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1 Antibody Staining Methods
1.6 Protocol for staining early embryos by Bruce Bowerman
1. Collecting early embryos.
Synchronize a culture of worms by doing a hatchoff: fter hypochloriting a population
of adult N2 worms to isolate embryos, the embryos are allowed to hatch out as
arrested L1s overnight at room temperature in M9.. We hypochlorite by adding
0.5 ml of 5% NaHypochlorite (Sigma; Safeway will do) and 0.2 ml of 5 N KOH to
5 ml of worms in distilled water, using a 15 ml plastic conical tube. Once the
adults have dissolved, pellet the embryos (at 2000 rpm in a table top Beckman
centrifuge: stop the rotor once it reaches 2000 rpm and let brake to a stop),
and wash twice by resuspending in M9 and repelleting. The embryos are then transferred
to a flask of M9 and put on a rotating platform overnight to hatch. The L1s
are then put onto plates with food and allowed to grow up until they are just
becoming adults.
When worms are just becoming adults, harvest and hypochlorite as above. To speed
up hypochloriting, I shake the tube by hand for a minute or so, and then pellet
the worms and embryos as above. The pellet is resuspend in water and then add
fresh hypochlorite and KOH (check with dissecting scope or eyeball). Resuspending
in fresh hypochlorite definitely speeds things up; apparently the first batch
is partially neutralized by all the bacterial debris and various forms of worm
slime and goo. Add M9 and pellet as above. Resuspend in M9 and pellet again.
Resuspend in distilled water and put onto slides (see below).
I think that thorough hypochloriting is maybe important for weakening the eggshell
and facilitating permeabilization during fixation. This procedure allows for
collection of one cell embryos, usually after pronuclei are relatively close
together, and lots of 2-cell to 28-cell stage, and some later stage embryos.
A few dense plates (60 mm) will give you plenty of embryos for several slides.
2. Coating slides with poly-lysine.
Poly-L-lysine (in solution from Sigma: Cat. # P 8920). We use a capillary tube
to draw up solution by capillary action: place the full tip on surface of slide,
and allow a small drop to collect. Spread out on slide by using side of tube
to spread out evenly over entire slide. Slides should be reasonably clean--fresh
out of box or from box opened only to make these coated slides. It's not necessary
to acid wash, although that will improve sticking. If slides are clean, solution
should spread out evenly without beading up. Avoid getting any small pieces
of glass or debris on slide (these will support coverslips and prevent one from
adequately squashing embryos: see below). Place coated side up on a preheated
hot plate (on a fairly low setting: as long as it is warm when touched with
hand). By looking at the slide from an appropriate angle such that light reflects
from the coating, you can see when all of the water has evaporated. Remove immediately
and place in a slide rack. I keep the end of the slide hanging over edge of
hot plate to facilitate picking it up. These slides are good for at least two
days, but eventually the embryos will not stick well to them if they sit out
(in our experience). I typically prepare fresh slides before hypochloriting
worms for embryos. The solution from Sigma goes bad within about six months.
We store it at room temp. If frozen, it forms a precipitate. We routinely buy
a new bottle every three months to avoid having to figure why our embryos stop
sticking to slides.
3. Preparing paraformaldehyde (Mallinckrodt).
Make up a 5% solution (1.25g in 25 ml PBS is my usual) by adding paraformaldehyde
to PBS (standard recipe) and then boiling gently on a hot plate until paraformaldehyde
is dissolved. Allow to cool to room temp before using. Make up same day, before
hypochloriting worms for embryos. Prepare in hood. All to common a problem is
the loss of SKN-1 staining during fixation. The antibody is stable for months
at 4 degrees (with 0.05% azide). We tried putting the embryos in water on ice
before putting onto chilled slides for fixing, while working at a room temperature
dissecting scope. This didn't help. With much diligent practice, one can get
50-80% of slides to fix and stain at detectable levels. We've tried many fixation
protocols and still find this the only one that works.
4. Fixing embryos for staining.
After pelleting embryos from hypochlorited worms (see above), suction off the
water until you have a reasonably concentrated batch of embryos in 0.5 ml or
less of water. Add 30 to 50 ul the resuspended embryos to the middle of slide
(poly-lysine side up! I use frosted slides to make it easy to check which side
is which). Allow embryos to settle onto slide. They should be crowded but not
piling up at all (too many embryos make it difficult to squash them adequately).
About 100 to 300 embryos in 50 ul is a good number. If embryos are in M9 or
any other salt solution, they won't stick as well, although it isn't essential
to have them in water to get them to stick. Once embryos have settled (you can
tell if they're sticking because they won't move if you quickly jiggle the slide),
add about 150 ul of 5% paraformaldehyde with a pipetteman, coming in from the
side (not squirting directly on top of embryos). Remove most liquid (with pipetteman),
discard, and add another 150 ul of fixative. I do this three times to ensure
embryos are in paraformaldehyde. Embryos will remain stuck; a few will come
loose and be lost. Place a 22 x 22 mm cover slip over embryos (see drawing).
Brace back corner with finger of one hand and lower cover slip gently with other
hand (I use fancy forceps). Try to avoid trapping any air bubbles. Maintain
brace with finger of first hand to keep cover slip from floating off. Wick out
excess liquid with a kimwipe while watching embryos in dissecting scope. This
sometimes requires kimwipes on both sides (once most liquid is out you can quit
bracing with your finger and use both hands to line edges of kimwipes with the
edges of the coverslip to maximize the wicking out of the liquid. The embryos
should visibly flatten: to get good fixation, it is often necessary to flatten
them to the point that some of the older embryos (28-cell stage or so) begin
to ooze out blastomeres from their eggshells. When this starts to happen, I
stop wicking out liquid and put the slide in a humidity chamber to fix for about
15-30 minutes. I typically set up 4 to 5 slides consecutively, placing them
together in the humidity chamber as I finish each one. If the fixation is good,
the embryos will almost immediately become very white instead of the yolky-yellow/brown
color they are before squashing. If they remain yellow/brown, they probably
won't stain, although you can't count on the staining working just because they
turn whitish. It usually takes quite a bit of practice to get good at this,
and even then I'm usually successful in getting staining with only about 60-80%
of the slides I prepare in which the embryos turn whitish (I toss out slides
in which I don't get good squashing to the point of having the embryos turn
whitish). This squashing is a crucial step. Good squashing => 100% embryos
stain with SKN-1 antibodies. But such a slide can be rare unless one is routinely
doing this.
It is also possible to get staining by cuttin open worms on slides in PBS or
water, and then replacing the water with paraformaldehyde/PBS, and squashing.
The carcasses help prevent complete crushing of the relatively small number
of embryos. I usually do about 5-10 worms/slide. More than that, and the embryos
seem to not stick as well as you cut open more worms.
After 20 minutes of fixing, freeze slides by placing on a nice smooth surface of dry ice in an ice bucket (or an aluminum block embedded in crushed dry ice). Hold down with finger tips until frozen (you can see a wave of ice crystals go across as they freeze when viewing at proper angle.). Let sit for 5 to 20 minutes frozen. Remove slide from dry ice and quickly pop off cover slip and immediately put slide into room temp 100% Methanol (not necessary to use anydrous beads, but they probably don't hurt). Let sit for 4 minutes; transfer to room temp Tris Tween or PBS. Let sit 4 minutes, and then you can begin the staining. I've tested lots of other procedures (rehydrating through a series of methanol/water steps, different temperatures, different amounts paraformaldehyde, acetone (completely kills staining with SKN-1 antibodies), mixing paraformaldehyde with methanol at different temperatures etc.) and this definitely worked best for me. Let me know if you find that other conditions work better for you and your antibodies. In general, we find that large, structural or membrane associated proteins stain well with methanol/acetone protocol, but some small nuclear proteins we've tried required paraformaldehyde fixation, are killed by acetone, and stain best using this protocol. [Note: This procedure tends to irritate my eyes due to the paraformaldehyde. Probably best not to wear contact lenses (although I usually do).
5. Staining embryos.
Remove slide from Tris Tween (or PBS) and use a kimwipe to dry off bottom and
top, leaving a moist film covering the embryos. You can pre-block if you wish
(we use 1% BSA in PBS), but in general for MAb's in medium with serum, and for
dilutions of rabbit or mice serum in 1% BSA/PBS, we don't bother. But when we
do, we block for 30 minutes at room temp. Remove the BSA by wiping with a kimwipe
until a thin film of moisture remains, and add antibody (we typically use 20-25
ul of whatever dilution). Incubate in a humidity chamber for 1 hour at room
temp (or overnight at 4ęC if you prefer). If you dry the slide off everywhere
but where the antibody drop is, and keep the humidity chamber on a level surface,
the antibody drop will remain over the embryos. Alternatively, one can gently
lower an 18x18mm cover slip on top of the embryos, and then use a squirt bottle
of Tris Tween and squirt gently around edges of coverslip to help loosen it
and float it off when you are ready to wash. Place the slide in Tris Tween (we
use Coplin jars). Wash three times for 4 minutes each. Again, wipe off excess
moisture around embryos, and add secondary Ab (we like Tago antibodies very
much, FITC or rhodamine conjugated). Incubate in humidity chamber at room temp
for one hour. I typically use 1/100 dilutions (in 1%BSA in PBS) with 50 ul drops
on the slide, and we don't use a cover slip since getting cover slip off usually
causes more loss of embryos each time). I find almost all antibodies that are
going to stain do so within a half hour at room temp, and do one hour for peace
of mind. I often do overnight at 4ęC for primary antibody; 37ęC works fine if
that makes you feel better. Wash secondary as before, except after last wash
go into PBS with DAPI (we add 1 ul of 1 mg/ml DAPI to a Coplin jar (about 50
ml of PBS) and then into PBS without DAPI for 1 minute. Remove slide, use kimwipe
to wipe off excess liquid, put on a 22 x 22 coverslip, wick out excess liquid
(thoroughly), add mounting medium, and seal with fingernail polish.
Good luck.
Barbara Page modifications for SKN-1 fixation/staining:
e-mail: bpage@nsit-popmail.uchicago.edu
Here's the procedure I use and it works beautifully for me.
embryos are squashed in 4% paraform. buffered with 60mM pipes pH 6.8; 25mM hepes,
pH 6.9; 10mM EGTA; 2mM MgCl2
the slides are kept at room temp for 5 minutes to fix, then frozen on dry ice,
the coverslip cracked off and the slide is placed in -20 degree solution of
DMF for 3 minutes. Then the slide is washed in PBS, 3X and blocked with 10%
goat serum for 30 minutes. Extra fluid in drained off (but slide is not washed)
and anti-SKN-1 serum is added and incubated at 4 degrees for over night-the
rest you know.
This procedure allows me to get staining without as much squashing as other
methods I've tried. Also the little bit of remaining goat serum on the slide
helps block. (but you may want to order horse serum-I think the background is
even lower with that serum)
Part of the reason I see such good SKN-1 staining could be due to the secondary
I use. It's from Jackson lab;it's Cy3-conjugated anti-mouse IgG. The lot number
I have is 42729 and it is NOT specific to IgGs. I've tried others and it is
by far the best.