Worm Breeder's Gazette 7(1): 73

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.

Antibody Staining of C. elegans

D. Albertson, J. Sulston, E. Hedgecock

We have used two procedures for staining C.  elegans with florescent 
antibodies.  In the first procedure, which is especially useful for 
1) Young gravid adults (about six) are picked and placed into a 5-10 
l drop of M9 buffer on a microscope slide subbed with bovine serum 
albumen.  (Slides dipped in a 0.1% BSA solution, drained and dried in 
a vertical jar beforehand.) The animals are rinsed, if necessary, with 
M9 buffer then cut at the vulva with a razor blade.
2) A 12 x 12 mm coverslip is placed gently on top of the drop so 
that the eggs are not crushed.  The slide is immediately placed on dry 
ice for at least 10 min.
3) The coverslips are pried off with a razor blade and the slide 
placed in methanol at -20 C for 2 minutes followed by acetone at    -
20 C for 4 minutes.  Slides are then rehydrated through a series of 
alcohols (90, 70, 50, 30% ethanol; 2 minutes in each) and then into 
two changes of PBS (phosphate buffered saline).  They may be stored in 
the final jar of PBS.
4) Just prior to use, the excess PBS is wiped off the slide leaving 
a drop over the specimen and 5-10 l of antibody are pipetted onto that 
spot.  The slide is placed in a humidified chamber for 45-60 minutes, 
rinsed briefly by gentle dipping into a jar of PBS, and then for five 
minutes in a second jar of PBS.  A second antibody may be applied to 
the damp slide.  It is then incubated and rinsed as before.
This method gives reproducible staining of eggs using antitubulin 
and useful, though less consistent, penetration of the cut adults.  
Penetration of antibodies into whole mounts of eggs appears to be 
possible because the freezing and removal of the coverslips crack the 
eggshell and sometimes remove it from the egg.  The thickness of the 
liquid layer between the coverslip and the slide is probably most 
crucial and the correct amount will probably be found by practice.
To overcome penetration problems in larger animals, we have used a 
squashing procedure, followed by freezing and fracturing which yields 
flattened half-animals attached to each slide with their internal 
surfaces facing outward.  This procedure can be modified for antigens 
which do not survive the organic solvents used in step 3 above.
Washed, densely settled, worms are pipetted onto BSA subbed slides 
in 1 l drops of M9 buffer (up to 2 x 4 = 8 spots per slide).  A second 
subbed slide is lowered over and the two slides are pressed together 
with the thumbs without displacing the top slide laterally.  The pair 
of squashed slides is frozen on dry ice and then pried apart with a 
razor blade.  At this stage, the slides can be fixed in one of three 
ways, according to the antigen: 
1) sequential immersion in cold methanol and acetone as described 
above (slides are air dried and stored after acetone) 
2) heating for 2 minutes on a hot plate at 100 
or  3) air drying at room temperature.
The last procedure is the mildest but the fractured wormhalves are 
poorly fixed and may wash away during staining.
For humidified chambers, we use either a large petri dish containing 
water and the lid of a small petri plate to elevate the slide or, more 
simply, an inverted agar plate (containing 0.1% sodium azide to 
maintain sterility).