[Histonet] Autofluorescence on Retina Tissue

[James Watson]

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
jwatson <@t> gnf.org


-----Original Message-----
From: histonet-bounces <@t> lists.utsouthwestern.edu [mailto:histonet-bounces <@t> 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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