[Histonet] RE: Questions about IHC in Frozen Sections

Fri Dec 6 08:12:13 CST 2013

I agree with Liz,

We usually fix with acetone/ethanol 5-10 min then go right into buffer, but occasionally use 2.0% NBF for some antibodies.  Our buffer contains 0.1% Tween and our sections can be anywhere from 8-20um depending on the specific project. I think the 30min in acetone is messing up your morphology.

Brett

Brett M. Connolly, Ph.D.
Principal Scientist, Imaging Dept.
Merck & Co., Inc.
PO Box 4, WP-44K
West Point, PA 19486
brett_connolly <@t> merck.com
T- 215-652-2501
F- 215-993-6803

-----Original Message-----
From: histonet-bounces <@t> lists.utsouthwestern.edu [mailto:histonet-bounces <@t> lists.utsouthwestern.edu] On Behalf Of Elizabeth Chlipala
Sent: Thursday, December 05, 2013 5:59 PM
To: Lewis, Patrick; 'Histonet <@t> lists.utsouthwestern.edu'
Subject: [Histonet] RE: Questions about IHC in Frozen Sections

Patrick

Here is what we do for frozen IHC, this is based upon methods that I received from Gayle Callis.

Cut frozen sections and let air dry - at least 20-30 minutes post the last section cut.  If we are going to stain that same day or the following day we leave the slides at room temp (we are pretty dry here in Colorado) but if you have issues with humidity you can store them in a dessicator overnight.

If you need to store at -80 then we package the slides in smaller slide boxes and only package enough slides for one run to avoid freeze and thaw artifact.
So once the slides have dried we place them in slide boxes and in those slide boxes we add a small or medium nylon tissue bag that contains Silica Gel, 6-16 mesh (indicating) we just staple the nylon bag shut.  
We then use a food sealer to seal the slide box in one of those food sealing bags (we got ours at Cost Co they have them on sale every once and a while, along with the bags) and then that goes in the -80 for storage.

The day before we are going to stain we pull out the sealed slide box from the -80 and let it sit on the counter top until the next morning when we open up and then fix with the best method for the particular IHC that we are going to use it could be 10% NBF or 4% paraformaldehyde or one that Gayle recommended to us - its an ethanol/acetone mixture - the protocol is listed below.

1.  Fix for 5 minutes in solution made of 75% Acetone and 25% Absolute Ethyl Alcohol.

NOTE:  We purchase Absolute Ethyl Alcohol in the small bottles.  Both Acetone and Absolute Ethyl Alcohol are both stored in the flammable storage cabinet.

2.  Rinse in two buffer changes for at least 2 minutes.
3.  Continue with staining protocol.

Good Luck

Liz

Elizabeth A. Chlipala, BS, HTL(ASCP)QIHC
Premier Laboratory, LLC
PO Box 18592
Boulder, CO 80308
(303) 682-3949 office
(303) 682-9060 fax
(303) 881-0763 cell
liz <@t> premierlab.com
www.premierlab.com

Ship to Address:

Premier Laboratory, LLC
1567 Skyway Drive, Unit E
Longmont, CO 80504

-----Original Message-----
From: histonet-bounces <@t> lists.utsouthwestern.edu [mailto:histonet-bounces <@t> lists.utsouthwestern.edu] On Behalf Of Lewis, Patrick
Sent: Thursday, December 05, 2013 3:29 PM
To: 'Histonet <@t> lists.utsouthwestern.edu'
Subject: [Histonet] Questions about IHC in Frozen Sections

Hi Everyone.

I am trying to troubleshoot  my IHC on frozen sections.

My sections are human tonsil at 7 uM. On charged Superfrost slides.

They are stored at -80 after drying for 1 hour.

When I use them for IHC, I take them out of the -80 and let them air dry for 1 hour before placing them in cold acetone for 30 minutes to fix.

Question:
If I place them directly in H20 or TBST pH 8.0 after fixation, will that cause cell lysis?

Should I dry the slides after acetone fixation before washing them?

If so, for how long?

My problem seems to be that the tissue is getting digested on the slide, I am trying to trouble shoot which step is causing my tissues to disintegrate.

So far I have tried thicker sections 10, 15 uM (That made the problem worse, I am consider going back to 4 uM sections)

I also Changed the concentration of H2O2 for my H202 block from 3% to 0.3%, (In my next IHC attempt I will try to examine the slide at each step to see if I can see loss of integrity)

Also in my next attempt I plan to eliminate any H20 washes and dry the slide post acetone fixation before washing in TBST.

Also I plan to decrease the amount of Tween20 in my Wash buffer from 0.2% to 0.02%.

Any advice would be helpful.

Patrick.

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