Sounds like 10% dirty/oily slides, unless you clean them.
Frederick C. Monson, PhD
Technical Director
Light, Electron, X-Ray and Scanning Probe Imaging and Analysis Center
(LEXSPIAC)
Large Scientific Instrument Core
West Chester University
S. Church St. and W. Rosedale Ave.
West Chester, PA, 19320
610-738-0437
fmonson@wcupa.edu
URL: http://lexspiac.wcupa.edu/casi
==========================3D==================
I have chosen to celebrate,
during this Christmas season,
the salvation of the Grinch as described by Dr. Seuss!
==========================3D==================
-----Original Message-----
From: histonet-bounces@lists.utsouthwestern.edu
[mailto:histonet-bounces@lists.utsouthwestern.edu] On Behalf Of Bauer,
Karen
Sent: Tuesday, December 04, 2007 1:20 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Tissue Sections
Hi to all...
I know we've all talked about this before, and I've spent the last hour
searching the archives for any information that I could use. Since
there were no real answers to help me out, I'm turning to the pros for
assistance.
Lately we have been experiencing a lot of headaches with sections
falling off, bad sections, and folding around the tissue edges. We have
not changed anything that we have been doing and it's not all the
slides. I asked the pathologist how many slides he's finding like that
and he stated probably 15 to 20 slides a day. 15-20 out of 200 plus are
coming out crappy, when all of the others are fine. Everything is
processed, embedded, cut, stained and coverslipped the same, but some
come out beautiful and some come out very bad. Whenever they ask for
recuts on the bad slides, they say they are always beautiful. A lot of
times, the recuts are cut by the same tech that cut it the first time.
I've talked to the staff a lot about this occurrence and know all are
knowledgeable of proper cutting techniques. They all turn in quality
work. We have really slowed down our cutting in the morning in order to
hand in that "perfect" slide. They macroscopically look great when we
put them on the slide, but when the doctor gets them, crappy ones show
up.
We've thought about water trapped under the section and have let them
drain vertically and air dry longer before we put them in the oven.
We've let them cool after the oven to make sure the sections are adhered
securely before putting the slides in xylene. We stain by hand, so we
make sure we gently dip the slides and not dip aggressively. We
coverslip by hand also. While coverslipping, there are many times that
I see the edges of tissues move slightly when I put the coverslip on, so
I know the tissue is not adhering.
We have a slide etcher, so we use the ColorMark slides with the black
backing. We use charged slides for all of our prostate cores, breast
cores, and any small core biopsies that usually require special stains
or IPs. We use plain ColorMark slides for all other routine Histology
cases. We've tried using charged slides for all tissues, but were still
getting bad slides here and there.
As I said, all tissues are treated the same, but only 15-20 slides out
of 200+ are coming out bad. Our techs rotate cutting, so I know it's
not just cutting techniques. It's happening to all of us. It's gotten
to the point that our doctors are focusing on the few bad slides and
it's driving them crazy.
I'm open for any suggestions. We are thinking about trying different
slides or maybe putting an adhesive in the waterbath (yep, I know about
adhesive and not using charged slides). Vendors who have different
black backed slides than ColorMark for etching are welcome to reply.
Thanks in advance,
Karen Bauer HT(ASCP)
Histology Supervisor
Luther Hospital
Eau Claire, WI
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