PROTOCOLS       

Fixing whole-mount worms for antibody staining.

There are a number of techniques for fixing worms in preparation for antibody staining. Different antibodies may require different types of fixation. The following fixation technique, however, works well for many antibodies. It is taken from Bettinger et al. (1996) Development 122: 2517-2527. This version of the protocol is from Dr. Simon Tuck.

The problem with worms when it comes to antibody staining is that they are small, and after methanol/formaldehyde treatment, quite fragile. The key to success for good staining is to treat the fixed worms as gently as possible. It is also better not to try to take off all the supernatant after each incubation/wash. There are so many steps in the protocol that if you loose just a few worms at each stage you will end up with nothing.

  1. Before you start place a bottle containing 2XRFB in a ice/water bath (not just an ice bath). Wash worms off the plate(s) with M9 buffer into a 15ml Falcon tube. Take all the worms and all the bacteria. Spin down the worms at 1400rpm in the bench top IEC centrifuge for 1min 15s. Remove the supernatant (being careful not to disturb the pellet) and wash the worms with 5mls of M9. Do 3 more washes in the same way over the course of about 45 mins (this allows the worms to digest the bacteria in their intestines).
  2. After the final M9 wash do one 5ml wash in just water. Remove the supernatant and then adjust the total volume of the suspension to 900ml by adding more water. The best way to measure the volume of the suspension is to pipette 900ul of water into a second tube and then add water to the tube with worms until the volumes look the same. If you pipette the worms themselves you will loose many of them (they stick to the insides of plastic pipette tips) and also risk breaking them. Place the worms suspension into the ice/water bath for at least 5 mins to cool. Make sure the tube is already labeled both on the top and side with genotype and date.
  3. While the worms are cooling fetch some liquid nitrogen (you can use one of the green ice-buckets). Secondly place a bottle of formaldehyde into the hood together with a 200ul pipette man.
  4. Add 1 ml of ice-cold 2XRFB to the cooled worm suspension and mix gently. Working in the liquid add 110ul of 36.5% formaldehyde to the worms and mix gently again. Tighten the lid and wrap parafilm around the top to seal it and then immediately snap freeze the worms in the liquid nitrogen.
  5. The frozen worms can be conveniently stored at -80oC until several different samples are ready. If the worms are to be used immediately, however, they must still be frozen in liquid nitrogen before the next stage. The freezing is part of the protocol it helps permeabalize the cuticle to allow antibody access.
  6. Remove the worms from the -80oC freezer and thaw them under a stream of warm water from the tap. Don’t let the worms themselves warm up. As soon as the suspension has thawed mix it gently and then place it into an ice/water bath.
  7. Incubate the worms in the ice bath for 3.5 hrs. Gently mix the worms by inverting the tube several times every 30 mins. Don’t mix after the final 30 mins.
  8. When the 3.5 hrs are over spin the worms down in the cooled Sorval 6000D centrifuge at 1K for 1 min at 4oC. Remove the supernatant and then wash the worms twice (at RT) with 2mls of TTE. Spin the worms at 1 K for 1 min in the benchtop IEC between washes.
  9. After the second wash remove the supernatant and add 2mls of freshly prepared TTE + 1% beta-mercaptoethanol. (Do this in the hood.)
  10. Incubate the worms in a 37oC water bath for 4hrs. Mix gently as before every 30 mins. Alternatively put tubes on a nutator in 37 oC air incubator for 4hrs. Either way works well.
  11. Don’t mix the worms after the final 30 mins but instead spin them down at 1K for 1 min in the bench top IEC. Remove the supernatants (in the hood) and then wash the worms once with 2mls of 1XBO3 buffer (at RT). Spin, remove the supernatants and add 2mls of freshly prepared 1XBO3 buffer + 10mM DTT. Incubate in a 37oC water bath for 15 mins.
  12. Spin, remove the supernatants and then wash with 2mls of 1XBO3 buffer. Spin again, remove the supernatants and then add 2mls of freshly prepared 1XBO3 buffer +0.3% H2O2. Incubate for 15 mins at RT. The timing of this last step is critical. The worms will be destroyed if they are left in this solution too long. It is also important that the H2O2 is not too old. You should be able to see bubbles forming in the tubes after 5 mins or so. The temperature of the room can affect this step. If the room is very hot, this step should be shortened. Using a 25oC worm incubator for this step can solve this problem.
  13. When the 15 mins are over pellet the worms by spinning as before (1K, 1 min @RT). Remove the peroxide solution and then wash the worms with 2mls of 1XBO3. Spin again, remove the supernatant and add 3mls of PTC. Leave the worms to rock GENTLY at RT for 30 mins. Spin down the worms (1K, 1 min). Remove supernatants and add PTB to a final volume of 2mls. You should have around 100-250ml of packed worms. These worms are stable for several months if kept at 4oC.
Buffers 50ml working stock

2XRFB is:

160mM KCl
40mM NaC
20mM EGTA
10mM Spermidine
30mM PIPES
50% Methanol 

4.0 ml 2M KCl
0.4 ml 5M NaCl
4.0 ml 0.25M EGTA
0.5 ml 1M Spermidine
5.0 ml 0.3M PIPES
25ml Methanol

Mix in a tube 5mls of 0.2M EGTA, 0.5mls 1.0M Spermidine, 5mls 0.3M PIPES, 8mls 1M KCl, 2mls 1ml NaCl.. Carefully pH the resulting solution to 7.4 with HCL (use conc. HCl to get the pH to 8 and 1 M to adjust it to 7.4). Adjust the volume to 25mls and then add 25mls of methanol [Merk 1.06009 (Kebo #5.2524-25)]. After addition of methanol the pH will be approx. 7.25. Store solution at 4oC.
TTE is: 100mM Tris. Cl pH 7.4
1mM EDTA

1% (v/v) Triton X-100

  

Mix in a bottle 10mls of 1M Tris.Cl (pH7.4), 200ul 0.5M EDTA and 89mls H2O. Using a blue pipette tip with the end cut off pipette up 1 ml of Triton X-100 (Sigma) and add to the bottle. When as much of the Triton has been expelled from the tip as possible eject the whole tip into the solution. Stir until the Triton is properly dispersed in the solution (approx. 1 hr).

100XBO3 buffer is 1M H3BO3 pHed to 9.2 with NaOH.

10mM DTT in 1XBO3. Make a 1:100 dilution of the 100XBO3 buffer. To 10mls of the diluted solution add 15.2 mgs of solid DTT (Boehringer Mannheim 708984). Use the solution fresh. Throw out anything you don’t use.

0.3% H2O2 in 1XBO3. To 10mls of 1XBO3 buffer add 100ml of 30% H2O2. Use fresh.
PTB is: 1X PBS

1% BSA (Boehringer Mannheim 735078)
1mM EDTA
0.5%Triton X-100
0.05% sodium azide

  

First make up 1X PBS. Dissolve 8g NaCl, 0.2g KCL, 1.42g anhydrous Na2HPO4 and 0.24g KH2PO4 in 800mls MilliQ water. Adjust the pH to 7.2 with IM HCl ( or 1M H3PO4). Adjust the volume to 1 liter. It is important that you make up PBS in this way and not make up 10X buffer and then dilute it. The pH of the diluted solution is not correct. To make up 50mls of PTB pour 47mls of 1X PBS into a bottle and add 0.5g BSA, 100ml of 0.5M EDTA and 2.5 mls of sodium azide. When the BSA has dissolved add 250ml of Triton X-100 as for TTE. Stir until the Triton is completely dispersed. Solution can be stored at RT.

PTC is the same as PTB except that it contains 0.1% BSA. To make up 200mls (you will need more of it than PTB) mix in a bottle 188mls of 1X PBS, 0.4mls of 0.5M EDTA, 10mls of 2% sodium azide and 0.2g BSA. Stir until the BSA has dissolved and then add 1ml of Triton X-100 as per usual. Stir until the Triton is dispersed.