[Histonet] RE... Frozen section fixation problems

Thu Apr 30 18:19:22 CDT 2015

I have been following this with interest both now and in the past.  

A word of caution about the acetone/ethanol fixation.  I did NOT use the
acetone/alcohol fixative cold, but at RT (as it was taught to me by an IHC
expert).   That is a bonus since you don't have to maintain A/A fixative in
a refrigerator.   It could be that in Brett's hands, A/A at -20C works well
so I can't argue with a successful variation for this fixative.    A major
caveat:  A/A is used for rodent CD markers and cannot be used for human CD4
or CD8 as reported by the late Dr. Chris van der Loos.  He and I
collaborated about frozen section fixatives many times along with trying
each other's method.   He always had success with 4C acetone in very humid
The Netherlands but was careful to air dry the sections overnight in front
of a fan.    These two human CD markers do not tolerate ethanol
consequently, I wouldn't use A/A for any human CD marker work.    We  have
used it exclusively for murine and rat CD markers and Q-fever organisms.   A
good rule it to have a panel of fixation methods in order to optimize
fixation for any given antigen.   

I do not understand why Patrick has such problems using cold acetone
fixation which leads to poor sections.    We air dried frozen sections for a
minimum of 30 min before A/A fixation.   Most of the time, frozen sections
were cut and immediately dried at RT for up to 4 hours, then stored in a box
containing only one day's worth of staining.  The unfixed sections are
stored at -80C with a bag of silica gel in the box (25 slide capacity).  The
slide box can be taken out the night before staining, or even the day of
staining with lid on to NOT GET WATER CONDENSATION ON THE SECTION.   Water
condensation can damage morphology and antigens.   I would NEVER use an
acetone gradient for fixation since the increase in water could be a cause
of the damage.  Water is not going to maintain isotonic conditions and
prevent damage.   If you want to blow away a frozen section after acetone
fixation, just rinse with water........a sure way to damage the morphology.
After acetone fixation only (10 min at 4C), air dry section for 15 min, then
go into  PBS or TBS.     

Before fixation, use barrier pen i.e. ImmEdge (vortexed to mix components
before drawing around section) from Vector around section, then fix in A/A
10 min @RT and then go immediately from A/A into pure PBS for 3 changes.
The 4th change is PBS/0.2% Tween 20 to equilibrate the section for IHC
buffer conditions. 

What I suspect, after so many continued problems, is the snap freezing of
the tissue may be done improperly and the damage could be excessive freezing
artifact.   Something is amiss and it may be BEFORE FIXATION with the
acetone.   

In general, I have found methanol to be a poor fixative for IHC, and should
be totally avoided for any CD marker work since it causes protein hydrolysis
of the epitope causing weak, poor staining.   4% paraformaldehyde @ 4C
without antigen retrieval can give weak staining and antigen retrieval with
frozen sections has to be done carefully to maintain delicate sections on
the slide.    95%, even 100%, ethanol can also result in weak staining.   

You did not say what epitopes you are trying to preserve and stain for?   I
don't think the plus charge slides are the culprit since I had labs using
acetone fixation of FS on plain glass slides before Plus charge was so
popular.  Are you sure your PBS or TBS is correctly made?   Incorrectly made
PBS  caused morphology havoc to completely blow away my frozen sections.
This led to purchasing Sigma Dulbecco PBS which never gave problems.      

Maybe you can describe more of what you are doing from the time you receive
and snap freeze the tissues, species, etc., including manual or automated
staining in order to have other help you chase away these annoying
"gremlins".  

Gayle M. Callis

HTL/HT/MT(ASCP)  

Patrick,

We do a lot of frozen section IHC work. Years ago Gayle Callis turned me on
to fixing in cold acetone:ethanol (3:1) . We keep it at -20C and I fix for
10  min. on the bench then wash in PBS and proceed with the IHC. We do dry
slides for at least 30 min before fixing.  This has worked well in our hands
for many different antibodies.

Brett

Brett M. Connolly, Ph.D.

Principle 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 Lewis,
Patrick

Sent: Tuesday, April 28, 2015 5:56 PM

To: (Histonet <@t> lists.utsouthwestern.edu)

Subject: [Histonet] Acetone fixation problems with OCT Tissues

Hi Everyone,

I am still having issues with my IHCs with Acetone fixation.

If I fix in 100% Acetone, I get IHC staining, but my tissues are 50-90%
destroyed.

If I fix in 4% paraformaldehyde, or 10% NBF or (95%  Etoh and/or Methanol
with Acetone) I lose the epitopes I either get no staining or very  weak
staining, but the tissue morphology look fine.

I just tried an acetone gradient where I cut the tissues at 5 uM and dried
them overnight, then fixed for 10 minutes in 100% acetone, then fixed in 95%
acetone for 1 minute, then fixed in 70% acetone for 30 seconds, then quick
rinsed in H20, then washed as normal in DPBS pH 7.4.

I did 4 slides, 2 slides with one company's Charged slides ,and 2 slides
with another company's charged slides.

One company's slides look completely destroyed, the others may turn out, it
was hard to tell how much damage there was.  I'll know tomorrow when I
finish staining and Hemotoxylin them.

Patrick Lewis

Research Associate II Bench

Seattle Childrens Research Institute

206-884-1115