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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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 two absolute requirements of the method: gastight seal and environmental stability. Maintaining a gastight environment is straightforward but requires care. Schlenk flasks and tubes are ideally suited as respirometry chambers. Glass construction avoids gas permeability present in many types of plastic19, which is especially critical in prolonged experiments. The standard joints allow gastight connections with the plugs containing the sensors and electronic connections. Stopcocks provide reliable seals and can be opened or closed as needed during experiments. To ensure gastight seals for each experiment, stopcocks were inspected, joints were cleaned thoroughly, judicious amounts of clean silicone grease were applied, and joints were secured with Keck clamps.
The sensitivity of the method is limited by the stability of temperature and humidity within the chamber. Changes in either parameter will result in pressure fluctuations that interfere with the VO2 signal. Use of an environmental chamber12 or circulating water bath provides more stable temperature control. Previous work by others has demonstrated that the method can measure O2 consumption at the nmol&#183;h-1 level when temperature was maintained within &#177;0.01 &#176;C20. To stabilize humidity, pieces of cotton roll immersed in water maintained humidity at nearly 100%. Temperature and humidity were allowed to stabilize for at least 90 min before chambers were pressurized with electrolytically generated O2 and recordings begun. All experiments included one chamber that was assembled with all components but did not contain flies, in order to monitor environmental conditions.
Constantly monitoring the temperature, pressure and humidity regularly greatly simplifies troubleshooting. For example, the artificially high apparent VO2 caused by a damaged stopcock (Figure 3B) or the apparent loss of VO2 due to unstable chamber humidity (Figure 3C) were able to be detected and corrected.
The coulometric microrespirometer described here has several advantages. First, it is relatively inexpensive, with an approximate total cost of $100 per channel (comprising chamber, sensor and controller). Second, because each channel acquires and transmits data independently, the system is scalable, with the number of chambers only limited by the size of the environmental controller (water bath or environmental chamber) and the number of available USB ports in the computer (generally expandable to &#62;100). Also, because the chambers and controllers are independent, failure of any of them will not affect the others. Third, controllers can be easily reprogrammed to adapt to ambient pressure or to adjust sensitivity as needed. Related to this, the pressure inside the chambers is set to a fixed value, so experimental pressure will be constant across experiments regardless of ambient barometric pressure. Fourth, the steady stream of data regarding time, temperature, pressure, humidity, and current allows detailed analysis of these parameters for each chamber, facilitating troubleshooting or analysis of temporal changes in VO2 in longer experiments. Finally, the environmental conditions inside the chamber can be maintained at a constant level for many hours or days, because consumed O2 is continually replaced. In the present study, chamber pressure fluctuated from 1016 to 1017 hPa, which is less than 0.1%, with the resulting variation in PO2 being less than 0.5%. Based on the amount of CuSO4 in each O2 generator, which can generate 28 mL of O2, and the average VO2 of groups of flies in this study, 1.15 mL/day (mean = 21.8 flies per chamber), metabolic rate can be studied for up to 24 days at a time. Assuming the flies are provided with adequate nutrition, this means that metabolism can be studied continuously for most of a fly&#39;s lifespan.
At present, most studies of metabolism in D. melanogaster are based on gas analysis or manometry. Methods that rely on gas analysis, such as stop-flow and continuous flow respirometers2,3,6 have the advantage that the method is well-established and equipment is available commercially. However, they require expensive and complex equipment, including manifolds and molecular flow sensors to regulate flow, and gas analyzers to measure gas concentrations. Alternatively simple manometers9 are far less expensive. In the simplest form, commonly in use for D. melanogaster, flies are anesthetized and placed in a small chamber which contains CO2 absorbent material and which is connected to a fluid reservoir by a capillary tube. As O2 is consumed, and the resulting CO2 absorbed, pressure drops in the chamber, which draws fluid into the capillary tube. The height of the fluid is then proportional to the amount of O2 consumed. Because consumed O2 is not replaced during the experiment, PO2 decreases continuously over the course of the experiment. Although routine experiments may not reduce PO2 sufficiently to negatively affect VO2, prolonged experiments, or experiments at elevated temperatures could deplete O2 and reach the critical PO2 at which VO2 declines sharply3. Another issue is that the weight of the fluid in the capillary exerts a downward force, reducing the height of the fluid in proportion to the downward force exerted, resulting in potential errors. Methods to compensate for this effect have been described4,21, but are cumbersome and are not in widespread use. Direct comparison of gas analysis and manometry indicated that the two methods produced significantly different results, but the authors were unable to produce a satisfactory explanation for the dicrepancy22.
Despite its advantages, the coulometric microrespirometer also has limitations. First, unlike some stop-flow gas analysis systems, it is not commercially available. The components are easy to acquire and neither the design nor the programming are complicated, but it is potentially more complex to construct than the simple manometric systems9. Second, acquisition of accurate data must include multiple cycles of O2 generation, so each experiment must last for several hours. This is also true of experiments using manometry. Finally, again like manometry, temperature and humidity inside the chambers must be stable. Nonetheless, the coulometric microrespirometer offers a middle ground with regard to cost and complexity, is robust and reliable once assembled, and measures O2 consumption precisely.
The experiments with CASK&#916;18 mutants&#160;demonstrate both negative (mass-specific VO2) and positive results (fly-specific VO2). If the &#62;50% reduction in walking of CASK&#916;18 mutants resulted from compromised metabolism, it might be predicted that VO2 would be reduced by a similar amount. Yet the change in mass-specific O2 consumption was modest (9.6 % reduction in median VO2), and not statistically significant. This negative result is consistent with CASK&#916;18 mutants having relatively normal metabolism, with the behavioral phenotype resulting from decreased locomotory drive. Although it is possible that the method lacks sufficient sensitivity, the reduction of VO2 caused by a relatively small difference in size (15.5% reduction in median mass) was highly significant.
Walking is associated with significantly increased energy expenditure5,23, so why is VO2 relatively normal in CASK mutants? It is possible that that excessive grooming in CASK mutants16 could counterbalance the reduction in walking, or the walking phenotype is less apparent when flies are tested in groups in small tubes, but these hypotheses await experimental verification. Nonetheless, we conclude that mutations in the CASK locus do not have strong, direct effects on metabolism, and that the coulometric microrespirometer is an effective tool for studying metabolism in D. melanogaster.
Because this apparatus is easily constructed from common components, measures O2 consumption precisely, is highly portable, can be used in any environment with stable temperature, and has been used with organisms as small as 0.5 mg flies (the present study) and as large as 500 mg scorpions (DJS unpublished), it adds a versatile tool for studying the metabolism of diverse organisms.

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><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>O&lt;sub&gt;2&lt;/sub&gt; generator</td></tr><tr><td>2-Pin Connector</td><td>Zyamy</td><td>40PIN-RFB10</td><td>O&lt;sub&gt;2&lt;/sub&gt; 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&amp;nbsp;</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>O&lt;sub&gt;2&lt;/sub&gt; 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&lt;br/&gt; 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 O&lt;sub&gt;2&lt;/sub&gt; 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>O&lt;sub&gt;2&lt;/sub&gt; 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>CO&lt;sub&gt;2&lt;/sub&gt; 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 average amplitude of each pulse is 50.1 mA, the duration is 16.1 s, yielding a charge transfer of about 0.81 coulombs (C) per pulse. The total charge transfer for this chamber was 3.28 C over a total time of 240.0 min. Using the calculations described in Procedures with the mass and number of flies (23 flies weighing 14.9 mg total), O2 consumption for the group in this chamber was 3.19 mL h-1 g-1 or 2.07 &#181;L h-1 fly-1.
The equipment can be set up easily, with a minimum of training, and performs reliably for many cycles of assembly and shutdown. Nonetheless, equipment must be maintained and inspected regularly, and experimental conditions must be controlled carefully. For example, the loss of a gastight seal, due to failure of a joint or stopcock, can lead to rapid pressurization cycles and spuriously high VO2 (Figure 3B). Additionally, temperature and humidity must remain stable inside the chamber. If temperature or humidity decreases, the resulting pressure drop will be interpreted erroneously as a O2 being consumed. Conversely, upward drift in temperature or humidity will counteract the pressure decrease caused by O2 consumption, and artificially reduce or eliminate the VO2 signal (Figure 3C).
The method was used to test VO2 of CASK&#916;18 mutants, which were generated by imprecise excision of a transposable element from the CASK locus14, and in which locomotion is drastically reduced14,16. In wild-type w(ex33) controls, generated by precise excision of the transposable element, average mass-specific O2 consumption was 3.65 &#177; 0.24 mL&#183;g-1&#183;h-1 (n = 16 chambers; Figure 4A).
Despite their visibly reduced locomotion, CASK&#916;18 mutants&#39; VO2 was slightly but not significantly lower than that of controls (mean &#177; s.e.m.= 3.23 &#177; 0.13 mL&#183;g-1&#183;h-1; n = 11 groups; P = 0.08 Mann-Whitney u-test).
Because the validity of expressing metabolic rate in terms of body mass has been questioned18, O2 consumption was also analyzed on a per-fly basis (Figure 4B). Using this analysis, VO2 was significantly reduced in CASK&#916;18 compared to wild-type controls (ex33: 2.22 &#177; 0.13 &#181;L&#183;fly-1&#183;h-1; CASK&#916;18: 1.58 &#177; 0.10 &#181;L&#183;fly-1&#183;h-1; P = 0.0003, Mann-Whitney u-test). However, the mean mass of CASK&#916;18 flies was &#62;20% lower than that of ex33 controls (Figure 4C; ex33 0.61 &#177; 0.01 mg; CASK&#916;18&#160;0.51&#177; 0.02 mg; P = 0.0005, Mann-Whitney u-test), so the difference in metabolic rate between genotypes is probably due to the difference in their sizes.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65379/65379fig01.jpg" /> 
Figure 1: Respirometer setup. (A) Diagram of sensor plug (above) and 50 mL chamber (consisting of a 50 mL Schlenk tube, below) before assembly. Note the locations of the 19/22 ground glass joints that will connect the chamber and sensor plug, and that must be cleaned before each experiment. The stopcock, which is necessary for opening or sealing the chamber, is also indicated. (B) Diagram of chamber and components, assembled and ready for the experiment, showing: wet cotton roll, polypropylene tube containing flies, plugged with a cotton stopper, soda-lime cartridge, and O2 generator filled with CuSO4. (C) Photograph of the assembled chamber. The Keck clamp securing the plug to the chamber is partially obscured by the ring stand clamp holding the chamber. (D) Photograph of controller showing switch controlling current through O2 generator and window for viewing OLED display. (E) Assembled chambers in a water bath. Seven chambers are shown, with three containing mutants, three with wildtype controls, and one chamber containing all components except flies. <a href="https://www.jove.com/files/ftp_upload/65379/65379fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65379/65379fig02.jpg" /> 
Figure 2: Data acquisition setup. (A) PuTTY interface for selecting the serial port for data acquisition. COM3 has been selected, with a BAUD rate of 9600 to match the output of the controller. (B) PuTTY interface for setting up log file. &#34;Printable output&#34; is selected to enable logging of data to a text file, data folder is selected using the &#34;Browse&#34; button, and a filename is created. (C) PuTTY log file during an experiment. Data are acquired approximately twice per second, and each line contains the following comma-delimited information: Sensor Number, time (ms) since the beginning of acquisition, chamber temperature (&#176;C), chamber pressure (hPa), Humidity (percent relative), and current (mA). (D) Data logging during a typical experiment, with seven windows for experimental chambers, plus one channel recording bath temperature, ambient air temperature, pressure, and humidity. <a href="https://www.jove.com/files/ftp_upload/65379/65379fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65379/65379fig03.jpg" /> 
Figure 3: Data from microrespirometer. (A) Pressure (grey line, left axis) and current across the O2 generator (black line, right axis) in a single respirometer chamber containing 23 w(ex33) flies. At the beginning, a long current pulse is required to pressurize the chamber from ~992 hPa to the OFF threshold of 1017 hPa. As the flies consumed O2, pressure dropped until it reached the ON threshold of 1016 hPa, which activated current through the O2 generator, which re-pressurized the chamber to 1017 hPa. The process was repeated six times in this experiment. (B) An example of a leaky chamber caused by a damaged stopcock, taken from a different series of experiments. The chamber failed to maintain pressure (grey line), resulting in constant cycling of electrolytic current (black line). Note different timescale from panel A. (C) Effect of drift in humidity. O2 consumption by the 20 mg lady beetle (Hippodamia convergens) in the chamber should have produced a cycling pattern of pressure similar to Figure 3A, but the steady increase in humidity (black line) caused an artifactual increase in chamber pressure (grey line) that masked VO2. <a href="https://www.jove.com/files/ftp_upload/65379/65379fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65379/65379fig04.jpg" /> 
Figure 4: Quantitative data from wild-type and CASK mutant D. melanogaster. (A) Mass-specific VO2 for wild-type (w(ex33)) and mutant (CASK&#916;18) flies. In all plots, bottoms and tops of boxes indicate first and third quartiles, respectively, whiskers indicate extreme values, and the sample sizes (numbers of chambers, each containing 17-24 flies) are given in parentheses above the genotypes.&#160;CASK&#916;18&#160;flies are not statistically significantly different from ex33 (median: ex33: 3.420 mL&#183;g-1&#183;h-1, CASK&#916;18: 3.029 mL&#183;g-1&#183;h-1; p = 0.08 Mann-Whitney u-test). (B) Fly-specific VO2. CASK&#916;18 flies consumed significantly less O2 (median 1.650 mL&#183;fly-1&#183;h-1) than w(ex33) (2.078 mL&#183;fly-1&#183;h-1; p = 0.0003, Mann-Whitney u-test; significance indicated by asterisks). (C) Mass differed between CASK&#916;18 and the wildtype (median: 0.526 mg, ex33: 0.623 mg; p = 0.0005, Mann-Whitney u-test; significance indicated by asterisks). <a href="https://www.jove.com/files/ftp_upload/65379/65379fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Table 1. Survey of Drosophila respirometry data from wildtype flies at 25 &#176;C. With one exception18, studies are limited to those measuring O2 consumption. In most cases, it was necessary to estimate VO2 from graphs, and figure numbers from the original papers are provided. Although all genotypes were considered to be &#34;wildtype&#34; the sources and propagation methods varied. <a href="https://www.jove.com/files/ftp_upload/65379/Table 1_rev.xlsx" target="_blank">Please click here to download this Table.</a>

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

Nanyang Technological University

Research

JoVE Journal

Neuroscience

982 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 ...

Measurement of Mitochondrial Oxygen Consumption in Permeabilized Fibers of Drosophila Using Minimal Amounts of Tissue

The fruit fly, Drosophila melanogaster, represents an emerging model for the study of metabolism. Indeed, drosophila have structures homologous to human organs, possess highly conserved metabolic pathways and have a relatively short lifespan that allows the study of different fundamental mechanisms in a short period of time. It is, however, surprising that one of the mechanisms essential for cellular metabolism, the mitochondrial respiration, has not been thoroughly investigated in this model. It is likely because the measure of the mitochondrial respiration in Drosophila usually requires a very large number of individuals and the results obtained are not highly reproducible. Here, a method allowing the precise measurement of mitochondrial oxygen consumption using minimal amounts of tissue from Drosophila is described. In this method, the thoraxes are dissected and permeabilized both mechanically with sharp forceps and chemically with saponin, allowing different compounds to cross the cell membrane and modulate the mitochondrial respiration. After permeabilization, a protocol is performed to evaluate the capacity of the different complexes of the electron transport system (ETS) to oxidize different substrates, as well as their response to an uncoupler and to several inhibitors. This method presents many advantages compared to methods using mitochondrial isolations, as it is more physiologically relevant because the mitochondria are still interacting with the other cellular components and the mitochondrial morphology is conserved. Moreover, sample preparations are faster, and the results obtained are highly reproducible. By combining the advantages of Drosophila as a model for the study of metabolism with the evaluation of mitochondrial respiration, important new insights can be unveiled, especially when the flies are experiencing different environmental or pathophysiological conditions.

In this paper, a method to measure oxygen consumption using high-resolution respirometry in permeabilized thoraxes of Drosophila is described. This technique requires a minimal amount of tissue compared to the classic mitochondrial isolation technique and the results obtained are more physiologically relevant.

The fruit fly, Drosophila melanogaster, represents an emerging model for the study of metabolism. Indeed, drosophila have structures homologous to human ...

Biochemistry

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 ...

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

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

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

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

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

Behavior

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 ...

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

This protocol describes a method for measuring the metabolism in Drosophila melanogaster larval and adult brains. Quantifying metabolism in whole organs provides a tissue-level understanding of energy utilization that cannot be captured when analyzing primary cells and cell lines. While this analysis is ex vivo, it allows for the measurement from a number of specialized cells working together to perform a function in one tissue and more closely models the in vivo organ. Metabolic reprogramming has been observed in many neurological diseases, including neoplasia, and neurodegenerative diseases. This protocol was developed to assist the D. melanogaster community&#39;s investigation of metabolism in neurological disease models using a commercially available metabolic analyzer. Measuring metabolism of whole brains in the metabolic analyzer is challenging due to the geometry of the brain. This analyzer requires samples to remain at the bottom of a 96-well plate. Cell samples and tissue punches can adhere to the surface of the cell plate or utilize spheroid plates, respectively. However, the spherical, three-dimensional shape of D. melanogaster brains prevents the tissue from adhering to the plate. This protocol requires a specially designed and manufactured micro-tissue restraint that circumvents this problem by preventing any movement of the brain while still allowing metabolic measurements from the analyzer&#39;s two solid-state sensor probes. Oxygen consumption and extracellular acidification rates are reproducible and sensitive to a treatment with metabolic inhibitors. With a minor optimization, this protocol can be adapted for use with any whole tissue and/or model system, provided that the sample size does not exceed the chamber generated by the restraint. While basal metabolic measurements and an analysis after a treatment with mitochondrial inhibitors are described within this protocol, countless experimental conditions, such as energy source preference and rearing environment, could be interrogated.

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

We present a protocol for measuring oxygen consumption and extracellular acidification in Drosophila melanogaster larval and adult brains. A metabolic analyzer is utilized with an adapted and optimized protocol. Micro-tissue restraints are a critical component of this protocol and were designed and created specifically for their use in this analysis.

This protocol describes a method for measuring the metabolism in Drosophila melanogaster larval and adult brains. Quantifying metabolism in whole organs ...

Measurement of Metabolic Rate in Drosophila using Respirometry

Metabolic disorders are a frequent problem affecting human health. Therefore, understanding the mechanisms that regulate metabolism is a crucial scientific task. Many disease causing genes in humans have a fly homologue, making Drosophila a good model to study signaling pathways involved in the development of different disorders. Additionally, the tractability of Drosophila simplifies genetic screens to aid in identifying novel therapeutic targets that may regulate metabolism. In order to perform such a screen a simple and fast method to identify changes in the metabolic state of flies is necessary. In general, carbon dioxide production is a good indicator of substrate oxidation and energy expenditure providing information about metabolic state. In this protocol we introduce a simple method to measure CO2 output from flies. This technique can potentially aid in the identification of genetic perturbations affecting metabolic rate.

Measurement of Metabolic Rate in Drosophila using Respirometry

Metabolic disorders are among one of the most common diseases in humans. The genetically tractable model organism D. melanogaster can be used to identify novel genes that regulate metabolism. This paper describes a relatively simple method which allows studying the metabolic rate in flies by measuring their CO2 production.

Metabolic disorders are a frequent problem affecting human health. Therefore, understanding the mechanisms that regulate metabolism is a crucial ...

Biology

Measurement of Carbon Dioxide Production from Radiolabeled Substrates in Drosophila melanogaster

The power of Drosophila genetics is increasingly being applied to questions of hormone signaling and metabolism and to the development of models of human disease in this organism. Sensitive methods for measurements of parameters such as metabolic rates are needed to drive the understanding of physiology and disease in small animals such as the fruit fly. The method described here assesses fuel oxidation in small numbers of adult flies fed food containing trace amounts of 14C-labeled substrates such as glucose or fatty acid. After the feeding period and any additional experimental manipulations, flies are transferred to short tubes capped with mesh, which are then placed in glass vials containing KOH-saturated filter paper that traps exhaled, radiolabeled CO2 generated from oxidation of radiolabeled substrates as potassium bicarbonate, KHCO3. This radiolabeled bicarbonate is measured by scintillation counting. This is a quantitative, reproducible, and simple approach for the study of fuel oxidation. The use of radiolabeled glucose, fatty acids, or amino acids allows determination of the contribution of these different fuel sources to energy metabolism under different conditions such as feeding and fasting and in different genetic backgrounds. This complements other approaches used to measure in vivo energy metabolism and should further the understanding of metabolic regulation.

Measurement of Carbon Dioxide Production from Radiolabeled Substrates in Drosophila melanogaster

This paper describes a method for the measurement of fuel oxidation in Drosophila melanogaster in which trace amounts of specific radiolabeled metabolic substrates are fed to flies. The exhaled radiolabeled CO2 that is a produced from fuel oxidation is collected and measured.

The power of Drosophila genetics is increasingly being applied to questions of hormone signaling and metabolism and to the development of models of human ...

Measuring O₂ Consumption in Drosophila melanogaster Using Coulometric Microrespirometry

Summary

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Chapters in this video

Magnetic Resonance Spectroscopy of live Drosophila melanogaster using Magic Angle Spinning

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Measurement of Mitochondrial Oxygen Consumption in Permeabilized Fibers of Drosophila Using Minimal Amounts of Tissue

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

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

Determination of the Spontaneous Locomotor Activity in Drosophila melanogaster

Straightforward Assay for Quantification of Social Avoidance in Drosophila melanogaster

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

Measurement of Metabolic Rate in Drosophila using Respirometry

Measurement of Carbon Dioxide Production from Radiolabeled Substrates in Drosophila melanogaster