gcallis <@t> montana.edu
Thu Nov 9 11:28:01 CST 2006
I presume you are referring to the publication in J of Histochemistry
Cytochemistry on this UV light exposure method to get rid of
autofluorescence? I have seen a message on Histonet where they tried this
and did not have success.
I will be sending you a pdf on review of autofluorescence privately if you
want it. Also, go to IHC World website, find discussions on fluorescence
and autofluorescence - there are also other ways to attempt elimination
of the problem. I did a copy/paste document from Histonet archives of a a
superb discussion on ways to get rid of autofluorescence, and am forever
grateful to the nice gentleman who put this together. Please excuse
repetitive statements, it may have been composite of two messages(?).
We use 100 mM glycine, in Dulbeccos PBS at 7.4 on rehydrated tissue
sections, NOT frozen sections. With frozen sections we never prefix the
tissue, they are cryosectioned in fresh state, then solvent fixed for
immunofluorescence staining including anti GFP work (our fixative ruins the
GFP) - this way we avoid any possiblity of aldehyde induced
autofluorescence. Narrow band filters help as do spectral imaging
instruments which can remove autofluorescence, as one can do on a confocal
laser scanning microscope i.e. "gate it out" so to speak. Glycine method
can also be used on a tissue coming out of fixative (soak in 100 mM glycine
1 hour, then rinse well with buffer, proceed to processing and maybe even
before cryoprotection/snap freezing. We have not tried the latter.
Your assessment on too thick sections is correct.
1. After dehydrating your paraffin embedded slides and before blocking for
protein incubate them in 100 mM glycine for 20 minutes. This will quench
autofluorescence caused by free aldehydes. Works like a charm, I use it.
2. I've pretty much moved away from doing IF on paraffin
embedded stuff(ICC yes) but I have had some experience trying to knock
back autofluorescence (endogenous and fix related) in tissue sections.
I've had some luck pre-treating fixed sections w/ one of the following:
1. 50mM Ammonium chloride in PBS for 10 min.
2. 0.1M Glycine in PBS, pH 7.4 for 5-10 min.
3. 1% Sodium borohydride in PBS for 10-20 min.
It varies from sample to sample which method works the best but I've had
the most success w/ the borohydride and NH4Cl methods.
Autofluorescence can be brought on by certain endogenous tissue
constituents, ie. fibronectin, lipofuscin and elastin, as well as by
fixation in aldehydes.
You don't say if your sections are fixed or not. If so, you should look
at using sodium borohydride (0.5mg/ml in PBS) for 5 minutes
(glutaraldehyde)or PBS plus a few drops of 1M glycine(formaldehyde) to
block any reactive groups. ***Sodium borohydride is flammable on contact
with water, and harmful by ingestion, inhalation etc. Take adequate
Another thing to consider is reducing the section thickness, if possible,
as the intensity of autofluorescence is related to this.
You also don't mention what fluorochromes you are using. It may be
worthwhile trying a fluorochrome of a longer wavelength as there is less
likelihood of any spectral overlap with the endogenous material. As I
mentioned in an earlier posting today, we have had good results switching
to the Alexa dyes (Molecular Probes).
There are a couple of simple things you can do to help reduce
Some of the chemical reactions causing autofluorescence occur most rapidly
with higher temperatures and on exposure to light. Therefore, performing
the labeling at 4 C in the dark can help reduce this problem.
Autofluorescence intensity is related to section thickness. You may want to
try thinner sections if at all possible. Sometimes using fluorophores
excited at longer wavelengths can help diminish autofluorescence.
If autofluorescence is still an issue, there are a few preincubation steps
you could try.
A Tris-glycine mixture (adjust 0.1M glycine to pH 7.2-7.4 with 1M Tris
base)will saturate free aldehyde groups. (15-30 minutes at room temp in
Tris-glycine. Wash well in PBS
The use of 1% sodium borohydride in PBS helps reduce any free aldehyde
groups in the tissue, making them non-reactive. Incubate sections for 30
minutes in borohydride and then wash well(minimum 15 minutes) in several
changes of PBS. Proceed with labeling.
These techniques can be used alone or sequentially. If the tissue is
fragile though, only use the Tris-glycine method. Please note that sodium
borohydride is very reactive and is flammable on contact with
water. Another technique to block unreacted groups is to incubate sections
for 5 minutes in 50mM NH4Cl, and rinse in PBS before labeling.
At 08:43 AM 11/9/2006, you wrote:
>My name is Stephen Clark and i work in the neuro lab at Eastern Illinois
>University. We use fluorescent immunohistochemistry to stain the
>Olfactory Epithelium of mice and have recently had a problem with
>autofluorescence of our tissues. They are fixed in 4% paraformaldehyde in
>PBS and i have already tried sodium borohydrate with little success. I
>have read about photobleaching using UV, neon, and lights of specific
>wavelengths (488 and 633nm). I am just wondering if anyone utilizes
>photobleaching via a light source and where it would be possible to
>purchase them. I am also wondering if the 18um sections might be a little
>too thick and whether that would have an increased affect on
>autofluorescence. Any tips would be appreciated.
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