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:
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.
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 ml 40X 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-A PBST-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 Brew Tris 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).
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(-azide-EDTA), 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.