Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

To perform electrophysiological recordings in drosophila embryos, they must first be harvested, coordinated, and dissected controlled dissection is made possible with water polymerizing surgical glue, which stabilizes the embryos to cover slips. The internal organs are then removed from stabilized embryos to expose the ventral CNS, the peripheral nerves and the somatic musculature. Hi, I am Dave Featherstone from the Department of Biological Sciences here at the University of Illinois at Chicago.

Hi, I’m Ton from the Lab of Dave Featherstone in the Department of Biological Sciences at University of Illinois at Chicago. Today we’re going to show you how to collect and manually dissect late stage embryos from Joof. We use this procedure General lab to prepare for electro alternative testing of joof neuromuscular system development and function, which I’ll show you another upcoming Jo Ova presentation.

So let’s get started. This protocol begins with embryo collection, a common procedure used in a variety of joof experiments, which we will now review. Establish a mix of 20 to 40 young male and female flies in a laying pot.

House them in a humidified chamber at 25 degrees Celsius. To accurately estimate the developmental stage of the embryo, the temperature and humidity should be consistent. Although commercial laying pots are available, they can be constructed from perforated 100 mil trip poor plastic beakers that fit onto 60 millimeter apple auger plates.

The apple auger recipe can be found in the written protocol. Perforations in the beaker can be made with a heated probe and should be smaller than adult flies. The plates are supplemented with a bolus of yeast paste, which will stimulate the females to lay many eggs, mix yeast and water to a peanut butter consistency, and then smear on the plates with a spatula.

Change the plate twice daily, Tapping the flies off the plate. And now I can put on a new plate and that can go back to the incubator. But meanwhile, I could check to see how many eggs have been laid here By the third day.

A good pot should produce 100 to 200 eggs per hour with the highest egg output between dawn and late morning. To simplify collection, scratch, or chisel origin into the collection plate auger, most eggs will be laid in or next to the scratches for this procedure. The flies are allowed to lay eggs for one to two hours, so late stage embryos can be harvested, but this timeframe will vary depending on the experiment.

Embryo collection methods will also vary depending on the number of embryos needed for the experiment. For dissection and electrophysiology, relatively few embryos are needed, in which case they can be picked gently from the plates with fine forceps. To remove the outer corion, rinse the embryos with distilled water and place them in a dish containing fresh 50 to 100%bleach each monitor the embryos under a dissection microscope.

After 30 to 120 seconds, the corion will dissolve exposing the shiny transparent vital membrane. Because bleach can rapidly destroy the vitalized tissues, the dechlorinated eggs should be briefly rinsed or placed in saline. Place the dechlorinated eggs in a plastic culture dish containing distilled water.

At this time, they can be visually genotyped by checking for GFP markers with a fluorescence dissection microscope, and appropriate filters after sorting out the of interest, proceed with the dissection. Because embryos are too small for dissection pins, some advanced preparation is required. Older embryos with developed cuticles must be glued to sogar coated cover slips using cyanoacrylate surgical or veterinary glue cyanoacrylate surgical glue, also known as histo aal.

Blue works well because it polymerizes in contact with solution containing divalent ions. Begin the dissection by transferring an embryo of interest to a cigar coated cover slip under a drop of saline with its dorsal side up. Remove the vital membrane by making a small incision in the membrane with a glass micro pipette or a tungsten needle.

Then gently free the embryo from the membrane. Early stage embryos will attach directly to clean glass or glass coated with poly L lysine. For older embryos, attach them to sard coated cover slips by delivering surgical glue through a small glass pipette attached to a rubber tube.

The glue flow is controlled by mouth. Use small amounts of glue to firmly attach the head and tail. Do this carefully because the glue mizes quickly in the saline.

To dissect the stabilized embryo, gently perforate a line along the dorsal midline with a glass electrode or a sharpened tungsten needle. Using the same tool make an incision by cutting through the perforations. It is critical that the dissection needle is always at a shallow angle to avoid damaging the ventral neuro musculature.

Next, remove the internal organs using gentle mouth suction from a glass pipette attached to rubber tubing. Spread the embryo flat by using more glue to attach the sides of the incision to the cover slip. Once all the internal organs are vacuumed away, the ventral CNS peripheral nerves and somatic musculature are exposed for experimentation.

At this point, the embryo is ready for electrophysiological recordings or it can be fixed for immunohistochemistry and subsequent microscopy using conventional techniques. We’ve just shown you how to isolate and dissect the later stage to softener embryos. The application of surgical glue and dissection of the embryos will be frustrating at first, but there are no secret tricks.

You’ve gotta develop the fine motor skills with practice over several weeks, but gradually the percentage of successful dissections will increase. Please have a look at our other geo video, which details recording configurations from muscle cells and the motor neurons. So that’s it.

Thanks for watching and good luck with your experiments.