Antibody Staining

Antibody Staining of C. elegans

by Michael Koelle, modified from Michael Finney

 

                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:

•     While waiting for your rabbits to make antibodies, construct 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. 

•     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 100 µl  in the tube.  Place on ice to chill.  Add  200 µl cold 2X witches brew, and  100 µl  10% paraformaldehyde (2.5 % final).  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.

3.  Incubate in 1 ml 1% ßME/tris-triton for 1-2 hours at 37° rotating.

4.  Wash in ~1 ml 1X borate buffer.

5.  Incubate in 1 ml 10 mM DTT/1X borate buffer for 15 min. at room temp.

6.  Wash in ~1 ml borate buffer.

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 ~1 min. in borate buffer.

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 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 Cappel (catalog # 1212-0081).  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.

 

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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 Cappel).  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 (MRWB)                                                                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

 

Add BME to 2% (fresh)

 

20% formaldehyde (see 10% paraformaldehyde)

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.

 10% paraformaldehyde ( Make fresh each time  as lasts about a week and old stuff doesn’t work as ). In a 15mL conical: -10 mL dH2O,  1g paraformaldehyde, - 50 uL 10M NaOH dissolve in 65° water bath (just crank up the 42° water bath, but remember to turn back down). It takes about 30 minutes to dissolve so make sure you complete this step before starting the protocol!)

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).

 

Ian’s Embryo Staining Protocol

 

1)      Wash several plates of worms off with M9.  Use microfuge tubes to spin worms. Wash 2X with  1ml M9 buffer.  Pool worms into 1 or 2 microfuge tubes. Use the tubes that have siliconized to prevent loss of embryos.

2)      Add 1 ml of Bleach solution to worm pellet.  Vortex occasionally and let sit for 3min. 

3)      Spin embryos at full speed for 30 sec.  Aspirate off bleach solution and add 1ml of new bleach solution. Again vortex and let set for 3min. Try not to exceed greater than 10 min in the bleach solution as the embryos tend to get damaged.

4)      Wash 3X with M9 buffer (1ml each time). Be careful not to loose the embryos. They tend to stick to the tube. If they are sticking to the side of the tube try spinning at 6000 rpm for 2 min.

5)      On the final spin aspirate off all but 30ul of the M9 buffer.

6)      Add 200 ul 2X witches brew (with 2% BME). Mix.

7)      Add 70ul 10% paraformaldehyde solution. Mix.

8)      Freeze immediately in liquid Nitrogen for at least 10 min. At this stages embryos may be kept at –80C for several weeks until ready to use.

9)      Thaw on ice for for at least 20 min.

10)  Wash 1X with Tris Triton Buffer (2min)

11)  Wash 2X with Antibody Buffer A (10min each wash)

12)  Add primary antibody (usually 1:100- 1:500 dilutions)

13)  Let sit for 4 hours or overnight without rocking.

14)  Wash 4X with Antibody Buffer B (1ml each with 10min between each wash). \

15)  Add 500 ul of Antibody Buffer A 0.5 to 1ul of secondary antibody (ie. FITC anti-rabbit)

16)  Let sit for at leas 2 hours at room temperature in the dark. 

17)  Wash 3X with Antibody Buffer B.

18)  Resuspend embryos in 10% glycerol (25-50 ul) plus anti-fading reagent.

19)  Put 5-10ul on a slide to visualize under the microscope.

 

Bleach Solution:

1ml bleach

1ml 10 N NaOH

8ml H20

Make fresh each time.