The FlyBar: Administering Alcohol to Flies

The overall goal of this procedure is to measure alcohol induced behaviors in Drosophila, using an inexpensive and easily assembled setup well suited for undergraduate research or researchers. New to circadian studies. This is accomplished by first assembling the fly bar, allowing simultaneous comparisons of multiple groups of flies.

The next step of the procedure is to culture drosophila under specified light dark cycles and collect age-matched flies for the experiment. Then the flies are exposed to defined concentrations of alcohol vapor and the data is collected. Ultimately, measuring the loss of writing reflex or sedation can show differences in alcohol sensitivity between groups, either based on time of day or genetics.

Visual demonstration of this method is critical because the observation steps are difficult to learn due to the challenges of observing behavior and the dim red light, and determining which flights have lost their writing reflex and which ones have not. First assemble the fly bar. Start with assembling the airflow.

Connect a short piece of flexible silicone tubing to either a building air tap or an aquarium aerator to provide airflow. Split the airflow with a Y connector. Connect the first branch to a regulator.

It will control the total amount of air through the system. Connect the second branch to a quick connector. This will allow the Airstream to be interrupted as needed without affecting the calibrated airflow.

Add a Y connector and connect each branching tube to an airflow regulator. Then connect each regulator to an airflow meter. Now section one milliliter glass pipettes and melt a right angle into each section.

These will function as rigid elbow tubes in the setup. Insert the straight glass pipette for the air inlet through one hole and extend it into the fluid to one centimeter above the bottom. Insert an elbow section of glass through the other hole until it just enters the bottle.

Use these outlets to join the airflow from each bottle at a Y connector. Using flexible tubing, connect the airflow meters to the straight section of a pipette. These will serve as the air inlets into the two solution bottles.

Water or alcohol, keep both solution bottles in a water bath Set two degrees higher than ambient air temperature. Stop the bottles with rubber stoppers that have two holes with silicone tubing. Attach the other end of the Y to a bottle with a two hold rubber stopper.

The airstream from this mixing flask will feed vaporized alcohol and water to the observation vials. Split the outlet airstream from the mixing flask two or three times to obtain four or eight smaller streams of air. One for each observation vial.

Keep using silicone tubing. The observation vials are empty Fly vials sealed with a two hole rubber stopper, an inlet and outlet for the alcohol vapor. Glass tubes fit the stopper holes.

Cover the ends of the glass tubing with netting. Keep the netting in place using a small piece of flexible plastic tubing. Insert this tube through the first hole until it extends to approximately half the length of the vial.

If needed, use Teflon tape to obtain a snug fit. Insert the second glass tube with netting until it is flush with the inside edge of the rubber stopper. Mix the appropriate fractions of the two Airstreams bubbled through alcohol or water.

Monitor the air pressure continuously and make adjustments as needed to maintain the desired blend of alcohol and water vapor in the mixing bottle. For this sort of experiment, always ensure that the room is adequately ventilated. To minimize variation between replicas, utmost attention needs to be paid to the rearing conditions of the flies.

Flies need to be cultured under low crowding conditions with a very strange synchronized so that the flies of the same age and cohort can be tested against each other. Approaching the flies dusk. Collect freshly enclosed animals and store them for 24 hours in holding vials with a small amount of high agar concentration food at the same time.

On the next day, subdivide the collected flies into cohorts of approximately 30 flies using an aspirator and transfer the cohorts to fresh vials to test for circadian modulation of behavior. Prepare enough cohorts to test each condition at six evenly spaced time points on the testing day to test circadian modulation behavior. Also maintain the flies under constant darkness at 25 degrees Celsius for two days prior to testing.

Regardless of experimental design, at least one hour prior to the behavioral assay, transfer the flies to the experimental behavior room lit only by dim red light. Run the experiments under dim red light so photo stimulation does not confound the results. Before the experiment run air through the system for at least 10 minutes and calibrate the air flows.

Under the observation vials. Place a piece of white paper to increase the fly's visibility with added contrast. Once the airflow has stabilized, disconnect the quick release and load one 30 fly cohort into each observation vial.

During each run of the fly bar, test each experimental condition. This will increase the robustness of the experiment and minimize the impact of daily variations. Once all the observation vials are loaded, reconnect the airflow and start the timers.

Use one timer to keep track of the total time of alcohol exposure and use the other to count down five minute intervals throughout the trial. Monitor the airflows and make necessary adjustments so they remain constant. Generally, the airflows stabilize every five minutes using a red light lamp.

Count the number of flies that have lost their writing reflex. Keep the lamp at least 12 inches from the vials. Apply a firm tap to the vial and count how many flies fail to write themselves within approximately four seconds.

These flies may still move their legs and wings, but they will not be able to turn themselves upright. If the fly does not move any limbs, it is completely sedated. This can be scored as a different criterion from a lack of writing reflex.

The session will last about an hour at its end count the total number of flies in each file using the described protocol. A circadian modulation in alcohol sensitivity was measured. The lack of writing reflex was measured at six time points during the second day of darkness in Canton S the time of day had a significant effect on time.

Half the flies failed to write themselves. The percentage of cantons flies sedated after 40 minutes with 30%alcohol. Vapor was measured at CT five subjective day and CT 17 night.

It was found that significantly fewer flies were sedated during the subjective day compared to the night. Employing a time series illustrated differences in writing reflex and sedation when Canton s flies were compared with flies carrying the white 1 1 1 8 mutation. In the Canton S background, the white 1 1 1 8 flies showed better writing reflex in response to ethanol vapor than Canton S.Differences between white 1 1 1 8 mutants and wild type flies were also found in the rate of sedation.

The difference is most likely due to the altered levels of biogenic enemies with the white 1 1 1 8 mutation. Thus, it is necessary to control for behavioral responses due to the white 1 1 1 8 mutation present in many transgenic lines. After watching this video, you should have a good understanding of how to administer alcohol to flies using the fly barn.

It is essential that the flies are collected in a methodical manner and that the observations in the dim red light are standardized as much as possible.