- Collect live Drosophila embryos and remove their non-transparent chorions. Then, onto a glass coverslip, apply a line of heptane glue, which is tape adhesive dissolved in liquid heptane. To be able to handle the now dechorionated embryos, carefully align the specimens in a row onto the adhesive.
Transfer the coverslip into a dehydration chamber to partially desiccate the embryos. This will prevent cytoplasmic leakage during or after injections. Next, remove the coverslip and cover the embryos with halocarbon oil to prevent further dehydration while still allowing oxygen to diffuse into the embryos.
To perform the microinjection, use a microinjection system on a compatible microscope. By looking through the eyes pieces, identify both the embryos and the needle preloaded with injection solution. Break open the needle and ensure that it produces consistent and appropriately-sized liquid drops. Then, bring the needle inside the embryo and inject the appropriate volume. Bring out the needle, and proceed to the next one, repeating the injection for the remaining embryos.
In the example protocol, we will see a demonstration of Drosophila embryo microinjection for live imaging studies of cell division.
- Before setting up the coverslip, make heptane glue by unrolling double-sticky tape, and placing it in a 100 milliliter bottle. Add about 50 milliliters of heptane, seal the bottle, and rock it for several days.
Before preparing the embryos, make a dehydration chamber that the embryos will be placed in prior to injection. To do this, put one part of a 35 millimeter dish inside of a 100 millimeter Petri dish to make a "table". Add Drierite, which is anhydrous calcium sulfate, around it so that the height of the Drierite is no higher than the "table", and cover.
To begin preparing the embryos, first collect them from the lay cage by changing the grape juice plate with yeast every hour. The embryos should be imaged about two hours after the start of the collection. To prepare the coverslip, place it on one side of a microscope slide and tape the four corners down, so it doesn't move.
Using a cotton-tipped applicator, put one layer of heptane glue in a line on the coverslip. The glue should not be viscous and should dry in a few seconds. If it is too thick, add more heptane.
Finally, put a piece of double-sticky tape on the slide next to the coverslip. With a moistened brush, carefully pick up the embryos from the grape juice plate and place them on the double-sticky tape on the slide.
Next, use one half of a tweezer to roll the embryos over the double-sticky tape until the chorion breaks open. Pick up the embryo by gently rolling it over the chorion, so it sticks to the tweezer, and place it on the heptane glue on the coverslip with the long side of the embryo parallel to the long side of the coverslip.
Place 10 to 20 embryos in one row. Remove the coverslip and place it in the dehydration chamber for three to eight minutes. The time depends on the local humidity and the amount to be injected. Next, place the coverslip on a metal chamber with vacuum grease. Finally, cover the embryos with halocarbon oil to avoid further dehydration. The embryos are now ready for injection.
First, find the embryos under a 16X objective. Move the embryos away, and find the needle without moving the focal plane. Center the needle and move it up without moving it in the X or Y direction.
To open the needle, put the edge of the coverslip into the field of view, but not where a needle will be, and lower the needle to the same focal plane. Very carefully, move the coverslip until it hits the needle and gently breaks it open. Move the needle up.
Now bring the embryos into view. Lower the needle into the oil, and make sure to obtain nice liquid drops from the needle. Steadily move the embryo into the needle, inject a drop into the embryo, and then move the embryo away. After all the embryos have been injected, they are ready for observation on a confocal microscope with a 60 or a 100X objective.