Measurement of Carbon Dioxide Production from Radiolabeled Substrates in Drosophila melanogaster

The overall goal of this protocol is to measure carbon dioxide produced by intact fruit flies, set radiolabeled substrates such as palmitic acid or glucose. This method can help answer key questions for Drosophila researchers, such as whether a given mutation may affect energy metabolism. The main advantage of this technique is that it allows quantitative, sensitive, and reproducible measurements of fuel oxidation in small numbers of fruit flies.

Visual demonstration of this method is critical as the fly pod assembly steps require quick manipulation of small parts to ensure proper construction of the experimental apparatus before flies recover from anesthesia. Carbon-14 is a low energy beta emitter and has a short range in air, but care must still be taken to prevent spilling of radiolabeled molecules or accidental ingestion by the experimenter. Mix one to two microcurie radiolabeled substrate with 15 microliters of FD&C Number One blue food dye and water to obtain a total volume of 25 microliters.

Pipe head all of the radiolabeled substrate and blue dye mix into the bottom of an empty fly food vial. Since fly food vials tip over easily, be sure to house them in a container that will prevent them from tipping. Heat standard fly food in a microwave oven until the food is just liquefied.

15 to 20 seconds at a high power level is usually sufficient for one vial containing 10 milliliters of food. Add a 975 microliters of the molten food to the radiolabeled substrate and blue dye mix, and quickly swirl while monitoring the uniformity of the blue color to ensure complete mixing. Pipe head tips may need to be cut with the scissors to widen the narrow end, thereby allowing accurate pipe heading of the viscus molten food.

Allow the food to cool and solidify completely at room temperature for 20 to 30 minutes. A carbon dioxide anesthetizing apparatus consisting of porous polyethylene fused to an acrylic base and connected to a carbon dioxide tank with a pressure regulator is used to anesthetize adult flies. Flow carbon dioxide into the anesthetizing apparatus at five liters per minute.

Transfer the anesthetized flies to vials containing radiolabeled blue dyed food. Cap each vial with a foam stopper and rest it horizontally until the flies wake up. Transfer the vials to a lidded acrylic container.

For short-term feeding, starve flies for 18 to 24 hours before their transfer to radiolabeled food to ensure that they will eat during the two to three hour feeding step. For long-term feeding over the course of five to seven days, flies need not be starved, but the experimenter should monitor the water content of the radiolabeled food in the vials, and use a needle and syringe to add water to vials with dry food. When the period of radiolabeled substrate feeding is over, transfer the flies to unlabeled food by tapping them into a new vial.

An initial chase period of two to four hours allows flies to clean radiolabeled food particles from their cuticles, and permits digestion of radiolabeled food remaining in the gut. Monitor the progress of food through the gut by visual inspection of the flies to look for blue abdomens. Subsequently, transfer the flies to different food types, or ambient temperatures.

The length of this chase period will vary with the experiment being performed. Prior to making the carbon dioxide collection apparatus, assemble materials to construct fly pods, which are mesh-capped tubes that will contain flies labeled with C-14 glucose, or C-14 palmitic acid, and remain open to the atmosphere. Clamp a razor blade with a hemostat, heat it in the flame of a Bunsen burner until red hot, and use the heated blade to cut off the top 50 millimeters of 12 millimeter by 75 millimeter polypropylene tubes, leaving 25 millimeter long round bottomed tubes.

Prepare 35 millimeter by 35 millimeter squares of 130 micron nylon mesh. Cut several one and a half to two inch long pieces of transparent tape. Next, assemble materials for the carbon dioxide collection apparatus.

For each fly pod, assemble a 20 milliliter glass scintillation vial, a rubber topped stopper with an off center hole, a center well, and grade GFB glass microfiber filter paper. Insert the center well through the hole in the top stopper. Prepare 5%potassium hydroxide fresh on the day of use.

Just before anesthetizing flies for transfer to the fly pod, fold and place the circular GFB filter paper into the center well that is threaded through the hole of the rubber top stopper, and saturate it with 100 microliters of 5%potassium hydroxide. Transferring anesthetized flies into the fly pods is the most difficult step of this protocol. To ensure success, have all the needed materials at close reach, and work quickly to limit the flies'exposure to carbon dioxide, and to avoid flies waking up and escaping before the mesh has been securely affixed to the tube.

Following the incubations on unlabeled media, anesthetize the flies with carbon dioxide. Brush the anesthetized flies into the cut off polypropylene tube, cap with the nylon mesh, and use transparent tape to adhere the mesh to the tube. An equal number of flies should be used in each fly pod for a particular experiment.

10 to 20 flies per fly pod produce sufficient radiolabeled carbon dioxide for measurement by scintillation counting. Transfer the fly pod to the 20 milliliter glass scintillation vial. Cap the glass vial with the rubber topped stopper, holding a center well containing potassium hydroxide saturated GFB filter paper.

Fold the wide top of the rubber topped stopper over the lip of the glass scintillation vial. Set the glass vials containing the flies in a lidded acrylic container, and incubate for varying periods of time. At the end of the incubation, uncap the glass vial, and transfer the potassium hydroxide saturated GFB filter paper to a six milliliter plastic scintillation vial containing 4 milliliters of scintillation cocktail.

Prepare additional scintillation vials, one containing unused GFB filter paper to serve as background, and one to two other vials containing 0.1 to 0.5 microcurie radiolabeled substrate to determine counts per minute per microcurie. Using a scintillation counter, measure counts per minute for each sample according to the manufacturer's protocol. Subsequently, calculate total carbon dioxide production as described in the protocol text.

Flies were fasted for 18 hours, fed radiolabeled palmitic acid for two hours, and then tested for radiolabeled carbon dioxide production in groups of 12 or 25 animals for three or six hour incubations. As shown in this graph, the amount of carbon dioxide produced by 25 flies was nearly double the amount produced by 12 flies, and the amount of each group doubled when the incubation time was increased from three to six hours. The picomolar radiolabeled carbon dioxide per fly per hour was nearly equivalent in each group.

Since fasting stimulates the breakdown of fatty acids by beta oxidation, carbon dioxide production from fatty acid oxidation should be elevated in fasted animals compared with fed animals. To test this, 18 hour fasted flies were fed radiolabeled palmitic acid for two hours, divided into two groups, and transferred to unlabeled fly food, or 1%agar. In two separate experiments, flies chased on agar produced significantly more radiolabeled carbon dioxide than flies chased on food, indicating that beta oxidation was increased in the fasted animals.

Following this procedure, complimentary methods such as measurement of total carbon dioxide production, and total oxygen consumption, and calculation of the respiratory exchange ratio can be performed in order to answer additional questions such as what the preferred fuel source is. After watching this video, you should have a good understanding of how to use this protocol to determine the effect of a given mutation or experimental manipulation on the ability to use a specific fuel as an energy source. Don't forget that working with radioactivity can be hazardous, and precautions such as wearing gloves, lab coats, and goggles, and using proper shielding, should always be taken while performing this procedure.