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 _______________________________________________ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet