using a respiratory chamber to measure metabolic rate in drosophila melanogaster

Adapting Microrespirometry for &lt;em&gt;Drosophila&lt;/em&gt; - Insights into CASK Mutant Metabolism

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.

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

Measuring O2 Consumption in Drosophila melanogaster Using Coulometric Microrespirometry

Our research is focused on how temperature and activity affect insect metabolism. This is essential to understand how species have evolved to survive in their surroundings, and how they will continue to adapt in a changing world. Understanding the physiology and behavior of individual organisms is crucial to this endeavor.Metabolic rate can be measured directly by measuring heat production, or indirectly by measuring the consumption of oxygen or the production of carbon dioxide. Although it is technically possible to measure heat production by an organism as small as a fruit fly, indirect methods are used more commonly. The current challenges to measuring metabolism on this scale will include cost, environmental stability, and reliability.Commercially available systems are reliable, stable, and relatively easy to purchase, but they can be complex and expensive. DIY manometers are inexpensive, but can introduce error and require fabrication for each experiment. The coulometric respirometer is relatively inexpensive, easy to use, provides a convenient readout of environmental parameters, and all components are reusable.In addition, the regular replacement of oxygen maintains a stable environment for hours or even days. In the lab, we'll be studying the metabolic variation between established control strains of drosophila, and the effects of hyper and hypoactive mutants on oxygen consumption. In the field, we'll begin studying the metabolic rates and temperature tolerance of wild, desert-adapted beetles.

Watch this Scientific Journal Video about Using a Respiratory Chamber to Measure Metabolic Rate in Drosophila melanogaster at JoVE.com

Using a Respiratory Chamber to Measure Metabolic Rate in Drosophila melanogaster  

Using a Respiratory Chamber to Measure Metabolic Rate in Drosophila melanogaster

Begin by maintaining the flies at a temperature of 25 degrees Celsius, in narrow vials filled with standard drosophila food. Turn on the water bath and adjust the temperature as required. Use a laboratory wipe sprayed with 70%ethanol to clean the ground glass joints of the chambers and sensor plugs, and then use a fresh wipe to remove any remaining ethanol.Next, place a one centimeter piece of a cotton roll, soaked in purified water, at the bottom of the chamber to create a stable, humidity environment. Add sufficient water to form a modest pool at the base of the cotton roll. Ensure the joint of the chamber is dry by wiping any spilled water.Carefully transfer the flies into labeled polypropylene tubes using a funnel, and plug the tubes with a cotton roll. Now, add a ventilated tube containing flies into each respirometer chamber. Fill the soda lime cartridges and position them at the top of the tube that contains flies inside the chamber.Then fill the oxygen generators with a saturated solution of copper sulfate, ensuring that the level remains below the vent holes. Connect the filled oxygen generator to the two-pin connector located on the sensor plug. Place two small dabs of clear silicone grease on opposite sides of the ground glass joint of the sensor plug.Insert the plug into the chamber and gently rotate it with moderate pressure to evenly distribute the grease within the joint. Wipe off any excess grease using a laboratory wipe. Then secure the chamber plugs by snapping on the plastic keck clamps.Place the assembled chambers into a rack in the water bath, ensuring the stopcocks are open. Maintain stopcocks in the open position and let the chambers equilibrate for approximately 30 minutes. Ensure that the switches supplying current to the oxygen generators are off.Connect the controllers to the respirometer chambers using the six conductor cables, and verify that the organic light emitting diode displays on the controllers actively display environmental parameters. Now activate the oxygen generators using the switch on the controller. Once the current value rises from zero to between 35 and 55 milliamperes, the controller and chamber are ready for the experiments.After turning off the oxygen generators, identify the communication ports that the controllers are using. Open the PuTTY software on the desk desktop and set up a log file for each channel of the respirometer by selecting the communication port for the controller by entering the number of that port in the serial line box. Then click logging.Choose principal output in the session logging and under the log file name, click browse. In the chosen folder, create a file name that contains descriptive information. Then click save.Click open, and a window showing common delimited data being logged will open. Repeat this process for all other controllers being used. Each communication port input will be displayed in a separate window.After equilibrating the chambers for 30 minutes, seal them by closing the stopcocks. Cover the bath and the chambers with a polystyrene foam box to create a stable environment and equilibrate for an hour. Initiate the current to the oxygen generator of each chamber using the switch located on the controller box.Once the oxygen generators are activated, verify that the pressure increases to the predefined off pressure. After the experiment, turn off the oxygen generators on all controllers and open the stopcocks to unseal the chambers. Allow the PuTTY windows to remain open for five to 15 minutes to establish a final baseline, and then end the recordings by closing the PuTTY window for each controller.Disconnect the sensors from the cables and transport the chambers to the dry rack. One by one, remove the sensor plugs from the chambers. Disconnect the oxygen generators and carefully place them in the tube rack.Then wipe the grease off the sensor plug and place it back in the rack. Clean any grease from the chamber joints and remove the tubes containing flies and soda lime. Finally log the weight and the number of flies for each tube.Quantitative oxygen consumption data for wild type and cask delta 18 mutant flies showed that mass specific oxygen consumption was not significantly different among wild type controls and cask delta 18 mutants. Oxygen consumption analyzed on a per fly basis showed that the oxygen consumption was significantly reduced in cask delta 18 compared to wild type controls. However, the mean mass of cask delta 18 flies was lower than that of wild type controls.

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133 Views.  University of Maryland, College Park, Universities at Shady Grove. Begin by maintaining the flies at a temperature of 25 degrees Celsius, in narrow vials filled with standard drosophila food. Turn on the water bath and adjust the temperature as required. Use a laboratory wipe sprayed with 70%ethanol to clean the ground glass joints of the chambers and sensor plugs, and then use a fresh wipe to remove any remaining ethanol.Next, place a one centimeter piece of a cotton roll, soaked in purified water, at the bottom of the chamber to create a stable, hum...