[Histonet] Re: Questions about IHC on Frozen Sections

Fri Dec 6 16:29:05 CST 2013

Dear Brett, Liz and Patrick,

I agree with Brett and Liz having been in contact with them over the years.
However if your tissue is of human origin and you want to do CD4 and/or CD8
staining, then the acetone/alcohol fixative should NOT be used. The alcohol
will ruin Human CD4 and CD8 antigens but does not harm mouse or rat CD4 or
CD8 antigen.  I learned this from Dr. Chris van der Loos who is now going to
be sorely missed by the immunostaining community.  For human CD4 and CD8,
the sections should be air dried, then fixed in cold 4C reagent grade
acetone for 10 minutes, then air dried to let the acetone evaporate before
going into the buffer.   I never rinsed my solvent fixed frozen sections in
water, and if the buffer is not made correctly, as I learned the hard way,
the sections can look horrible.   Being a purist, my acetone fixed FS were
rinsed in 3 changes of pure buffer before equilibrating with in protocol
rinse buffer/0.05% Tween 20.   IF YOU DO USE the acetone/alochol fixation,
the sections are fixed at RT in this mixture and then go directly into
buffer for 3 changes.   DO NOT AIR DRY AFTER THIS Acetone/alcohol
mixture/fixative.  

If you fix a long, long time in acetone, you can get lesser staining of the
antigen.   Another clever trick is doing a double cold acetone fixation of
air dried frozen sections. This stabilizes the section so that it stays on
the slide better, and doesn't harm the antigens.   It would work for murine
and human FS.   Procedure is:  Fix air dry section for 10 min in 4C acetone,
remove and air dry section for 10 minutes, then return for fix again for 10
min in 4C acetone, then air dry these sections to evaporate away the acetone
approx 10 to 15 minutes, rinse in pure buffer, proceed with staining.   

Do NOT rinse your solvent fixed ( or air dried, unfixed)  frozen sections
with water (the enemy!), you want to use buffer to maintain isotonicity and
cellular integrity of the solvent fixed FS.   At the end of a chromogenic
protocol (after the chromogen is developed), you can even rinse with pure
buffer, then immerse the stained sections into NBF to post fix the section
for 10 minutes, rinse gently with running water and then counter stain with
hematoxylin.    This is also a van der Loos trick to improve the cellular
morphology of the nuclei in solvent fixed FS,  and doesn't harm the
chromogen.    

Why do you use TBSTw at pH 8?   That pH seems to be a big high for IHC, as
the norm tends to be pH 7.6? 

You can also make up this endogenous peroxidase block that will NOT chew
your sections up.   Solvent fixed frozen sections do NOT like strong
hydrogen peroxide concentrations, and this one worked perfectly for us.  It
is also a published method.  

PEROXIDASE BLOCK (0.03% hydrogen peroxide)

5 mls DPBS (Dulbeccos, Sigma), pH 7.4 - 7.4

5 ul 30% hydrogen peroxide 

50 ul 10% sodium azide

Make up, put in a dropper bottle,  and use for 1 week, refrigerate.  Discard
after 1 week or make up fresh daily.

Add to section, incubate for 10 - 15 minutes at RT, rinse well after
blocking.  If you wish, you can drain off the block, and add new half way
through the block if the tissue is particularly bloody. 

If you think the peroxidase block is still too strong, simply do Alkaline
phosphatase methods instead.  

Always let your unfixed frozen sections just taken from -80C freezer,
equilibrate for 20 minutes or more to RT before opening a box as water
condensation is the enemy to both antigens and morphology.   

I am sure I have repeated a great deal of what Liz and Brett presented, but
it does drive home some points. 

Take care

Gayle Callis

HTL,HT/MT (ASCP)

-----Original Message---------------------

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

 <http://lists.utsouthwestern.edu/mailman/listinfo/histonet> brett_connolly
<@t> merck.com

T- 215-652-2501

F- 215-993-6803

-----Original Message-----

From:  <http://lists.utsouthwestern.edu/mailman/listinfo/histonet>
histonet-bounces <@t> lists.utsouthwestern.edu [mailto:
<http://lists.utsouthwestern.edu/mailman/listinfo/histonet> histonet-bounces
<@t> lists.utsouthwestern.edu] On Behalf Of Elizabeth Chlipala

Sent: Thursday, December 05, 2013 5:59 PM

To: Lewis, Patrick; '
<http://lists.utsouthwestern.edu/mailman/listinfo/histonet> 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

 <http://lists.utsouthwestern.edu/mailman/listinfo/histonet> liz <@t>
premierlab.com

www.premierlab.com

Ship to Address:

Premier Laboratory, LLC

1567 Skyway Drive, Unit E

Longmont, CO 80504

-----Original Message-----

From:  <http://lists.utsouthwestern.edu/mailman/listinfo/histonet>
histonet-bounces <@t> lists.utsouthwestern.edu [mailto:
<http://lists.utsouthwestern.edu/mailman/listinfo/histonet> histonet-bounces
<@t> lists.utsouthwestern.edu] On Behalf Of Lewis, Patrick

Sent: Thursday, December 05, 2013 3:29 PM

To: ' <http://lists.utsouthwestern.edu/mailman/listinfo/histonet> 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.