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.

Update on Microinjection into C. elegans Gonads

J. Kimble

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).