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's 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 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. 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 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 printable 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 comma 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.
