Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

This video demonstrates how to perform single syn recordings in Drosophila, melanogaster, and an awfully gambier prepared odorants are pipetted onto paper strips and are inserted into pasture pipettes. Live insects are mounted to immobilize the olfactory organs. The insect is placed under a microscope and a reference electrode is inserted into the insect's eye.

The recording electrode is then inserted into an olfactory centum. An odor is puffed from the pipette to the centum and neuronal activity is recorded. Hi, I am Leslie Vasal from the Sal Laboratory at Rockefeller University, And I'm Mariza Pellegrino, also from the Vasal lab.

And I'm from the, Today we will show you a procedure to record from olfactory sensory neurons of the vinegar fly esophagal melanogaster, and the malaria mosquito and nais gambi. We use this procedure in our laboratory to study how the sensor smell works in insects and how neurons respond to volatile chemicals. So let's get started.

Begin this procedure by sharpening the electrodes to be used for the recordings. Using a drill, make a small hole about two millimeters in diameter on the wall of a 20 liter syringe, one centimeter from the tip. Fill the syringe with filtered 0.5 molar potassium hydroxide and clamp it horizontally on a stand under a dissecting microscope with the tip in field of view, push an electric wire into the hole on the syringe, ensuring that it does not block the syringe tip and connect it to the anode of a power supply.

On the right side of the microscope, attach an electrode holder shaft to a manual micro manipulator. Then using a crocodile clip, attach an electric cable at the base of the electrode holder shaft. Connect the cable to the cathode of the power supply.

Insert a five centimeter piece of tungsten wire into the electrode holder and screw it to the shaft. On the manipulator. Set the power supply to six folds.

While viewing under the dissecting microscope, insert an electrode into the tip of the syringe such that 90%of the electrodes length is in the solution. After one minute, slowly withdraw the electrode, then insert the electrode again. This time, only 50%of the electrode should be in contact with the solution.

After 30 seconds, withdraw the electrode again. Repeat this process 10 times each time inserting the needle such that 50%of the electrode is in contact with the potassium hydroxide. Once the electrode has been sharpened, it is ready to be used for experiments.

Begin the preparation of the odorant delivery system by cutting chromatography paper into three millimeter by five centimeter strips onto each strip pipette 30 microliters of the desired odorant. Then place the strip inside a glass past your pipette. Attach a piece of airline tubing about three centimeters in length to the pipette.

Then use a connector to close the tubing and place the prepared odorants aside. Next, using a razor blade cut a 10 milliliter plastic serological pipette at the four milliliter mark from the top, discard the pointed end. Then using a small drill, create two holes, one at the minus 1.5 milliliter mark, and one at the minus 0.5 milliliter mark.

Next, insert a 200 microliter pipette tip into a barbed coupler, and push the coupler into the cut end of the 10 milliliter pipette. Clamp the pipette to a magnetic stand and position it near the microscope. Then connect the serological pipette to an air pump using airline tubing and place the pipettes containing the odorants next to the serological pipette ready to use.

Next, screw the electrodes in the electrode holder and the probe connected to the manipulators on the microscope setup. Place a piece of cover glass on top of dental wax at a 30 degree angle, ensuring that the dental wax does not cover the upper part of the glass. Then place the piece of dental wax in the middle portion of a microscope.

Slide now place an additional smaller piece of wax on the slide as shown here on top of it. Place a glass capillary pulled with a vertical puller. Construct a flight aspirator by inserting a cut 200 microliter pipette tip into one end of a 120 centimeter piece of airline tubing.

Cover the other end of the tubing with a 1.5 centimeter by 1.5 centimeter piece of mesh and secure it into place by capping it with a cut. One milliliter pipette tip. Now use the fly aspirator to pick up an adult fly from a bottle.

Trap the fly by capping the tip of the fly aspirator with a 200 microliter tip. Then blow into the tube to push the fly towards the end of the 200 microliter tip. Using a razor blade trim, the wide end of the pipette tip a few millimeters away from the fly.

Quickly insert dental wax to trap the fly under a dissecting microscope. Trim the pipette tip near the head of the fly, taking care not to injure it with a small pipette tip. Push the wax until about half of the fly's eyes extrude from the tip.

Ensure that the legs do not come out as they may interfere with the recordings. Mount the fly on a piece of wax, its head facing upwards. Then place the wax on the slide adjacent to the cover glass as shown here.

Nudge the wax slightly so that the antenna extend and rest on the glass. Fix the antenna by positioning the glass capillary in the groove between the second and third antenna segments. Once the fly has been positioned on the slide, transfer it to the microscope used with the recording and odorant setups at low magnification.

Position the antenna roughly in the middle of the field of view. Gently lower the reference electrode and position it adjacent to the eye. Then lower the recording electrode and position it adjacent to the antenna.

Increase the magnification and reposition the antenna in the middle of the field of view. Place the odor delivery device close to the head of the fly pointing at the antenna. Switch back to low magnification using a micro manipulator.

Insert the reference electrode into the eye of the fly. Lower the recording electrode over the antenna without touching its surface. Then switch back to high magnification ification using the micro manipulator.

Insert the recording electrode into a syn at any point along its length when the electrode has been properly placed, spontaneous activity in the cells of the sens can be detected. If needed, the electrode can be pushed farther in or even all the way through the sens to obtain a better signal to noise ratio. To record responses to odorants, connect the pasture pipette to the air pump using airline tubing and puff the odorant in this case, methyl acetate by activating the air pump for about one second at about 220 milliliters per minute.

Using an electric aspirator, collect 40 to 63 to five day old mosquitoes in a small plastic cage. Place the cage on ice for about 15 minutes to anesthetize the mosquitoes. Once the mosquitoes have stopped moving, transfer four to six mosquitoes to the dissecting microscope stage.

Both male and female. Mosquitoes can be used for recording. In males.

The antennae appear feather like While in females, they appear filament like using fine forceps, remove the wings and legs of the mosquitoes to immobilize them. Once the wings have been removed, place the mosquitoes in a humidified plastic cup to prevent them from Desiccating. Place two one centimeter long pieces of double-sided tape parallel to each other and approximately one centimeter apart at the center and at the side of a glass slide under the dissecting microscope, use forceps to place one mosquito on its side.

On the central tape. One eye will stick to the tape. Adjust the position of the maxillary pulps so that both extend parallel on the tape.

Fix the maxillary pulps by placing thin strings such as human hairs at the base and at the tip of the pulps. Once the mosquito has been positioned on the slide, transfer it to the recording setup. Under low magnification position, the maxillary pulps roughly in the middle of the field of view and the head at the top rotate the stage until one of the pulps is at a right angle.

With the recording electrode adjust to the height of the electrodes so that the reference electrode is positioned just above the eye of the mosquito. And the recording electrode is near the maxillary pulps. Place the odor delivery device so that it is as close as possible to the pulps.

Insert the reference electrode into the eye. Switch to the 100 x objective and insert the recording electrode into the peg on the palp. Once the electrode is in the sentum, the spontaneous activity of the cells can be detected.

As with the drosophila record, odorant induced responses by puffing the odorant in this case, one OCT three all by connecting the posterior pipette containing the odor to the air pump, and then activating the air pump. Now we'll show some representative recordings from both drosophila and oph olfactory sensory neurons. Depending on the sun and the quality of the recording, one can distinguish different numbers of olfactory neurons within a single trace in the large base iconic silla of drosophila melanogaster, for example, between two and four cells, that difference spike amplitude should appear during the recording.

This figure shows recordings made from two drosophila AB two lum cells. Recordings from the A cell are shown in blue, and recordings from the B cell are shown in green, while neither cell is activated during application of paraffin oil. As shown in the figure on the left only the A cell response to methyl acetate in the maxillary pal of OPIS gambier, the grooved pegs and soum contains three cells, but only two are easily discriminated.Here.

The recordings shown in green indicate the B cell response to the odorant one, OCT three. All We've just shown you how to record from olfactory sensory neurons of the vinegar fly drosophila melanogaster, and the malaria mosquito and novelly gambi. When doing this procedure, it's important to remember to have a stable isset preparation and a fine tanks and electrode to successfully perform recordings.

So that's it. Thanks for watching and good luck with your experiments.