Retrograde Labeling of Drosophila Motor Neurons Using Fluorescent Lipophilic Dyes

Begin with a dissected Drosophila embryo fixed on a glass slide with tape around it.

The fly’s motor neurons are fluorescently labeled for visualization. 

Using a microscope, visualize the embryo and position a green-fluorescent lipophilic dye-filled micropipette near the embryo. 

Then, switch to fluorescence mode and locate the motor neuron's axonal tip at the neuromuscular junction, where the motor neuron connects to the muscle. 

Switch to brightfield mode, contact the target axon’s tip with the pipette, release a dye drop, and then incubate.

Due to their lipophilic nature and axonal vicinity, the dyes selectively enter the targeted axon’s lipid-rich membrane.

Over time, the dye travels retrogradely, moving backward along the axonal lipid bilayer toward the neuronal body, labeling the neuronal projections. 

Remove the tape from the slide and place a coverslip. 

Observe the slide under a confocal microscope. 

The retrograde-labeled target neuron appears in green fluorescence, displaying detailed neuronal morphology and projections.

Use a glass needle under a dissection microscope to drag the embryo out from the vitelline membrane from the tape onto the glass, taking care not to damage the interior tissues of the embryo. Then cut through the midline of a single embryo at its surface from its posterior to anterior end. Flip the epithelial tissues from the center and attach the epidermal edge onto the surface of the glass slide.

Use a tube connected needle with the tip opening of about 300 micrometers to aspirate or blow air to remove the dorsal longitudinal tracheal trunks, as well as any remaining guts. Use 4% PFA and PBS to fix the embryos for 5 minutes at room temperature on an orbital shaker. Then wash them three times with PBS.

Stain the embryos with 1 microliter of anti-horseradish peroxidase antibody conjugated with cyanine 3 dye and 200 microliters of PBS for one hour on the orbital shaker. And repeat the washes in PBS. To fill the injection micropipette, place it into the capillary holder. Next, place the dye slide onto stage and use the micromanipulator to position it over the slide.

Then adjust the stage to place the micropipette onto the dye. Collect the dye and the micropipette by setting the injection pressure to between 200 and 500 hectopascal, the injection time between 0.1 and 0.5 seconds, and the compensation pressure to 0 for 5 minutes. Once the dye has been collected, remove the dye slide and place the sample onto the microscope stage.

Then increase the compensation pressure to a range of 30 to 60 hectopascal and lower the micropipette into the sample. Use the 10 times objective lens to locate the embryo and align the micropipette with the embryo. Change the objective lens to a water immersion 40 times lens, and submerge the lens into the PBS.

Use fluorescence microscopy to check the neuronal morphology marked by anti-HRP Cy3 and determine the injection site. When the embryo is in focus, change the position of the micropipette to make gentle contact with the tip of the axon of interest. Then use bright field microscopy during the injection to see the dye droplet. Drop the dye in a right abdominal hemisegment at the neuromuscular junction of ACC or RP3 with either DiD or DiO.

Use the hand control to release the dye and remove the micropipette. Then move on to the next injection site. Incubate the sample for 1 hour at room temperature on an orbital shaker prior to imaging. Using fine forceps, remove the tapes from the glass slide.

To mount the sample, prepare a coverslip with a small amount of vacuum grease at the four corners. Carefully place it on the sample, making sure to avoid air bubbles. Push down the coverslip to adjust the space from the sample for allowing working distance between the objective lens and the slide.

Remove any excess PBS with task wipes. Completely seal the edges of the coverslip with nail polish. Image at 10 times and 100 times magnification with a confocal microscope and process the images with ImageJ software.