Stephen,

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.





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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 
precautions***
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 
autofluorescence.
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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