IMMUNOPEROXIDASE STAINING
Base solution: 4% BSA in PBS with 5% Non Fat Dry Milk (NFDM).
Blocking solution: 10% serum in base solution. The choice of serum is dictated by the
secondary antibody. Use serum from whatever species the secondary was raised in (i.e. horse
serum for biotinylated horse anti-mouse).
Rinsing or wash solution: 0.5% BSA in PBS.
Primary and Secondary (biotinylated 2° Abs from Jackson ImmunoResearch or Vector Labs)
solutions are almost always made in the full blocking solution, the exception being some
lectin or anti-CHO antibodies whose staining is blocked in the presence of NFDM. 2°
used at 1:50-1:150. If staining rat tissue, use 2% normal rat serum in blocking solution
for 2°, and if using a monoclonal on rat, use the special rat tissue-adsorbed version of
bHaM from Vector.
Tertiary, i.e. avidin–biotinylated HRP (Vector labs, Elite kit) is made up in the Base solution 30
minutes in advance at a concentration of 1:50; add avidin to solution and mix, then add
bHRP and mix.
DAB tetrahydrochloride (potential carcinogen): 2 mg/ml in 0.1 M citrate-HCl buffer, pH 5.1
50 ml/coplin jar: 1.47 g sodium citrate/50 ml Double distilled water -200 ml/staining dish: 5.88 g sodium citrate/200 ml dd water -adjust pH to between 5.05-5.15 with concentrated HCl, dissolve 100 mg or 400 mg DAB
in solution, respectively, vacuum filter with P2 filter paper.
Just before use add 16.8 µl of 30% hydrogen peroxide/50 ml of DAB-citrate or 67 µl hydrogen
peroxide/200 ml.
PROCEDURE:
Paraffin sections: deparaffinize and hydrate to water (4 changes of Xylenes 4 minutes each, 2 X
100% EtOH, 95%, 70%, 3 minutes each , 2 changes of tap H2O, then hold in distilled
water)
Frozen sections: general: 5 minutes in PBS to remove OCT then up and down in graduated
alcohols (70%, 95%, 100%, 95%, 70% -- 1 minute each), PBS -- 5 minutes.
Check protocol for pretreatments if any. There are various pretreatments for different antibodies.
Block for 10 minutes, Room Temperature (R.T.)
Primary Ab: 1.5 hour, R.T. or overnight at 4°C.
Rinse 3 x 5 minutes in BSA-wash
Secondary Ab: 1 hour R.T.
Rinse 3 x 5 minutes
Tertiary reagent: 45 minutes R.T.
Rinse 3 x 5 minutes
DAB: 8 minutes
Rinse well with water
Dehydrate through alcohols and clear with xylene
Coverslip with DPX
Notes Regarding Procedure:
1) Fixation of the tissue: We routinely use three types of material in the lab.
(i) Bouin’s fluid-fixed (75 ml saturated picric acid [Sigma], 25 ml formalin
[commercially prepared 37% formaldehyde, with preservatives], 5 ml acetic acid:
Perfusion fixation (push and drip), some postperfusion immersion fixation (4-8 hrs),
either cryostat or paraffin sectioned, also some whole mount stuff. Very good for OMP,
GAP-43 and TuJ-1 staining, particularly on cryostat sections for the last two. Best for
routine histology of the olfactory epithelium.
(ii) Periodate-lysine-paraformaldehyde (PLP)-fixed (with the concentration of
paraformaldehyde varying between 0.5 and 2%; McLean and Nakane, 1974, J.
Histochem. Cytochem., 22: 1077): Perfusion fixation (push and drip), cryostat
sectioning. Very good for general screening. Most antigens stand up to this fixative at
0.75% formaldehyde and allow some degree of staining (the exception being some antikeratin antibodies), including vimentin and neurofilament antibodies (maximal sensitivity
is seen with organic solvent fixed and cryostat sectioned in these cases, but the fixation is
good enough that the tissue looks OK). It also permits decalcification with saturated
EDTA (see below) prior to OCT embedding and cryostat sectioning. Very good for
NCAM, other cell surface antigens.
(iii) Organic solvent-fixed (either Carnoy’s solution [60 ml absolute EtOH, 30 ml
chloroform, and 10 ml glacial acetic acid] or a commercial preparation called Omni-Fix II
from An-Con Genetics, Melville, NY), paraffin-embedded: Push some fixative through
after clearing out blood with PBS, then immersion fix for 4-18 hours, depending on the
size of the tissue. Used for BrdU staining, some intermediate filaments (with
pretreatments, see below). For those very refractory intermediate filament antigens, can
sometimes get away with organic solvent fixation, rehydration, decalcification and
cryostat sectioning, and the outcome is better than trying to cut undecalcified, unfixed
tissue, but still not great. This tissue doesn’t hold together very well.
2. Decalcification: We routinely strip the heavy frontal and nasal bones off the nasal skeleton
and olfactory bulb after fixation and then use two forms of decalcification to decalcify the
smaller turbinate bones that remain.
(i) Formic acid-sodium citrate solution (Armed Forces Institute of Pathology Manual -AFIP): used for Bouin’s-fixed tissue and after rehydration of the organic-solvent fixed
tissue in the case of some antigens). 8-24 hours, depending on the age of the animal.
(ii) Saturated, neutral EDTA: per 900 ml of ddH2O, add 120 g of disodium EDTA, then
SLOWLY add 30 g of NaOH pellets to a pH of 7.3. Everything should go into solution.
Requires 3-5 days at 4°C to decalcify the turbinates, but needed for those antigen that
won’t stand up to acid decalcification.
(iii) RDO: Commercially available Rapid decalcification solution from . A
After decal, tissue is either cryoprotected with 30% sucrose, until the tissue sinks or 48 hours, or
dehydrated for paraffin-embedding. For the cryostat material, the freezing step is critical for
good histology (if too slow, you get a bad “Swiss cheese” freezing artifact). Because orientation
and good preservation of structure is important to us, we form an aluminum cup by wrapping a
thin layer of foil around a 12 mm tube, and sealing the bottom by twisting. The tissue (maximum
size of the block is 20-25 mm long, by about 7-10 mm in diameter -- i.e. the size of the bulb and
nasal skeleton of a rat up to the anterior tips of the 2nd endoturbinates) in the cup is surrounded
by only as much OCT as is needed to cover it over, then immersed to the level of the OCT in
liquid nitrogen. Do not immerse completely, or else the block will crack. Watch to make sure
that the top is the last to freeze. It should, at the end, look a bit like the top of a soft ice cream
cone, with a swirl that is extruded as the deeper part of the OCT freezes and expands. Remove
the cup from the liquid nitrogen and put in a -80° freezer when the top is not completely frozen,
i.e. when the center 3 mm or so is still liquid at the top. Too long in the liquid nitrogen causes
cracking, too little causes freeing artifact.
3. Sectioning -- routine is 8 m for cryostat and 5 m for paraffin, can go down to 3-4 and 2,
respectively. Mount either on “Plus” slides from Fisher.
4. Pretreatments of the tissue prior to staining.
(i) The alcohol dehydration and rehydration of cryostat sections improves penetration of
antibodies into the tissue and lowers background by extracting lipids. Alternatively can
use acetone for more complete delipidation. Do not use if staining for a ganglioside
antigen.
(ii) Methods to restore antigenicity of tissue that is too harshly fixed or paraffinembedded have generally involved protease treatment (proteinase K at 20 g/ml for 5-20
min). In many cases we have found a simple 20 min pretreatment with 0.1-1 M NaOH to
be as or more efficacious, besides being faster, easier and cheaper (presumably breaking
some of the cross links and producing some degree of proteolysis -- like in the Lowry
protein assay). For example, staining of solvent-fixed, paraffin-embedded sections with
anti-cytokeratin 5/6 antibodies is very poor without this NaOH step and very good with it
(see JCN 359: 15-37 and 363: 129-146). More recently other labs have turned to some
microwave techniques to open up the tissue.
(ii) Steamer method of antigen retrieval
5. Modifications for immunofluorescence.
The protocol is much the same for immunofluorescence staining as for IP. We have
found that the optimum concentration of the primary antibody for IF staining using tagged avidin
or streptavidin is 4 times that used for IP staining. If double-labeling is required, use the Jackson
IR secondaries that have been raised in donkey and are specifically cross-adsorbed for these
purposes.
Most recently we have found it very useful to have a double filter set cube (available
directly from Chromega in Burlington, VT or via your microscope rep) that allows you to directly
visualize Texas red and fluorescein simultaneously. This is obviously useful for double-labeling
experiments, but it also very good for low-intensity staining with a single marker because the
background autofluorescence is a different color than the specific staining (generally it also
reduces the level of tissue autofluorescence to near nil). In addition, with this dual cube you can
actually do double labeling with antibodies raised in the same species (like two mouse
monoclonals) by sequentially cycling them through the two different tagged avidins (Ab1
through to Av1 then Ab2 through to Av2). The first round deposits Av1 (FITC, for example)
around Ab1, the second round deposits Av2 (Texas red) around Ab1 and Ab2. With this dual
cube, Ab1 visualizes as yellow and Ab2 visualizes as red, which are discriminable by
photography. Obviously, this won’t work if the antigens co-localize at a cellular level, but is
quite useful if you need to identify more than 1 cell population on the same slide. A word of
caution, although they do sell triple cubes, for simultaneous viewing of FITC, Texas red and UV,
UV visualization is really lousy, because the extra glass in the filter causes a huge decrease in
UV signal intensity.
The other difference is in mounting. IF slides cannot be dehydrated, cleared and covered
with DPX. There are a variety of anti-fadent mountants. Vectashield, from Vector is very good
for FITC and Texas red (does a better job of suppressing fading than anything I’ve used before).
However, the anti-fadent fluoresces with UV wavelength illumination blocking the visualization
of any blue signal from AMCA-labeled Abs, DAPI or Hoechst. If you need a blue label, you can
use para-phenylenediamine as the antifadent in 10% PBS-90% glycerin (1 g/ml final
concentration; Johnson and Nogueria Araujo, 1981, J. Immunol. Meth. 43: 349).
6. Sequential double labelling with IP
We have found that you can achieve reasonable double-labeling wit IP material if both
antigens are robust. The first round of staining proceeds to Av:bHRP and DAB; the second
round proceeds to Av:bHRP and a commercial substrate, either SG or VIP, from Vector. The
deposition of DAB blocks access of antibodies and other reagents during the subsequent round of
staining. The deposition of a thin coating of DAB also reduces the intensity of the staining with
the second antibody when compared to the second antibody used alone (therefore, do the weaker
staining antibody first). The slides must be rapidly dehydrated and coverslipped with DPX at the
end because both SG and VIP products are weakly water soluble. DAB must be used first, for
the same reason (the SG and VIP will fade during a subsequent cycle of staining). You cannot
treat the sections with NaOH or alcohols after the DAB in between the rounds of staining as this
exposes the underlying antigen-antibody complex for deposition of the reagents used during the
second round of staining.
Renee Mezza/Jim Schwob
Jan 19, 1996
revised April 30, 1999