Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

The overall goal of the following experiment is to record subcellular events during early embryogenesis Insino Ratu elegance. This is achieved by using transgenic strains with multiple fluorescently labeled proteins that allow the visualization of DNA as well as subsets of vesicles In the early embryo, specific organelles are labeled by a microinjection of fluorescent dyes, such as Lyo tracker or MIT tracker into the gonad of the adult worm. Next embryos are cut out of adult worms and observed with a confocal microscope to obtain time-lapse recordings of the movements of the labeled proteins in organelles.

Ultimately, the protocol reveals interesting movement of lysosomes and GFP tagged vesicles during early development in CL allergans. My name is Connie Hedger, and today I'll be showing you how we perform time-lapse microscopy in our lab, the Boyd lab at the University of Alabama in Huntsville. For this experiment, gro nematodes on NGM agri plates, seated with an OP 50 bacterial lawn at 25 degrees Celsius the day before the experiment, pick at least 40 L four larvae onto seated plates and incubate them overnight at 25 degrees Celsius.

These worms will be young adults. For the experiment. Make an injection pad by sandwiching, two drops of 2%aros in water from a past pipette between two cover glass slips, and applying pressure to shape the agros into a 1.5 centimeter diameter circle.

Remove the top cover slip and dehydrate the pad by placing it in an 80 degree Celsius oven for one hour, or allow it to sit on the benchtop overnight. On the day of the injection, prepare a diluted MIT tracker or lyo tracker dye solution and pull injection needles from a 1.2 od glass capillary. Backfill the needles with a diluted dye solution and store them in a dark humidified chamber mount injection needle onto the micro manipulator and connect the needle to a pressure regulator with a 1.2 millimeter internal diameter tube.

Now place two drops of heavy mineral oil onto the injection pad. Place the injection pad onto the microscope and lower the needle into the oil. Apply injection pressure and observe whether fluid flows out of the needle.

If not, you will need to gently break the end of the needle by gently driving the tip into a small bit of broken cover glass placed on the injection pad. After the needle is flowing, proceed with the injection a pro one hour prior to viewing embryos. Use a pic to transfer young adult worms into the oil drop on the injection pad.

Arrange three to 10 worms in parallel and a little less than one worm length apart. It is now important to inject the worms quickly before the worms desiccate. Move the pad with worms onto the microscope stage.

Focus onto the central part of the distal gonad. Then use the micro manipulator to move the tip of the needle into the same focal plane. Puncture the worm by moving the stage horizontally.

When the needle tip is inside the cus apply pressure to fill the gonad with the dye mixture. The gonad will expand with the volume of the injection. Proceed with injecting.

Both go net arms of each worm after injection. Cover the worms with 0.5 milliliters of egg buffer applied. Using a pulled pasture pipette, then allow the worms to recover for one to two hours at room temperature in a light free humidified chamber.

Begin by preparing a new agros pad for viewing the injected embryos with a past pipette. Put three drops of molten agros onto a clean microscope slide and place that slide between two other slides wrapped in one layer of labeling. Tape the tape controls the thickness of the pad.

Next, press the clean microscope slide down onto the agros drops so it rests on the tape slides. Do not remove the top slide until the embryos are ready to mount. Now drop 20 microliters of egg buffer onto the center of a cover glass and transfer the injected worms into the buffer Under a dissecting microscope, cut the worms open near the vulva.

Using a 26 gauge hypodermic needle, the embryos will spill out of the worm. Now remove the top slide of the imaging slide and lower the aros pad upside down onto the cover glass with the embryos. If the embryos will be imaged over an extended period of time, seal the edges of the cover glass with petroleum jelly to prevent desiccation.

Now, begin imaging by locating embryos under the 10 x objective to find early stage embryos, search for embryos that are in the process of completing maternal meiosis. Myotic embryos can be distinguished from later embryos because they have not undergone shortening. After completing meiosis, there's a significant gap between the cell membrane and the eggshell at the anterior end of the embryo.

Once an appropriately staged embryo is identified, move to a higher magnification lens for recording of embryogenesis. This embryo expresses both M cherry histone two B and GFP ubiquitin tagged proteins. The maternal and paternal DNA is visible via the M cherry H two B, and thus shows the stages of early development.

This embryo was injected with OC Tracker blue and expresses GFP Ubiquitin. The images were collected using a 63 x 1.4 NA lens with 4 88 and 5 55 lasers set at minimal power and maximum gain. Typically, Zacks are collected every 10 to 15 seconds for up to 60 time points before photobleaching occurs.

For this TimeLapse video, the Z stacks were collapsed into maximum projection images While performing time-lapse. Microscopy embryos should begin pronuclear migration within a few minutes of completing myosis two, and within 20 minutes. The first cytokinesis should begin.

If you observe arrested development of your embryos, try decreasing the laser intensity or use a thicker augurous pad pad.