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