Staining protocol, zebrafish embryos
The following protocol worked with most of my antibodies. The oldest stage I stained was 6
days, but I think there´s not much difference between a six and a seven day old embryo.
You should not worry about the embryo´s thickness, one can see very well through, even at
this stage. To inhibit zebrafish pigmentation we keep 0.2mM PTU (phenylthiourea) in the
medium (water, embryo medium, etc) from 15 hpf on (Westerfield et al, 1994). Be careful,
PTU is very toxic!
Some of the steps can be modified, so you should play around a little bit in order to find the
best conditions for your antibody "a" and "b" designate alternative steps. I do most steps in
2ml tubes on a rotating wheel (slow!). To prevent the embryos from sticking to pipettes or
tubes, I coat plastic and glassware with PBS/BSA/DMSO solution (see step No 10). You just
need to quickly pipette this solution and your tips/pipettes will be coated. Same with the
tubes, just pipette this solution in and out and they are ready. Ok, here is the protocol:
1a:
Put the embryos for 30-60 sec in methanol at -20°C (Advantage: a better penetration of the
antibody in the tissue; disadvantage: sometimes the tissue shrinks during this step)
Wash 2-3 times in PBS
OR:
1b:
Anesthesize the Embryos in 0,03% MS222 in water (Sigma) for 2 min.
2a:
Fixation: 4h in 4% PFA-Fixation buffer at 4°C.
[250 ml recipe for Fixation buffer: 10g PFA + 150ml dist H2O  60°C (not higher T!) add
0,5ml 1M NaOH the solution should become clear filtrate through a paperfilter add
62,5ml phosphate buffer (Phosphate buffer, 0,37M: 5,32g NaH2PO4 x 1H2O + 52,11g
Na2HPO4 x 12H2O at 0,5l H2O dest, pH 7,4) add dist H2O to 250ml adjust pH to 7,4
(with HCl, most times not necessary)]
OR:
2b:
Like in 2a, use the same Fixationbuffer, but including 4% Saccharose and 0,15mM CaCl2
(some people say it´s better, but I see no difference…except lots of bacteria in my buffer after
a few days).
3:
Wash in Phosphate buffer (take the phosphate buffer from above and dilute it 1:4 in dist. H2O
 Final concentration: 0,09M) twice for 5 min.
4:
Rinse the embryos for 5-10 sec in dist. H2O.
5:
Permeabilize the embryos with acetone at –20°C, depending on age:
20hpf: 2min; 30hpf: 2-3min …7days: 7-12 min.
Take the time to find the right conditions here, because this is the most critical step for a
good staining.
6:
rinse in H2O for 5-10 sec
7:
wash two times for 5min in Phosphate buffer (0,09M)
8:
Incubate 3x for 5min in 1µg/ml NaBH4 (in PBS), prepare this solution fresh for each step.
This step decreases the background fluorescence caused by PFA. The concentration of the
Sodium Tetraborate is not that critical, I simply add a grain ("bead", "bit", "crumb"???) to
about 10 ml of PBS in a 6-well tissue culture plate.
9:
Wash 3-4 times in PBS, no bubbles should be seen or produced anymore.
10:
(Blocking is an optional step to decrease the backround staining, wich depends on the
unspecific binding properties of your secondary antibody, I leave this step out)
Block (depending on your secondary antibody) in 2% (vol/vol)donkey- or goat serum in PBS
+ 1%(w/v)BSA + 1%(vol/vol)DMSO, for 1h at 37°C.
11:
Primary antibody incubation: dilute your antibody in the upper blocking solution (or, if you
didn´t block, in PBS / 1%BSA / 1%DMSO) and incubate overnight at 4°C. Here I use 250µl
PCR-Tubes.
12:
Wash 4 times in PBS/BSA/DMSO for 15min.
13:
Dilute the Secondary antibody in blocking-solution (or, if you didn´t block, in PBS / 1%BSA
/ 1%DMSO) and incubate 1-2h at 37°C
14:
Wash 3 times in PBS/BSA/DMSO and one time in PBS.
15:
If necessary, remove the yolk with two thin needles.
16:
Embed the Embryos in Mowiol or Glycerol.