I have had good success with the yw line of flies. Set the flies up in the small clear cylindrical cages and maintain them on grape plates supplemented with dried yeast. Sprinkle a generous layer of yeast on the grape plate. Change the tray at least once per day. The flies take several days after eclosing before they lay well, and then they will continue to lay at an acceptable level for about 5 days. On the day of injection, change the grape plate once per hour for several changes so that the flies lay held eggs.
Before doing any injections, I strongly suggest you determine what fraction of the embryos survive all the treatments in the absence of injections. In other words, follow all the procedures described below but skip the injection step. You should have better than 50% survival. Since the injection procedure is likely to kill numerous embryos, survival from the other manipulations must be high to have a good chance of getting transformants. If your survival is low in the absence of injections, there are several things to consider. Are the lights illuminating the embryos during the dechorianation to close or to bright - this can heat the embryos. Are you being too rough during the dechorianation - freshly dechorianated embryos should appear like jelly beans with a smooth surface. Are the embryos becoming too dry - perhaps reduce the number of embryos you are trying to dechorianate so that they aare covered with oil more quickly. Lastly, is your fly stock healthy? I gave up on our rosy stock because many of the flies seemed to be laying a large fraction of dead embryos. Ideally, the flies should be between 3 and 7 days old and taken from bottles that are not too crowded.
The injection mix should already be prepared and have been stored in the freezer. Thaw the mix and microfuge it for 5’ to pellet any insoluble material. I leave the DNA in the tube and I draw up solution from the top to avoid insoluble debris. A few microliters of DNA is loaded into an injection needle. Chris Bell is preparing the injection needles on Ordway’s needle puller. A setting of 500 is giving excellent needles that taper rapidly to a sharp point. The needle is loaded by drawing the DNA solution up into a 50 ul capillary tube that has been drawn to a long fine tip in a flame. The fine tip is carefully inserted into the back of the injection needle and the DNA is slowly expelled. Sometimes the DNA solution closes off the needle about 1 cm up from the tip. There is a fine glass hair in the injection needle that guides the solution into the tip. Stick the needle in an upright position on the red clay in the injection room so that gravity assists in bringing the DNA into the tip. It will take about 5 to 10 minutes for the DNA to accumulate in the tip.
Your goal is to inject the embryos within 90’ after fertilization. The embryo is a syncitium during this time and injected DNA is taken up by nuclei without having to pass through a plasma membrane. You are injecting the DNA into the posterior end of the embryo where the germ cells will form.
In the summer, in the injection room is 64 oC and the humidity is between 65 and 70%. It is important to note the humidity as this effects how rapidly the embryos dehydrate. I begin by placing a fresh grape plate under the flies. The grape plate has a small amount of yeast in one region. The cage is placed on the floor just outside the injection room. This seems to work better to encourage egg laying than placing the cage in the incubator in the fly room. After about 40 minutes, change the grape plate and bring the plate with eggs into the injection room. Two pieces of double stick tape are placed side-by-side on a clean glass slide. On piece is about 1mm by 1cm - this is where dechorionated embryos are lined up. The other piece is about 5mm by 1cm. The placement of the tape varies depending on which microscope you are using for the injections. With tweezers, gently transfer about 40 embryos to the larger piece of double stick tape. Sometimes, you can scoop up clumps of several embryos and other times individuals are collected by grabbing the small protrusions sticking out from the posterior end of the embryos.
Nudge the embryos with the one prong of the tweezers to break the shell. I usually do this by placing the tip of the tweezers near where the embryo contacts the tape and gently rolling the embryo. Crack the shells on all the embryos before proceeding farther. All the embryos should be cracked within 5’ so the desiccation process is synchronized. This is particularly important when the humidity in the room is low as is the case in the winter. Then transfer the embryos in a line to the second tape. The embryos are snuggled up next to each other so that their posterior ends extend in the same direction over the edge of the tape by about 1/3 of the body length. To make each transfer, gently pry the embryo out of the cracked chorion with one prong of the tweezers and roll the embryo around on the chorion until the embryo sticks to the prong. If the embryos are not sticking to the tweezer prong, try wiping the prong with a kimwipe moistened with ethanol. Then stick the embryo to the second tape. Transfer as many embryos as possible during a 10 to 15 minute period. Finally, remove the larger piece of tape that has the residual chorions stuck to it.
The embryos are now ready for desiccation. If the humidity in the room is between 65 and 70%, place the slide in a jar containing blue colored desiccant for 3 to 4 minutes. I tip the jar on its side and place the slide directly on the desiccant. Screw the cap on while the embryos are being dehydrated. The exact time for dehydration will vary so you may have to adjust the time if the embryos look too dry or bleed frequently during the injections. Also, in the winter when the humidity can drop to 20%, there may be no need to place the embryos in the desiccant or the time might be shortened significantly. The best guide is to consider what happens when you do the injections. If none of the embryos bleed or the embryos have a shriveled appearance, then they are too dry. If all the embryos are bleeding, then they need to be dehydrated more (or you have a very bad needle).
After the embryos have been dehydrated, immediately overlay them with halocarbon oil. I usually bring a fresh aliquot of about 0.5 ml with me to the injection room. I cut the tip of a blue pipette tip, draw up 0.5 ml of oil from the stock in the lab and place the tip with oil in a 1.5 ml microfuge tube. I transfer oil to the embryos by lifting the blue tip out of the microfuge tube and dribbling the oil onto the embryos. I prefer this to using oil that has sat around in the injection room because there is no way of knowing what the oil has been subjected to.
To do the injections, the needle is carefully mounted in the needle holder. By now, the DNA solution should have flowed into the tip. The needle is placed at about a 300 angle relative to the stage so that the needle holder clears the slide and the slide holder. Be very careful to avoid knocking the tip of the needle against something, otherwise, the tip breaks and a new needle will be needed. Place the slide of embryos on the stage and bring the needle tip and posterior ends of the embryos into view. The tip of the needle should taper to a point and not appear broken. Apply gentle pressure to the syringe that is attached to the needle and proceed to pierce the first embryo. Usually, DNA does not flow until the needle pierces the first embryo - I think this breaks open the tip of the needle.
You should attempt to introduce a small but visible amount of DNA into the very tip of the embryo. In ideal cases, there is a crescent-shaped space of fluid at the end of the embryo where you can see a small amount of DNA enter. More often, the space is absent or the fluid originally in the space spills out of the embryo when the needle penetrates. Don’t despair; go ahead by injecting a small amount of solution into the end of the embryo.
Continue injecting all the embryos on the slide. After penetrating the first embryo, you should be able to see solution emitting from the tip of the needle when pressure is applied and the needle is outside the embryo. An ideal needle will produce a stream of small aqueous drops when the needle is moved rapidly through the oil in a direction away from the embryos. If no drops are produced, the needle may be clogged. Sometimes, the needle can be unclogged by piercing another embryo but often you will need to replace the needle. If large droplets form and you see that the tip fails to taper to a fine point, the needle is probable broken too much. A replacement is needed.
After completing the injections, the light on the scope is dimmed and the slide is left on the stage with the needle immersed in the oil. I leave the needle immersed in the oil so it doesn’t dry out. I now start preparing the next batch of embryos. It should take about 45 minutes to go through an entire cycle of collecting, dechorionating, desiccating and injecting embryos. If it takes over one hour, reduce the number of embryos you work with until you have mastered the technique. If it takes less time, you can try increasing the number you process for each injection cycle.
When the next batch of the embryos is ready for injection, transfer the previous slide of embryos into a humid Tupperware container and keep them in the injection room. Before removing the slide from the stage, raise the needle out of the oil and be careful not to bump the tip. Continue doing injections as long as you like. In general, I alternate between two grape plates and remove the excess eggs on the plate by wiping the surface with a moist brush.
Approximately 18 to 24 hours after completing the injections, carefully transfer the strips of tape containing the embryos to the surface of a clean grape plate. Do not add any more oil to the embryos. Put the plate in the humid chamber and leave it in the injection room. Approximately 72 hours after doing the injections, inspect the plate for hatched larvae. Carefully transfer the larvae to a vial of fly food that has been sprinkled with several grains of yeast. I prefer to pick up the larvae with an eyelash that has been taped to a handle such as a straightened paperclip. I touch the eyelash to residual oil on the grape plate and then poke a larva until it sticks to the eyelash. I then swipe the eyelash on the surface of the food in the tube. Record on the side of the tube the number of larvae, the injected DNA and the date of the injections. The larvae are allowed to mature at room temperature in the lab.
About 12 to 14 days after the injections, adults will eclose. These are referred to as Go flies and they will have white eyes. If the eyes are colored, you should be concerned that the yw injection stock may have been contaminated. Put Go individuals into separate vials and mate them with 3 or 4 yw flies. The yw females must be virgins. It is also best if Go females are collected as virgins as this reduces the chance of mating two transformants with inserts in separate locations. Newly eclosed flies take about 8 hours or more at room temperature before they reach sexual maturity. Therefore, you should be checking the flies twice a day when the Go flies are eclosing and placing the Go pupae at 18OC in the evening to reduce the chances that two transformants mate.
About 10 days after mating the Go and yw flies, the vials should be cleared of adults. "G1" adults will begin to eclose after about 12 to 14 days. Inspect these carefully for flies with colored eyes - these should be transformants. The color can range widely from a very subtle yellow to an orange or red. Any transformant should be mated individually with 3 or more yw flies. Use virgin yw females and also try to collect the G1 females when they are virgins. Keep track of which vial the transformants come from, labeling ones from a common vial with a common number and a different letter of the alphabet (e.g. 1A, 1B, and so on). This is important since transformants from the same vial might have the same insert. Transformants from different vials are likely to have different inserts unless mating occurred between two transformed Go flies.
The flies produced from the G1 adults are called G2 flies. There should be many colored eye flies in the G2 generation - all heterozygous. At this point, you are ready to begin characterizing these transformants. Possible courses of action are described in my directions called "What to do once you have a transformant?".