therefore O2 consumption, would be dramatically reduced compared to controls.
O2 consumption was measured in CASK&#916;18 and their wildtype congeners, w(ex33). Groups of flies were placed into respirometry chambers, O2 consumption was measured, O2 consumption was calculated and expressed on both a mass-specific and per-fly basis. The apparatus recorded VO2 in wildtype flies that was consistent with previous studies, and it could differentiate between the per-fly O2 consumption of wildtype and CASK mutant flies.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>19/22 Thermometer Adapter</td>
			<td>Wilmad-Labglass</td>
			<td>ML-280-702</td>
			<td>Sensor Plug</td>
		</tr>
		<tr>
			<td>2 ml Screwcap Tubes</td>
			<td>Fisher</td>
			<td>3464</td>
			<td>O2 generator</td>
		</tr>
		<tr>
			<td>2-Pin Connector</td>
			<td>Zyamy</td>
			<td>40PIN-RFB10</td>
			<td>O2 generator: cut to 2-pin</td>
		</tr>
		<tr>
			<td>4-Pin Female Connector</td>
			<td>TE Connectivity</td>
			<td>215299-4</td>
			<td>Sensor Plug</td>
		</tr>
		<tr>
			<td>5 ml Polypropylene Tube</td>
			<td>Falcon</td>
			<td>352063</td>
			<td>Cut to 5.5 cm and perforated&#160;</td>
		</tr>
		<tr>
			<td>50 ml Schlenk Tube 19/22 Joint</td>
			<td>Laboy</td>
			<td>HMF050804</td>
			<td>Chamber</td>
		</tr>
		<tr>
			<td>6-Conductor Cable</td>
			<td>Zenith</td>
			<td>6-Conductor 26 ga</td>
			<td>Cable</td>
		</tr>
		<tr>
			<td>6-Pin Female Bulkhead Connector</td>
			<td>Switchcraft</td>
			<td>17982-6SG-300</td>
			<td>Controller</td>
		</tr>
		<tr>
			<td>6-Pin Female Connector</td>
			<td>Switchcraft</td>
			<td>18982-6SG-522</td>
			<td>Sensor plug</td>
		</tr>
		<tr>
			<td>6-Pin Male Connector</td>
			<td>Switchcraft</td>
			<td>16982-6PG-522</td>
			<td>Cable</td>
		</tr>
		<tr>
			<td>800 ul centrifuge tube</td>
			<td>Fisher</td>
			<td>05-408-120</td>
			<td>Soda Lime Cartridge</td>
		</tr>
		<tr>
			<td>ABS Plastic Enclosure</td>
			<td>Bud Industries</td>
			<td>PS-11533-G</td>
			<td>Controller</td>
		</tr>
		<tr>
			<td>Arduino Nano Every</td>
			<td>Arduino LLC</td>
			<td>ABX00028</td>
			<td>Controller</td>
		</tr>
		<tr>
			<td>BME 280 Sensor</td>
			<td>DIYMall</td>
			<td>FZ1639-BME280</td>
			<td>Sensor Plug</td>
		</tr>
		<tr>
			<td>Circuit Board</td>
			<td>Lheng</td>
			<td>5 X 7 cm</td>
			<td>Controller</td>
		</tr>
		<tr>
			<td>Copper Sulfate</td>
			<td>BioPharm</td>
			<td>BC2045</td>
			<td>O2 Generator</td>
		</tr>
		<tr>
			<td>Computer</td>
			<td>Azulle</td>
			<td>Byte4</td>
			<td>Data Acquisition</td>
		</tr>
		<tr>
			<td>Cotton Rolls</td>
			<td>Kajukajudo</td>
			<td>#2</td>
			<td>Cut in half to plug fly tubes 
			Cut in quarters for humidity</td>
		</tr>
		<tr>
			<td>Environmental Chamber</td>
			<td>Percival</td>
			<td>I30 VLC8</td>
			<td>Fly Care</td>
		</tr>
		<tr>
			<td>Epoxy</td>
			<td>JB Weld</td>
			<td>Plastic Bonder</td>
			<td>Secure Electrodes in O2 Generator</td>
		</tr>
		<tr>
			<td>Fly Food</td>
			<td>Lab Express</td>
			<td>Type R</td>
			<td>Fly Care</td>
		</tr>
		<tr>
			<td>Keck Clamps</td>
			<td>uxcell</td>
			<td>a20092300ux0418</td>
			<td>Secures glass joint of chamber to plug</td>
		</tr>
		<tr>
			<td>Low-Viscosity Epoxy</td>
			<td>Loctite</td>
			<td>E-30CL</td>
			<td>Sensor Plug</td>
		</tr>
		<tr>
			<td>OLED Display</td>
			<td>IZOKEE</td>
			<td>IZKE31-IIC-WH-3</td>
			<td>Controller</td>
		</tr>
		<tr>
			<td>Platinum Wire 24 ga</td>
			<td>uGems</td>
			<td>14349</td>
			<td>O2 generator</td>
		</tr>
		<tr>
			<td>Silicone grease</td>
			<td>Dow-Corning</td>
			<td>High Vacuum Grease</td>
			<td>Seals chamber-plug connection</td>
		</tr>
		<tr>
			<td>Soda Lime</td>
			<td>Jorvet</td>
			<td>JO553</td>
			<td>CO2 absorption</td>
		</tr>
		<tr>
			<td>Toggle Switch</td>
			<td>E-Switch</td>
			<td>100SP1T1B1M1QEH</td>
			<td>Controller</td>
		</tr>
		<tr>
			<td>USB Cable</td>
			<td>Sabrent</td>
			<td>CB-UM63</td>
			<td>Controller</td>
		</tr>
		<tr>
			<td>USB Hub</td>
			<td>Atolla</td>
			<td>Hub 3.0</td>
			<td>Connect controllers to computer</td>
		</tr>
		<tr>
			<td>Water bath</td>
			<td>Amersham</td>
			<td>56-1165-33</td>
			<td>Temperature Control</td>
		</tr>
		<tr>
			<td>Water Bath Tank</td>
			<td>Glass Cages</td>
			<td>15-liter rimless acrylic</td>
			<td>Bath for Respirometers</td>
		</tr>
	</tbody>
</table>

measuring o2 consumption in drosophila melanogaster using coulometric microrespirometry

Coulometric microrespirometry is a straightforward, inexpensive method for measuring the O2 consumption of small organisms while maintaining a stable environment. A coulometric microrespirometer consists of an airtight chamber in which O2 is consumed and the CO2 produced by the organism is removed by an absorbent medium. The resulting pressure decrease triggers electrolytic O2 production, and the amount of O2 produced is measured by recording the amount of charge used to generate it. In the present study, the method has been adapted to Drosophila melanogaster tested in small groups, with the sensitivity of the apparatus and the environmental conditions optimized for high stability. The amount of O2 consumed by wildtype flies in this apparatus is consistent with that measured by previous studies. Mass-specific O2 consumption by CASK mutants, which are smaller and known to be less active, was not different from congenic controls. However, the small size of CASK mutants resulted in a significant reduction in O2 consumption on a per-fly basis. Therefore, the microrespirometer is capable of measuring O2 consumption in D. melanogaster, can distinguish modest differences between genotypes, and adds a versatile tool for measuring metabolic rates.

The pressure and current outputs of the respirometer controller are shown for one chamber in one experiment in Figure 3A. The first, long current pulse pressurized the chamber from ambient pressure (approximately 992 hPa) to the pre-set OFF threshold of 1017 hPa. As the flies consumed O2 and CO2 was absorbed, pressure decreased slowly until it reached the ON threshold of 1016 hPa, which activated current through the O2 generator. In the example shown, the ave...

Watch this Scientific Journal Video about Influence of Temperature on Drosophila melanogaster and Desert-Adapted Beetles at JoVE.com

Author Spotlight: Influence of Temperature on Drosophila melanogaster and Desert-Adapted Beetles 

Coulometric respirometry is ideal for measuring the metabolic rate of small organisms. When adapted for Drosophila melanogaster in the present study, measured O2 consumption was within the range reported for wildtype D. melanogaster by previous studies. Per-fly O2 consumption by CASK mutants, which are smaller and less active, was significantly lower than the wildtype.

Measuring O2 Consumption in Drosophila melanogaster Using Coulometric Microrespirometry

Coulometric microrespirometry is a straightforward, inexpensive method for measuring the O2 consumption of small organisms while maintaining a stable ...

measuring-o2-consumption-drosophila-melanogaster-using-coulometric

Research

JoVE Journal

Neuroscience

805 Views.  University of Maryland, College Park, Universities at Shady Grove. Coulometric respirometry is ideal for measuring the metabolic rate of small organisms. When adapted for Drosophila melanogaster in the present study, measured O2 consumption was within the range reported for wildtype D. melanogaster by previous studies. Per-fly O2 consumption by CASK mutants, which are smaller and less active, was significantly lower than the wildtype. 

Video: Author Spotlight: Influence of Temperature on Drosophila melanogaster and Desert-Adapted Beetles 

Magnetic Resonance Spectroscopy of live Drosophila melanogaster using Magic Angle Spinning

High-Resolution Magic Angle Spinning (HRMAS) proton magnetic resonance spectroscopy (1H-MRS) is a novel non-destructive technique that improves spectral line-widths and allows high-resolution spectra to be obtained from extracts, intact cells, cell cultures, and more importantly intact tissue to investigate relationships between metabolites and cellular processes. In vivo HRMAS 1H-MRS studies have yet to be reported in the live fruit fly Drosophila melanogaster. Drosophila, as a simpler genetic organism, allows the multiple biological functions and various evolutionarily conserved signaling pathways to be examined at the whole organism level and it is a useful model for investigating genetics and physiology. To this end, we developed and implemented an in vivo HRMAS 1H-MRS method to investigate live Drosophila at 14.1 T. Here, we outline an HRMAS 1H-MRS protocol for the molecular characterization of Drosophila with a conventional MR spectrometer equipped with an HRMAS probe. This technique is a novel, in vivo, non-destructive Drosophila metabolite measurement approach, which enables the identification of disease biomarkers and thus may contribute to novel therapeutic development.

Magnetic Resonance Spectroscopy of live Drosophila melanogaster using Magic Angle Spinning

This technique enables the use of high-resolution magic angle spinning proton MR spectroscopy (HRMAS 1H-MRS) for molecular characterization of live Drosophila melanogaster with a conventional 14.1 tesla spectrometer equipped with an HRMAS probe.

High-Resolution Magic Angle Spinning (HRMAS) proton magnetic resonance spectroscopy (1H-MRS) is a novel non-destructive technique that improves spectral ...

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed in sensory hairs called sensilla, which cover the surface of olfactory organs. The surface of each sensillum is covered with tiny pores, through which odorants pass and dissolve in a fluid called sensillum lymph, which bathes the sensory dendrites of the OSNs housed in a given sensillum. The OSN dendrites express odorant receptor (OR) proteins, which in insects function as odor-gated ion channels4, 5. The interaction of odorants with ORs either increases or decreases the basal firing rate of the OSN. This neuronal activity in the form of action potentials embodies the first representation of the quality, intensity, and temporal characteristics of the odorant6, 7.
Given the easy access to these sensory hairs, it is possible to perform extracellular recordings from single OSNs by introducing a recording electrode into the sensillum lymph, while the reference electrode is placed in the lymph of the eye or body of the insect. In Drosophila, sensilla house between one and four OSNs, but each OSN typically displays a characteristic spike amplitude. Spike sorting techniques make it possible to assign spiking responses to individual OSNs. This single sensillum recording (SSR) technique monitors the difference in potential between the sensillum lymph and the reference electrode as electrical spikes that are generated by the receptor activity on OSNs1, 2, 8. Changes in the number of spikes in response to the odorant represent the cellular basis of odor coding in insects. Here, we describe the preparation method currently used in our lab to perform SSR on Drosophila melanogaster and Anopheles gambiae, and show representative traces induced by the odorants in a sensillum-specific manner.

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Electrophysiological responses of olfactory sensory neurons to odorants can be measured in insects using single sensillum recordings. In this video article we will demonstrate how to perform single sensillum recordings in the antennae of the vinegar fly (Drosophila melanogaster) and the maxillary palps of the malaria mosquito (Anopheles gambiae).

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed ...

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

A growing number of genetically encoded tools are becoming available that allow non-invasive manipulation of the neural activity of specific neurons in Drosophila melanogaster1. Chief among these are optogenetic tools, which enable the activation or silencing of specific neurons in the intact and freely moving animal using bright light. Channelrhodopsin (ChR2) is a light-activated cation channel that, when activated by blue light, causes depolarization of neurons that express it. ChR2 has been effective for identifying neurons critical for specific behaviors, such as CO2 avoidance, proboscis extension and giant-fiber mediated startle response2-4. However, as the intense light sources used to stimulate ChR2 also stimulate photoreceptors, these optogenetic techniques have not previously been used in the visual system. Here, we combine an optogenetic approach with a mutation that impairs phototransduction to demonstrate that activation of a cluster of loom-sensitive neurons in the fly's optic lobe, Foma-1 neurons, can drive an escape behavior used to avoid collision. We used a null allele of a critical component of the phototransduction cascade, phospholipase C-&beta;, encoded by the norpA gene, to render the flies blind and also use the Gal4-UAS transcriptional activator system to drive expression of ChR2 in the Foma-1 neurons. Individual flies are placed on a small platform surrounded by blue LEDs. When the LEDs are illuminated, the flies quickly take-off into flight, in a manner similar to visually driven loom-escape behavior. We believe that this technique can be easily adapted to examine other behaviors in freely moving flies.

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

Genetically encoded optogenetic tools enable noninvasive manipulation of specific neurons in the Drosophila brain. Such tools can identify neurons whose activation is sufficient to elicit or suppress particular behaviors. Here we present a method for activating Channelrhodopsin2 that is expressed in targeted neurons in freely walking flies.

A  growing number of genetically encoded tools are becoming available that allow non-invasive  manipulation of the neural activity of specific neurons in ...

Determination of the Spontaneous Locomotor Activity in Drosophila melanogaster

Drosophila melanogaster has been used as an excellent model organism to study environmental and genetic manipulations that affect behavior. One such behavior is spontaneous locomotor activity. Here we describe our protocol that utilizes Drosophila population monitors and a tracking system that allows continuous monitoring of the spontaneous locomotor activity of flies for several days at a time. This method is simple, reliable, and objective and can be used to examine the effects of aging, sex, changes in caloric content of food, addition of drugs, or genetic manipulations that mimic human diseases.

Determination of the Spontaneous Locomotor Activity in Drosophila melanogaster

Drosophila melanogaster are useful in studying genetic or environmental manipulations that affect behaviors such as spontaneous locomotor activity.&#160; Here we describe a protocol that utilizes monitors with infrared beams and data analysis software to quantify spontaneous locomotor activity.&#160;

Drosophila melanogaster has been used as an excellent model organism to study environmental and genetic manipulations that affect behavior. One such ...

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 ...

Conditions Affecting Social Space in Drosophila melanogaster

The social space assay described here can be used to quantify social interactions of Drosophila melanogaster &#8212; or other small insects &#8212; in a straightforward manner. As we previously demonstrated 1, in a two-dimensional chamber, we first force the flies to form a tight group, subsequently allowing them to take their preferred distance from each other. After the flies have settled, we measure the distance to the closest neighbor (or social space), processing a static picture with free online software (ImageJ). The analysis of the distance to the closest neighbor allows researchers to determine the effects of genetic and environmental factors on social interaction, while controlling for potential confounding factors. Diverse factors such as climbing ability, time of day, sex, and number of flies, can modify social spacing of flies. We thus propose a series of experimental controls to mitigate these confounding effects. This assay can be used for at least two purposes. First, researchers can determine how their favorite environmental shift (such as isolation, temperature, stress or toxins) will impact social spacing 1,2. Second, researchers can dissect the genetic and neural underpinnings of this basic form of social behavior 1,3. Specifically, we used it as a diagnostic tool to study the role of orthologous genes thought to be involved in social behavior in other organisms, such as candidate genes for autism in humans 4.

Conditions Affecting Social Space in Drosophila melanogaster

The effect of genes and environment on social space of Drosophila melanogaster can be quantified through a powerful but straightforward analytical paradigm. We show here different factors that can affect this social space, and thus need to be taken into consideration when designing experiments in this paradigm.

The social space assay described here can be used to quantify social interactions of Drosophila melanogaster &#8212; or other small insects &#8212; in a ...

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 ...

A Behavioral Assay for Mechanosensation of MARCM-based Clones in Drosophila melanogaster

Because of the structural and functional homology to the hair cells of the mammalian inner ear, the neurons that innervate the Drosophila external sense organs provide an excellent model system for the study of mechanosensation. This protocol describes a simple touch behavior in fruit flies which can be used to identify mutations that interfere with mechanosensation. The tactile stimulation of a macrochaete bristle on the thorax of flies elicits a grooming reflex from either the first or third leg. Mutations that interfere with mechanotransduction (such as NOMPC), or with other aspects of the reflex arc, can inhibit the grooming response. A traditional screen of adult behaviors would have missed mutants that have essential roles during development. Instead, this protocol combines the touch screen with mosaic analysis with a repressible cell marker (MARCM) to allow for only limited regions of homozygous mutant cells to be generated and marked by the expression of green fluorescent protein (GFP). By testing MARCM clones for abnormal behavioral responses, it is possible to screen a collection of lethal p-element mutations to search for new genes involved in mechanosensation that would have been missed by more traditional methods.

A Behavioral Assay for Mechanosensation of MARCM-based Clones in Drosophila melanogaster

In order to identify novel mutations affecting mechanosensation, we designed an assay that measures the behavioral response to tactile stimulation of fly bristles in mutant clones generated by the MARCM method. The combination of techniques allows for the identification of mechanosensitive mutations that would otherwise be lethal.

Because of the structural and functional homology to the hair cells of the mammalian inner ear, the neurons that innervate the Drosophila external sense ...

Foraging Path-length Protocol for Drosophila melanogaster Larvae

The Drosophila melanogaster larval path-length phenotype is an established measure used to study the genetic and environmental contributions to behavioral variation. The larval path-length assay was developed to measure individual differences in foraging behavior that were later linked to the foraging gene. Larval path-length is an easily scored trait that facilitates the collection of large sample sizes, at minimal cost, for genetic screens. Here we provide a detailed description of the current protocol for the larval path-length assay first used by Sokolowski. The protocol details how to reproducibly handle test animals, perform the behavioral assay and analyze the data. An example of how the assay can be used to measure behavioral plasticity in response to environmental change, by manipulating feeding environment prior to performing the assay, is also provided. Finally, appropriate test design as well as environmental factors that can modify larval path-length such as food quality, developmental age and day effects are discussed.

Foraging Path-length Protocol for Drosophila melanogaster Larvae

We provide a detailed protocol for a Drosophila melanogaster foraging path-length assay. We discuss the preparation and handling of test animals, how to perform the assay and analyze the data.

The Drosophila melanogaster larval path-length phenotype is an established measure used to study the genetic and environmental contributions to behavioral ...

A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

We describe a protocol for measuring ethanol self-administration in fruit flies (Drosophila melanogaster) as a proxy for changes in reward states. We demonstrate a simple way to tap into the fly reward system, modify experiences related to natural reward, and use voluntary ethanol consumption as a measure for changes in reward states. The approach serves as a relevant tool to study the neurons and genes that play a role in experience-mediated changes of internal state. The protocol is composed of two discrete parts: exposing the flies to rewarding and nonrewarding experiences, and assaying voluntary ethanol consumption as a measure of the motivation to obtain a drug reward. The two parts can be used independently to induce the modulation of experience as an initial step for further downstream assays or as an independent two-choice feeding assay, respectively. The protocol does not require a complicated setup and can therefore be applied in any laboratory with basic fly culture tools.

A Simple Way to Measure Alterations in Reward-seeking Behavior Using Drosophila melanogaster

We describe a protocol for inducing rewarding and nonrewarding experiences in fruit flies (Drosophila melanogaster) using voluntary ethanol consumption as a measure for changes in reward states.

We describe a protocol for measuring ethanol self-administration in fruit flies (Drosophila melanogaster) as a proxy for changes in reward states. We ...