Preparing Individual Drosophila Egg Chambers for Live Imaging

The aim of the following experiment is to record dynamic processes in living drosophila U sites. This is achieved by first isolating intact ovaries from well fed two day old female flies. As a second step individual aerials are extracted from the ovaries to image the egg chambers.

Then if they express an appropriate fluorescent protein, individual egg chambers may be imaged directly or may undergo further manipulations, such as microinjection of fluorescent markers prior to imaging. Ultimately, through analysis of the movie sequences, the dynamic interaction of intracellular components can be revealed. This method has helped us answer some key questions in the field of RNA transport and localized translation.

For example, how is the expression of proteins that set up the body plan regulated in time and space in the living oversight? Generally, individuals new to this method will struggle because it requires considerable speed and dexterity. Both practice better results will be achieved.

Visual demonstration of this method is critical as the manipulations of the egg chambers are difficult to convey text and because patient's technical expertise and sure hands are needed. Begin this procedure by placing a small drop of halo carbon oil on a cover slip and set it aside. Then anesthetize the prepared flies with carbon dioxide.

Use a sharp fine nose forceps to grasp a fat female fly at the anterior of the abdomen. Do not grasp in the center of the abdomen. Hold the female roughly two to three centimeters above the prepared cover slip.

Using a second pair of forceps, remove the tissue at the extreme posterior of the female. This will usually include gut tissues with clean forceps. Gently massage the abdomen from the anterior to the posterior end, like squeezing toothpaste from a tube.

The ovaries look like large opaque structures and will stick on the end of the forceps. Collect the ovaries by touching them to the halo carbon oil on the slide. Then repeat the process until four to six ovaries are collected.

Prior to isolating the aerials. Adjust the light source on the microscope so that the illumination strikes the slide at a shallow angle and creates contrasting shadows on the tissues. When dissecting the flies, it's very important to use the right tools.

The pointer we use is not too sharp so it damages the tissue, but fairly blunt so we can capture the the tweezers we use. Again, not too sharp so they damage the tissues, but just fine enough so we can manipulate things without bending the tool. It's also very important to have clean tools, so we clean them regularly throughout the procedure With closed forceps.

Separate the two ovaries by removing the connecting tissue at the posterior end. The posterior end has the most mature egg chambers. Orient an ovary horizontally with the youngest egg chambers towards your dominant hand.

Then gently grasp the posterior end by the remaining connective tissue and use a dissecting pro or tungsten needle to tease out individual aerials individual egg chambers from the Jamar stage to stage 10 will separate from each other, but older egg chambers will stay attached to the ovary. With the probe carefully drag 15 to 25 separated aerials to the center of the oil drop. If more than one ovary needs to be dissected, choose one from a different fly.

Discard any excess ovarian tissue and work quickly as cytes will begin to display phenotypic changes due to stress. After 40 minutes of being dissected from the ovary, remove the excess ovarian tissue from the oil Using clean instruments for the injection of mid stage cytes, orient the aerials perpendicularly to the long axis of the cover slip. To isolate late stage egg chambers.

A different method is required with the posterior of the ovary grasp as described earlier. Introduce the dissecting probe into the posterior end of the ovary near the forceps. To avoid puncturing the egg chambers, insert the probe between late stage egg chambers.

Pull the dissecting needle through the ovary until it exits at the early stage egg chambers. When done correctly, the probe will remove the connective tissue holding the aerials and late stage egg chambers in place. Repeat this step until the aerials are spread out on the cover slip.

When egg chambers are no longer stacked on top of each other, this step is complete. Using a dissecting probe, separate the late stage egg chambers to facilitate imaging to position Stage 14. Neck chambers use forceps to grasp their dorsal appendages.

Prepare the solutions to be injected by briefly centrifuging the mutt high speed. This will pellet any solid deposits that might block the injection needle. Then load the injection needle with a loading tip and carefully attach the loaded injection needle to the injector.

Now place a small piece of glass adjacent to the samples and mount the slide on the microscope stage. The glass will serve as a wall to break or unc. Unplug the needle.

Prepare the microscope and injection apparatus for the experiment. Taking care not to break the needle tip. If possible, use an automated stage that has point visiting capability to mark all the stage eight and stage nine cytes for injection.

Now, lower the injection needle until it touches the oil. Select the first point on the marked points list and focus on the cyte. Manually lower the needle until it is in the same plane of focus as the cyte.

Using the manipulator controls, move the point of the injection needle until it is inside the U site. A fast stabbing motion is often more successful than a slow one. Once inside, eject a small volume from the needle and visually assess the effects.

Repeat is required. If the injection is successful, the cytoplasm inside the cyte will be displaced. The precise amount of fluid injected is difficult to quantitate.

To increase the injected volume, either adjust the pressure or duration of the injection pulse. Break the needle tip to increase the needle opening slightly, or simply use multiple injection pulses before selecting the next mark.Cyte. Raise the injection needle high enough in the oil so that it does not bump into other cytes.

Extensive stabbing or lingering with the needle inside the cyte severely damages the egg chamber. So be quick. The injection should take less than 10 seconds.

If a mistake is made, move to the next mark to a site. Do not waste time on damaged to sight. When the needle gets clogged, move the needle to the broken piece of glass.

With the glass and needle in the same focal plane, gently ram the needle into the glass. Test that the needle is unclogged by pushing the injection button and seeing if any fluid exits the tip of the needle. When all the US sites have been injected, manually move the needle outta the oil and orient it outta the beam path for the microscope.

Make sure that it is in a safe orientation to avoid eye damage, lights are turned off. To facilitate fluorescence imaging speed is an imperative. If an hour or more has elapsed between egg chamber isolation and cyte injection, the cytes will be difficult to inject and exhibit phenotypic changes associated with stress.

Similar issues can arise with rough handling of the cyte or injection of excess volumes. Egg chambers expressing trenchgenic fluorescent proteins can be imaged without the need for injection. MS two labeling of endogenous RNA can be imaged with good results using the tissue isolation procedures using the complete procedure in vitro synthesized.

Alexa, 5 46 kin, RNA was injected into an ME 31 BGFP egg chamber, the RNA localized to the dorsal anterior of the cyte and formed a cap around the nucleus Once mastered. This technique can be done in about 10 minutes from the time of dissection to the start of imaging While attempting this procedure take care of the tissue by working quickly, but carefully. After watching this video, you should have a good understanding of how to record dynamic processes in living Drosophila cytes.