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(Paraformaldehyde
fixed tissue, see below for non-fixed tissue*)
Notes before you begin:
In our hands, for best results, samples should be paraformaldehyde
fixed and frozen using 2-methylbutane cooled with liquid nitrogen (see
Tissue Fixation and Freezing protocol).
After freezing, samples should be stored at -80oC. Samples to be
transferred between labs or waiting sectioning should be stored on dry
ice or in the cryostat chamber and never allowed to thaw. Thawing will
affect section ability, tissue quality and staining.
Unlabeled, frozen sections should be stored at -80oC or -20oC. Long
term storage (over 2 weeks, especially in a frost-free -20 °C
freezer) is not recommended as samples tend to lyophilize (freezer
burn).
Controls: All
experiments should include a sample that has been treated to control
for non-specific primary and secondary binding. Some controls include:
*Secondary antibody
only for secondary antibody non-specificity.
*Non-immune serum,
isotype control or non-immune IgG as the primary control antibody.
*Infrequently, some
tissues display autofluorescence, you may need to have a non-processed
sample to control for this. Examples: elastin in elastic and muscular
arteries autofluoresce in the green channel. Paneth Cells in intestinal
crypts autofluoresce in the red channel.
Protocol
1.) Obtain cryostat sections (~6 microns) and put onto glass slides.
Some tissues (e.g. blood vessels, brain, fat) may require gelatin
coated or poly-lysine coated slides to remain attached to the slide
after multiple wash steps. An expensive, but good solution is to
use Superfrost Plus Stain Slides (from, e.g., Fischer)
2.)Keep slides at –20oC until ready for use (see above).
3.)Rehydrate tissue sections with 2 washes of 1xPBS, keep in liquid at
all times until sample has a gelvatol adhered cover glass
4.)Depending on your protein or cell type of interest, detergent
permeabilization may be necessary (10 minutes with PBS + 0.1%Triton
X-100).
5.)BLOCK with 2% BSA for 45 minutes. Alternative blocks are using 20%
serum in PBS+0.5% BSA (PBB) from the species in which your secondary
antibodies are made (e.g. if you secondary antibody is made in goat,
use 20% goat serum. Likewise if your secondary is made in donkey, use
20% donkey serum).
6.) Wash sections with 5 times with PBS+0.5% BSA (PBB).
7.)Primary antibody: Dilute to the desired concentration in PBB (vortex
gently, spin down for 5 min at 10-12K RPM to get rid of aggregates).
Gently drop 70-100 µl antibody solution over the section (so the
section is all covered). Incubate for 60 minutes. Longer primary
antibody incubations may be necessary and must be determined
empirically in your system. Initial antibody optimization should
be accomplished in a range up to 5ug/ml antibody in PBB. If
longer incubation times are necessary, place a piece of wet paper towel
in your slide box (or chamber) with your samples to minimize
evaporation of your primary antibody solution. In some cases
overnight incubations at 4oC are necessary, you must put your samples
in a humid environment (wet paper towel within slide box), avoid
disturbing the sample volume on your section. Primary antibodies can be
added together, but the host must be from 2 different species (unless
the primary antibodies are directly conjugated with the fluorophore).
8.) Wash sections 5 times with PBB.
9.)Secondary antibody: Add secondary antibody to the section for 60
minutes (made in PBB, vortex, spin down at 10-12K RPM for 5 minutes to
pellet any aggregates – very important). The secondary
antibody step should be kept to within an hour. You may add
fluor-conjugated phalloidin (for F-actin counterstain) or Draq5 (a
far-red nuclear stain from Biostatus) to your secondary solution.
Secondaries can be added together if using more than one, but they must
be against two separate species.
10.)Wash sections with 5 times with PBB.
11.)Wash sections with 5 times with PBS to remove the BSA that would
bind Hoechst indiscriminately.
12.)Add Hoechst stain 30 seconds to stain the nuclei (Hoechst stain=1
mg/100 ml dH2O, Sigma CatNo.B-2883). Caution! This substance is
carcinogenic and toxic to skin!!
13.)Wash sections 3 times with PBS.
14.) Adhere cover glass over your sample with gelvatol, place
slides horizontal in slide box, and allow cover glass to
adhere to slide overnight at 4oC in the dark. Gelvatol is water
soluble, if needed coverslip may be gently removed by submerging sample
in water and allowing coverslip to lift without resistance from the
slide. Recipe for Gelvatol on protocol website.
BLOCK=PBS+2% BSA = 2g BSA per 100 mL 1xPBS
PBB=PBS+0.5% BSA = 0.5g BSA per 100 mL 1xPBS
In some cases
non-fixed tissue is used for frozen sectioning. After sectioning,
tissue can be fixed in 2% Paraformaldehyde in PBS for 30 min-1 hr*.
Alternatively sections can be fixed in cold
100% methanol or acetone (-20oC), which will precipitate the proteins
in the tissue. Protocol below:
1. Immerse sections in cold (-20oC) solvent for
10 min.
2. Remove and let air dry.
3. Store these at 4oC (1-2 days) or -20oC (long-term)
until processing.
4. Rehydrate tissues with PBS and wash 3 times in PBB.
5. Proceed as above for paraformaldehyde fixed
tissues.
Note: Tissues fixed in this manner will NOT counterstain with
phalloidin.
There are many other permutations and other options available when it
comes to immunofluorescence protocols. The ones described above are
basic and work for most applications. However if you have any
questions, or are experiencing problems, please contact a CBI staff
member for assistance.
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