Antibody Staining of C. elegans
by Michael Koelle, modified from Michael Finney, 1/98
This protocol works for many antibodies. A much longer protocol involving collagenase
treatment of the worms is necessary for some antisera (e.g. anti-serotonin). Dauers are not
permeabilized by this protocol and thus don't stain. Animals fixed this way can be stained
with X-gal, and GFP fluorescence is supposedly still present.
The critical parameters are extents of fixation, time of reduction steps, pH of the borate
buffer, and antibody concentration. Be especially careful with the borate pH; some earlier
protocols called for a lower pH which gives very poor staining. When trying an antibody
for the first time, you should try a few different fixation times and a few different
antibody dilutions. It's easy to process many such variants simultaneously and then
compare them to determine the optimum conditions.
Initially getting an antibody to work for stains:
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While waiting for your rabbits to make antibodies, contruct a worm strain carrying
an integrated array of extra copies of the gene encoding your antigen. Use this
overexpressor strain to optimize the staining conditions.
Identify a protein null mutant by sequencing alleles or performing western blots on
extracts of mutants. This mutant can be used as a negative control for staining.
Also, the primary antiserum can be preabsorbed against fixed mutant worms
before being used for stains; this will eliminate some background.
A general note on handling worms without losing them. This protocol involves many
transfers and washes of the worms. A common problem of beginning stainers is to lose
some worms at each step, and thus end up with none at the end of the procedure.
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Transfers: Worms stick to plastic pipetteman tips unless detergent is present. Thus
the first transfers described below must be done with a glass pasteur pipette, but
plastic pipetteman tips can be used for the later transfers when the worms are in
buffers containing triton.
Washes: Always spin the worms down gently, at ~3000 RPM for ~30 sec. in a
variable speed microfuge. The very best way is to use a swinging bucket
microfuge; this avoids losing worms that are spun up against the wall of the tube. I
use a Shelton Scientific model VS-13. After spinning remove most of the liquid
with a 1 ml pipetteman blue tip, and the remaining small amount of liquid with a
small tip. Costar makes some white platic tips with a very small orifice that are
good for removing liquid without losing worms. During the many washes between
and after the antibody incubations, it is better to just leave the last ~20 µl of liquid
in the tube than to risk losing any worms; you're doing enough washes that this
residual liquid gets diluted out and won't matter. The biggest loss of worms should
come early in the procedure, at the reduction/oxidation step, which make the
worms sticky. All the washes should be done in 0.5 ml of liquid with the tubes on
a rotator (e.g. "Labquake" brand).
1. Fixing the worms. Wash worms off an unstarved plate with M9, spin them down in a
clinical centrifuge, and resuspend/spin with dH20 to wash out most of the bacteria.
Transfer the worms to a microfuge tube with a pasteur pipette, spin 3K for 30 sec. and
remove some supernatant to leave 0.5 ml in the tube. Place on ice to chill. Add 0.5 ml
cold 2X witches brew, and 20% formaldehyde to a final concentration of 1-4% (1% is
typical). Mix well, and freeze in liquid nitrogen (may place in a -80° freezer indefinitely
at this point), and thaw quickly in a 70° water bath (remove just before all the ice in the
tube thaws). Incubate at 4° with occasional agitation for 30 min to overnight (30 min to a
couple of hours is typical).
Adjust the fixation extent as necessary to optimize the staining. e.g. UNC-86 antigen is
sensitive and staining goes away if the worms are fixed too long; other antigens are
stabilized by longer fixation.
Theory: methanol precipitates proteins, reducing diffusion before fixation. Spermidine
and formaldehyde together crosslink proteins. Chilling the worms hypercontracts their
muscles; initally fixing them in this state makes the worms physically stronger so that
they survive the procedure without falling apart. Freezing cracks egg shells, letting
fixatives in.
2. Wash the worms twice in tris-triton buffer (each wash is for 1 minute on a rotator in 1
ml).
3. Incubate in 1 ml 1% ßME/tris-triton for 1-2 hours at 37° rotating.
4. Wash in ~1 ml 1X borate buffer for 1 minute.
5. Incubate in 1 ml 10 mM DTT/1X borate buffer for 15 min. at room temp.
6. Wash in ~1 ml borate buffer for 1 minute.
7. Incubate in 0.3% H2O2/1X borate buffer 15 min. at room temp. rotating. Be carefull
here, since oxygen released from the solution may cause loose fitting tube caps to pop
off! Either use a clip ("LidLock") to hold the caps on, a screw cap tube, or else don't
rotate the tube and leave it upright.
Theory: the above reduction/oxidation steps help permeabilize the worm by disrupting
the cuticle, which is extensively crosslinked by disulfide bonds. The prolonged ßME
treatment at 37° also helps kill worms enzymes like proteases, peroxidases, and DNAses.
8. Wash in 1 ml borate buffer for 1 minute.
9. Incubate 15 min. in PBST-B. At this point the worms are stable and can be stored in
PBST-B in the refridgerator indefinitely.
Theory: The BSA in PBST-B blocks non-specific binding of antibody.
10. Primary antibody incubation. Transfer a suspension of worms containing the
equivalent of ~5µl of packed worms to a 0.5 ml tube, spin and remove as much liquid as
possible, and add ~20 µl antibody diluted in PBST-A. Mix by pipetting up/down.
Incubate at room temperature overnight (agitate occasionally if you can).
Try a few different antibody dilutions the first time you do stains. A ballpark estimate is
to use a 10-fold higher concentration than what works well on westerns. A decent crude
serum usually works on westerns at about 1:2000.
11. Wash the worms 4 times for 25 minutes each on a rotator at room temperature in
PBST-B.
12. Incubate 1-2 hours at room temperature in 20 µl 2° antibody diluted in PBST-A,
agitating occasionally. I've been using a 1:25 dilution of FITC conjugated goat anti-rabbit
IgG purchased from ICN (catalog # 55646). The FITC can bleach, keep the tubes covered
in foil or in a dark box when possible. Keep this 2° antibody solution after use; it can be
used again, and will give a lower background on the second use.
13. Wash the worms 4 times for 25 minutes each on a rotator at room temperature in
PBST-B.
14. The stained worms can be stored for months at 4° in the dark.
15. Viewing: place 3 µl worm suspension on a microscope slide. Add 3 µl antibleaching
solution and mix by stirring/pipetting. Drop on an 18X18 mm coverslip. Seal the edges
by applying a strip of clear nail polish all around the coverslip (Revlon clear nail enamel
is good). Slides thus prepared can be stored in the freezer for months.
Hints on optimizing signal/noise in antibody stains:
1. A common trick is to preabsorb the 1° and 2° antibodies against fixed worms to reduce
the background staining. The 1° antibody should be preabsorbed if possible against a
mutant that lacks the antigen of interest. This sometimes makes it possible to use a crude
serum for stains instead of an affinity purified antibody.
2. A weak signal may be amplified by adding a 3° antibody step. After incubating with
the FITC-conjugated goat anti-rabbit 2° antibody and washing four times, incubate with
an FITC-conjugated rabbit anti-goat antiserum, using the same condtions and washes as
for the 2° antibody (this 3° antibody can also be purchased from ICN). After the 3°
antibody step the background will be significantly higher, but the signal/noise ratio may
be better than after the 2° antibody alone.
10X PBS/200 ml40X BO3 buffer
16 g NaCl 1 M H3BO3
0.4 g KCl 0.5 M NaOH
2.3 g Na2HPO4.7H20 Very important: check that pH >9.5
0.4 g KH2PO4 add more NaOH if required
PBST-APBST-B
1X PBS Same as PBST-A except 0.1% BSA
1% BSA (Pentax Fraction V)
0.5 % Triton X 100
5 mM sodium azide
1 mM EDTA
2X Witches BrewTris Triton buffer
160 mM KCl 100 mM Tris Cl pH 7.4
40 mM NaCl 1% Trion X-100
20 mM Na2EGTA 1 mM EDTA
10 mM spermidine HCl
30 mM Na Pipes, pH 7.4
50% methanol
20% formaldehyde
Weigh somewhat more dry paraformaldehyde than you need (<300 mg) and put it in a
microfuge tube. Multiply the weight in mg by 4.5 and add that volume in microliters of 5
mM NaOH. Place in a 65° water bath for 30 minutes with occasional mixing. Spin for 1
min. to pellet any undissolved paraformaldehyde. Use the supernatant immediately.
Antibleaching solution
1 mg/ml phenylenediamine
10% PBS
90% glycerol
This is carcinogenic and should be stored at -20° in the dark.
Note: some people use other antibleaching reagents, such as 2% n-propyl gallate or 2%
Dabco™ 33-LV (Aldrich catalog #29,073-4).
Preliminary Protocol for Metal Enhanced DAB Staining
using HRP 2° Antibodies
This protocol is far from optimized yet. I haven't attempted to maximize its sensitivity.
Protocols for DAB staining Drosophila are more elaborate, and probably more sensitive.
They generally use an "ABC" triple sandwich amplification method (1° antibody,
followed by a biotinylated 2° antibody, and finally HRP conjugated avidin). You can buy
reagents for this sort of thing from Vectastain or Pierce. The Drosophila protocols also
use fancier blocking procedures. It is also possible to suppress endogenous peroxidase
activity, which might lower the background (e.g. Pierce sells a "peroxidase suppressor",
catalog #35000).
1. Carry out the fixation, permeabilization, and primary antibody incubation steps exactly
as described above for the FITC detection method. You should carry out a separate
dilution series of the 1° antibody to optimize its concentration for DAB detection: I ended
up using 4-fold less 1° antibody for DAB than for FITC detection.
2. For the remainder of the procedure, solutions will be in PBST lacking sodium azide
and lacking EDTA. (Azide inhibits HRP, and I left the EDTA so as not to chelate the
metal ions in the developing solution).
3. After the 1° antibody incubation, wash 4X25 min. in PBST-B (-azide-EDTA) at room
temp on a rotator.
4. Incubate 2 hours at room temperature in 20 µl 2° antibody diluted in PBST-A(-azideEDTA), agitating occasionally. I've been using a 1:60 dilution of HRP conjugated goat
anti-rabbit IgG purchased from Biorad.
5. Wash the worms 4 times for 25 minutes each on a rotator at room temperature in
PBST-B (-azide-EDTA).
6. Development. Wear gloves and be careful here; DAB is a suspected carcinogen. I've
been using a commercial DAB preparation from Pierce to develop the stain
("ImmunoPure Metal Enhanced DAB substrate Kit", catalog #34065). Store the 10X
DAB solution at -20°, and the 1X peroxide solution at 4°. Just before use, mix the DAB
solution to resuspend the metals, and combine 1 part DAB solution with 9 parts peroxide
solution. Spin the woms down and remove as much supernatant as possible. Add 400 µl
of DAB/peroxide developing solution, and incubate on a rotator at room temp. 1-20
minutes. Can remove an aliquot of the developing worms to a glass depression slide and
watch them develop under a dissecting scope to decide when to stop the reaction. Most of
the staining occurs very quickly (1-2 minutes), and very little occurs after that. I've
routinely been developing for 8 minutes.
7. To stop the reaction: spin the worms down, remove the supernatant. Wash 1 min. on a
rotator in PBST-B(-azide-EDTA), followed by three more 5 minute washes.
8. Mount 3 µl stained worms, with 3 µl 80% glycerol, under a 18 mm square coverslip
sealed with nail polish. The stained worms are stable in the fridge for at least a week.