Retrograde Tracing of Drosophila Embryonic Motor Neurons Using Lipophilic Fluorescent Dyes

The tracing technique described in this protocol helps us study neuronal morphogenesis and snap the connectivity in the motor neuron system of Drosophila. The lipophilic dye can rapidly diffuse to every part of the cellular membrane with an extremely high density. This is why our protocol efficiently resolves to find structures of a labeled neuron.

If an axon terminal is accessible with an injection micropipette, this technique could be applied to any neuron in the larval and adult stages of flies or in other organisms. If you have no experience dissecting or using a microinjector precise the operation on the specimen, can be challenging. Understanding the anatomy of the embryo, will help with guidance to proper portioning of tools to carry out dissection or a dinjection.

To start, prepare and layout all embryo collecting materials. Prepare the filtration apparatus by severing a 50 milliliter tube, and cutting open a hole in the cap. Then set a mesh filter with 100 micrometer pores in between the tube and the cap.

Prepare the dye injection micropipette and dissection needle from the same capillary tubing with an inner diameter of 1.2 millimeters. Pull the tubing with a micropipette, puller at 7%from 170 volt maximum output to create a sharp needle with a taper of about 0.4 centimeters in length. To prepare the dye injection micropipette, soak the grinder with a wetting agent and place the needle on the micropipette clamp at 25 to 30 degrees.

Then lower the tip onto two thirds of the radius out from the center of the beveling surface. Grind the needle while a syringe with tubing pushes air into it. Mark the micropipette with a fine tip permanent marker to indicate the position of the opening at the tip after beveling because it can be challenging to locate the narrow opening that is formed at an angle.

Allow the flies to lay eggs overnight at room temperature and collect the pipe with the eggs in the morning. To collect the embryos, dechlorinate the eggs on the plate with 50%bleach for five minutes. Once the chlorions have cleared, pour the contents of the plate through the filtration apparatus or cell strainer to isolate the embryos.

Use a squeeze bottle of water to dilute the bleach left on the plate and gather as many embryos as possible by decanting the mixture into the filter. Wash the embryos three to four times with water until the bleach odor dissipates. Remove the filter from the apparatus and wash them onto another clean plate with water.

Then decant the water from the new plate that the embryos are on. Use fine forceps to select five to 10 embryos at 15 hours after egg laying and place them on double sided tape with the dorsal side facing up. Add insect ringer saline to the dissection pool to protect the embryos from desiccation.

Use a glass needle under a dissection microscope to drag the embryo out from the middling membrane from the tape onto the glass, taking care not to damage the interior tissues of the embryo. Then cut through the mid line of a single embryo at it's 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 a 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 five minutes at room temperature on an orbital shaker. Then wash them three times with PBS.

Stain the embryos with one microliter of anti horseradish peroxidase antibody conjugated with Cyanine3 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 the 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 in 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 zero for five 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 the 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 one 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 cover slip with a small amount of vacuum grease at the four corners. Carefully placed it on the sample, making sure to avoid air bubbles.

Push down the cover slip 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 cover slip with nail polish.

Image at 10 times and 100 times magnification with a confocal microscope and process the images, with the imageJ software. This protocol was used to retrograde label the aCC motor neuron innervating muscle one and the RP3 motor neuron innervating muscles six and seven. Dendritic branches from the ACC and RP3 motor neurons show extensive overlap.

Both neurons are bipolar, establishing two different populations of dendrites. To demonstrate the quantitative capabilities of this method, the total number of dendrite tips was counted in wild type and Dscam1 mutants. The ipsilateral dendrites of aCC are shown for the wild type, but few ipsilateral dendrites were observed for the Dscam1 mutant.

When attempting this procedure, it is important to remember that the size of the dye droplet is crucial, which is approximately 10 to 20 micrometers. Test the size on the double sided tape and then apply it to the injection site. Using this procedure, we can take advantage of the photo switchable property of DiD probe to obtain super resolution images of the plasma membrane.

This way we can study the ultra structural characterization of the synaptic membranes at the neuromuscular junction. We did the click. We performed phenotypic analysis of the aCC dendrites in mutant strain, and discover that a Dscam1-Dock-Pak injection defines the site of the dendrite outgrowth in aCC.