Anti-Serotonin protocol with detailed notes

• Note: One 60 mm plate with a dense, well-fed, monoxenic culture has plenty of worms for a staining experiment. Starved worms have greater autofluorescence. Some bacterial contaminants also create considerable background fluorescence.
• Note: The entire procedure can be done in the same tube. For this reason, it is good to add a durable label such as a white ‘tough tag’ or ‘tough spot’ rather than just writing on the tube alone. Over the course of the experiment, the writing can rub off.
• Note: Because of steps 5 and 8, you can use a 1.5 ml tube with a ‘locking’ feature to prevent the lid popping open during 37C incubations.

2) Rinse 2-3x with M9 (if needed), spinning down and removing supernatant

• Note: If the worm plate has little bacteria, the M9 rinse may not be necessary, or could be reduced to a single solution change.
• Note: One pasteur pipet-full is ~1.5 ml. Remove supernatant consistent with retaining as many worms as possible. In this protocol, "Rinse" always means adding the maximum fluid that will fit in the tube, i.e., one pipet-full.

3) Fix overnight (ON) at 4 degrees C in 500 - 750 µl of 4% paraformaldehyde / 1x PBS.

• Note: Paraformaldehyde is somewhat toxic and probably carcinogenic. Work with it in the hood with gloves and lab coat.
• Note: Over-fixation kills staining. Staining seems to work better with 12-16 hr fixation vs. 24 hr. Two days of fixation abolishes staining.
• Note: 4% paraformaldehyde can be kept in the freezer for months. It can be thawed and used for about 2 weeks if kept at 4^o C. If in a hurry, thaw tube of 4% paraformaldehyde in hot water; be sure to cool on ice before putting on worms. Hot fix kills all staining! (Also, if a precipitate has formed, vortex the solution vigorously. Heating the solution again to 60-70 C may be necessary if substantial precipitate remains.) Note that one can also make 4% PFA solution fresh from refrigerated 32% or 36% concentrate (stabilized form, i.e. from EMS, see below).

4) Rinse 3x in 0.5% Triton X-100 / 1x PBS (PBSTx).

• Note: First, remove most PFA with a pasteur pipet and put into a labeled PFA waste container in the hood.
• Note: After this rinse, the fixed worms can be kept at 4^o C for several days if necessary before proceeding, perhaps up to weeks. For preps with marginal staining, however, it is probably better not to wait.

5) Incubate ON at 37^o C in 500 µl of freshly made 5% beta-ME in TrisTx (1% TX-100/0.1 M Tris, pH 7.4). Usually done rocking on Nutator in small incubator. (You can use microfuge tubes with a 'locking' feature to avoid the rare circumstance of popping open during 37C incubations.)

• Note: beta-ME (beta-mercaptoethanol aka 2-ME) is extremely smelly and somewhat toxic. Work with it in the hood with gloves and lab coat. Container with the b-ME solution must be kept tightly closed or b-ME will evaporate. Make small batches in a glass container; if staining stops working it could be from loss of potency of this solution from evaporation.
• Note: Overtreatment at this step (e.g., 2 days) will also abolish staining. Like with the fixation, it may be better to do closer to 12 hr than 24 hr - i.e., set up in the evening, and rinse first thing the next morning.

6) Rinse 2x in TrisTx

• Note: Leave used pipets and tips in the hood for a day or two before discarding (the beta-ME will evaporate in that time).

7) Rinse 1x in collagenase buffer (1mM CaCl₂in TrisTx)

8) Incubate 15 min - 8 hr at 37^o C in 2000 Units/ml Collagenase type IV (SIGMA C5138) in collagenase buffer. Usually done rocking on Nutator in small incubator. (You can use microfuge tubes with a 'locking' feature to avoid the rare circumstance of popping open during 37C incubations.)

• Note: This is likely the most critical step in getting good staining. Too little digestion and staining is weak or absent. Too much digestion and worms will be destroyed. Worms can be quite fragile after this step.
• Note: Usual volume is ~100-300 µl, but this is usually adjusted depending on mass of collagenase aliquot used. Batches of collagenase range from ~100 - 500 U/mg. We weigh out small aliquots (e.g., 5-30 mg) in 1.5 ml tubes (stored desiccated at -20C) and add the volume of buffer necessary to get 2000 U/ml. [Our current batch of collagenase has 390 U/mg, and typically works well with a 30 min incubation for both C. elegans and P. pacificus.] The volume of collagenase solution used can be lower for a smaller number of worms in the tube. Use a higher volume with tubes containing a lot of worms.
• Note: Digestion time is determined by examining 1-2 µl drops of worms (under the dissecting microscope) taken from the tube at regular intervals, starting as early as 15 min. When a few gravid adult hermaphrodites begin to break open (usually at the vulva), stop treatment. Once a time is determined for a batch of collagenase, one needn't necessarily monitor digestion, however, it is always best to monitor. You never have exactly the same number & stages of worms or ultimately the same concentration of collagenase.) Digestion time may need to be adjusted for different nematode species; this is sometimes also necessary for different C. elegans mutant strains. To get optimal head staining, one can do a more ‘severe’ digestion that fragments most worms. Heads will tend to stay intact, and give better staining, and may more reliably show faint serotonergic cells. It’s not a bad idea to split a tube of worms to do a short digest and a long digest.

9) Rinse 3x in PBSTx

10) Incubate for 1 hr at RT in 100 µl of 1% BSA in PBSTx (or Ab Soln A)

• Note: This step is intended to block non-specific Ab binding. It can be extended for hours to overnight; 1 hr is a minimal time.
• Note: BSA: Bovine serum albumen, Fraction V (Sigma A-9306), essentially immunoglobulin free for antibody solutions.
• Note: Boris Shtonda (Leon Avery's lab) recommended also including 10% goat serum (e.g., Sigma G9023) in the block and antibody steps. We have also tried this and it may reduce background staining; 1% GS is likely enough to achieve the same effect.

11) Incubate ON at RT in 1:100 anti-serotonin in 1% BSA in PBSTx (or Ab Soln A).

• Note: Although this incubation is typically performed ON at RT, this step can also be shortened to 1-4 hr at 37^o C.
• Note: It is important for best staining in C. elegans to use an antiserum made to PFA-conjugated serotonin antigen. We have been using for many years a Sigma antiserum (catalog S5545), one of the few available made to a PFA-conjugated serotonin.) Keep aliquots at -80C for long term. Do not refreeze once thawed. It can safely be stored for weeks to months (perhaps even years) at 4^o C. When finishing a tube, wash it out with the diluent to get as much Ab as possible.

12) Rinse 1x in PBSTx

13) Rinse 3-4x over 1-2 hr in 0.1% BSA/ PBSTx (or Ab Soln B)

14) Incubate ON in the dark at RT (or 4^o C) with 50 µl of 1:100 TRITC-conjugated goat anti-rabbit IgG in 1% BSA in PBSTx (or Ab Soln A).

• Note: Use your favorite secondary antibody. We prefer red fluorophores like TRITC because they typically fade much more slowly than others.
• Note: This incubation can also be performed for 1-4 hr at 37^o C.
• Note: Keep secondary antibody covered (e.g., with foil) at 4^o C for weeks. Aliquots are frozen at -70^o C for long-term storage. Do not refreeze once thawed.
• Note: TRITC-conjugated Abs fade much more slowly than those with FITC.

15) Rinse 1x in PBSTx

16) Rinse 3-4x over 1-2 hr in 0.1% BSA/ PBSTx (or Ab Soln B)

• Note: If in a hurry one can skip this step and look immediately after the PBSTx rinse, with higher background; if stopping here, do add 0.1% BSA solution.)

• Note: After the collagenase step, worms are often very fragile. Slightly sliding the coverslip after mounting on agarose pad can completely destroy the worms, leaving only shreds and pharynges.
• Note: Worms can typically be kept in tubes at 4^o C in the dark after staining and viewed for several days or even weeks later without significant fading.
• Note: We often add 1 µl of '100x' DAPI in water (10 µg/ml, so final conc is ~0.1 µg/ml) to the tube prior to mounting to see nuclei, and to assess quality of permeabilization. Bad/uneven DAPI staining indicates insufficient permeabilization for good antibody staining (although good DAPI staining is not a guarantee of sufficient permeabilization for antibody staining). Keep stock DAPI solution at 4^o C or frozen and protected from light.

Other Notes:
In general, when rinsing, remove as much supernatant as possible without losing worms. Err on the side of removing less to retain worms. One exception is prior to adding the primary antibody, when you want to remove as much supernatant as possible to have the highest effective concentration of Ab.

You can add Sodium Azide (NaN₃, final concentration 0.1 - 0.05%) to most or all solutions to keep things from growing (then they need not be autoclaved).

1M Tris, pH 7.4

• Tris HCl 131.9 g/liter
  
• Tris Base 19.7 g/liter
  (This solution should come out pretty close to pH 7.4, without further adjustment.)

1. Make up this stock by adding 10 ml Triton X-100 to 90 ml water.
   
2. Put stir bar in bottle with water, then add Triton very slowly while stirring.
   
3. Wash out pipet with solution at the end because much viscous Triton will be stuck to the inside of the pipet.

1. dissolve in ~800 ml ddH₂O
   
2. adjust pH to 7.4 with approx. 13 ml 1N NaOH
   (=13 mMoles/1liter 10x buffer or 1.3 mMoles//1liter 1x buffer)
   
3. adjust to 1 liter
   
4. aliquot and autoclave

   Na₂HPO₄.7H₂O = Sodium Phosphate, dibasic, heptahydrate (or if using Na₂HPO₄, use 11.94 g)
   NaH₂PO₄.H₂O = Sodium Phosphate, monobasic, monohydrate

1. Heat ~80 ml dd water to 55-65^o C in beaker stirring in hood (use dedicated beaker, stir bar and 'hood thermometer').
2. Add 4 g paraformaldehyde powder or flakes, stir until dissolved. Add a few drops of 1M NaOH if necessary to help the paraformaldehyde to go into solution.
3. Solution should become clear. Don't boil or heat longer than necessary.
   
4. Add 10 ml 10x PBS and make up to 100 ml.
   
5. Aliquot 10 ml each in 15 ml centrifuge tubes and store in -20^o C freezer.

Antibody solution A (Ab soln A)
1% BSA, 0.05% NaAzide, 1mM EDTA, 0.5% Triton X-100 in 1X PBS

Note that 0.5% Triton X-100 in 1X PBS is ‘PBSTx’ - one can start with PBSTx, add BSA solid (0.1 g / 10 ml) and other concentrated solutions (e.g., 20 µl 0.5M EDTA stock + 100 µl 5% NaAzide stock per 10 ml). NOTE: Sodium Azide is EXTREMELY TOXIC - handle with care.

Antibody Buffer B (Ab soln B)
Same as Ab soln A except 0.1% BSA. Much more soln B is needed - make 50-100 ml at a time.

Neurotransmitter antisera were "invented" by H.W.M. Steinbusch et al. (1988). This protocol for C. elegans is derived from that originally published by Desai et al. (1988) with minor modifications (Loer & Kenyon, 1993). I have continued to modify the above protocol, and provide notes on variations that you can or cannot get away with.

Desai C, Garriga G, McIntire SL, Horvitz HR (1988) A genetic pathway for the development of the Caenorhabditis elegans HSN motor neurons. Nature 336:638-46.

Loer CM, Kenyon CJ (1993) Serotonin-deficient Mutants and Male Mating Behavior in the Nematode Caenorhabditis elegans. J Neurosci 13: 5407-5417.

Steinbusch HWM, Wouterlood FG, deVente J, Bol JGJM, Berkenbosch F (1988) Immunohistochemical localization of monoamines and cyclic nucleotides. Acta Histochem S35:85-106.