Worm Breeder's Gazette 7(2): 50
These abstracts should not be cited in bibliographies. Material contained herein should be treated as personal communication and should be cited as such only with the consent of the author.
The microinjection technique reported in WBG 5 No.2 can be difficult to implement without a few words of advice and an update on techniques used. The notes I provide here are primarily for injection into the gonad where it is necessary that the gonad keep functioning after injection. Obviously, there are worm injections that are less demanding on technique (e.g. injection into fixed animals). For gonadal injection, both the state of the worm and the quality of the pipette are critical. If it is difficult to penetrate a wild type worm, one of the two, or both, must be at fault. Worms for gonadal injection should be taken from a healthy, uncontaminated plate. Odd though it is, worms from some plates are easier to inject than others. I select animals for injection in which the distal arm of the gonad is arranged dorsally in an easily visible white plaque. My attempts to prepare several animals at a time for injection have not been very successful, so now I prepare one animal at a time, using the same anaesthetic as previously described, but leaving the animal in anaesthetic for only about 15-30 sec. I don't bother with a tetramisole plate anymore for drying the animal off, but do add 0.1% phenoxypropanol to the Voltalef oil covering the animals for injection. Worms are manipulated with a platinum wire during the first stages, and with a flamed micropipette when they are under oil. Large, old hermaphrodites are the easiest animals to inject and therefore can be tried to begin with; usually they can be mated to produce progeny if these are desired. After injection, it is instructive to look at the site of injection with Nomarski optics to assess the damage incurred. The use of a fluorescent dye (e.g. fluorescein conjugated BSA) is also extremely helpful to learn where you've had the pipette. It is absolutely necessary to score worms with Nomarski 4-24 hours after injection; if young embryos are not present in the uterus, the injected gonad is not producing embryos and the animals should be discarded. (My success rate is about 50%). The pipettes are less fussy, but still critical. I siliconize the pipette glass before pulling, and clean the pipette routinely in chromic acid and rinse it well in injection buffer or distilled water ( drops on a siliconized well slide) before injection. The tip diameter I use now is 3-4 microns. Some people like to work with oil in their pipette all the way to the tip. I find this difficult and have oil only in as far as it is easy to use for drawing up some fluid into the tip of the pipette. DNA solutions are relatively easy to inject at concentrations of 0. 5mg/ml or lower. I draw out a 5 l capillary pipette, and keep 1/4 l aliquot of DNA from the stock solution in that pipette in the cold for day storage. To draw DNA into the micropipette, I dispense the small drop onto a siliconized slide for a short time. If the DNA does not enter the micropipette well, try moving the pipette tip laterally in and out of the small drop of DNA. If the pipette is unclogged, this should allow entry of the DNA. If it is clogged, clean it in chromic acid. If the tip is kept under oil most of the time, the DNA does not clog the pipette. If you have trouble seeing where you are injecting in the animal with the dissecting microscope, it is possible to inject under Nomarski optics at 400x (C. Kenyon, personal communication).