Not sure if this will help with your retinas but we use the copper sulfate
treatment on all our fluorescent stains and it eliminates autofluorescence in
the RBC completely and helps with other types of autofluorescence. The
treatment to be use depends on the cause of the autofluorescence. We counter
stain our slides with DAPI. Here are all of our present autofluorescent
treatments.
Autofluorescence Quenching
1) The best ways to address the issue of autofluorescence in order of
preference:
a) Avoid it
i) Not always possible.
b) Try to filter it out during image acquisition
i) Difficult due to the broad emission spectrum.
c) Chemically remove it
i) Can also reduce "real" signal.
2) 10 mM Copper Sulfate
a) This treatment is primarily for inhibition of autofluorescence
in Red Blood cells, but helps decrease autofluorescence in connective tissue.
b) 10 mM Copper Sulfate
i) Cupric Sulfate...............1.25 gm.
ii) 50 mM Ammonium acetate (pH5)........500.0 ml
iii) Adjust pH to 5.0 with 1.0 M NaOH
c) 50 mM Ammonium acetate (pH5)
i) Ammonium acetate.............1.93 gm.
ii) Distilled water................500.0 ml
iii) Adjust pH to 5.0 with 1.0 M HCl
D) Treatment Procedure
i) Rinse in PBS 2 times for 10 minutes each.
ii) Rinse in distilled water 5 minutes.
iii) Place slides in 10mM copper sulfate for 8 minutes.
iv) Return slides to distilled water and check for
autofluorescence with microscope.
v) If needed return slides to 10mM copper sulfate for a
couple of more minutes and check again.
vi) Rinse slides for 5 minutes in distilled water.
vii) Counterstain with DAPI 5 minutes.
viii) Rinse in PBS 2 times for 10 minutes each.
ix) Coverslip slides with appropriate mounting media.
3) 2.0 mM Glycine
a) Used primarily for autofluorescence caused by free aldehyde
groups.
b) 2.0 mM Glycine
i) Glycine...................3.9 gm.
ii) Distilled Water...............26.0 ml
c) Treatment Procedure
i) Deparaffinize and rehydrate slides to H2O
ii) Rinse in PBS 2 times for 10 minutes each.
iii) Rinse in distilled water 5 minutes.
iv) Place slides in 2.0 mM Glycine for 20-60 minutes.
v) Rinse slides for 5 minutes in distilled water.
vi) Rinse in PBS 2 times for 10 minutes each.
vii) Continue with normal staining procedure
4) Sodium Borohydride
a) The use of this reagent is particularly suited to reduce the
reversible Schiff's bases that are formed by the aldehyde-NH2 reaction and lead
to autofluorescence, especially when using glutaraldehyde. If you can use
paraformaldehyde for fixation, the reduction step is often unnecessary and
autofluorescence is low. This material has a high potential for
explosion and is very caustic.
b) The protocol was prepared by Jennifer Kramer and a similar
procedure is described by Beisker, et al. (Beisker et al. 1987).
i) Immediately before use
(1) Make up a 1 mg/ml solution of sodium
borohydride in a physiological buffer such as PBS.
(a) The solution will be fizzy like
carbonated water. Preparing this solution on ice and performing all subsequent
incubations on ice has also been recommended.
ii) Apply this solution immediately (while fizzing) to
cells or tissue sections.
(1) For glutaraldehyde fixed cell monolayers
incubate in the sodium borohydride solution for 4 minutes. Replace with fresh
sodium borohydride solution for another 4 minutes.
(2) For paraformaldehyde fixed paraffin embedded 7
μm sections incubate 3 times, 10 minutes each in sodium borohydride solution.
5) Sudan Black
a) This treatment is primarily for inhibition of autofluorescence
in Lipfuscin.
b) 0.3% Sudan Black (w/v) in 70% EtOH (v/v) stirred in the dark
for 2 hours
c) Apply to slide for 10 minutes after the secondary antibody
application.
d) Rinse quickly with PBS 8 times and mount
e) For FITC and Alexa 594 this does not reduce the emission signal
noticeably
James Watson HT ASCP
GNF Genomics Institute of the Novartis Research Foundation
Tel 858-332-4647
Fax 858-812-1915
jwat...@gnf.org
-----Original Message-----
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Allyse
Mazzarelli
Sent: Tuesday, October 15, 2013 11:44 AM
To: Histonet
Subject: [Histonet] Autofluorescence on Retina Tissue
Hi all!
Question for all you that may be more immuno-experienced than I:
I've consistently run immunofluorescence on pig retina and I seem to have a
severe case of autofluorescence/background. I've played around with primary and
secondary antibody ratios, but that doesn't seem to help my case. The primary
antibodies I use are goat-anti-FLT.1, and mouse-anti-rhodopsin. The secondary
antibodies I use are AlexaFluor donkey-anti-goat488 &
rabbit-anti-mouse555 (For my experiments, each slide contained only one primary
antibody, and it's corresponding secondary. For imaging purposes, I did not
double-label on these slides. E.g. FLT.1 was labeled with the 488 secondary,
and rhodopsin was labeled with the 555 secondary).I re-hydrate, conduct antigen
retrieval, and block as per normal IHC protocol. However, when imaging, I
noticed that both slides, labeled with either rhodopsin or
FLT.1 seem to "bleed" through to the next filter. For example,
mouse-anti-rhodopsin labeled with the 555 secondary works beautifully at a
1:600 ratio. However, when I switch to the FITC filter on my scope, all the
retinal tissue appears green on the slides, even though it has ONLY the 555
secondary and NO 488. I've noticed this for the FLT.1 antibody as well (i.e.
switch to red filter and tissue fluoresces even though no slide saw the 555
secondary antibody).
As I mentioned, I decreased the ratios of all antibodies, but that still
doesn't eliminate the problem.
If anyone has any ideas as to how I go about fixing this, please let me know.
I've only been in the field for a very short period of time, so if I missed
something in my description, don't hesitate to ask! Thanks for whatever help
you can direct my way!
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