Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

The overall goal of this procedure is to prepare oph and melanogaster embryos for whole mount visualization of the nervous system. This is accomplished by first collecting embryos from the fly strain of interest. The second step of the procedure is to remove the corium with bleach.

The third step of the procedure is to apply heptane and methanol to remove the vilin membrane. The final step of the procedure is to apply antibodies to the embryos. Ultimately, results can be obtained that show the structure of the drosophila embryo nervous system through immunofluorescence microscopy.

This method can help answer key questions in the field of neuroscience, such as what role mutations can play in the developing nervous system. Generally, individuals new to this method will struggle because they may accidentally pipe out embryos from the vial. Begin this protocol by preparing the reagents that will be needed, including P-E-M-F-A, buffer PBC 5%BSA with 0.01%sodium azide saline solution and formaldehyde fixative.

Prepare a cage for the drosophila strain of interest by knocking the flies out with carbon dioxide and then transferring at least 100 flies to a plastic bottle containing an adapter with an apple juice agar feeding plate. Place the cage of flies in the dark at room temperature for 72 hours. Change the apple juice agar feeding plates every eight to 12 hours so that the flies get acclimated to the cage and do not hold their eggs for the experiment.

After the 72 hour acclimation period, collect embryos 12 hours from the last change in apple juice agar feeding plate. Remove the apple juice agar feeding plate from the laying cage and replace it using saline solution. Rinse the apple juice agar feeding plate.

Collect the embryos in a fine sieve. Use a clean paint brush to dislodge any embryos that remain stuck to the plate. Rinse the embryos in the sieve with saline to remove any excess yeast using a spatula, gently pick up the embryos and place them in a small glass vial.

Add about one milliliter of saline solution. Then cap the vial and incubate for five Minutes to wash the embryos. Following the Collection of embryos.

Prepare the heptane fixed solution by combining 50%heptane and 50%fixative. The solution will be layered with heptane on top and the fixative on the bottom. After five minutes have passed, use a pasta pipette to remove the excess saline from the glass vial of embryos.

Rinse the embryos with deionized water by applying approximately one milliliter of deionized water to the glass vial containing the embryos. Allow the embryos to sit for one minute before removing the water at a 50%bleach solution. Shake the vial a few times by hand and then incubate for three minutes in order to coate the embryos.

The ated embryos will sink to the bottom of the vial. Embryonic CORs will float. Remove the bleach and the CORs after the three minute incubation.

Following the decoration, rinse the embryos thoroughly with deionized water applying approximately one milliliter of deionized water to the glass vial containing the embryos. Allow the embryos to sit for one minute before removing the water leftover CORs will float and should be removed. Add the heptane fix solution to the embryos and incubate them for 30 minutes with gentle shaking to fix them following fixation.

Use a pasta pipette to remove the fixative, which is the bottom layer, leaving the heptane in the tube. Then add methanol back to the bottom of the tube in place of the fixative and cap the tube shake vigorously by hand to remove the vitelline membrane. Following this step, the ized embryos will sink to the bottom of the tube.

Using a pasta pipette, remove the methanol and hets from the vial leaving behind the deionized embryos. Then rinse the embryos five times with methanol, removing the methanol with a pasta pipette between each rinse. For each rinse, incubate the embryos for Five minutes.

Place the embryos in a mixture of 50%PBT solution and 50%methanol for five minutes to rehydrate them. Then continue the rehydration process by transferring them to 100%PBT solution for five minutes. Next, block the embryos by incubating them in 5%BSA with 0.01%sodium azide solution for one hour at four degrees Celsius.

Next, prepare the primary antibody for embryo staining by diluting it to the appropriate concentration using 5%BSA with 0.01%Sodium azide solution. The antibodies used here are directed against the vesicular cysteine string protein and anti HRP antibody. Anti HRP recognizes a neural specific carbohydrate moiety present in the surface glycoproteins, which labels all neurons.

Incubate the embryos overnight at four degrees Celsius in the primary antibody solution. Wash the embryos 10 times with PBT over the course of an hour. By applying approximately one milliliter of PBT to the glass vial containing the embryos, allow the embryos to sit for six minutes before removing the PBT.

Repeat this process of applying PBT, incubating and removing PBT 10 times. Prepare the secondary antibody solution in the dark because it is light sensitive by diluting to the appropriate concentration. We use Alexa secondary antibodies at a concentration of one to 200.

Incubate the embryos for two hours at four degrees Celsius in the secondary antibody solution. Wrap the glass vial of embryos and secondary antibody solution in aluminum foil in order to avoid light exposure. Wash the embryos 10 times with PBT over the course of an hour.

Add vector shield mounting medium to the embryos and incubate them overnight at four degrees Celsius to mount. Use a pastel pipette to suck up the embryos along with some of the mounting medium. Apply the embryos and medium to a glass slide.

Apply a cover slip over the embryos and seal the cover slip with nail polish. Visualize embryos using a Fluorescence microscope here. Embryonic CNS is shown at stage 16 to 17.

Cystine string protein is shown in red. HRP neurons are shown in green. Note the distinct staining of the commissure in this representative image.

The embryonic PNS is shown at stage 16 to 17. Again, cystine string protein is shown in red, and neurons are shown in green. Note the repeated patterns of Peripheral neurons.

When Attempting This procedure, it's important to be mindful of the embryos so as to not accidentally remove them from the vial. After watching this video, you should have a good understanding of how to prepare oph fla embryos for whole mount visualization of the nervous system.