Re: [Histonet] Re: 100% ethanol in tissue processing

[Gudrun Lang via Histonet]

   If there is a cytology lab near you, they will take use of it.
   --
   Diese Nachricht wurde von meinem Android Mobiltelefon mit GMX Mail
   gesendet.

   Am 30.05.24, 01:21 schrieb Curt Tague via Histonet
   :

 My mistake, it’s methanol….
 No bueno for tissue?
 Curt
 Get Outlook for iOS<[1]https://aka.ms/o0ukef>
 
 From: Mac Donald, Jennifer 
 Sent: Wednesday, May 29, 2024 3:50:05 PM
 To: Curt Tague ;
 Histonet@lists.utsouthwestern.edu
 
 Subject: RE: 100% ethanol in tissue processing
 Absolutely can be used on the tissue processor or for staining.
 -Original Message-
 From: Curt Tague via Histonet 
 Sent: Wednesday, May 29, 2024 12:16 PM
 To: Histonet@lists.utsouthwestern.edu
 Subject: [Histonet] 100% ethanol in tissue processing
 EXTERNAL SENDER - Exercise caution with requests, links, and
 attachments.
 Hi all, stupid question (despite what all the college professors
 said, yes, I think there are some stupid questions and this is one
 of them, I should know)!
 Someone dropped off a bunch of pure ethanol, they don't need it, I
 said I'd help dispose it... can't be used for processing can it? I
 seem to recall damaging some tissue many years ago when using
 ethanol... your thoughts?
 Maybe just clean cycles on the processors...
 Thanks,
 Curt
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[Histonet] Re IF staining questions

[Amos Brooks via Histonet]

Hi,
 Direct IiF (a fluorescent conjugated primary) is certainly easier, but
there are some reasons one might prefer using an indirect method (using a
conjugated secondary). Indirect methods allow the use of a different
wavelength to be used simply by switching the secondary. It is also
possible for the fluorescent tag to fade or not be strong enough to start
with. The secondary could be used at a higher concentration or replaced in
the case of a faded primary.

Anos



Message: 1
Date: Thu, 19 Oct 2023 09:22:56 -0400
From: Charles Riley 
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] IF staining questions
Message-ID:

Content-Type: text/plain; charset="UTF-8"

Why would one decide to use a primary antibody along with a secondary
antibody rather than a primary antibody conjugated with the secondary?

Example.   I have a researcher who wants to do CD11C staining with
Alexafluor488

Is it better to buy and use a primary antibody CD11C conjugated with
Alexafluor488  or to do the CD11C primary and a Rabbit anti-rat (H_L) IgG
antibody secondary?
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Re: [Histonet] Re Protocol for DAPI staining on paraffin sections

[Hobbs, Carl via Histonet]

Dear Alida,
That is a good point.
I probably have only done HOECHST on Pwax sections after they have been 
heat-Antigen retrieved ( HIER)
However, H ( or DAPI) works on fixed cell monolayers: they don't get HIER 
treatment)
Nor do frozen sections ( well, some do) and the HOECHST/DAPI works fine on both
However, perhaps you can just take a Pwax section to water and apply HOECHST 
for 10 mins, wash off, mount in anti fade aq. mountant and have a look down a 
Fluorescence scope?
If you can wait, I will check this next Monday ( put H on a non HIER'd Pwax 
section  and get back to you by Monday later)

Unless, whoever that is doing the fluorescence DAPI/HOECHST nuclear staining is 
actually doing it on Pwax sections that have been HIER-ed?

Best wishes

Carl



Carl Hobbs FIBMS
Histology and Imaging Manager
Wolfson CARD
Guys Campus, London Bridge 
Kings College London
London
SE1 1UL
 


020 7848 6810
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[Histonet] Re Protocol for DAPI staining on paraffin sections

[Hobbs, Carl via Histonet]

Use DAPI/HOECHST at the same concentration you use for cells/FS
I use Hoechst H33258 Sigma 10mg/ml soln
I dilute by adding 0.5 microL to 100 microL buffer then add 1/100 to Alexa 
secondaries ( final 1/20K diln factor)
Some incubate in HOECHST or DAPI after secondary ab incubation, for IF
Sure, if only "staining" nuclei, incubate in 1/20K Hoechst in same buffer
This will vary for each lab
Or, buy mounting medium that contains DAPI

Good luck
Carl

Carl Hobbs FIBMS
Histology and Imaging Manager
Wolfson CARD
Guys Campus, London Bridge
Kings College London
London
SE1 1UL

020 7848 6813
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Re: [Histonet] Re-embed agar/FFPE tissue in plastic?

[Morken, Timothy via Histonet]

Kate, we regularly re-embed paraffin-embedded tissue in epoxy resin for 
electron microscopy. We are using very small tissue - kidney, liver, 1mm core 
biopsies. 

For that we run thru xylene 4 x 30 min at 60C, 100% ethanol 3 x 10 min, 95, 70, 
50% ethanol 5 min each, then to distilled water, then buffer. Then it goes on 
to processing for resin embedding. The wax is gone and will not interfere with 
the plastic. The agar will not be affected or affect plastic embedding. We use 
agar to embed cilia for EM with good results. 


You might want to do some test tissue to get the timing right before trying on 
your samples. 


Tim Morken
Supervisor, Electron Microscopy/Neuromuscular Special Studies
Department of Pathology
UC San Francisco Medical Center


-Original Message-
From: Kate Davoli via Histonet [mailto:histonet@lists.utsouthwestern.edu] 
Sent: Wednesday, May 29, 2019 8:12 AM
To: Histonet Usegroup
Subject: [Histonet] Re-embed agar/FFPE tissue in plastic?

I have a bunch of precious FFPE samples that were embedded in agar prior to
being embedded in wax (so as to orient the tissue easily).  The client now
wants these samples to be taken backwards out of wax and processed for
semithin plastic GMA (JB-4) sectioning.

Is that possible? Will the tissue having previously been embedded in wax
cause problems for the reagent infiltration or the plastic curing?  Will it
interfere with the catalyst?  I know that taking tissue back through
xylenes and alcohols is *supposed *to be able to remove all the wax, but
does it really?

I need the agar support surrounding the sample to STAY on the sample so
that it can stand upright during the plastic curing ... has anybody tried
to do double embedding with agar and then JB-4?  Does that work?

Please help!
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[Histonet] Re-embed agar/FFPE tissue in plastic?

[Kate Davoli via Histonet]

I have a bunch of precious FFPE samples that were embedded in agar prior to
being embedded in wax (so as to orient the tissue easily).  The client now
wants these samples to be taken backwards out of wax and processed for
semithin plastic GMA (JB-4) sectioning.

Is that possible? Will the tissue having previously been embedded in wax
cause problems for the reagent infiltration or the plastic curing?  Will it
interfere with the catalyst?  I know that taking tissue back through
xylenes and alcohols is *supposed *to be able to remove all the wax, but
does it really?

I need the agar support surrounding the sample to STAY on the sample so
that it can stand upright during the plastic curing ... has anybody tried
to do double embedding with agar and then JB-4?  Does that work?

Please help!
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Re: [Histonet] re-processing help

[HARRISON,Sharon via Histonet]

Dear Lauren,
Please leave samples in formalin for about 48 hours.
Some tissues will be recovered others may be a bit more difficult to recover.
Then reprocess after the soak and see what happens.
Regards.
Sharon Harrison
Chief Medical Technologist in charge of Histopathology
UWI Mona, Dept of Pathology

From: Lauren Sweeney via Histonet [histonet@lists.utsouthwestern.edu]
Sent: Monday, February 11, 2019 3:03 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] re-processing help

Hello Histonet,

We have issues- please advise! Long story short- some tissues did not get 
processed over the weekend, they went through the process but were not 
submerged in ANY (we think) of the reagents. Now they are shriveled up and 
brittle (they are intestinal). Is there any way to save them? Can we re-process 
them? Has anyone ever had this problem and had success recovering?

Gratitude,
Lauren
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[Histonet] re-processing help

[Lauren Sweeney via Histonet]

Hello Histonet,

We have issues- please advise! Long story short- some tissues did not get 
processed over the weekend, they went through the process but were not 
submerged in ANY (we think) of the reagents. Now they are shriveled up and 
brittle (they are intestinal). Is there any way to save them? Can we re-process 
them? Has anyone ever had this problem and had success recovering?

Gratitude,
Lauren
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Re: [Histonet] Re guinea pig IHC

[Patsy Ruegg via Histonet]

good advise Carl and you asked some of the things I was wondering.

Patsy Ruegg, HT(ASCP)QIHC
Ruegg IHC Consulting
40864 E Arkansas Ave
Bennett, CO 80102
H 303-644-4538
C 720-281-5406
prueg...@hotmail.com



From: Hobbs, Carl 
Sent: Saturday, February 2, 2019 12:38 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Re guinea pig IHC



Ms Ruegg, as usual , gives excellent advice: avoid HRP.
Use Alk phos?
Block end. alk phos by using levamisole.

However, Ms Shivers does not state the fixation status of her FS.
Is the Gpig tissue perfused-fixed then frozen orfrozen as unfixed...then 
fixed?
Alsowhy 0.3% H2O2?
Use 3%kill the enzyme, not feed it?
NOT aq for FS
Make up in IMS ( 74OP)
No tissue disruption
However: you state that you get loadsa bubbles...so what? Is your section still 
attached to your slide?
Can you then carry out successful IF/IHC?
If yesno problem.
Sure, there's the argument that using a coagulant ( alcohol) in block is a No No
I never had a problemprovided that  the Formalin fixation was sufficient 
for unfixed crosections ( 15 mins)
I do STILL severely dislike FS ( sure, I spent many years in Diagnostic 
Histopath doing Operative FS)
 Why not use Pwax sections, Ms Shivers?

Curious-illy

Carl



Carl Hobbs FIBMS
Histology and Imaging Manager
Wolfson CARD
Guys Campus, London Bridge
Kings College London
London
SE1 1UL

020 7848 6813



From: histonet-requ...@lists.utsouthwestern.edu 

Sent: 02 February 2019 18:00
To: histonet@lists.utsouthwestern.edu
Subject: Histonet Digest, Vol 183, Issue 2


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Today's Topics:

   1. Re: cytology listserv (Webster, Thomas S.)
   2. Re: guinea pig IHC (Patsy Ruegg)


--

Message: 1
Date: Fri, 1 Feb 2019 18:28:30 +
From: "Webster, Thomas S." 
To: "'histonet@lists.utsouthwestern.edu'"

Subject: Re: [Histonet] cytology listserv
Message-ID: <93fc6a1cc62f41f5ad5890786ab04...@crh.org>
Content-Type: text/plain; charset="us-ascii"

Haven't seen any cytology listservs except the one for members of the ASC.  
There are some cytology facebook pages where you could get questions answered. 
This Histonet listserv is very informative.


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information.  If you are not the intended recipient, please contact the
sender by reply e-mail immediately.  Please destroy all copies of the
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appreciated.
Columbus Regional Hospital
2400 East 17th Street
Columbus, Indiana 47201

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Message: 2
Date: Fri, 1 Feb 2019 18:39:54 +
From: Patsy Ruegg 
To: Jan Shivers , histonet

Subject: Re: [Histonet] guinea pig IHC
Message-ID:



Content-Type: text/plain; charset="iso-8859-1"

Especially a very blood tissue like GP spleen.

Patsy Ruegg, HT(ASCP)QIHC
Ruegg IHC Consulting
40864 E Arkansas Ave
Bennett, CO 80102
H 303-644-4538
C 720-281-5406
prueg...@hotmail.com



From: Patsy Ruegg 
Sent: Thursday, January 31, 2019 11:51 AM
To: Jan Shivers; histonet
Subject: Re: [Histonet] guinea pig IHC

In my experience it is not that GP have a higher peroxidase level, it is frozen 
sections in general that cannot be blocked with h202, unless they are fixed for 
a long time in formalin.  What are others experiences with h202 blocking on 
frozen sections.  I always  used an IHC detection system that did not require 
h202 blocking for frozen sections.

Patsy Ruegg, HT(ASCP)QIHC
Ruegg IHC Consulting
40864 E Arkansas Ave
Bennett, CO 80102
H 303-644-4538
C 720-281-5406
prueg...@hotmail.com



From: Jan Shivers 
Sent: Tuesday, January 29, 2019 12:58 PM
To: histonet
Subject: [Histonet] guinea pig IHC

Has anyone ever performed IHC on frozen sections of guinea pig tissue?  I
am experiencing an enormous amount of bubbling when doing the peroxidase
blocking step,

[Histonet] Re guinea pig IHC

[Hobbs, Carl via Histonet]

Ms Ruegg, as usual , gives excellent advice: avoid HRP.
Use Alk phos?
Block end. alk phos by using levamisole.

However, Ms Shivers does not state the fixation status of her FS.
Is the Gpig tissue perfused-fixed then frozen orfrozen as unfixed...then 
fixed?
Alsowhy 0.3% H2O2?
Use 3%kill the enzyme, not feed it?
NOT aq for FS
Make up in IMS ( 74OP)
No tissue disruption
However: you state that you get loadsa bubbles...so what? Is your section still 
attached to your slide?
Can you then carry out successful IF/IHC?
If yesno problem.
Sure, there's the argument that using a coagulant ( alcohol) in block is a No No
I never had a problemprovided that  the Formalin fixation was sufficient 
for unfixed crosections ( 15 mins)
I do STILL severely dislike FS ( sure, I spent many years in Diagnostic 
Histopath doing Operative FS) 
 Why not use Pwax sections, Ms Shivers?

Curious-illy

Carl

  
 
Carl Hobbs FIBMS 
Histology and Imaging Manager 
Wolfson CARD 
Guys Campus, London Bridge  
Kings College London 
London 
SE1 1UL 
  
020 7848 6813



From: histonet-requ...@lists.utsouthwestern.edu 

Sent: 02 February 2019 18:00
To: histonet@lists.utsouthwestern.edu
Subject: Histonet Digest, Vol 183, Issue 2
  

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When replying, please edit your Subject line so it is more specific
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Today's Topics:

   1. Re: cytology listserv (Webster, Thomas S.)
   2. Re: guinea pig IHC (Patsy Ruegg)


--

Message: 1
Date: Fri, 1 Feb 2019 18:28:30 +
From: "Webster, Thomas S." 
To: "'histonet@lists.utsouthwestern.edu'"
    
Subject: Re: [Histonet] cytology listserv
Message-ID: <93fc6a1cc62f41f5ad5890786ab04...@crh.org>
Content-Type: text/plain; charset="us-ascii"

Haven't seen any cytology listservs except the one for members of the ASC.  
There are some cytology facebook pages where you could get questions answered. 
This Histonet listserv is very informative.


CONFIDENTIALITY NOTICE:
This e-mail message, including all attachments, is for the sole use of the
intended recipient(s) and may contain confidential and privileged
information. You may NOT use, disclose, copy or disseminate this
information.  If you are not the intended recipient, please contact the
sender by reply e-mail immediately.  Please destroy all copies of the
original message and all attachments. Your cooperation is greatly
appreciated.
Columbus Regional Hospital
2400 East 17th Street
Columbus, Indiana 47201

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Message: 2
Date: Fri, 1 Feb 2019 18:39:54 +
From: Patsy Ruegg 
To: Jan Shivers , histonet
    
Subject: Re: [Histonet] guinea pig IHC
Message-ID:
    

    
Content-Type: text/plain; charset="iso-8859-1"

Especially a very blood tissue like GP spleen.

Patsy Ruegg, HT(ASCP)QIHC
Ruegg IHC Consulting
40864 E Arkansas Ave
Bennett, CO 80102
H 303-644-4538
C 720-281-5406
prueg...@hotmail.com



From: Patsy Ruegg 
Sent: Thursday, January 31, 2019 11:51 AM
To: Jan Shivers; histonet
Subject: Re: [Histonet] guinea pig IHC

In my experience it is not that GP have a higher peroxidase level, it is frozen 
sections in general that cannot be blocked with h202, unless they are fixed for 
a long time in formalin.  What are others experiences with h202 blocking on 
frozen sections.  I always  used an IHC detection system that did not require 
h202 blocking for frozen sections.

Patsy Ruegg, HT(ASCP)QIHC
Ruegg IHC Consulting
40864 E Arkansas Ave
Bennett, CO 80102
H 303-644-4538
C 720-281-5406
prueg...@hotmail.com



From: Jan Shivers 
Sent: Tuesday, January 29, 2019 12:58 PM
To: histonet
Subject: [Histonet] guinea pig IHC

Has anyone ever performed IHC on frozen sections of guinea pig tissue?  I
am experiencing an enormous amount of bubbling when doing the peroxidase
blocking step, even though I'm only using a 0.3% concentration of H2O2.
And when I say 'enormous', I mean it's like continuous champagne bubbles
rising out of the tissue, even after 20 minutes in the H2O2 solution.

I can't find anything in the literature that mentions guinea pigs having a
higher peroxidase content in their tissues.

Thanks for any help that anyone can provide.

Jan Shivers
Senior Scientist
IHC/Histology 

[Histonet] re; cell blocks and frozen for IHC

[Greg Dobbin via Histonet]

No pretreatments for anything that is not formalin fixed. I think 95% ETOH
for a fixative but but others may have a better idea than I.
Greg

-- 
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1205 Pleasant Grove Rd
RR#2 York,
PE  C0A 1P0


*Everything in moderation...even moderation itself**!*
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[Histonet] Re-attaching slide film to slide

[Renee H. Workman via Histonet]

Hey Histonetter,

How can I re-attach the film with the tissue back to the blank slide?  Have 
some older slides and the slide film has popped off.

Renee H. Workman
Histology Supervisor
Virginia Urology
9105 Stony Point Drive
Richmond, VA  23235
W: 804-527-1316 | F: 804-270-0917
rhwork...@uro.com | www.uro.com





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[Histonet] Re-embedment of tissue for TEM?

[daniel blackburn via Histonet]

We had an unsuccessful embedding run, using Embed 812 resin (an Epon 
replacement, from Electron Microscopy Services).  After curing in the oven, the 
resin blocks came out “gummy” (flexible), perhaps due to a carryover of water 
in the paper labels.  Would it be possible to re-embed the tissue, e.g., by 
running it through propylene oxide and 100% ethanol washes and back into new 
resin?  We hope so, since the tissue is valuable to us.  Thanks for any advice.

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[Histonet] Re #2: μm H staining, WHOOPS!!

[Gayle Callis via Histonet]

Sorry about hitting send too soon. 

 

Repeat of things to try: 

 

a.  Do not use regressive hematoxylin and eosin where hematoxylin
can be overly differentiated i.e. removed from too thin sections.  Use
progressive hematoxylin i.e. Gill II or Gill III type formulation. DO NOT
use acid alcohol differentiation with progressive hematoxylin.  Try staining
longer, i.e. 10 min in Gill III, and use acetic acid clarifier only 1 or 2
dips or skip clarifying solution entirely. 

b.  Never use acid alcohol differentiation even with your
hematoxylin

c.   Use progressive hematoxylin, and do not clarify or use acid
alcohol differentiation solution.  Wash well for 1 minute in running tap
water then blue. 

d.   Increase the thickness of sections to see if this satisfies the
post-docs.  Start with 3 μm and 4 μm but stain these sections with
progressive H  If you don't need 2 μm, then go to a more routine  4 μm or
5 μm thickness.   You need to explain to these post docs about too thin
sections do NOT have enough tissue/cell left to stain well enough. 

e.   Treat sections with FRESH MADE 1% periodic acid for 10 min,
rinse well and stain with progressive H  This periodic acid technique is
found in Sheehan and Hrapchak Theory and Practice of Histotechnology book.
PA treatment might improve the staining with your sections by making more
groups on DNA available to hematoxylin.   However, I didn't find it improved
my thin section staining as much as I wanted.   The sections were just too
thin. 

f.Try Eosin-phloxine mixture, start dehydration in a few quick
dips in 95%, then proceed to 100% alcohols.   Eosin-phloxine is available as
ready to use or make up in the lab.   Sheehan and Hrapchak is also a source
of this eosin formulation.  

 

Good luck

Gayle M. Callis HT/HTL/MT(ASCP)

GCallis Histology Service, LLC.  

 

 

 

   

 

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[Histonet] re best glass coverslipper

[Steven Weston via Histonet]

I've been using the Dako coverslipper for over three years now.  Once i had got 
used to ensuring the sliding mechanism was always kept mountant free and 
regularly changed the xylene in the u tube we have had very few problems with 
this machine. Its footprint is without question the smallest coverslipper i 
have seen and generally with good maintenance  (yearly PM) it functions very 
well. Occasionally we get small bubbles under the coverslips but they are 
easily removed. Highly recommend looking at one, also the price is very good.

regards



steve weston
lab manager
Breathe-Well CRE
UTAS-SOM


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[Histonet] re automatic cover slippers

[Steven Weston via Histonet]

We use the Dako cover slipper and provided the moving arm is kept free from 
dried mounting medium we have found this to be a terrific machine. It also has 
a really small footprint and so fits nicely into our fume hood. Can do about 
400 slides an hour and produces very few bubbles.

Steve Weston
Breathe-Well CRE
Lab Manager
0408990859




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[Histonet] Re. Decalcification with formic acid sodium

[Tyrone Genade via Histonet]

It isn't very clear, but for the Formic Acid/sodium Citrate buffer you have
to use trisodium citrate. 10 g of disodium or monosodium citrate wouldn't
work as well...

--
Tyrone Genade
Orange City, Iowa
tel: (+1) 712 230 4101
http://tgenade.freeshell.org

Romans 6:23: The gift of God is eternal life through Christ Jesus our Lord.
To find out how to receive this FREE gift visit http://www.alpha.org.
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[Histonet] Re. Decalcification with formic acid sodium

[Gayle Callis via Histonet]

Dorothy and Carl, 

 

Comments about your Histonet replies on formic acid decalcification. 

 

The Morse solution referred to by Dorothy can be picked up online by typing
in the DOI number:  10.1.1.4689.3439.pdf  or title,Morse A.  Formic
acid-sodium citrate decalcification and butyl alcohol dehydration of teeth
and bones for sectioning in paraffin. 1945  J Dental Res 1945:24:143.   You
will find the reference to Evans and Krajian paper on formic acid/sodium
citrate along with the original recipe for their solution (equal parts of
85% (stock) formic acid and 20% sodium citrate).   Morse modified the Evans
Krajian method (1 part diluted stock formic acid i.e. 90% diluted 1:1 with
water for 45% formic acid) plus 1 part 20% sodium citrate.   The Morse paper
was excellent and well worth reading.   Interestingly, in 1962, our lab used
the Morse solution  for decalcifying teeth although it was never referred to
by that name but simply  formic acid/sodium citrate.   The formic acid
content in Morse's solution is half the concentration of formic acid in
Evans/Krajian solution.  It seems both work equally well and the higher
concentration of formic acid should increase the decalcification rate
somewhat.   Morse also did chemical decalcification endpoint testing.  

 

Carl is correct about not mixing citric acid with formic acid as citric acid
is not going to act as a buffer salt.   However, you will find in the
literature that citric acid is very mild and has been used as a decalcifying
agent for EM studies.   Carl is also correct in that sodium formate can be
used as a buffering salt instead of sodium citrate.  We have worked with
both of the buffering salts/formic acid formulations and found they works
equally well for decalcification.  I have some publications on file
comparing acid versus EDTA for cartilage and IHC studies and learned some
researchers referred to buffered formic acid methods as acidic buffers .
The latter terminology could be confusing to people in the business of
decalcifying bones and teeth. but no more so than the acronyms manufacturers
give their solutions.   It pays to read the MSDS for any decalcifying
solution,  and even compare this information to what is in histology
textbooks as part of our education.   

 

I have found the discussions on this topic enlightening.I will be happy
to send the pdf of the Morse method to those interested in reading it.   I
have not been able to access the 1930 Evans Krajian method yet.  What is
important is knowing these older, classic formic acid methods are still
tried and true with the added advantage of being available commercially for
our convenience. 

 

Thanks everyone

 

Gayle Callis 

 

 

 

 

 

 

 

 

You Wrote:  

 

There was a paper
http://www.genedetect.com/Merchant2/ExampleRefs/Decalcifying_protocols.pdf
http://www.genedetect.com/Merchant2/ExampleRefs/Decalcifying_protocols.pdf

Talking about formic acid (Morse solution) can get as good result as EDTA in
ISH. 

FYI.

 

Dorothy Hu

 

Mouse knee joints:

done lots of decalcified FFPWS for assessment of articular cartilage
degeneration models.

See Histonet images for a Tol blue image.

Decal in 10 % EDTA for 3 days on a rocker at RT.

Sure5days if you are worried.

No difference in Immuno-reactivity, imho.

If you want to use buffered Formic acid, use Formic acid; sodium formate.

Use of citric acid with Formic acid does not make a buffer.

It's just mixing two relatively mild acids.

However, I am sure that Prof Kiernan can further enlighten us.

 

Respectfully,

Carl Hobbs FIBMS 

Histology and Imaging Manager 

Wolfson CARD 

Guys Campus, London Bridge  

Kings College London 

London 

SE1 1UL 

  

020 7848 6813

 

 

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Re: [Histonet] Re. Decalcification with formic acid sodium

[Hobbs, Carl via Histonet]

Mouse knee joints:
done lots of decalcified FFPWS for assessment of articular cartilage 
degeneration models.
See Histonet images for a Tol blue image.
Decal in 10 % EDTA for 3 days on a rocker at RT.
Sure5days if you are worried.
No difference in Immuno-reactivity, imho.
If you want to use buffered Formic acid, use Formic acid; sodium formate.
Use of citric acid with Formic acid does not make a buffer.
It's just mixing two relatively mild acids.
However, I am sure that Prof Kiernan can further enlighten us.

Respectfully,
 
Carl Hobbs FIBMS 
Histology and Imaging Manager 
Wolfson CARD 
Guys Campus, London Bridge  
Kings College London 
London 
SE1 1UL 
  
020 7848 6813
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Re: [Histonet] Re. Decalcification with formic acid sodium citrate

[Merissa via Histonet]

Thank you, Gayle! This is exactly what I was looking for and we are willing to 
make this in house. We are trying just for if acid and water, but the buffering 
salt should be added. I will try de calcifying mouse knees next week with this 
protocol. Thank you for the reference, I appreciate your help!

Sincerely,
Merissa



 On Jul 25, 2015, at 10:24 AM, Gayle Callis via Histonet 
 histonet@lists.utsouthwestern.edu wrote:
 
 Merissa and Tim,  
 
 
 
 This formic acid decalcifying solution is basically the classic Evans and
 Krajian fluid (Sheehan and Hrapchak,   Theory and Practice of
 Histotechnology, 2nd edition, P.92).  Shandon has added other ingredients
 for some reason, and has kept those concentrations proprietary.   You really
 don't need to add a surfactant or PVP emulsifier when making up this
 decalcifying agent.   Simply use the classic recipe for successful
 decalcification.   This is also referred to as buffered formic acid and in
 some publications an acidic buffer.  It is excellent if IHC is needed and
 less damaging, obviously, than a strong mineral HCL acid decalcifiers.  
 
 
 
 Sodium citrate crystals (a buffering salt) 10 g 
 
 90% formic acid stock25 ml  
 
 Distilled water75 ml   
 
 
 
 One can calculate the concentration of formic acid i.e. approx. 4.5% since
 is it made from 90% formic acid stock.  
 
 
 
 Don't bother with the surfactants or PVP.  
 
 
 
 Enjoy an excellent in house formic acid decalcifying solution.  I also
 suggest you read Sheehan and Hrapchak textbook chapter on bone as a way to
 familiarize yourself with decalcifiying solutions that manufacturers now
 supply with some modifications.  Some manufacturers will refer to these
 methods but probably prefer not to do this since they want you to buy their
 commercial product that is obviously a time saver with elimination of having
 to store stock acid solutions.   The classic methods made in house are
 excellent if you have time to make them up.   Formic acid with sodium
 formate is another popular buffered formic acid.   I suggest you look for
 another source/manufacturer of the your favorite decalcifier in question as
 more than one company will make it.  Decal Corp, recently sold to Stat Lab,
 could also be the source as Shandon isn't the only game in town.   Others
 are Newcomer Supply, Poly Scientific.  Not having to make it up may remain
 your preference. 
 
 
 
 Gayle M. Callis 
 
 HTL/HT/MT(ASCP) 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 Written by Tim and Merissa:   
 
 
 
 Merissa,
 
 
 
 Water77-80   solvent
 
Formic acid  21-23   active ingredient
 
 Fluorad  1   surfactant  - a
 wetting agent to make the solution wet the bone more easily
 
Sodium citrate   1   emulsifier , buffer
 
 Polyvinyl pyrrolidone1   emulsifier 
 
 
 
 They say less than one percent of the last three, but you really have no
 idea whether that is 1%, .1% or .01%. It could be any of those.
 
 
 
 But all those surfactants and emulsifiers are meant to keep the solution
 viable for long periods on the shelf. When you make it fresh you don't
 really need them.
 
 
 
 You can either buy a different decalcifier, or make your own. Making your
 own with just the water and acid will work just fine. 
 
 
 
 
 
 Tim Morken
 
 Pathology Site Manager, Parnassus 
 
 Supervisor, Electron Microscopy/Neuromuscular Special Studies
 
 Department of Pathology
 
 UC San Francisco Medical Center
 
 
 
 -Original Message-
 
 From: M.O. via Histonet [mailto:
 http://lists.utsouthwestern.edu/mailman/listinfo/histonet histonet at
 lists.utsouthwestern.edu] 
 
 Sent: Wednesday, July 22, 2015 1:24 PM
 
 To:  http://lists.utsouthwestern.edu/mailman/listinfo/histonet histonet at
 lists.utsouthwestern.edu
 
 Subject: [Histonet] understanding reagents in decalcifier; making it
 in-house
 
 
 
 Hello Histonet
 
 
 
 The supplier for our decalcifier, TBD-2 from Shandon, is having issues with
 getting the product out and we will not be receiving it for at least another
 month.  Our samples are piling up and I don't know what I should do, but
 maybe I can make the decalcifier in-house.  I am wondering if I can make my
 own based on the reagents they listed and their percentages and if certain
 reagents are not actually necessary.
 
 
 
 The samples we typically decalcify are mouse knees (decal time = 2 days),
 mouse spines (3 days), human bone slabs about 7mm in thickness (7-12 days).
 Fixation is in zinc buffered formalin, then decalcification, then 70% EtOH.
 Our choice to use TBD-2 is due to the gentle decalcification for IHC and we
 get GREAT results.
 
 
 
 Composition of Shandon TBD-2 Decalcifier:
 
 ComponentWeight %
 
 Water  77-80
 
 Formic acid 21-23
 
Fluorad  

[Histonet] Re. Decalcification with formic acid sodium citrate

[Gayle Callis via Histonet]

Merissa and Tim,  

 

This formic acid decalcifying solution is basically the classic Evans and
Krajian fluid (Sheehan and Hrapchak,   Theory and Practice of
Histotechnology, 2nd edition, P.92).  Shandon has added other ingredients
for some reason, and has kept those concentrations proprietary.   You really
don't need to add a surfactant or PVP emulsifier when making up this
decalcifying agent.   Simply use the classic recipe for successful
decalcification.   This is also referred to as buffered formic acid and in
some publications an acidic buffer.  It is excellent if IHC is needed and
less damaging, obviously, than a strong mineral HCL acid decalcifiers.  

 

Sodium citrate crystals (a buffering salt) 10 g 

90% formic acid stock25 ml  

Distilled water75 ml   

 

One can calculate the concentration of formic acid i.e. approx. 4.5% since
is it made from 90% formic acid stock.  

 

Don't bother with the surfactants or PVP.  

 

Enjoy an excellent in house formic acid decalcifying solution.  I also
suggest you read Sheehan and Hrapchak textbook chapter on bone as a way to
familiarize yourself with decalcifiying solutions that manufacturers now
supply with some modifications.  Some manufacturers will refer to these
methods but probably prefer not to do this since they want you to buy their
commercial product that is obviously a time saver with elimination of having
to store stock acid solutions.   The classic methods made in house are
excellent if you have time to make them up.   Formic acid with sodium
formate is another popular buffered formic acid.   I suggest you look for
another source/manufacturer of the your favorite decalcifier in question as
more than one company will make it.  Decal Corp, recently sold to Stat Lab,
could also be the source as Shandon isn't the only game in town.   Others
are Newcomer Supply, Poly Scientific.  Not having to make it up may remain
your preference. 

 

Gayle M. Callis 

HTL/HT/MT(ASCP) 

 

  

 

 

 

 

 

 

Written by Tim and Merissa:   

 

Merissa,

 

 Water77-80   solvent

Formic acid  21-23   active ingredient

 Fluorad  1   surfactant  - a
wetting agent to make the solution wet the bone more easily

Sodium citrate   1   emulsifier , buffer

 Polyvinyl pyrrolidone1   emulsifier 

 

They say less than one percent of the last three, but you really have no
idea whether that is 1%, .1% or .01%. It could be any of those.

 

But all those surfactants and emulsifiers are meant to keep the solution
viable for long periods on the shelf. When you make it fresh you don't
really need them.

 

You can either buy a different decalcifier, or make your own. Making your
own with just the water and acid will work just fine. 

 

 

Tim Morken

Pathology Site Manager, Parnassus 

Supervisor, Electron Microscopy/Neuromuscular Special Studies

Department of Pathology

UC San Francisco Medical Center

 

-Original Message-

From: M.O. via Histonet [mailto:
http://lists.utsouthwestern.edu/mailman/listinfo/histonet histonet at
lists.utsouthwestern.edu] 

Sent: Wednesday, July 22, 2015 1:24 PM

To:  http://lists.utsouthwestern.edu/mailman/listinfo/histonet histonet at
lists.utsouthwestern.edu

Subject: [Histonet] understanding reagents in decalcifier; making it
in-house

 

Hello Histonet

 

The supplier for our decalcifier, TBD-2 from Shandon, is having issues with
getting the product out and we will not be receiving it for at least another
month.  Our samples are piling up and I don't know what I should do, but
maybe I can make the decalcifier in-house.  I am wondering if I can make my
own based on the reagents they listed and their percentages and if certain
reagents are not actually necessary.

 

The samples we typically decalcify are mouse knees (decal time = 2 days),
mouse spines (3 days), human bone slabs about 7mm in thickness (7-12 days).
Fixation is in zinc buffered formalin, then decalcification, then 70% EtOH.
Our choice to use TBD-2 is due to the gentle decalcification for IHC and we
get GREAT results.

 

Composition of Shandon TBD-2 Decalcifier:

ComponentWeight %

 Water  77-80

Formic acid 21-23

Fluorad   1

Sodium citrate   1

Polyvinyl pyrrolidone  1

 

If you have any input on what reagents I should use and the percentages for
making a decalcifier myself, it would be much appreciated!

 

Thank you for you help,

Merissa

 

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Re: [Histonet] RE..... murine CD4, CD8 and CD68 for FFPE tissue

[Hobbs, Carl]

I use the anti CD68 clone FA-11 ( obtained from Abcam).
It works very well on frozen sections but, I have been unable to get any 
positivity on Pwax sections ( using +/- Citric acid pH6 HIER)

Carl
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Re: [Histonet] : Re: HE Stainer Question

[Simmons, Christopher]

Depending on your LIS integration, the block barcode is scanned, there is a 
delay that you can specify, then the slide(s) ordered on that particular block 
are printed in the order they are ordered.
SlideMates have 2 versions, one sends a text file to the server to print the 
slide, the other sends a picture file that the slidemate copies, you will have 
to determine which one you want.
They are a great time-saver, cost-saver, and CAP compliant machine.

Chris Simmons B.S., A.S., HTL(ASCP)
Supervisor, UPP Dermatopathology
412.864.3880 office
412.612.0881 cell

-Original Message-
From: Histology Technician [mailto:histology81...@att.net] 
Sent: Wednesday, May 13, 2015 12:06 PM
To: kathy.mach...@lpnt.net; histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] : Re: HE Stainer Question

Does anyone have a Thermo PrintMate that you'd like to share the SOP on?  
Thanks! 


 On Tuesday, May 12, 2015 1:50 PM, kathy.mach...@lpnt.net 
kathy.mach...@lpnt.net wrote:
   

 We just got rid of our Leica XL.  It was not working right after about 10 
years, little things started going wrong. And then it just quit reading the 
white clip that we used for counterstaining IHC.  The cost for someone to come 
to the lab to fix it was ridiculous.  The customer service from Leica was not 
the best.
We now have a Gemini from Thermo and love it.  It has two ovens and stains 
similar to the Leica.  So far we have not had any problems and we LOVE our 
Thermo/Fisher reps.  They have been wonderful.

Kathy Machado, HTL
Danville Regional Medical Center
Danville, VA
kathy.mach...@lpnt.netmailto:kathy.mach...@lpnt.net
434-799-3867

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[Histonet] RE..... murine CD4, CD8 and CD68 for FFPE tissue

[Gayle Callis]

You wrote: 

 

Is anyone of you familiar with the possibility of applying CD4, CD8 and CD68
antibodies on formalin fixed paraffin embedded liver of mice?

 

Thanks in advance.

Joost Bruijntjes

 

**

Thankfully and at long last after years of frustration by many


 

eBioscience has both the CD8a and CD4 which is now available for FFPE murine
tissues.  Read the data sheets carefully as the CD4 indicated this antibody
was to be used on FFPE and not frozen sections.  

Anti-Mouse CD8a Purified Catalog Number: 14-0808 Also known as:
CD8 alpha, Ly-2, Ly-35, Lyt-2

Anti-Mouse CD4 Purified Catalog Number: 14-9766 Also known as:
L3T4, Ly-4 

 

Serotec has a rat anti mouse CD68 where the data sheet says can be used on
FFPE murine tissue.  This clone must be available from other companies as
well.  

 

Gayle M. Callis

HTL/HT/MT(ASCP) 

 

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Re: [Histonet] RE..... murine CD4, CD8 and CD68 for FFPE tissue

[Joseph Hardin]

Has anyone tried these Abs? If so, please send me your beautiful photos and 
protocol.

Joseph Hardin
Senior Research Specialist
UWCCC Experimental Pathology
WIMR I Rm. 4012
 Highland Av.
Madison, WI 53705
(608)262-1836



From: Gayle Callis gayle.cal...@bresnan.net
Sent: Wednesday, May 13, 2015 12:34 PM
To: Histonet
Subject: [Histonet] RE.  murine CD4, CD8 and CD68 for FFPE tissue

You wrote:



Is anyone of you familiar with the possibility of applying CD4, CD8 and CD68
antibodies on formalin fixed paraffin embedded liver of mice?



Thanks in advance.

Joost Bruijntjes



**

Thankfully and at long last after years of frustration by many




eBioscience has both the CD8a and CD4 which is now available for FFPE murine
tissues.  Read the data sheets carefully as the CD4 indicated this antibody
was to be used on FFPE and not frozen sections.

Anti-Mouse CD8a Purified Catalog Number: 14-0808 Also known as:
CD8 alpha, Ly-2, Ly-35, Lyt-2

Anti-Mouse CD4 Purified Catalog Number: 14-9766 Also known as:
L3T4, Ly-4



Serotec has a rat anti mouse CD68 where the data sheet says can be used on
FFPE murine tissue.  This clone must be available from other companies as
well.



Gayle M. Callis

HTL/HT/MT(ASCP)



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[Histonet] : Re: HE Stainer Question

[Kathy.Machado]

We just got rid of our Leica XL.  It was not working right after about 10 
years, little things started going wrong. And then it just quit reading the 
white clip that we used for counterstaining IHC.  The cost for someone to come 
to the lab to fix it was ridiculous.  The customer service from Leica was not 
the best.
We now have a Gemini from Thermo and love it.   It has two ovens and stains 
similar to the Leica.  So far we have not had any problems and we LOVE our 
Thermo/Fisher reps.  They have been wonderful.

Kathy Machado, HTL
Danville Regional Medical Center
Danville, VA
kathy.mach...@lpnt.netmailto:kathy.mach...@lpnt.net
434-799-3867

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[Histonet] RE: Question

[Morken, Timothy]

Bernice, You have to buy the 10N solution. You can only dilute a given normal 
solution, you cannot concentrate them.


Tim Morken
Pathology Site Manager, Parnassus 
Supervisor, Electron Microscopy/Neuromuscular Special Studies
Department of Pathology
UC San Francisco Medical Center






-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Bernice 
Frederick
Sent: Thursday, April 30, 2015 12:35 PM
To: Histonet@lists.utsouthwestern.edu
Subject: [Histonet] Question

All,
I have a procedure here that call for and I quote 1.25 ml NaOH 10N in 1L of 
water. I know how to get 1 N, but how do I get 10. Having rarely hd the 
opportunity to make many Normal solutions ,my brain is not computing. Is it an 
error?
Bernice

Bernice Frederick HTL (ASCP)
Senior Research Tech
Pathology Core Facility
Robert. H. Lurie Cancer Center
Northwestern University
710 N Fairbanks Court
Olson 8-421
Chicago,IL 60611
312-503-3723
b-freder...@northwestern.edumailto:b-freder...@northwestern.edu

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[Histonet] RE: IHC billing question

[Mike Pence]

Ouch!

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Cartun, Richard
Sent: Thursday, April 30, 2015 4:34 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] IHC billing question

Effective January 1, 2015, our LIS team removed all of the CPT 88342 codes for 
IHC from our CoPath stain dictionary since you couldn't tell whether a 
Cytokeratin-7 was being performed as an 88341 or as an 88342.  Now, as you 
might have expected,  none of the inpatient IHC testing has been accounted 
for (the outpatient IHC has been billed manually from the pathology report), 
and they want someone to go back and enter all the CPT codes into the system 
(hopefully, not me!).  Has anyone else encountered this problem?  Thanks (I 
think).

Richard

Richard W. Cartun, MS, PhD
Director, Histology  Immunopathology
Director, Biospecimen Collection Programs Assistant Director, Anatomic 
Pathology Hartford Hospital
80 Seymour Street
Hartford, CT  06102
(860) 972-1596
(860) 545-2204 Fax


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[Histonet] RE... Frozen section fixation problems

[Gayle Callis]

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 watera 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 

[Histonet] RE: Acetone fixation problems with OCT Tissues

[Connolly, Brett M]

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_conno...@merck.com
T- 215-652-2501
F- 215-993-6803



-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Lewis, Patrick
Sent: Tuesday, April 28, 2015 5:56 PM
To: (Histonet@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

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Re: [Histonet] RE: Nuclear Artifact

[WILLIAM DESALVO]

What type of tissue cassette is being used? What type of insert or wrap is 
used. If one cassette processes correctly and the next to it does not, hard to 
say tissue processor is causing issue.
Sounds like a water problem and it could be water trapped in cassette. Check 
the rest of you process before moving to the processor. 

Sent from my iPhone

 On Apr 29, 2015, at 9:13 AM, Burnett, Brandy bburn...@capecodhealth.org 
 wrote:
 
 We are having similar issues with our tissue. 
 Any troubleshooting insight would be greatly appreciated!
 Thanks!
 
 
 From: histonet-boun...@lists.utsouthwestern.edu 
 [histonet-boun...@lists.utsouthwestern.edu] on behalf of Sue 
 [suetp...@comcast.net]
 Sent: Tuesday, April 21, 2015 7:55 PM
 To: Lisa Roy
 Cc: histonet@lists.utsouthwestern.edu
 Subject: Re: [Histonet] Nuclear Artifact
 
 OMG we are experiencing the same issue. At first it was just GI and now we 
 are seeing it on prostate. One pathologist said it looks like the tissue has 
 been cooked. The only issue is we can have two biopsies right next to one 
 another in the basket one looks good and one looks bad. My director also 
 thinks it is the processors. I had Thermo out and they could find nothing. We 
 changed out all the reagents and the biopsies were fine than two days later 
 we had some bad ones. I know in July Fisher had a formalin recall associated 
 to the mixture of buffer, water and formalin. We thought that might be it but 
 it is now almost a year later and all the bad formalin should be gone. The 
 histotechs say the tissue is crunchy and they are right. I am running a test 
 tonight of a small needle biopsy that I made from a colon. I placed it is 
 straight formaldehyde overnight and am processing it on our biopsy cycle 
 tonight. My director also wanted us to only put three levels on our Thermo, 
 but he wanted the middle level to have empty baskets. I stopped that today 
 because I think the other issue is that the poor biopsies may be on the top 
 level and as the reagents are used the level changes, and also due to 
 displacement with the middle level being empty the reagent levels may not 
 reach the top. We just do not have the manpower to inspect every reagent 
 every day, we have 6 processor and it would take a tech all day. We actually 
 take a digital picture when they come out of the processor. I want to check 
 my problems cases tomorrow. We do not use sponges but the only other like was 
 the PA who was wrapping the blue paper very tight around the tissue. I really 
 do not think this is the issue though.. Any other insight would be greatly 
 appreciated.
 
 Susan T. Paturzo
 TJUH
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[Histonet] RE: Nuclear Artifact

[Burnett, Brandy]

We are having similar issues with our tissue. 
Any troubleshooting insight would be greatly appreciated!
Thanks!


From: histonet-boun...@lists.utsouthwestern.edu 
[histonet-boun...@lists.utsouthwestern.edu] on behalf of Sue 
[suetp...@comcast.net]
Sent: Tuesday, April 21, 2015 7:55 PM
To: Lisa Roy
Cc: histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] Nuclear Artifact

OMG we are experiencing the same issue. At first it was just GI and now we are 
seeing it on prostate. One pathologist said it looks like the tissue has been 
cooked. The only issue is we can have two biopsies right next to one another in 
the basket one looks good and one looks bad. My director also thinks it is the 
processors. I had Thermo out and they could find nothing. We changed out all 
the reagents and the biopsies were fine than two days later we had some bad 
ones. I know in July Fisher had a formalin recall associated to the mixture of 
buffer, water and formalin. We thought that might be it but it is now almost a 
year later and all the bad formalin should be gone. The histotechs say the 
tissue is crunchy and they are right. I am running a test tonight of a small 
needle biopsy that I made from a colon. I placed it is straight formaldehyde 
overnight and am processing it on our biopsy cycle tonight. My director also 
wanted us to only put three levels on our Thermo, but he wanted the middle 
level to have empty baskets. I stopped that today because I think the other 
issue is that the poor biopsies may be on the top level and as the reagents are 
used the level changes, and also due to displacement with the middle level 
being empty the reagent levels may not reach the top. We just do not have the 
manpower to inspect every reagent every day, we have 6 processor and it would 
take a tech all day. We actually take a digital picture when they come out of 
the processor. I want to check my problems cases tomorrow. We do not use 
sponges but the only other like was the PA who was wrapping the blue paper very 
tight around the tissue. I really do not think this is the issue though.. Any 
other insight would be greatly appreciated.

Susan T. Paturzo
TJUH
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Re: [Histonet] Re: H. Pylori Testing

[Garreyf]

In the past when using giemsa stain, 
I came across two human cases of very long helicobacter organisms.. I was 
stumped the first time since I had never seen one previously. I reflexed both 
to immuno and both were positive with the h pylori antibody. I assume they were 
both heilmani. I think it was called gastrospirillum in the past.I agree the 
immuno stain is much faster (easier)to look at.  I've never studied it but it's 
sensitivity is probably just a little better than giemsa. It's those cases with 
very few organisms that are more apt to  be missed using a non-immuno type of 
stain.  Agree about the contaminants as well  we alway went on the seagull 
shaped morphology though.


Garrey

Sent from my iPhone

 On Apr 29, 2015, at 2:04 PM, Bob Richmond rsrichm...@gmail.com wrote:
 
 Nancy Stedman observes:
 
 I believe IHC is more sensitive than the special stains too. One caveat
 for anyone who works with veterinary samples - the H. pylori antibodies are
 specific for H. pylori, so I have not found these antibodies to be helpful
 for evaluating other species with helicobacter-associated gastritis. One
 exception is the antibody made by Biocare which seems to stain some of the
 feline helicobacters, and maybe others too (have not tried).
 
 As far as I know, the only Helicobacter species other than H. pylori
 reported as a human pathogen is H. heilmanii - I've seen it exactly once,
 with a dye method - supposedly more common in Japan, often with a history
 of close association with cats. Supposedly H. heilmanii marks with the
 commercial IHC antibodies also.
 
 I don't think the data exist to show that IHC is more sensitive than the
 older dye methods, in terms of detecting clinical disease. As I noted
 before, the IHC is much faster for the pathologist to read. Also, many
 pathologists report any bacteria seen with dye methods as H. pylori,
 including the bacteria brought down by the endoscope from the oral cavity.
 
 Bob Richmond
 Samurai Pathologist
 Maryville TN
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[Histonet] RE: Recycling of alcohol

[Morken, Timothy]

Jim, yes, in my experience you are going to use this for 70 to 95% alcohol 
steps, not 100, unless you get a water absorber (BR has one, at least used 
to). 


Tim Morken
Pathology Site Manager, Parnassus 
Supervisor, Electron Microscopy/Neuromuscular Special Studies
Department of Pathology
UC San Francisco Medical Center



-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Vickroy, James
Sent: Wednesday, April 29, 2015 12:11 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Recycling of alcohol

In the past I have plenty of years of experience recycling formalin and 
clearing agents, however I don't have much experience with recycling reagent 
grade alcohols.  If we recycle waste alcohol that is 95% - 100 %, I am told 
that our recycled product will be 99% or above.   With that said how are others 
using this alcohol?   Are they using it as 100% on the tissue processors?   Are 
they using it in the strainers for 100%?  Are others using it as 95% instead of 
100%.   Obvioulsy I am concerned that my last alcohol station on the VIP is as 
close to 100% as possible and the same is true for the automated stainer.

Obviously we can use the recycled alcohol for cleaning reagents on the tissue 
processors.   Please share with me if you are recycling alcohol to what extent 
are you using the recycled product.

Jim

Jim Vickroy
Histology Manager
Springfield Clinic, Main Campus, East Building
1025 South 6th Street
Springfield, Illinois  62703
Office:  217-528-7541, Ext. 15121
Email:  jvick...@springfieldclinic.commailto:jvick...@springfieldclinic.com



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Re: [Histonet] Re: H. Pylori Testing

[wsimons]

I prefer (and my pathologist) the Warthin Starry.  I performed that routinely, 
and affordably, on the Dako Artisan at my last employer.
At my current small lab we perform a Giemsa for h.pylori.   I've seen two cases 
of  H. heilmanii  in the last 2 years.  Both stained with Giemsa.  

When my lab expands I have the Artisan on my wishlist for both IHC and Warthin 
Starry.  Thank you everyone for
the comments.

Wanda K. Simons, HT (ASCP)
Athens Gastroenterology Association
3320 Old Jefferson Road, Bldg.400
Athens, GA 30607
www.histosearch.com/gsh/


  ---Original Message---
  From: Garreyf garr...@gmail.com
  To: Bob Richmond rsrichm...@gmail.com
  Cc: Histonet@lists.utsouthwestern.edu histonet@lists.utsouthwestern.edu
  Subject: Re: [Histonet] Re: H. Pylori Testing
  Sent: Apr 29 '15 14:21
  
  In the past when using giemsa stain,
  I came across two human cases of very long helicobacter organisms.. I was 
 stumped the first time since I had never seen one previously. I reflexed both 
 to immuno and both were positive with the h pylori antibody. I assume they 
 were both heilmani. I think it was called gastrospirillum in the past.I agree 
 the immuno stain is much faster (easier)to look at.  I've never studied it 
 but it's sensitivity is probably just a little better than giemsa. It's those 
 cases with very few organisms that are more apt to  be missed using a 
 non-immuno type of stain.  Agree about the contaminants as well  we alway 
 went on the seagull shaped morphology though.
  
  
  Garrey
  
  Sent from my iPhone
  
   On Apr 29, 2015, at 2:04 PM, Bob Richmond rsrichm...@gmail.com wrote:
  
   Nancy Stedman observes:
  
   I believe IHC is more sensitive than the special stains too. One caveat
   for anyone who works with veterinary samples - the H. pylori antibodies are
   specific for H. pylori, so I have not found these antibodies to be helpful
   for evaluating other species with helicobacter-associated gastritis. One
   exception is the antibody made by Biocare which seems to stain some of the
   feline helicobacters, and maybe others too (have not tried).
  
   As far as I know, the only Helicobacter species other than H. pylori
   reported as a human pathogen is H. heilmanii - I've seen it exactly once,
   with a dye method - supposedly more common in Japan, often with a history
   of close association with cats. Supposedly H. heilmanii marks with the
   commercial IHC antibodies also.
  
   I don't think the data exist to show that IHC is more sensitive than the
   older dye methods, in terms of detecting clinical disease. As I noted
   before, the IHC is much faster for the pathologist to read. Also, many
   pathologists report any bacteria seen with dye methods as H. pylori,
   including the bacteria brought down by the endoscope from the oral cavity.
  
   Bob Richmond
   Samurai Pathologist
   Maryville TN
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Re: [Histonet] Re: H. Pylori Testing

[Bob Richmond]

Helicobacter heilmannii (sorry I misspelled it before) was named
Gastrospirillum hominis when it was first described. Tight cylindrical
spirals, unlike the gull-wing (like you learned to draw birds in the sky
when you were in the fourth grade) morphology of H. pylori. I'm not sure
you could see those tight cylindrical spirals clearly using IHC, and tell
them from gull-wings.

Sensitivity on Giemsa staining depends on microscopic technique. Real men
don't use oil immersion in pathology, but you need an oil lens to make the
morphologic confirmation, and to search the occasional slide that has
neutrophils in the pyloric lamina propria (chronic active gastritis), and
this is time-consuming and messy, and few pathologists do it.

Bob Richmond
Samurai Pathologist
Maryville TN

On Wed, Apr 29, 2015 at 2:18 PM, Garreyf garr...@gmail.com wrote:

 In the past when using giemsa stain,
 I came across two human cases of very long helicobacter organisms.. I was
 stumped the first time since I had never seen one previously. I reflexed
 both to immuno and both were positive with the h pylori antibody. I assume
 they were both heilmani. I think it was called gastrospirillum in the
 past.I agree the immuno stain is much faster (easier)to look at.  I've
 never studied it but it's sensitivity is probably just a little better than
 giemsa. It's those cases with very few organisms that are more apt to  be
 missed using a non-immuno type of stain.  Agree about the contaminants as
 well  we alway went on the seagull shaped morphology though.


 Garrey

 Sent from my iPhone

  On Apr 29, 2015, at 2:04 PM, Bob Richmond rsrichm...@gmail.com wrote:
 
  Nancy Stedman observes:
 
  I believe IHC is more sensitive than the special stains too. One caveat
  for anyone who works with veterinary samples - the H. pylori antibodies
 are
  specific for H. pylori, so I have not found these antibodies to be
 helpful
  for evaluating other species with helicobacter-associated gastritis. One
  exception is the antibody made by Biocare which seems to stain some of
 the
  feline helicobacters, and maybe others too (have not tried).
 
  As far as I know, the only Helicobacter species other than H. pylori
  reported as a human pathogen is H. heilmanii - I've seen it exactly once,
  with a dye method - supposedly more common in Japan, often with a history
  of close association with cats. Supposedly H. heilmanii marks with the
  commercial IHC antibodies also.
 
  I don't think the data exist to show that IHC is more sensitive than the
  older dye methods, in terms of detecting clinical disease. As I noted
  before, the IHC is much faster for the pathologist to read. Also, many
  pathologists report any bacteria seen with dye methods as H. pylori,
  including the bacteria brought down by the endoscope from the oral
 cavity.
 
  Bob Richmond
  Samurai Pathologist
  Maryville TN
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[Histonet] Re: H. Pylori Testing

[Bob Richmond]

Nancy Stedman observes:

I believe IHC is more sensitive than the special stains too. One caveat
for anyone who works with veterinary samples - the H. pylori antibodies are
specific for H. pylori, so I have not found these antibodies to be helpful
for evaluating other species with helicobacter-associated gastritis. One
exception is the antibody made by Biocare which seems to stain some of the
feline helicobacters, and maybe others too (have not tried).

As far as I know, the only Helicobacter species other than H. pylori
reported as a human pathogen is H. heilmanii - I've seen it exactly once,
with a dye method - supposedly more common in Japan, often with a history
of close association with cats. Supposedly H. heilmanii marks with the
commercial IHC antibodies also.

I don't think the data exist to show that IHC is more sensitive than the
older dye methods, in terms of detecting clinical disease. As I noted
before, the IHC is much faster for the pathologist to read. Also, many
pathologists report any bacteria seen with dye methods as H. pylori,
including the bacteria brought down by the endoscope from the oral cavity.

Bob Richmond
Samurai Pathologist
Maryville TN
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[Histonet] RE: URGENT: microwave oven for staining and antigen retrieval

[Liette Tougas]

Thank you so much you to the ones who already replied to me!  However I did not 
receive any comment about the H1850 or H2250 from Energy Beam Science (EBS).  
Does anyone have experience with any of these?

Thanks again in advance for a most appreciated prompt reply!

Liette Tougas, RT, B.Sc., M.Sc.
Biomedical Laboratory Technology Department
Dawson College
514-931-8731, ext 1519

From: histonet-boun...@lists.utsouthwestern.edu 
[histonet-boun...@lists.utsouthwestern.edu] on behalf of Liette Tougas 
[ltou...@dawsoncollege.qc.ca]
Sent: April 28, 2015 11:23 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] URGENT: microwave oven for staining and antigen retrieval

Hi all,



We need to purchase a vented laboratory microwave oven that will be used only 
for staining, secondary fixation (Bouin) and antigen retrieval (no processing, 
drying of slides or routine fixation).  It will be used only during 2 months of 
the year in the histotechniques course.  However, during the days it will be 
used, as students are going to be using it one after the other, ventilation has 
to be efficient (hooked up to central system).



I would appreciate any advice as to brand and model suggested ASAP.



Thanks so much in advance,



Liette Tougas, RT, B.Sc., M.Sc.
Biomedical Laboratory Technology Department
Dawson College, Montreal, Qc
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[Histonet] RE: Tri-State Histology Symposium

[WIAPP-MB-hymclab]

Last chance to get on the bandwagon and join our Tri-State histology meeting 
next week  It promises to be a great meeting!!  Go to 
www.whs.wildapricot.org to sign up or see full brochure.


-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of hymclab
Sent: Monday, March 02, 2015 11:07 AM
To: 'Teri Johnson'; colleen_herr...@bshsi.org
Cc: 'histonet@lists.utsouthwestern.edu'
Subject: [Histonet] Tri-State Histology Symposium


Dear Histonetters:  You are invited to join the histology societies of 
Wisconsin, Iowa and Minnesota as we celebrate Hats Off to Histology at the 
2015 Tri-State Histology Symposium, May 6-8 at The Madison Concourse Hotel and 
Governors Club in Madison, Wisconsin.

For program, registration and vendor/exhibit information contact the following 
representatives:

Wisconsin:  Kathryn Stoll kst...@mcw.edu

Iowa:  Judi Stasko judith.sta...@ars.usda.gov

Minnesota: Lois Rowe rowe.l...@mayo.edu

Vendor/Exhibit:  Dawn Schneider dawn.schnei...@ministryhealth.org




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[Histonet] Re: H. Pylori Testing

[Bob Richmond]

From the pathologist's viewpoint, immunohistochemical stains for
Helicobacter are much faster to read than are the old dye methods such as
Giemsa.

Ventana claims that you can break even with one of their stainers if you
produce 600 billable slides a year (unless they've changed that number - a
rep can tell you.

It's likely that we are going to get a lot more resistance in the future
from the third parties about paying for a Helicobacter stain done in
advance of the pathologist's looking at the slide. I'd think about that
before investing very much in anything.

Bob Richmond
Samurai Pathologist
Maryville TN
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[Histonet] RE: Centrifuge for cytospins

[Cooper, Brian]

We make our BAL cytospins directly--no centrifuging first.

Brian

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Piche, Jessica
Sent: Tuesday, April 28, 2015 8:58 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Centrifuge for cytospins

Hi Everyone,

Just asking a quick question for our Cyto department. Are BAL slides 
centrifuged and then made in to cytospin slides or do you just make the 
cytospin slides with no centrifuging?

Thank you,

Jessica Piche, HT(ASCP)



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[Histonet] Re: Fixation of frozen section

[Bob Richmond]

Fixation of frozen sections in surgical pathology: about the only purpose
of the fixation step is to protect the hematoxylin from contamination.
Alcohol works fine for this purpose.

Bob Richmond
Samurai Pathologist
Maryville TN
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[Histonet] RE: Centrifuge for cytospins

[Joe W. Walker, Jr.]

In all of my experiences with cytology preparations, the first step would be to 
centrifuge the specimen.  This concentrates the cells for the next step; 
cytocentrifuge, which is used for the presentation of the cells for evaluation. 
 Here is an excerpt from our procedure:

3.  Label a 50 ml centrifuge tube with the cytology accession label.
4.  Mix the specimen using a vortex mixer for 10-30 seconds.
5.  Pour contents of specimen from the original container into the 50 ml 
centrifuge tube and cap the tube.
6.  Centrifuge the specimen at 600g for 10 minutes (program 1) or 1200g for 
5 minutes (program 2).
7.  Pour off the supernatant and re-suspend the sediment in 0.5 to 1ml of 
Cytospin (green) fluid, depending on the cell button size.

Evaluating the number of drops you might need for your cytocentrifuge can be 
performed by taking a crop from the sediment, placing it on a slide, placing a 
24x50mm cover glass on the specimen and then observing the cellularity of the 
specimen.  I have document that illustrates this but I don't think the list 
serve will allow attachments.  You can email me privately for this document.

Joe W. Walker, Jr. MS, SCT(ASCP)CM
Manager of Anatomical Pathology, Microbiology and Reference
Rutland Regional Medical Center
160 Allen Street, Rutland, VT 05701
P: 802.747.1790  F: 802.747.6525
Email joewal...@rrmc.orgwww.rrmc.org

Our Vision:
To be the Best Community Healthcare System in New England

Rutland Regional...Vermont's 1st Hospital to Achieve Both ANCC Magnet 
Recognition® and the Governor's Award for Performance Excellence


-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Cooper, Brian
Sent: Tuesday, April 28, 2015 12:10 PM
To: Piche, Jessica; histonet@lists.utsouthwestern.edu
Subject: [Histonet] RE: Centrifuge for cytospins

We make our BAL cytospins directly--no centrifuging first.

Brian

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Piche, Jessica
Sent: Tuesday, April 28, 2015 8:58 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Centrifuge for cytospins

Hi Everyone,

Just asking a quick question for our Cyto department. Are BAL slides 
centrifuged and then made in to cytospin slides or do you just make the 
cytospin slides with no centrifuging?

Thank you,

Jessica Piche, HT(ASCP)



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RE: [Histonet] Re: H. Pylori Testing

[Stedman, Nancy]

I believe IHC is more sensitive than the special stains too.  One caveat for 
anyone who works with veterinary samples - the H. pylori antibodies are 
specific for H. pylori, so I have not found these antibodies to be helpful for 
evaluating other species with helicobacter-associated gastritis.  One exception 
is the antibody made by Biocare which seems to stain some of the feline 
helicobacters, and maybe others too (have not tried).

-Nancy Stedman

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Bob Richmond
Sent: Tuesday, April 28, 2015 12:33 PM
To: Histonet@lists.utsouthwestern.edu
Subject: [Histonet] Re: H. Pylori Testing

From the pathologist's viewpoint, immunohistochemical stains for
Helicobacter are much faster to read than are the old dye methods such as 
Giemsa.

Ventana claims that you can break even with one of their stainers if you 
produce 600 billable slides a year (unless they've changed that number - a rep 
can tell you.

It's likely that we are going to get a lot more resistance in the future from 
the third parties about paying for a Helicobacter stain done in advance of the 
pathologist's looking at the slide. I'd think about that before investing very 
much in anything.

Bob Richmond
Samurai Pathologist
Maryville TN
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[Histonet] Re: IHC and Oven Temperatures

[richardboen]

The specific pages in the Dako Education Guide: Immunohistochemistry 
Staining Methods, Fifth Edition are: Discussion on page 32 and 
references on page 33. It's in the Fixation and Processing Chapter and 
says no part of the process should have temperatures above 60C.


Rick Boen, BS, HTL (ASCP)
Histology Lab
St. Luke's Hospital
Duluth, Mn

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RE: [Histonet] Re: IHC and Oven Temperatures

[Sebree Linda A]

That's the reference I have always gone by since I first started doing IHC back 
in 19..

Linda A. Sebree

From: histonet-boun...@lists.utsouthwestern.edu 
[histonet-boun...@lists.utsouthwestern.edu] on behalf of 
richardb...@charter.net [richardb...@charter.net]
Sent: Sunday, April 26, 2015 10:42 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Re: IHC and Oven Temperatures

The specific pages in the Dako Education Guide: Immunohistochemistry
Staining Methods, Fifth Edition are: Discussion on page 32 and
references on page 33. It's in the Fixation and Processing Chapter and
says no part of the process should have temperatures above 60C.

Rick Boen, BS, HTL (ASCP)
Histology Lab
St. Luke's Hospital
Duluth, Mn

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RE: [Histonet] Re: IHC and Oven Temperatures

[Joelle Weaver]

Thank you. I knew it was discussed in that reference. I guess my memory isn't 
totally gone!


Joelle Weaver MAOM, HTL (ASCP) QIHC


  

 
 Date: Sun, 26 Apr 2015 11:42:04 -0400
 From: richardb...@charter.net
 To: histonet@lists.utsouthwestern.edu
 Subject: [Histonet] Re: IHC and Oven Temperatures
 
 
 The specific pages in the Dako Education Guide: Immunohistochemistry 
 Staining Methods, Fifth Edition are: Discussion on page 32 and 
 references on page 33. It's in the Fixation and Processing Chapter and 
 says no part of the process should have temperatures above 60C.
 
 Rick Boen, BS, HTL (ASCP)
 Histology Lab
 St. Luke's Hospital
 Duluth, Mn
 
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[Histonet] RE: Nuclear Artifact

[Tony Henwood (SCHN)]

This seems like a classic case of drying of biopsies prior to fixation. This 
can occur if biopsies are placed on absorbent paper (or on disinfecting alcohol 
swabs, heaven forbid).

From: histonet-boun...@lists.utsouthwestern.edu 
[histonet-boun...@lists.utsouthwestern.edu] on behalf of Sue 
[suetp...@comcast.net]
Sent: Friday, 24 April 2015 10:51 AM
To: Lisa Roy
Cc: histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] Nuclear Artifact

Hi All

So I have been seeing the same issue as I stated in past e-mails. I did one 
test and fixed a colon biopsy in formaldehyde and left it in overnight and 
processed next day. I was hoping that I could reproduce the artifact. The 
tissue was beautiful. At my pathologists requests we changed the paraffin 
temperature tonight is the first night. I do not think this i the issue. We are 
going to transfer our biopsies to another tissue processor just for test. I 
brought up in the past that i think the issue starts prior to the histo lab, my 
pathologist tended to disagree, but I think he is chaining his mind since my 
one common detonator is a PA. I do not think that she wets her blue wrap paper 
enough and the tissue sits on the paper dry she also fold the paper so tight 
that it is possible for the small biopsies to get stuck in a fold. My 
pathologists actually came in and said he thought I may be right. Wow. That is 
my next test. We are requiring our staff to do so much work that they tend to 
rush and as I have stated in the past grossing sets the tone for every step the 
nistotech is responsible for and if it is not prepared correctly in the gross 
lab the histologist cannot fix it. An old adage we are not magicians.

Sue Paturzo
TJUH
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[Histonet] RE: NY State Histology License

[Anne Murvosh]

There is a grandfather clause if you were a histotech by april 24.1995.  I 
asked ClIA recently when they came to inspect me, as I fell into this category 
but didn't know if you had to have the licence by then or just have been 
performing high complexity test at that time (my licence was later than that).  
The CLIA person couldn't decipher their own rules and was going to ask her 
supervisor.  I never heard back.  As for IHC any HT in our state can do it and 
CAP never brought it up when I worked at the hospital.

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Gail Marcella
Sent: Tuesday, April 21, 2015 8:48 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] NY State Histology License

Hi  - I've been a Histotech for 20+ years and got my Clinical Laboratory 
License in NY State when they required getting it. I don't have an Associates 
or Bachelor's degree but a Pathologist signed off for me. I have my HTASCP. I 
was told when I went for an interview in NY State that I couldn't gross small 
specimens or do IHC without an Associates  or Bachelor degree in biology. I was 
not aware of these restrictions. I don't see anything on the NYS website.  I 
was wondering if anyone else heard of this? Thanks - Gail
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[Histonet] RE: Histonet Digest, Vol 137, Issue 30

[Mayer,Toysha N]

Gail,
The regulation is from CLIA '88.  They list the requirements for that, and I 
have never heard of a grandfather clause.  I could be wrong about though.
NY state did not license HTL.  I had a student who wanted to move there, and 
did not because he was going to have an HTL, not HT.  He would only have been 
able to work clinical if he took both exams (HT and HTL).  I even contacted the 
state society for clarification.  Hopefully it has changed.
I believe it is because they do not have any HTL programs in the state, so it 
was not included in the licensure bill. 

Sincerely,

Toysha N. Mayer, D.H.Sc., MBA, HT (ASCP)
Instructor/Education Coordinator
Program in Histotechnology
School of Health Professions
UT M.D. Anderson Cancer Center
713.563-3481


Message: 8
Date: Thu, 23 Apr 2015 18:34:13 +
From: Joelle Weaver joellewea...@hotmail.com
Subject: RE: [Histonet] NY State Histology License
To: Gail Marcella gmarce...@nj-urology.com,
histonet@lists.utsouthwestern.edu
histonet@lists.utsouthwestern.edu
Message-ID: snt149-w8666703aac97b9793fdbc9d8...@phx.gbl
Content-Type: text/plain; charset=iso-8859-1

Yes, CLIA stipulation. I think that there may be a grandfather clause, but not 
sure of the time frame. You could check the regulation on that.


Joelle Weaver MAOM, HTL (ASCP) QIHC


  

 
 From: gmarce...@nj-urology.com
 To: histonet@lists.utsouthwestern.edu
 Date: Tue, 21 Apr 2015 11:47:58 -0400
 Subject: [Histonet] NY State Histology License
 
 Hi  - I've been a Histotech for 20+ years and got my Clinical Laboratory 
 License in NY State when they required getting it. I don't have an Associates 
 or Bachelor's degree but a Pathologist signed off for me. I have my HTASCP. I 
 was told when I went for an interview in NY State that I couldn't gross small 
 specimens or do IHC without an Associates  or Bachelor degree in biology. I 
 was not aware of these restrictions. I don't see anything on the NYS website. 
  I was wondering if anyone else heard of this? Thanks - Gail
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RE: [Histonet] RE: Question on IHC billing

[Joelle Weaver]

Yes, that is what we do. It is per specimen. First IHC 88342, additional are 
88341


Joelle Weaver MAOM, HTL (ASCP) QIHC


  

 
 From: joyce.we...@emoryhealthcare.org
 To: jvick...@springfieldclinic.com; histonet@lists.utsouthwestern.edu
 Date: Thu, 23 Apr 2015 20:31:09 +
 CC: 
 Subject: [Histonet] RE: Question on IHC billing
 
 Correct
 
 Joyce Weems
 Pathology Manager
 678-843-7376 Phone
 678-843-7831 Fax
 joyce.we...@emoryhealthcare.org
 
 
 
 www.saintjosephsatlanta.org
 5665 Peachtree Dunwoody Road
 Atlanta, GA 30342
 
 This e-mail, including any attachments is the property of Saint Joseph's 
 Hospital and is intended for the sole use of the intended recipient(s).  It 
 may contain information that is privileged and confidential.  Any 
 unauthorized review, use, disclosure, or distribution is prohibited. If you 
 are not the intended recipient, please delete this message, and reply to the 
 sender regarding the error in a separate email.
 
 
 -Original Message-
 From: histonet-boun...@lists.utsouthwestern.edu 
 [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Vickroy, James
 Sent: Thursday, April 23, 2015 4:19 PM
 To: histonet@lists.utsouthwestern.edu
 Subject: [Histonet] Question on IHC billing
 
 
 Let me see if I have this straight:If a pathologist orders an Hpylori 
 stain on 2 blocks from the same specimen C1 and C2 we can only bill one 88342.
 
 If this correct.Obviously if he ordered addition different IHC stains we 
 could change additional 88341's.
 
 Jim Vickroy
 Histology Manager
 Springfield Clinic, Main Campus, East Building
 1025 South 6th Street
 Springfield, Illinois  62703
 Office:  217-528-7541, Ext. 15121
 Email:  jvick...@springfieldclinic.commailto:jvick...@springfieldclinic.com
 
 
 
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 prohibited. If you have received this electronic message in error, please 
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[Histonet] RE: Can you please add me to this mailing list

[Weems, Joyce K.]

Hi Chris,

You must go to http://lists.utsouthwestern.edu/mailman/listinfo/histonetand 
sign yourself up..

Welcome!!!


Joyce Weems
Pathology Manager
678-843-7376 Phone
678-843-7831 Fax
joyce.we...@emoryhealthcare.org



www.saintjosephsatlanta.org
5665 Peachtree Dunwoody Road
Atlanta, GA 30342

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regarding the error in a separate email.


-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Chris Ringler
Sent: Thursday, April 16, 2015 4:57 PM
To: Histonet@lists.utsouthwestern.edu
Subject: [Histonet] Can you please add me to this mailing list

cring...@ironwooddermatology.com



Respectfully,



Chris



Chris Ringler

CEO

Ironwood Dermatology PC

1735 E Skyline Dr

Tucson AZ  85718

520-618-1630 ext. 106   I   Fax 520-618-1636



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[Histonet] RE: Nuclear Artifact

[Morken, Timothy]

One pathologist said it looks like the tissue has been cooked.  Which could 
also be drying artifact after bx, before formalin.

The only issue is we can have two biopsies right next to one another in the 
basket one looks good and one looks bad. My director also thinks it is the 
processors.

Same processor but two results? Solving intermittent problems takes time to 
check variables - time almost no one wants to spend to check out every 
possibility. But if one variable is the same for both, and the result is 
different, then most likely it is a different variable causing the problem.

I think the other idea suggested of checking the handling of the tissue at the 
source of the biopsy is more likely to shed some light on this issue. 


Tim Morken
Pathology Site Manager, Parnassus 
Supervisor, Electron Microscopy/Neuromuscular Special Studies
Department of Pathology
UC San Francisco Medical Center





-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Sue
Sent: Tuesday, April 21, 2015 4:55 PM
To: Lisa Roy
Cc: histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] Nuclear Artifact

OMG we are experiencing the same issue. At first it was just GI and now we are 
seeing it on prostate. One pathologist said it looks like the tissue has been 
cooked. The only issue is we can have two biopsies right next to one another in 
the basket one looks good and one looks bad. My director also thinks it is the 
processors. I had Thermo out and they could find nothing. We changed out all 
the reagents and the biopsies were fine than two days later we had some bad 
ones. I know in July Fisher had a formalin recall associated to the mixture of 
buffer, water and formalin. We thought that might be it but it is now almost a 
year later and all the bad formalin should be gone. The histotechs say the 
tissue is crunchy and they are right. I am running a test tonight of a small 
needle biopsy that I made from a colon. I placed it is straight formaldehyde 
overnight and am processing it on our biopsy cycle tonight. My director also 
wanted us to only put three levels on our Thermo, but he wanted the middle 
level to have empty baskets. I stopped that today because I think the other 
issue is that the poor biopsies may be on the top level and as the reagents are 
used the level changes, and also due to displacement with the middle level 
being empty the reagent levels may not reach the top. We just do not have the 
manpower to inspect every reagent every day, we have 6 processor and it would 
take a tech all day. We actually take a digital picture when they come out of 
the processor. I want to check my problems cases tomorrow. We do not use 
sponges but the only other like was the PA who was wrapping the blue paper very 
tight around the tissue. I really do not think this is the issue though.. Any 
other insight would be greatly appreciated. 

Susan T. Paturzo 
TJUH 
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RE: [Histonet] RE: Nuclear Artifact

[Arbaugh, Roberta]

We have had the same problem. We come to the conclusion that it was water 
droplets. We had the problem when our humidity was high in the lab, or our 
weekend run. The changes we made where :
1. We no longer process over the weekend . We cannot count on our heating and 
cooling in our building.
2. We place a towel and a thick piece of cardboard on top of the processor lid.
3.We do not use the really fine mesh cassettes. We will use formalin soaked 
sponges or perm papers.
4. We do not over pack cassette basket.
We had our processor check every time we saw the artifact and they could never 
find a problem with the processor.
Hope this help, Roberta

-Original Message-
From: Morken, Timothy [mailto:timothy.mor...@ucsf.edu]
Sent: Wednesday, April 22, 2015 11:16 AM
To: Sue; Lisa Roy
Cc: histonet@lists.utsouthwestern.edu
Subject: [Histonet] RE: Nuclear Artifact

One pathologist said it looks like the tissue has been cooked.  Which could 
also be drying artifact after bx, before formalin.

The only issue is we can have two biopsies right next to one another in the 
basket one looks good and one looks bad. My director also thinks it is the 
processors.

Same processor but two results? Solving intermittent problems takes time to 
check variables - time almost no one wants to spend to check out every 
possibility. But if one variable is the same for both, and the result is 
different, then most likely it is a different variable causing the problem.

I think the other idea suggested of checking the handling of the tissue at the 
source of the biopsy is more likely to shed some light on this issue.


Tim Morken
Pathology Site Manager, Parnassus
Supervisor, Electron Microscopy/Neuromuscular Special Studies Department of 
Pathology UC San Francisco Medical Center





-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Sue
Sent: Tuesday, April 21, 2015 4:55 PM
To: Lisa Roy
Cc: histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] Nuclear Artifact

OMG we are experiencing the same issue. At first it was just GI and now we are 
seeing it on prostate. One pathologist said it looks like the tissue has been 
cooked. The only issue is we can have two biopsies right next to one another in 
the basket one looks good and one looks bad. My director also thinks it is the 
processors. I had Thermo out and they could find nothing. We changed out all 
the reagents and the biopsies were fine than two days later we had some bad 
ones. I know in July Fisher had a formalin recall associated to the mixture of 
buffer, water and formalin. We thought that might be it but it is now almost a 
year later and all the bad formalin should be gone. The histotechs say the 
tissue is crunchy and they are right. I am running a test tonight of a small 
needle biopsy that I made from a colon. I placed it is straight formaldehyde 
overnight and am processing it on our biopsy cycle tonight. My director also 
wanted us to only put three levels on our Thermo, but he wanted the middle 
level to have empty baskets. I stopped that today because I think the other 
issue is that the poor biopsies may be on the top level and as the reagents are 
used the level changes, and also due to displacement with the middle level 
being empty the reagent levels may not reach the top. We just do not have the 
manpower to inspect every reagent every day, we have 6 processor and it would 
take a tech all day. We actually take a digital picture when they come out of 
the processor. I want to check my problems cases tomorrow. We do not use 
sponges but the only other like was the PA who was wrapping the blue paper very 
tight around the tissue. I really do not think this is the issue though.. Any 
other insight would be greatly appreciated.

Susan T. Paturzo
TJUH
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Re: [Histonet] RE: Nuclear Artifact

[Sue]

we do not use freeze spray in the lab at all. this issue has really gotten me 
bummed out since it is so sporadic. I am going to try to see if it is 
associated to one day in particular. it is one PA but multiple techs have 
identified the issue. 
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RE: [Histonet] RE: Nuclear Artifact

[Jennifer MacDonald]

Our pathologists also complained about some of the GI biopsies looking 
burnt.  We tracked all of the problem cases back to one histotech.  The 
histotech was causing the burn artifact with excessive use of freezing 
spray. 



From:   Arbaugh, Roberta rarba...@csdermatology.com
To: 'Morken, Timothy' timothy.mor...@ucsf.edu, Sue 
suetp...@comcast.net, Lisa Roy ro...@labcorp.com
Cc: histonet@lists.utsouthwestern.edu 
histonet@lists.utsouthwestern.edu
Date:   04/22/2015 11:32 AM
Subject:RE: [Histonet] RE: Nuclear Artifact
Sent by:histonet-boun...@lists.utsouthwestern.edu



We have had the same problem. We come to the conclusion that it was water 
droplets. We had the problem when our humidity was high in the lab, or our 
weekend run. The changes we made where :
1. We no longer process over the weekend . We cannot count on our heating 
and cooling in our building.
2. We place a towel and a thick piece of cardboard on top of the processor 
lid.
3.We do not use the really fine mesh cassettes. We will use formalin 
soaked sponges or perm papers.
4. We do not over pack cassette basket.
We had our processor check every time we saw the artifact and they could 
never find a problem with the processor.
Hope this help, Roberta

-Original Message-
From: Morken, Timothy [mailto:timothy.mor...@ucsf.edu]
Sent: Wednesday, April 22, 2015 11:16 AM
To: Sue; Lisa Roy
Cc: histonet@lists.utsouthwestern.edu
Subject: [Histonet] RE: Nuclear Artifact

One pathologist said it looks like the tissue has been cooked.  Which 
could also be drying artifact after bx, before formalin.

The only issue is we can have two biopsies right next to one another in 
the basket one looks good and one looks bad. My director also thinks it is 
the processors.

Same processor but two results? Solving intermittent problems takes time 
to check variables - time almost no one wants to spend to check out every 
possibility. But if one variable is the same for both, and the result is 
different, then most likely it is a different variable causing the 
problem.

I think the other idea suggested of checking the handling of the tissue at 
the source of the biopsy is more likely to shed some light on this issue.


Tim Morken
Pathology Site Manager, Parnassus
Supervisor, Electron Microscopy/Neuromuscular Special Studies Department 
of Pathology UC San Francisco Medical Center





-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu [
mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Sue
Sent: Tuesday, April 21, 2015 4:55 PM
To: Lisa Roy
Cc: histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] Nuclear Artifact

OMG we are experiencing the same issue. At first it was just GI and now we 
are seeing it on prostate. One pathologist said it looks like the tissue 
has been cooked. The only issue is we can have two biopsies right next to 
one another in the basket one looks good and one looks bad. My director 
also thinks it is the processors. I had Thermo out and they could find 
nothing. We changed out all the reagents and the biopsies were fine than 
two days later we had some bad ones. I know in July Fisher had a formalin 
recall associated to the mixture of buffer, water and formalin. We thought 
that might be it but it is now almost a year later and all the bad 
formalin should be gone. The histotechs say the tissue is crunchy and they 
are right. I am running a test tonight of a small needle biopsy that I 
made from a colon. I placed it is straight formaldehyde overnight and am 
processing it on our biopsy cycle tonight. My director also wanted us to 
only put three levels on our Thermo, but he wanted the middle level to 
have empty baskets. I stopped that today because I think the other issue 
is that the poor biopsies may be on the top level and as the reagents are 
used the level changes, and also due to displacement with the middle level 
being empty the reagent levels may not reach the top. We just do not have 
the manpower to inspect every reagent every day, we have 6 processor and 
it would take a tech all day. We actually take a digital picture when they 
come out of the processor. I want to check my problems cases tomorrow. We 
do not use sponges but the only other like was the PA who was wrapping the 
blue paper very tight around the tissue. I really do not think this is the 
issue though.. Any other insight would be greatly appreciated.

Susan T. Paturzo
TJUH
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DISCLAIMER: The information in this message is confidential and may be 
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Re: [Histonet] RE: Nuclear Artifact

[mucram11]

We don't even allow freeze spray in the lab.  

Sent from my Verizon 4G LTE Smartphone


-- Original message--
From: Jennifer MacDonald
Date: Wed, Apr 22, 2015 5:34 PM
To: Arbaugh, Roberta;
Cc: histonet@lists.utsouthwestern.edu;Lisa 
Roy;histonet-boun...@lists.utsouthwestern.edu;'Morken, Timothy';
Subject:RE: [Histonet] RE: Nuclear Artifact

Our pathologists also complained about some of the GI biopsies looking 
burnt.  We tracked all of the problem cases back to one histotech.  The 
histotech was causing the burn artifact with excessive use of freezing 
spray. 



From:   Arbaugh, Roberta 
To: 'Morken, Timothy' , Sue 
, Lisa Roy 
Cc: histonet@lists.utsouthwestern.edu 

Date:   04/22/2015 11:32 AM
Subject:RE: [Histonet] RE: Nuclear Artifact
Sent by:histonet-boun...@lists.utsouthwestern.edu



We have had the same problem. We come to the conclusion that it was water 
droplets. We had the problem when our humidity was high in the lab, or our 
weekend run. The changes we made where :
1. We no longer process over the weekend . We cannot count on our heating 
and cooling in our building.
2. We place a towel and a thick piece of cardboard on top of the processor 
lid.
3.We do not use the really fine mesh cassettes. We will use formalin 
soaked sponges or perm papers.
4. We do not over pack cassette basket.
We had our processor check every time we saw the artifact and they could 
never find a problem with the processor.
Hope this help, Roberta

-Original Message-
From: Morken, Timothy [mailto:timothy.mor...@ucsf.edu]
Sent: Wednesday, April 22, 2015 11:16 AM
To: Sue; Lisa Roy
Cc: histonet@lists.utsouthwestern.edu
Subject: [Histonet] RE: Nuclear Artifact

One pathologist said it looks like the tissue has been cooked.  Which 
could also be drying artifact after bx, before formalin.

The only issue is we can have two biopsies right next to one another in 
the basket one looks good and one looks bad. My director also thinks it is 
the processors.

Same processor but two results? Solving intermittent problems takes time 
to check variables - time almost no one wants to spend to check out every 
possibility. But if one variable is the same for both, and the result is 
different, then most likely it is a different variable causing the 
problem.

I think the other idea suggested of checking the handling of the tissue at 
the source of the biopsy is more likely to shed some light on this issue.


Tim Morken
Pathology Site Manager, Parnassus
Supervisor, Electron Microscopy/Neuromuscular Special Studies Department 
of Pathology UC San Francisco Medical Center





-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu [
mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Sue
Sent: Tuesday, April 21, 2015 4:55 PM
To: Lisa Roy
Cc: histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] Nuclear Artifact

OMG we are experiencing the same issue. At first it was just GI and now we 
are seeing it on prostate. One pathologist said it looks like the tissue 
has been cooked. The only issue is we can have two biopsies right next to 
one another in the basket one looks good and one looks bad. My director 
also thinks it is the processors. I had Thermo out and they could find 
nothing. We changed out all the reagents and the biopsies were fine than 
two days later we had some bad ones. I know in July Fisher had a formalin 
recall associated to the mixture of buffer, water and formalin. We thought 
that might be it but it is now almost a year later and all the bad 
formalin should be gone. The histotechs say the tissue is crunchy and they 
are right. I am running a test tonight of a small needle biopsy that I 
made from a colon. I placed it is straight formaldehyde overnight and am 
processing it on our biopsy cycle tonight. My director also wanted us to 
only put three levels on our Thermo, but he wanted the middle level to 
have empty baskets. I stopped that today because I think the other issue 
is that the poor biopsies may be on the top level and as the reagents are 
used the level changes, and also due to displacement with the middle level 
being empty the reagent levels may not reach the top. We just do not have 
the manpower to inspect every reagent every day, we have 6 processor and 
it would take a tech all day. We actually take a digital picture when they 
come out of the processor. I want to check my problems cases tomorrow. We 
do not use sponges but the only other like was the PA who was wrapping the 
blue paper very tight around the tissue. I really do not think this is the 
issue though.. Any other insight would be greatly appreciated.

Susan T. Paturzo
TJUH
___
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DISCLAIMER: The information in this message is confidential and may be 
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RE: [Histonet] RE: Nuclear Artifact

[Joelle Weaver]

When I had this occur recently and sporadically, it was a collection issue. 


Joelle Weaver MAOM, HTL (ASCP) QIHC


  

 
 From: lbla...@digestivespecialists.com
 To: ro...@labcorp.com; histonet@lists.utsouthwestern.edu
 Date: Tue, 21 Apr 2015 13:48:07 -0400
 CC: 
 Subject: [Histonet] RE: Nuclear Artifact
 
 The first place I would look is to what may be happening before they reach 
 me.  If it's only one site with an issue, it sounds more like an issue at 
 collection.
 Linda
 
 -Original Message-
 From: histonet-boun...@lists.utsouthwestern.edu 
 [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Roy, Lisa
 Sent: Tuesday, April 21, 2015 1:41 PM
 To: histonet@lists.utsouthwestern.edu
 Subject: [Histonet] Nuclear Artifact
 
 Hi HistoNetters:
 I have run into quite a problem.  My lab currently processes all tissue types 
 from 3 different sites.  Recently, we have been getting complaints from one 
 of the sites that the biopsies have a nuclear artifact.  It is described as 
 washed out or poor to no nuclear detail.  Pictures have been uploaded 
 (Nuclear Artifact).  The Medical Director at said site is convinced that a 
 processor error is occurring.  Our site is not seeing this on any of our 
 slides.  Biopsies from all three sites are processed and embed together.  We 
 have done all trouble shooting that we can think of.  Leica service has come 
 to inspect our Peloris processor and all areas checked out as functioning 
 properly.  The problem is not consistent daily.  Seems to be worst toward the 
 end of the week.
 We have been running the same processing protocol, staining protocol and 
 cutting protocols for years now.  This problem has just developed over the 
 last 2 months.  Any ideas, no matter how far-fetched, would be greatly 
 appreciated at this point.
 
 Lisa Roy, HT(ASCP)
 Histology Supervisor
 LabCorp at St. Vincent Hospital
 123 Summer St
 Worcester, MA
 (508)363-9420
 
 -This e-mail and any attachments may contain CONFIDENTIAL information, 
 including PROTECTED HEALTH INFORMATION. If you are not the intended 
 recipient, any use or disclosure of this information is STRICTLY PROHIBITED; 
 you are requested to delete this e-mail and any attachments, notify the 
 sender immediately, and notify the LabCorp Privacy Officer at 
 privacyoffi...@labcorp.com or call (877) 23-HIPAA / (877) 234-4722. 
 ___
 Histonet mailing list
 Histonet@lists.utsouthwestern.edu
 http://lists.utsouthwestern.edu/mailman/listinfo/histonet
 
 ___
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 Histonet@lists.utsouthwestern.edu
 http://lists.utsouthwestern.edu/mailman/listinfo/histonet
  
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[Histonet] RE: (no subject)

[Mayer,Toysha N]

-I think you can use the other tissue controls. 
 Even CDC (Hansen's Disease Center) will send out mycobacterium leprae controls 
free of charge, and I think they are armadillo.
The best control I have ever used for mast cells is canine mast cell tumors.  I 
request them from Vet Schools regularly.
Also think of antibodies, aren't  most antibodies animal?
The separation of the specimens I think is during processing.  They should be 
processed separately from routine human tissues when they are being used for 
clinical human tests.
I have heard of them running on the same processor, just on a different run 
when the solutions have been changed.

Toysha
-

Message: 10
Date: Mon, 20 Apr 2015 18:50:15 -0400
From: Garrey Faller garr...@gmail.com
Subject: Re: [Histonet] (no subject)
To: koelli...@comcast.net
Cc: histonet@lists.utsouthwestern.edu
histonet@lists.utsouthwestern.edu
Message-ID:
CAF2sxrVD7o96Wz84nDBR6uxf=93tpqpmueqwog57e4fetcc...@mail.gmail.com
Content-Type: text/plain; charset=UTF-8

Here is the CAP checklist requirement:
ANP.21450
All  histochemical stains are of adequate quality, and daily controls are
demonstrated on each day of use for the tissue components or organism for
which they were designed.

Ray...you should call the CAP and ask for guidance on this.
My interpretation of this requirement is that it should be OK to use a
fungus from an orange peel. An orange peel fungus should have the same
staining characteristics as a candida or aspergillus etc.  Similarly a
bacteria is a bacteria. If you can produce a control that has both gram
positives and negatives, it should be OK. But, don't quote me on this.

Call the CAP for a definitive answer. I am interested in their response.
Garrey

On Sun, Apr 19, 2015 at 9:06 PM, koelli...@comcast.net wrote:

 I asked about this in a different vein months ago.  Has anyone shown a
 strawberry or ground meat or slim jim or orange peel as a bacteria/fungus
 control used for diagnostics to an inspector inspecting the lab and was
 there any comment from the inspector either positive or negative. Never
 heard back anything.
 Ray, Lake Forest Park, WA

 - Original Message -

 From: tjfinney2...@gmail.com
 To: histonet@lists.utsouthwestern.edu
 Sent: Sunday, April 19, 2015 5:24:53 PM
 Subject: [Histonet] (no subject)

 GMS controls
 From my understanding we can't use non human controls on patients. I
 could be wrong, but you may want to look into it.

 Happy Connecting.  Sent from my Sprint Phone.

 ___
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--


Sent from my iPhone

 On 21 Apr 2015, at 8:51 am, Garrey Faller garr...@gmail.com wrote:

 Here is the CAP checklist requirement:
 ANP.21450
 All  histochemical stains are of adequate quality, and daily controls are
 demonstrated on each day of use for the tissue components or organism for
 which they were designed.

 Ray...you should call the CAP and ask for guidance on this.
 My interpretation of this requirement is that it should be OK to use a
 fungus from an orange peel. An orange peel fungus should have the same
 staining characteristics as a candida or aspergillus etc.  Similarly a
 bacteria is a bacteria. If you can produce a control that has both gram
 positives and negatives, it should be OK. But, don't quote me on this.

 Call the CAP for a definitive answer. I am interested in their response.
 Garrey

 On Sun, Apr 19, 2015 at 9:06 PM, koelli...@comcast.net wrote:

 I asked about this in a different vein months ago.  Has anyone shown a
 strawberry or ground meat or slim jim or orange peel as a bacteria/fungus
 control used for diagnostics to an inspector inspecting the lab and was
 there any comment from the inspector either positive or negative. Never
 heard back anything.
 Ray, Lake Forest Park, WA

 - Original Message -

 From: tjfinney2...@gmail.com
 To: histonet@lists.utsouthwestern.edu
 Sent: Sunday, April 19, 2015 5:24:53 PM
 Subject: [Histonet] (no subject)

 GMS controls
 From my understanding we can't use non human controls on patients. I
 could be wrong, but you may want to look into it.







--

Message: 14
Date: Tue, 21 Apr 2015 13:49:21 + (UTC)
From: koelli...@comcast.net
Subject: Re: [Histonet] controls to lengthy off topic
To: Garrey Faller garr...@gmail.com
Cc: histonet@lists.utsouthwestern.edu
Message-ID:
2017966350.7376432.1429624161902.javamail.zim...@comcast.net
Content-Type: text/plain; charset=utf-8

Hello Garrey, 
Curious myself, CAP contact info seems to be greyed out on website unless I 
officially log in and for now my concerns are with the Washington State Science 

[Histonet] RE: Nuclear Artifact

[Podawiltz, Thomas]

I had this issue a couple of years back. Found out that there was a delay of 
over an hour from the time the specimen was collected to the time it was being 
put into fixative. 

Tom 


Tom Podawiltz HT (ASCP)
AP  Section Head 
LRGHealthcare
 




-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Roy, Lisa
Sent: Tuesday, April 21, 2015 1:41 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Nuclear Artifact

Hi HistoNetters:
I have run into quite a problem.  My lab currently processes all tissue types 
from 3 different sites.  Recently, we have been getting complaints from one of 
the sites that the biopsies have a nuclear artifact.  It is described as 
washed out or poor to no nuclear detail.  Pictures have been uploaded 
(Nuclear Artifact).  The Medical Director at said site is convinced that a 
processor error is occurring.  Our site is not seeing this on any of our 
slides.  Biopsies from all three sites are processed and embed together.  We 
have done all trouble shooting that we can think of.  Leica service has come to 
inspect our Peloris processor and all areas checked out as functioning 
properly.  The problem is not consistent daily.  Seems to be worst toward the 
end of the week.
We have been running the same processing protocol, staining protocol and 
cutting protocols for years now.  This problem has just developed over the last 
2 months.  Any ideas, no matter how far-fetched, would be greatly appreciated 
at this point.

Lisa Roy, HT(ASCP)
Histology Supervisor
LabCorp at St. Vincent Hospital
123 Summer St
Worcester, MA
(508)363-9420

-This e-mail and any attachments may contain CONFIDENTIAL information, 
including PROTECTED HEALTH INFORMATION. If you are not the intended recipient, 
any use or disclosure of this information is STRICTLY PROHIBITED; you are 
requested to delete this e-mail and any attachments, notify the sender 
immediately, and notify the LabCorp Privacy Officer at 
privacyoffi...@labcorp.com or call (877) 23-HIPAA / (877) 234-4722. 
___
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THIS MESSAGE IS CONFIDENTIAL.  
This e-mail message and any attachments are proprietary and confidential 
information intended only for the use of the recipient(s) named above. If you 
are not the intended recipient, you may not print,distribute, or copy this 
message or any attachments.  If you have received this communication in error, 
please notify the sender by return e-mail and delete this message and any 
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[Histonet] RE: Nuclear Artifact

[Blazek, Linda]

The first place I would look is to what may be happening before they reach me.  
If it's only one site with an issue, it sounds more like an issue at collection.
Linda

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Roy, Lisa
Sent: Tuesday, April 21, 2015 1:41 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Nuclear Artifact

Hi HistoNetters:
I have run into quite a problem.  My lab currently processes all tissue types 
from 3 different sites.  Recently, we have been getting complaints from one of 
the sites that the biopsies have a nuclear artifact.  It is described as 
washed out or poor to no nuclear detail.  Pictures have been uploaded 
(Nuclear Artifact).  The Medical Director at said site is convinced that a 
processor error is occurring.  Our site is not seeing this on any of our 
slides.  Biopsies from all three sites are processed and embed together.  We 
have done all trouble shooting that we can think of.  Leica service has come to 
inspect our Peloris processor and all areas checked out as functioning 
properly.  The problem is not consistent daily.  Seems to be worst toward the 
end of the week.
We have been running the same processing protocol, staining protocol and 
cutting protocols for years now.  This problem has just developed over the last 
2 months.  Any ideas, no matter how far-fetched, would be greatly appreciated 
at this point.

Lisa Roy, HT(ASCP)
Histology Supervisor
LabCorp at St. Vincent Hospital
123 Summer St
Worcester, MA
(508)363-9420

-This e-mail and any attachments may contain CONFIDENTIAL information, 
including PROTECTED HEALTH INFORMATION. If you are not the intended recipient, 
any use or disclosure of this information is STRICTLY PROHIBITED; you are 
requested to delete this e-mail and any attachments, notify the sender 
immediately, and notify the LabCorp Privacy Officer at 
privacyoffi...@labcorp.com or call (877) 23-HIPAA / (877) 234-4722. 
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[Histonet] RE: (no subject)

[Cynthia Robinson]

A few years ago we got cited for not having the fungal control be in tissue. 
The citation was from a HQIP survey we participated in from CAP. We were using 
a cultured fungal specimen in a cell button at that time. Since then I have 
collected and shared a number of cases we have seen with fungal infections in 
feet. 

Just my 2 cents

Cindi Robinson, HT(ASCP)
Dunes Medical Laboratories
350 W Anchor Dr
Dakota Dunes SD 57049


From: histonet-boun...@lists.utsouthwestern.edu 
[histonet-boun...@lists.utsouthwestern.edu] on behalf of Mayer,Toysha N 
[tnma...@mdanderson.org]
Sent: Tuesday, April 21, 2015 2:12 PM
To: 'histonet@lists.utsouthwestern.edu'
Subject: [Histonet] RE: (no subject)

-I think you can use the other tissue controls.
 Even CDC (Hansen's Disease Center) will send out mycobacterium leprae controls 
free of charge, and I think they are armadillo.
The best control I have ever used for mast cells is canine mast cell tumors.  I 
request them from Vet Schools regularly.
Also think of antibodies, aren't  most antibodies animal?
The separation of the specimens I think is during processing.  They should be 
processed separately from routine human tissues when they are being used for 
clinical human tests.
I have heard of them running on the same processor, just on a different run 
when the solutions have been changed.

Toysha
-

Message: 10
Date: Mon, 20 Apr 2015 18:50:15 -0400
From: Garrey Faller garr...@gmail.com
Subject: Re: [Histonet] (no subject)
To: koelli...@comcast.net
Cc: histonet@lists.utsouthwestern.edu
histonet@lists.utsouthwestern.edu
Message-ID:
CAF2sxrVD7o96Wz84nDBR6uxf=93tpqpmueqwog57e4fetcc...@mail.gmail.com
Content-Type: text/plain; charset=UTF-8

Here is the CAP checklist requirement:
ANP.21450
All  histochemical stains are of adequate quality, and daily controls are
demonstrated on each day of use for the tissue components or organism for
which they were designed.

Ray...you should call the CAP and ask for guidance on this.
My interpretation of this requirement is that it should be OK to use a
fungus from an orange peel. An orange peel fungus should have the same
staining characteristics as a candida or aspergillus etc.  Similarly a
bacteria is a bacteria. If you can produce a control that has both gram
positives and negatives, it should be OK. But, don't quote me on this.

Call the CAP for a definitive answer. I am interested in their response.
Garrey

On Sun, Apr 19, 2015 at 9:06 PM, koelli...@comcast.net wrote:

 I asked about this in a different vein months ago.  Has anyone shown a
 strawberry or ground meat or slim jim or orange peel as a bacteria/fungus
 control used for diagnostics to an inspector inspecting the lab and was
 there any comment from the inspector either positive or negative. Never
 heard back anything.
 Ray, Lake Forest Park, WA

 - Original Message -

 From: tjfinney2...@gmail.com
 To: histonet@lists.utsouthwestern.edu
 Sent: Sunday, April 19, 2015 5:24:53 PM
 Subject: [Histonet] (no subject)

 GMS controls
 From my understanding we can't use non human controls on patients. I
 could be wrong, but you may want to look into it.

 Happy Connecting.  Sent from my Sprint Phone.

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--


Sent from my iPhone

 On 21 Apr 2015, at 8:51 am, Garrey Faller garr...@gmail.com wrote:

 Here is the CAP checklist requirement:
 ANP.21450
 All  histochemical stains are of adequate quality, and daily controls are
 demonstrated on each day of use for the tissue components or organism for
 which they were designed.

 Ray...you should call the CAP and ask for guidance on this.
 My interpretation of this requirement is that it should be OK to use a
 fungus from an orange peel. An orange peel fungus should have the same
 staining characteristics as a candida or aspergillus etc.  Similarly a
 bacteria is a bacteria. If you can produce a control that has both gram
 positives and negatives, it should be OK. But, don't quote me on this.

 Call the CAP for a definitive answer. I am interested in their response.
 Garrey

 On Sun, Apr 19, 2015 at 9:06 PM, koelli...@comcast.net wrote:

 I asked about this in a different vein months ago.  Has anyone shown a
 strawberry or ground meat or slim jim or orange peel as a bacteria/fungus
 control used for diagnostics to an inspector inspecting the lab and was
 there any comment from the inspector either positive or negative. Never
 heard back anything.
 Ray, Lake Forest Park, WA

 - Original Message -

 From: tjfinney2...@gmail.com
 To: histonet@lists.utsouthwestern.edu
 Sent: Sunday

[Histonet] RE: IHC and oven temperature

[Morken, Timothy]

The95 for HIER is in liquid, The 82in the oven is dry slides. While wet high 
temps enhance HIER, It seems the high dry temp does harm epitopes (there is a 
paper somewhere on this but I don't have access to it right now).



Tim Morken
Supervisor, Histology, Electron Microscopy and Neuromuscular Special Studies
UC San Francisco Medical Center
San Francisco, CA

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-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Preiszner, 
Johanna
Sent: Monday, April 20, 2015 8:47 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] IHC and oven temperature

Hi Netters,

is there something wrong with this logic:

If the tissue needs 95C for HIER, it's ok to dry the slides in an 82C oven.

Of course I'll test it before I try it on real specimens, but maybe someone 
else already knows the answer...

Thanks!

Hanna Preiszner
ETSU/QCOM


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RE: [Histonet] RE: IHC Billing Question

[Joelle Weaver]

No charge  to patient account for negative or positive controls. Only the 
patient test (s). The controls verify that the test ran accurately, and for 
interpretation. You just have to figure this into your cost/test. As many have 
already posted in this string it is why many labs aside from QC reasons,  have 
adopted the single slide for postive/patient and have also eliminated negatives 
when polymer detection is used. 


Joelle Weaver MAOM, HTL (ASCP) QIHC


  

 
 From: mw...@wakehealth.edu
 To: bburn...@capecodhealth.org; jmore...@sidra.org; 
 histonet@lists.utsouthwestern.edu
 Date: Thu, 16 Apr 2015 14:35:59 +
 CC: 
 Subject: [Histonet] RE: IHC Billing Question 
 
 We have not been charging for the negative control, assuming that it was just 
 a cost of doing business.  I would be interested to hear if anyone has been 
 charging for their negative controls as well.
  
 Martha Ward, MT (ASCP) QIHC
 Manager
 
 Molecular Diagnostics Lab
 Medical Center Boulevard  \  Winston-Salem, NC 27157
 p 336.716.2109  \  f 336.716.5890  
 mw...@wakehealth.edu  
  
  
 
 
 
 -Original Message-
 From: histonet-boun...@lists.utsouthwestern.edu 
 [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Burnett, 
 Brandy
 Sent: Thursday, April 16, 2015 9:48 AM
 To: 'Joana Moreira'; histonet@lists.utsouthwestern.edu
 Subject: [Histonet] RE: IHC Billing Question 
 
 We recently added HER2 IHC testing in our lab which we are required to use a 
 negative reagent control For each case. Is there a cpt code for negative 
 reagent control reimbursement? Any information on this Would be much 
 appreciated!
 Thanks
 
 Brandy Burnett
 Histotechnoligist, QIHC(ASCP)
 CCH Pathology/Histology
 
 
 Expert physicians. Quality hospitals. Superior care. 
 
 
 -Original Message-
 From: histonet-boun...@lists.utsouthwestern.edu 
 [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Joana Moreira
 Sent: Thursday, April 16, 2015 7:25 AM
 To: histonet@lists.utsouthwestern.edu
 Subject: [Histonet] IHC Billing Question 
 
 Greetings from Doha!
 
 This was much probably discussed before, but I was wondering if you could 
 help me with a query in regards to billing.
 For the sites that are still doing an IHC negative reagent control for each 
 patient specimen, do you bill for the negative control? Using code 88341?
 
 I believe it should be billed (since when and if performed correctly the 
 negative control follows a normal IHC technique) however I am completely new 
 to the billing subject. My previous experience is based in Portugal and UK 
 (where billing does not exist) and I've been introduced to this topic since I 
 joined my current institution that will be following the North American 
 Healthcare model. So... any help will be GREATLY appreciated!!
 
 Many Thanks in advance,
 Joana
 
 Joana Moreira
 Supervisor - Anatomical Pathology
 Department of Pathology
 
 Sidra Medical  Research Center
 PO Box 26999 | Doha, Qatar
 Direct Line  +974-4404-2036
 jmore...@sidra.org | www.sidra.org
 
 
 
 
 
 Disclaimer: This email and its attachments may be confidential and are 
 intended solely for the use of the individual to whom it is addressed. If you 
 are not the intended recipient, any reading, printing, storage, disclosure, 
 copying or any other action taken in respect of this e-mail is prohibited and 
 may be unlawful. If you are not the intended recipient, please notify the 
 sender immediately by using the reply function and then permanently delete 
 what you have received. Any views or opinions expressed are solely those of 
 the author and do not necessarily represent those of Sidra Medical and 
 Research Center.
 
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[Histonet] RE: IHC Billing Question

[Burnett, Brandy]

We recently added HER2 IHC testing in our lab which we are required to use a 
negative reagent control 
For each case. Is there a cpt code for negative reagent control reimbursement? 
Any information on this 
Would be much appreciated!
Thanks

Brandy Burnett
Histotechnoligist, QIHC(ASCP)
CCH Pathology/Histology


Expert physicians. Quality hospitals. Superior care. 


-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Joana Moreira
Sent: Thursday, April 16, 2015 7:25 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] IHC Billing Question 

Greetings from Doha!

This was much probably discussed before, but I was wondering if you could help 
me with a query in regards to billing.
For the sites that are still doing an IHC negative reagent control for each 
patient specimen, do you bill for the negative control? Using code 88341?

I believe it should be billed (since when and if performed correctly the 
negative control follows a normal IHC technique) however I am completely new to 
the billing subject. My previous experience is based in Portugal and UK (where 
billing does not exist) and I've been introduced to this topic since I joined 
my current institution that will be following the North American Healthcare 
model. So... any help will be GREATLY appreciated!!

Many Thanks in advance,
Joana

Joana Moreira
Supervisor - Anatomical Pathology
Department of Pathology

Sidra Medical  Research Center
PO Box 26999 | Doha, Qatar
Direct Line  +974-4404-2036
jmore...@sidra.org | www.sidra.org





Disclaimer: This email and its attachments may be confidential and are intended 
solely for the use of the individual to whom it is addressed. If you are not 
the intended recipient, any reading, printing, storage, disclosure, copying or 
any other action taken in respect of this e-mail is prohibited and may be 
unlawful. If you are not the intended recipient, please notify the sender 
immediately by using the reply function and then permanently delete what you 
have received. Any views or opinions expressed are solely those of the author 
and do not necessarily represent those of Sidra Medical and Research Center.

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[Histonet] RE: IHC Billing Question

[Martha Ward-Pathology]

We have not been charging for the negative control, assuming that it was just a 
cost of doing business.  I would be interested to hear if anyone has been 
charging for their negative controls as well.
 
Martha Ward, MT (ASCP) QIHC
Manager

Molecular Diagnostics Lab
Medical Center Boulevard  \  Winston-Salem, NC 27157
p 336.716.2109  \  f 336.716.5890  
mw...@wakehealth.edu  
 
 



-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Burnett, Brandy
Sent: Thursday, April 16, 2015 9:48 AM
To: 'Joana Moreira'; histonet@lists.utsouthwestern.edu
Subject: [Histonet] RE: IHC Billing Question 

We recently added HER2 IHC testing in our lab which we are required to use a 
negative reagent control For each case. Is there a cpt code for negative 
reagent control reimbursement? Any information on this Would be much 
appreciated!
Thanks

Brandy Burnett
Histotechnoligist, QIHC(ASCP)
CCH Pathology/Histology


Expert physicians. Quality hospitals. Superior care. 


-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Joana Moreira
Sent: Thursday, April 16, 2015 7:25 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] IHC Billing Question 

Greetings from Doha!

This was much probably discussed before, but I was wondering if you could help 
me with a query in regards to billing.
For the sites that are still doing an IHC negative reagent control for each 
patient specimen, do you bill for the negative control? Using code 88341?

I believe it should be billed (since when and if performed correctly the 
negative control follows a normal IHC technique) however I am completely new to 
the billing subject. My previous experience is based in Portugal and UK (where 
billing does not exist) and I've been introduced to this topic since I joined 
my current institution that will be following the North American Healthcare 
model. So... any help will be GREATLY appreciated!!

Many Thanks in advance,
Joana

Joana Moreira
Supervisor - Anatomical Pathology
Department of Pathology

Sidra Medical  Research Center
PO Box 26999 | Doha, Qatar
Direct Line  +974-4404-2036
jmore...@sidra.org | www.sidra.org





Disclaimer: This email and its attachments may be confidential and are intended 
solely for the use of the individual to whom it is addressed. If you are not 
the intended recipient, any reading, printing, storage, disclosure, copying or 
any other action taken in respect of this e-mail is prohibited and may be 
unlawful. If you are not the intended recipient, please notify the sender 
immediately by using the reply function and then permanently delete what you 
have received. Any views or opinions expressed are solely those of the author 
and do not necessarily represent those of Sidra Medical and Research Center.

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[Histonet] Re: IHC Billing Question

[Bryan Szpunar]

I have never heard of anyone billing for a negative control and would
highly recommend against doing so. As mentioned, it is a cost of doing
business--which is why when CAP clarified that polymer detection systems
did not require them, most cost-conscious labs quit running them.

Regards,

Bryan Szpunar, HT(ASCP)


 -Original Message-
 From: histonet-boun...@lists.utsouthwestern.edu [mailto:
 histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Joana Moreira
 Sent: Thursday, April 16, 2015 7:25 AM
 To: histonet@lists.utsouthwestern.edu
 Subject: [Histonet] IHC Billing Question

 Greetings from Doha!

 This was much probably discussed before, but I was wondering if you could
 help me with a query in regards to billing.
 For the sites that are still doing an IHC negative reagent control for
 each patient specimen, do you bill for the negative control? Using code
 88341?

 I believe it should be billed (since when and if performed correctly the
 negative control follows a normal IHC technique) however I am completely
 new to the billing subject. My previous experience is based in Portugal and
 UK (where billing does not exist) and I've been introduced to this topic
 since I joined my current institution that will be following the North
 American Healthcare model. So... any help will be GREATLY appreciated!!

 Many Thanks in advance,
 Joana

 Joana Moreira
 Supervisor - Anatomical Pathology
 Department of Pathology

 Sidra Medical  Research Center
 PO Box 26999 | Doha, Qatar
 Direct Line  +974-4404-2036
 jmore...@sidra.org | www.sidra.org





 Disclaimer: This email and its attachments may be confidential and are
 intended solely for the use of the individual to whom it is addressed. If
 you are not the intended recipient, any reading, printing, storage,
 disclosure, copying or any other action taken in respect of this e-mail is
 prohibited and may be unlawful. If you are not the intended recipient,
 please notify the sender immediately by using the reply function and then
 permanently delete what you have received. Any views or opinions expressed
 are solely those of the author and do not necessarily represent those of
 Sidra Medical and Research Center.


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[Histonet] RE: IHC Billing Questions Controls

[Hanson, Dawn]

Brandy  Joana

Controls are never billable, a control test does not produce a usable result 
based on the patient's specimen. The results of control tests - positive or 
negative only tell you that your reagents, stains, antibodies etc. are 
performing as expected.


| Dawn Hanson | Lab Manager | Outreach | Saint Agnes Hospital, Baltimore, MD |
| * p: 410-368-3083 | * dhan...@stagnes.orgmailto:dhan...@stagnes.org | ý 
www.stagnes.orghttp://www.stagnes.org/ |



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[Histonet] RE: perfusion stiffness

[James Watson]

Watch the liver, it will change color as the blood is washed out.  Are you 
perfusing first with PBS to rinse the blood out, if you start with formalin 
small blood vessels and capillaries can become blocked by the blood that has 
coagulated when exposed to the formalin.  We actually flush with 10% sucrose 
first since brains flushed with PBS can have up to 20% shrinkage.

I have not heard of a good theory behind body stiffness after perfusion yet, 
pretty sure it is not the effect of actual cross linking since that takes time. 
 Is someone knows the cause please let me know.

James Watson HT  ASCP
GNF  Genomics Institute of the Novartis Research Foundation
Scientific Technical Leader II, Histology
Tel    858-332-4647
Fax   858-812-1915
jwat...@gnf.org

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of David Wright
Sent: Wednesday, April 15, 2015 2:06 PM
To: histonet@lists.utsouthwestern.edu
Cc: Yves Heremans
Subject: [Histonet] RE: perfusion stiffness

Hi Yves  Histonet

It is certainly a good sign if limbs etc are stiff after perfusion, but maybe 
not a guarantee that the target organ is perfect given the short perfusions you 
describe. Definitely, if I don't see stiffness I worry, check for a torn aortic 
arch (you are doing it transcardially, I presume), adjust the needle placement 
and run more fixative until everything is stiff.

Do you harvest organs from all over the body? You can improve the efficiency of 
perfusion by limiting it to the regions of interest. For example, I perfuse 
rats for brain extraction and clamp off the descending vessels (clamped to the 
spine) at the level of the diaphragm. The lower half of the body then doesn't 
get rigid, but the upper half does so more fully/faster. (In rats, I perfuse 
for much longer than you describe and post-fix too.) Note there's two kinds of 
stiffness - an immediate, zombie-like outstretching of the forelimbs ( tail 
wiggling if you do the whole body) which is immediately satisfying as a sign of 
good needle placement but only happens with a very fresh cadaver, and a more 
generalized, slower rigidity. For my brains, I check for neck muscle rigidity 
as well as the forelimb zombie effect.

best wishes - David
==
David A. Wright, Ph.D.
University of Chicago
Section of Neurosurgery, MC3026

 Histonet Digest, Vol 137, Issue 17 Message: 8
Date: Tue, 14 Apr 2015 09:26:00 +0200
From: Yves Heremans yves.herem...@vub.ac.be
Subject: [Histonet] transcardial fixation
To: histonet@lists.utsouthwestern.edu
Message-ID: 51d52790-789d-4189-92b3-630b6aad9...@vub.ac.be
Content-Type: text/plain; charset=us-ascii

Dear Histonetters,

We routinely perfuse mice with fixative before taking out organs. Perfusion 
with fixative (4% PFA or 10% NBF) is done for only a few minutes (max. 2 min.). 
During that short timespan, the entire mouse becomes stiff. Can this stifness 
be taken as a sign of good initial fixation (we post-fix the organs overnight 
at room temp) or is this stifness not entirely related to fixation ?

Yves

End of
*

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[Histonet] RE: ORO tissue falling off

[Linda Prasad (SCHN)]

Usually with the fatty tissues, I pick them up on superfrost slides and let it 
air dry  for 2-3 days at room temperature and then perform the ORO stains. So 
far they seem to stay on.

Linda Prasad | Senior Scientist | Histopathology
t: (02) 9845 3306 | f: (02) 9845 3318 | e: linda.pra...@health.nsw.gov.au | w: 
www.schn.health.nsw.gov.au

Cnr Hawkesbury Road and Hainsworth Street, Westmead, NSW Australia
Locked Bag 4001, Westmead 2145, NSW Australia

♲  Please consider the environment before printing this email.

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Jo-Ann Bader, 
Ms.
Sent: Thursday, 16 April 2015 1:40 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] ORO tissue falling off

We are having difficulty with a particulate set of very, very fatty mouse 
livers.  The normal livers from this set stay on the slides the fatty livers 
fall off.  We have used different types of charged slides and we have even 
tried to drench the charged slides in Stay-On, dry them and then put the frozen 
tissues on (despirate times call for despirate measures).  No luck  Does anyone 
have any other ideas.  Help Help

Jo-Ann  Bader
Histology Coordinator
Goodman Cancer Research Center
1600 Pine Ave. W,
Room 312
Montreal Quebec, H3A 1A3
Email: jo-ann.ba...@mcgill.camailto:jo-ann.ba...@mcgill.ca
Office Tel:  514-398-5647
Lab:  Tel:  514-398-8270

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[Histonet] RE: ORO tissue falling off

[Grantham, Andrea L - (algranth)]

I used to be the Queen of ORO in my lab at the University of Arizona. I had 
some of the fattiest livers that could be possible from different research 
projects. I never had a problem with the livers staying on the slide. I used 
Epic coated slides or Stat Lab slides and the protocol was from Freida's second 
edition. The stain was from PolyScientific RD.
I cut the sections at 5 microns, let them sit in the cryostat for about 30 
minutes and then fixed in 37% Formaldehyde for about 10-30 minutes - never 
really timed it. All steps of staining were done gently, one slide at a time, 
no matter how many slides I had to stain.
Sections were beautiful - wish I could post a picture here.
Retired now - sometimes I miss doing things like this.

Andi Grantham

From: histonet-boun...@lists.utsouthwestern.edu 
[histonet-boun...@lists.utsouthwestern.edu] on behalf of Linda Prasad (SCHN) 
[linda.pra...@health.nsw.gov.au]
Sent: Wednesday, April 15, 2015 5:07 PM
To: 'Jo-Ann Bader, Ms.'; histonet@lists.utsouthwestern.edu
Subject: [Histonet] RE: ORO tissue falling off

Usually with the fatty tissues, I pick them up on superfrost slides and let it 
air dry  for 2-3 days at room temperature and then perform the ORO stains. So 
far they seem to stay on.

Linda Prasad | Senior Scientist | Histopathology
t: (02) 9845 3306 | f: (02) 9845 3318 | e: linda.pra...@health.nsw.gov.au | w: 
www.schn.health.nsw.gov.au

Cnr Hawkesbury Road and Hainsworth Street, Westmead, NSW Australia
Locked Bag 4001, Westmead 2145, NSW Australia

♲  Please consider the environment before printing this email.

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Jo-Ann Bader, 
Ms.
Sent: Thursday, 16 April 2015 1:40 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] ORO tissue falling off

We are having difficulty with a particulate set of very, very fatty mouse 
livers.  The normal livers from this set stay on the slides the fatty livers 
fall off.  We have used different types of charged slides and we have even 
tried to drench the charged slides in Stay-On, dry them and then put the frozen 
tissues on (despirate times call for despirate measures).  No luck  Does anyone 
have any other ideas.  Help Help

Jo-Ann  Bader
Histology Coordinator
Goodman Cancer Research Center
1600 Pine Ave. W,
Room 312
Montreal Quebec, H3A 1A3
Email: jo-ann.ba...@mcgill.camailto:jo-ann.ba...@mcgill.ca
Office Tel:  514-398-5647
Lab:  Tel:  514-398-8270

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This note also confirms that this email message has been virus scanned and 
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RE: [Histonet] RE: ORO tissue falling off

[Linda Prasad (SCHN)]

I Know fatty tissue is such a pain to cut. Bryan Llewellyn gave some really 
good techniques. Im going to try them out myself :)

Linda Prasad | Senior Scientist | Histopathology
t: (02) 9845 3306 | f: (02) 9845 3318 | e: linda.pra...@health.nsw.gov.au | w: 
www.schn.health.nsw.gov.au


Cnr Hawkesbury Road and Hainsworth Street, Westmead, NSW Australia
Locked Bag 4001, Westmead 2145, NSW Australia

♲  Please consider the environment before printing this email.


-Original Message-
From: Caroline Miller [mailto:mi...@3scan.com] 
Sent: Thursday, 16 April 2015 10:44 AM
To: Linda Prasad (SCHN)
Cc: Jo-Ann Bader, Ms.; histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] RE: ORO tissue falling off

+1 to Linda, but I have found no difference on overnight vs multiple days.

Fatty liver is hard to do on all counts! It is tough enough sometimes to get a 
decent section on the slide.

Thanks for the other suggestions, certainly something I would try in the future

Yours
Caroline

Caroline Miller (mills)
Director of Histology
3Scan, Inc
415-2187297

 On Apr 15, 2015, at 5:07 PM, Linda Prasad (SCHN) 
 linda.pra...@health.nsw.gov.au wrote:
 
 Usually with the fatty tissues, I pick them up on superfrost slides and let 
 it air dry  for 2-3 days at room temperature and then perform the ORO stains. 
 So far they seem to stay on.
 
 Linda Prasad | Senior Scientist | Histopathology
 t: (02) 9845 3306 | f: (02) 9845 3318 | e: 
 linda.pra...@health.nsw.gov.au | w: www.schn.health.nsw.gov.au
 
 Cnr Hawkesbury Road and Hainsworth Street, Westmead, NSW Australia 
 Locked Bag 4001, Westmead 2145, NSW Australia
 
 ♲  Please consider the environment before printing this email.
 
 -Original Message-
 From: histonet-boun...@lists.utsouthwestern.edu 
 [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Jo-Ann Bader, 
 Ms.
 Sent: Thursday, 16 April 2015 1:40 AM
 To: histonet@lists.utsouthwestern.edu
 Subject: [Histonet] ORO tissue falling off
 
 We are having difficulty with a particulate set of very, very fatty 
 mouse livers.  The normal livers from this set stay on the slides the 
 fatty livers fall off.  We have used different types of charged slides 
 and we have even tried to drench the charged slides in Stay-On, dry 
 them and then put the frozen tissues on (despirate times call for 
 despirate measures).  No luck  Does anyone have any other ideas.  Help 
 Help
 
 Jo-Ann  Bader
 Histology Coordinator
 Goodman Cancer Research Center
 1600 Pine Ave. W,
 Room 312
 Montreal Quebec, H3A 1A3
 Email: jo-ann.ba...@mcgill.camailto:jo-ann.ba...@mcgill.ca
 Office Tel:  514-398-5647
 Lab:  Tel:  514-398-8270
 
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Re: [Histonet] RE: ORO tissue falling off

[Caroline Miller]

+1 to Linda, but I have found no difference on overnight vs multiple days.

Fatty liver is hard to do on all counts! It is tough enough sometimes to get a 
decent section on the slide.

Thanks for the other suggestions, certainly something I would try in the future

Yours
Caroline

Caroline Miller (mills)
Director of Histology
3Scan, Inc
415-2187297

 On Apr 15, 2015, at 5:07 PM, Linda Prasad (SCHN) 
 linda.pra...@health.nsw.gov.au wrote:
 
 Usually with the fatty tissues, I pick them up on superfrost slides and let 
 it air dry  for 2-3 days at room temperature and then perform the ORO stains. 
 So far they seem to stay on.
 
 Linda Prasad | Senior Scientist | Histopathology
 t: (02) 9845 3306 | f: (02) 9845 3318 | e: linda.pra...@health.nsw.gov.au | 
 w: www.schn.health.nsw.gov.au
 
 Cnr Hawkesbury Road and Hainsworth Street, Westmead, NSW Australia
 Locked Bag 4001, Westmead 2145, NSW Australia
 
 ♲  Please consider the environment before printing this email.
 
 -Original Message-
 From: histonet-boun...@lists.utsouthwestern.edu 
 [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Jo-Ann Bader, 
 Ms.
 Sent: Thursday, 16 April 2015 1:40 AM
 To: histonet@lists.utsouthwestern.edu
 Subject: [Histonet] ORO tissue falling off
 
 We are having difficulty with a particulate set of very, very fatty mouse 
 livers.  The normal livers from this set stay on the slides the fatty livers 
 fall off.  We have used different types of charged slides and we have even 
 tried to drench the charged slides in Stay-On, dry them and then put the 
 frozen tissues on (despirate times call for despirate measures).  No luck  
 Does anyone have any other ideas.  Help Help
 
 Jo-Ann  Bader
 Histology Coordinator
 Goodman Cancer Research Center
 1600 Pine Ave. W,
 Room 312
 Montreal Quebec, H3A 1A3
 Email: jo-ann.ba...@mcgill.camailto:jo-ann.ba...@mcgill.ca
 Office Tel:  514-398-5647
 Lab:  Tel:  514-398-8270
 
 ___
 Histonet mailing list
 Histonet@lists.utsouthwestern.edu
 http://lists.utsouthwestern.edu/mailman/listinfo/histonet
 
 *
 This email and any files transmitted with it are confidential and intended 
 solely for the use of the individual or entity to whom they are addressed. If 
 you are not the intended recipient, please delete it and notify the sender.
 
 Views expressed in this message and any attachments are those of the 
 individual sender, and are not necessarily the views of The Sydney Children's 
 Hospitals Network.
 
 This note also confirms that this email message has been virus scanned and 
 although no computer viruses were detected, The Sydney Childrens Hospital's 
 Network accepts no liability for any consequential damage resulting from 
 email containing computer viruses.
 *
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[Histonet] RE: perfusion stiffness

[David Wright]

Hi Yves  Histonet

It is certainly a good sign if limbs etc are stiff after perfusion, but maybe 
not a guarantee that the target organ is perfect given the short perfusions you 
describe. Definitely, if I don't see stiffness I worry, check for a torn aortic 
arch (you are doing it transcardially, I presume), adjust the needle placement 
and run more fixative until everything is stiff.

Do you harvest organs from all over the body? You can improve the efficiency of 
perfusion by limiting it to the regions of interest. For example, I perfuse 
rats for brain extraction and clamp off the descending vessels (clamped to the 
spine) at the level of the diaphragm. The lower half of the body then doesn't 
get rigid, but the upper half does so more fully/faster. (In rats, I perfuse 
for much longer than you describe and post-fix too.) Note there's two kinds of 
stiffness - an immediate, zombie-like outstretching of the forelimbs ( tail 
wiggling if you do the whole body) which is immediately satisfying as a sign of 
good needle placement but only happens with a very fresh cadaver, and a more 
generalized, slower rigidity. For my brains, I check for neck muscle rigidity 
as well as the forelimb zombie effect.

best wishes - David
==
David A. Wright, Ph.D.
University of Chicago
Section of Neurosurgery, MC3026

 Histonet Digest, Vol 137, Issue 17 Message: 8
Date: Tue, 14 Apr 2015 09:26:00 +0200
From: Yves Heremans yves.herem...@vub.ac.be
Subject: [Histonet] transcardial fixation
To: histonet@lists.utsouthwestern.edu
Message-ID: 51d52790-789d-4189-92b3-630b6aad9...@vub.ac.be
Content-Type: text/plain; charset=us-ascii

Dear Histonetters,

We routinely perfuse mice with fixative before taking out organs. Perfusion 
with fixative (4% PFA or 10% NBF) is done for only a few minutes (max. 2 min.). 
During that short timespan, the entire mouse becomes stiff. Can this stifness 
be taken as a sign of good initial fixation (we post-fix the organs overnight 
at room temp) or is this stifness not entirely related to fixation ?

Yves

End of
*

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[Histonet] RE: Healthcare Specialist

[Emily Jacob]

 

 

Hi

 

Good day to you,

 

I am writing this email to check if you have got the below email and have
any update on the same.

 

Thanks and I will look forward to your response

 

Best Regards

Emily

 

From: Emily Jacob [mailto:emily.ja...@crystaldatalist.com] 
Sent: Monday, March 30, 2015 12:02 PM
To: 'histonet@lists.utsouthwestern.edu'
Subject: Healthcare Specialist 

 

Hi,


I hope you are the right person to discuss about Healthcare Specialist
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Please let me know your target audience and geographical area, so that we
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Looking forward to hear from you.

Regards,

 

Emily Jacob

 

| List acquisition | Technology Lists | Email/Data Appending | Search Engine
Optimization |

 

If you do not wish to receive future emails from us, please reply as 'leave
out'

 

 

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[Histonet] RE: OB/GYN Database

[Cynthia Hunter]

 

 

Hi

 

Good day to you,

 

I am writing this email to check if you have got the below email and have
any update on the same.

 

Thanks and I will look forward to your response

 

Best Regards

Cynthia

 

From: Cynthia Hunter [mailto:cynthia.hun...@crystaldatalist.com] 
Sent: Monday, March 30, 2015 11:06 AM
To: 'histonet@lists.utsouthwestern.edu'
Subject: OB/GYN Database

 

Hi,

 

I want to check if you're interested in OB/GYN Database?

 

Specialties- Pregnancy Care, Family Practice, Internal Med, Family Med,
Psychiatry, Independent Practices, Hospital Owned and many more..

 

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Online Marketing Executive

 

If you're not interested to receive further emails, please reply with the
subject line as Leave out

 

 

 

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[Histonet] Re: Paraffin embedding of whole mouse brain

[Samala, Ramakrishna]

Hello Histonet!

Thank you all for your time and wonderful suggestions. This is the very
first time, I have to work with paraffin embedded brain tissues.

Based on the suggestions from Histonet colleagues, I have decided to
obtain 200 um thick brain sections for processing and embedding. Since I
have about 30 mouse brains, I have to be very careful in labeling all
cassettes. So problem solved!

Again thank you all

Have a very good week

Sincerely

Ramakrishna Samala, Ph.D.
Senior Research Associate
Amarillo Research Building, School of Pharmacy, Texas Tech University
Health Sciences Center
1406 S. Coulter
Amarillo, TX 79106


The outcome of any serious research can only be to make two questions grow
where only one grew before ­Thorstein Veblen








On 4/13/15, 9:30 AM, Scouten, Charles
charles.scou...@leicabiosystems.com wrote:

Are you really embedding, or just encasing?  I have heard that paraffin
cannot penetrate that deeply, that small sections are necessary for the
paraffin to infiltrate the entire piece.  Is this not true?

Why not use frozen sections?  You can encase the brain in premade gelatin
molds see this link:
http://cp.mcafee.com/d/FZsS92gscyhJ5xdUQsILIfFCXCQQm4n73hOqenTzqqb2bwWZPhO
Orjhohssd79EVvd79J6ZT3hPtdBAsrzGKBoOEjxLkRnjZ1kVJAWmXQ6PsVJAWmXQ6PrOba11dZ
_HYyOCYUzvHTbFIFIYC---UCZORQX8FGEEKsG7DR8OJMddECSjt-hojuv78I9CzATsS02f-lJ8
iHgJivGQVv8_jZzoDAfrzmBFv2TVelad-nFZFOH2k2e8vfp7QNqdk6XjM_VmQKgzIE5O1Zr5vy
nH3YdS9X4_WtiRmV_057_aSA9lEmFfRqsKrsohudwIqid40mSeKKOwq8fl8uwxa14Qgltd41wD
k_Ph0HlKDCy0x7pgdIcCYtznMVY5iQrl
and section gelatin and all.  Every brain in the same plane of section.

Do you use fixation perfusion, or just extract the soft brain?  Animal
researchers routinely use perfusion for the better tissue quality.


Cordially,

Charles W. Scouten, Ph.D.
Applications Specialist
Leica Biosystems
charles.scou...@leicabiosystems.com
http://cp.mcafee.com/d/avndz9J5xdUQsILIfFCXCQQm4n73hOqenTzqqb2bwWZPhOOrjho
hssd79EVvd79J6ZT3hPtdBAsrzGKBoOEjxLkRnjZ1kVJAWmXQ6PsVJAWmXQ6PrOba11dZ_HYyO
CYUzvHTbFIFIYC---UCZORQX8FGEEKsG7DR8OJMddFCSjt-hojuv78I9CzATsS02lbgZKdjZ8s
KrIjS9_QWBGJP-0af-lJ8iHgJivGQVsSUMyYr1oQAq80JItttB0QguGgZ12k29EwGWq831eF_C
y1mHtfd412eOwropdIgvrKJ9Lwge
Ph.  630 964 0501
Cell  314 724 5920


-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Samala,
Ramakrishna
Sent: Friday, April 10, 2015 9:00 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Paraffin embedding of whole mouse brain

Hello Histonet!

I would like to paraffin embed whole mouse brain to obtain coronal
sections. I spoke with customer support teams of several companies, I
could able to get 10 mm deep molds,  but the mouse brain is about 13 mm
height. So, any help in this regard is much appreciated.

Sincerely

Ramakrishna Samala, Ph.D.
Senior Research Associate
Amarillo Research Building, School of Pharmacy, Texas Tech University
Health Sciences Center
1406 S. Coulter
Amarillo, TX 79106

The outcome of any serious research can only be to make two questions
grow where only one grew before -Thorstein Veblen


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8Ke6zAQsLCzASzuXxEVKCOOedNRniIpk9MTGqHF-wGsSOtbtW3pKsSOtbtW3pJV5B0wC-_R-hp
jushLRXBQSkSujvvvsjuVqWtAkRkknel3PWApmU6CSjr9K_8I9LfzAm4PhOrKr01DOFeDdPFlK
pcNZ9_67uwTwCHIcfBisEeRMDaAWwQToDIj_FRblrDY0kvYHqgBmxqA_lFOVJNx5US2NF8Qg1r
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[Histonet] Re: Histonet Digest, Vol 137, Issue 15

[Joanne Clark]

She can be here in two weeks. 

Sent from my iPhone

 On Apr 12, 2015, at 11:01 AM, histonet-requ...@lists.utsouthwestern.edu 
 histonet-requ...@lists.utsouthwestern.edu wrote:
 
 Send Histonet mailing list submissions to
histonet@lists.utsouthwestern.edu
 
 To subscribe or unsubscribe via the World Wide Web, visit

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 When replying, please edit your Subject line so it is more specific
 than Re: Contents of Histonet digest...
 
 
 Today's Topics:
 
   1. Pigs as models of human biology (Jorge A. Santiago-Blay)
 
 
 --
 
 Message: 1
 Date: Sun, 12 Apr 2015 09:45:20 -0400
 From: Jorge A. Santiago-Blay blayjo...@gmail.com
 Subject: [Histonet] Pigs as models of human biology
 To: histonet@lists.utsouthwestern.edu
 Message-ID:
cagbdduam+psofrstxunsksso8u88eqtjq4aqur-dfwraskz...@mail.gmail.com
 Content-Type: text/plain; charset=UTF-8
 
 Dear Histonetters:
 
 
 I often wonder what are the reasons why pigs seem to be used so often in
 studies of human physiology. For phylogenetic reasons, I would have thought
 chimps would be the preferred choice. Is it because of humane, $, or are
 there other considerations? Thank you.
 
 
 
 If you know, please send me an email to: blayjo...@gmail.com
 
 
 
 Sincerely (and apologies if you have received this message more than once),
 
 
 
 Jorge
 
 Jorge A. Santiago-Blay, PhD
 blaypublishers.com
 
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 End of Histonet Digest, Vol 137, Issue 15
 *



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RE: [Histonet] RE: whole mouse brains

[James Watson]

We also have embed mouse brains for coronal sections routinely in deep molds, 
yes it can be close at times.  With rat brains we actually embedded in deep 
molds with the Mega cassette upside down so part of the brain went above the 
mold and into the cassette.  If we needed to go through the whole brain once we 
got near the cassette we would re-embed the brain so the reminder was out of 
the cassette.

James Watson HT  ASCP
GNF  Genomics Institute of the Novartis Research Foundation
Scientific Technical Leader II, Histology
Tel    858-332-4647
Fax   858-812-1915
jwat...@gnf.org

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Catherine 
Simonson
Sent: Friday, April 10, 2015 10:38 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] RE: whole mouse brains

Hey there!

I embed whole mouse brains on a regular basis.  First, process on a longer 
schedule (about 1 to 1.5 hours per station, really.  Otherwise they are under 
processed and will not cut well and you will have problems getting them to stay 
on the slides during staining).  Use the deep molds.  Keep in mind that the 
tissue will shrink (about 20 - 30 %) during processing so they WILL fit for 
coronal sections.  If need be, trim some of the olfactory bulbs off (you 
probably would be facing those off on the microtome anyhow).

Hope this helps,

Catherine Simonson, HT (ASCP)
The Jackson Laboratory
Bar Harbor, ME
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[Histonet] Re: Levai-Laczko stain

[fernandom . munoz]

1. 30% H2O2 5 min. under movement.
2. wash in water 2x
3. 5% Acetic acid 1 min.
4. wash in water 2x
5. solution 1 20 min.
6. wash in water 2x
7. solution 2, piece by piece
8. wash in water and dry.

Solution 1: 1 part Azur II (1%) + 1 part Methylen blue (1%) + 2 parts  
Na2CO3 (1%).

Solution 2: Pararosanillin (1%).
All solutions in distilled water. NEVER alcohol or acetone in plastic  
sections.


Fernando María Muñoz Guzón
Facultad de Veterinaria
Campus Universitario s/n
27002 Lugo (España)
Tfno: (+34)600940225





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[Histonet] RE: Tri-State Histology Symposium

[hymclab]

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of hymclab
Sent: Monday, March 02, 2015 11:07 AM
To: 'Teri Johnson'; colleen_herr...@bshsi.org
Cc: 'histonet@lists.utsouthwestern.edu'
Subject: [Histonet] Tri-State Histology Symposium


Dear Histonetters:  You are invited to join the histology societies of 
Wisconsin, Iowa and Minnesota as we celebrate Hats Off to Histology at the 
2015 Tri-State Histology Symposium, May 6-8 at The Madison Concourse Hotel and 
Governors Club in Madison, Wisconsin.

For program, registration and vendor/exhibit information contact the following 
representatives:

Wisconsin:  Kathryn Stoll kst...@mcw.edu

Iowa:  Judi Stasko judith.sta...@ars.usda.gov

Minnesota: Lois Rowe rowe.l...@mayo.edu

Vendor/Exhibit:  Dawn Schneider dawn.schnei...@ministryhealth.org




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the sole use of the intended recipient(s) and contains information that is 
confidential and proprietary to Genoptix Medical Laboratory or its 
subsidiaries. Any unauthorized review, use, disclosure or distribution is 
prohibited. If you are not the intended recipient, immediately contact the 
sender by e-mail and destroy all copies of the original message.


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[Histonet] RE: Formaldehyde filters

[Shelly Coker]

I am not sure about the color change you are referencing, unless it is the 
change that occurs with potassium permanganate filters, which turn black when 
exhausted.

We have a countertop gross station with two Potassium Permanganate filters that 
we change every 6 months.  Keep in mind we do very small specimens and deal 
with very small amounts of formalin.  If you use the Potassium permanganate 
filters (which are no more expensive than charcoal through Mopec), then you 
should be able to come up with a changing schedule that will work with your 
workflow.
Try not to get the potassium permanganate anywhere on the floor or counter or 
your clothes thoughthe dust is a tan color, but reacts with moisture and 
turns purple!



Does anybody remember how to test whether a formaldehyde filter for a grossing 
station is exhausted?  If I remember right there is some sort of test where 
you remove a couple of granules and cut the granules in half.  The color of 
the cut service is used somehow.

Also I know that changing formaldehyde filters on ductless workstations needs 
to be done regularly.  Can anyone share how often they change the formaldehyde 
filters for a gross lab senior or a small station such as a HYperclean?

Jim

Jim Vickroy
Histology Manager
Springfield Clinic, Main Campus, East Building
1025 South 6th Street
Springfield, Illinois  62703
Office:  217-528-7541, Ext. 15121
Email:  jvick...@springfieldclinic.commailto:jvick...@springfieldclinic.com



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[Histonet] Re: Anyone need a Benchmark Ultra?

[Jeff McKenna]

Is anyone out there looking for an extra Ventana Benchmark Ultra for
IHC/ISH? I have one in good shape that is not being used anymore and I am
looking to free up some space. Email me if you are interested and I'll give
you more info on it.


Jeff Mack

dermack...@gmail.com

On Tue, Mar 17, 2015 at 1:55 PM, Jeff McKenna dermack...@gmail.com wrote:

 Is anyone looking for an extra Ventana Benchmark Ultra for IHC/ISH? I have
 one in good shape that is not being used anymore and I am looking to free
 up some space. Email me if you are interested and I'll give you more info
 on it.


 Jeff Mack

 dermack...@gmail.com

 (302) 827-3685

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[Histonet] Re: Laczko/Levai stain protocol

[Gayle Callis]

I found several pages from Hayat's book Stains and Cytochemical Staining
1993 (for EM) with original authors cited after a Google search, key words
Laczko and Levai staining protocol.   Go to this long link and read on hints
to improve staining plus the protocol. 

 

https://books.google.com/books?id=oGj7MLioFlQCpg=PA60lpg=PA60dq=Levai-Lac
zko+proceduresource=blots=5i96tYhIh1sig=iXOwXifgBUsaWGjdOGya_zjOPlAhl=en
sa=Xei=IjYkVcKIN5GrogStzYLQAwved=0CFIQ6AEwCTgK#v=onepageq=Levai-Laczko%2
0proceduref=false   

 

The Google search turned up many publications using this stain for
undecalcified bone in PMMA, but you would have to access the journals for
the publications, even the original publication for the stain by Laczko and
Levai.  

 

Cartilage and Bone  (Laczko and Levai, 1975)

Azure II Methylene blue Safranin O

Tissue was fixed with mixture of gluteraldehyde and
formaldehyde 2 -4 hr @ RT. I assume these are tiny (minced) pieces of
tissue for EM.   Bone was decalcified for 2 weeks at 4C, postfixed with
osmium tetroxide 2 -3 hr RT, embedded in Durcupan, an EM resin.   I doubt
people embedding mineralized bone in MMA/PMMA will want to fix in
gluteraldehyde, but rather NBF and not post fix in osmium tetroxide.  

 

The stain, as seen in many cited references was used for
thin sections of bone.  You can pick up those references via Google for more
on methods and materials for undecalcified bone in PMMA/MMA. 

 

Azure II/Methylene Blue Working Solution

Azure II 1%   10 ml

Methylene Blue 1%   10 ml

Sodium carbonate 1%  20 ml 

 

Protocol

1.  Stain in working solution  for 2 -5  min at 50 - 60C

2.  Rinse in 0.5% sodium carbonate followed by distilled
water

3.  Counterstain with 0.5% safranin aq.  3 min at 35 - 40C
and rinse with distilled water, dehydrate.  

Results

Chromatin, nucleoli  violet blue

Cytoplasm   blue 

Erythrocytes  dark blue

cartilage matrix light blue

bone matrix   bright red

The search found many photos of bone with this stain to give you examples. 

 Check out these links:
https://www.unizahnklinik-wien.at/en/science/tangl_laboratory/resources.php


 and geosoft.ru/stati/de%20Sanctis%202010_J%20Clin%20Perio_.pdf 

 by M de Sanctis - ‎2010 

Material and Methods: Eight beagle dogs received implants
randomly installed into . Levai. Laczko staining. Original magnification
x 2.5. Fig. 9. Most apical .

 

Good luck

 

Gayle Callis

HTL/HT/MT(ASCP)   

 

 

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RE: [Histonet] Re: Pam Marcum colleague losing bone sections from slides

[Cooper, Brian]

Have you ever tried blotting your slides dry before putting them into the oven? 
 The veteran who taught me this trick used it on brain tissues from the 
Coroner's office (in the early 80s--don't wanna offend anyone) which were 
grossed very thickly and were always poorly processed.   She never used charged 
slides or additives in her waterbath.  She claimed she was the only one in her 
lab who was allowed to cut this stuff because everyone else's slides had 
tissue loss!  I can tell you from my experience that it works well for toenail 
which is notorious for detaching from slides.  I've used it on many other 
tissues as well.

Anyway, press a slightly moistened clean L'Absorb or paper towel down onto the 
section after microtomy.  You don't want the paper towel soaking wet--just 
damp.  This will effectively wick the section of any excess moisture.  Then 
incubate and stain as usual.  

Good luck.

Brian D. Cooper, HT (ASCP)CM | Histology Supervisor 
Department of Pathology and Laboratory Medicine
Children's Hospital Los Angeles 
4650 Sunset Blvd MS#43- Los Angeles, CA 90027 
bcoo...@chla.usc.edu 

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Gayle Callis
Sent: Tuesday, April 07, 2015 2:57 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Re: Pam Marcum colleague losing bone sections from slides

From Pam:  I am currently trying to stain L6 vertebrae from rabbits. 
They
have been decalcified and paraffin processed properly. I've tried cutting at 
both 5 and 10 microns and my tissue is still not sticking to my slides. I know 
my sectioning is fine because I'm successful with every other tissue I've ever 
sectioned and stained. For some reason the bone I'm using won't stick to any 
slides. I was using charged slides and I even tried poly-L-lysine slides, but 
the bone keeps coming up even before I attempted to stain them. I've even tried 
leaving them in the incubator for more than the usual 48-72 hours. I know it's 
possible to do other stains beside HE on bone, but I think my main issue is 
just getting good contact between the tissue and slide. If you have any advice 
or thoughts, I would love to hear them. 

  

I will get the messages to him ASAP. 

  

Pam 

*

 

What was meant by incubator and at what temperature?   It helps to dry
sections FLAT, at 37 to 40C for several days.  Do NOT dry at 60C.   

 

If the sections are not staying on plus charge or poly L lysine coated slides,  
then use chrome gelatin subbing solution in a water bath OR by
pre-subbing clean microscope slides.

 

This is the Chrome gelatin protocol that worked for our huge decalcified bone 
sections and or problem bone sections. 

 

Chrome Gelatin Subbing Solution:  Section/Slide Coating Adhesive

 

0.1 g Chromium Potassium Sulfate (this is toxic.  Collect for proper disposal, 
not down the drain is you pre-sub the slides). 

1.0 g Gelatin:  100 bloom, Sigma.  For large bone sections, use 200 or 300
bloom gelatin, Sigma).   200 and 300 bloom gelatins are very large gelatin
molecules made from pig collagen.  100 bloom is a much smaller molecule than
200 bloom.   Do NOT use household (cooking)  gelatin used for
cooking. Buy the pure gelatins only. 

1 liter Distilled Water

 

Dissolve chromium potassium sulfate and gelatin in hot but not 
boiling water.  Cool subbing solution before use, and store in refrigerator.  
If gelatin gets growth, discard, make new.  A few crystals of Thymol in stock 
subbing solution can help prevent growth. 

 

DO NOT USE PLUS CHARGE SLIDES WITH SUBBING SOLUTION.   GELATIN COATS OVER A
PLUS CHARGE COATING AND NEGATES THE POSITIVE CHARGE.  

 

For presubbing glass slides, wash these by dipping in acetone, air dry before 
using the pre-subbing protocol to get rid of any greasy/oily residues
on glass surface.   If you put the subbing solution in a water bath,
uncoated,  glass slides will work fine without further washing.

 

You can do either of the following: 

 

1.Add 10 ml subbing solution to a warm water bath for paraffin
sections. Then mount sections onto the cleaned glass slide, drain, and air dry, 
store in a cool, dry place. 

2.Dip acetone washed, dry slides into subbing solution, air dry,
and store in a dust free area.  Box subbed slides and store until needed.  

 

If you get background staining with hematoxylin (hematoxylin stains gelatin) 
then dip  pre-subbed slides in NBF ~10 times, rinse with distilled water, air 
dry and store slides.  The aldehyde fixative cross links the gelatin to some 
degree, but still allows section to adhere without annoying background 
staining.  

 

Pick up sections from water bath drain and lay flat to dry at 40C for
several days.   You will not need extra subbing solution in the water bath
if using presubbed

FW: [Histonet] Re: Pam Marcum colleague losing bone sections from slides

[Carol Fields]

-Original Message-
From: Carol Fields 
Sent: Tuesday, April 07, 2015 3:37 PM
To: 'Cooper, Brian'
Cc: 'histonet-boun...@lists.utsouthwestern.edu'
Subject: RE: [Histonet] Re: Pam Marcum colleague losing bone sections from 
slides

I've used that also many times.  It really does work.
Carole Fields,  HT (ASCP)
Lead Histotechnologist, Pathology Laboratory Martin Luther King Jr. Community 
Hospital
 1680 E. 120th Street
Los Angeles, CA 90059

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Cooper, Brian
Sent: Tuesday, April 07, 2015 3:23 PM
To: gayle.cal...@bresnan.net; histonet@lists.utsouthwestern.edu
Subject: RE: [Histonet] Re: Pam Marcum colleague losing bone sections from 
slides

Have you ever tried blotting your slides dry before putting them into the oven? 
 The veteran who taught me this trick used it on brain tissues from the 
Coroner's office (in the early 80s--don't wanna offend anyone) which were 
grossed very thickly and were always poorly processed.   She never used charged 
slides or additives in her waterbath.  She claimed she was the only one in her 
lab who was allowed to cut this stuff because everyone else's slides had 
tissue loss!  I can tell you from my experience that it works well for toenail 
which is notorious for detaching from slides.  I've used it on many other 
tissues as well.

Anyway, press a slightly moistened clean L'Absorb or paper towel down onto the 
section after microtomy.  You don't want the paper towel soaking wet--just 
damp.  This will effectively wick the section of any excess moisture.  Then 
incubate and stain as usual.  

Good luck.

Brian D. Cooper, HT (ASCP)CM | Histology Supervisor Department of Pathology and 
Laboratory Medicine Children's Hospital Los Angeles
4650 Sunset Blvd MS#43- Los Angeles, CA 90027 bcoo...@chla.usc.edu 

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Gayle Callis
Sent: Tuesday, April 07, 2015 2:57 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Re: Pam Marcum colleague losing bone sections from slides

From Pam:  I am currently trying to stain L6 vertebrae from rabbits. 
They
have been decalcified and paraffin processed properly. I've tried cutting at 
both 5 and 10 microns and my tissue is still not sticking to my slides. I know 
my sectioning is fine because I'm successful with every other tissue I've ever 
sectioned and stained. For some reason the bone I'm using won't stick to any 
slides. I was using charged slides and I even tried poly-L-lysine slides, but 
the bone keeps coming up even before I attempted to stain them. I've even tried 
leaving them in the incubator for more than the usual 48-72 hours. I know it's 
possible to do other stains beside HE on bone, but I think my main issue is 
just getting good contact between the tissue and slide. If you have any advice 
or thoughts, I would love to hear them. 

  

I will get the messages to him ASAP. 

  

Pam 

*

 

What was meant by incubator and at what temperature?   It helps to dry
sections FLAT, at 37 to 40C for several days.  Do NOT dry at 60C.   

 

If the sections are not staying on plus charge or poly L lysine coated slides,  
then use chrome gelatin subbing solution in a water bath OR by
pre-subbing clean microscope slides.

 

This is the Chrome gelatin protocol that worked for our huge decalcified bone 
sections and or problem bone sections. 

 

Chrome Gelatin Subbing Solution:  Section/Slide Coating Adhesive

 

0.1 g Chromium Potassium Sulfate (this is toxic.  Collect for proper disposal, 
not down the drain is you pre-sub the slides). 

1.0 g Gelatin:  100 bloom, Sigma.  For large bone sections, use 200 or 300
bloom gelatin, Sigma).   200 and 300 bloom gelatins are very large gelatin
molecules made from pig collagen.  100 bloom is a much smaller molecule than
200 bloom.   Do NOT use household (cooking)  gelatin used for
cooking. Buy the pure gelatins only. 

1 liter Distilled Water

 

Dissolve chromium potassium sulfate and gelatin in hot but not 
boiling water.  Cool subbing solution before use, and store in refrigerator.  
If gelatin gets growth, discard, make new.  A few crystals of Thymol in stock 
subbing solution can help prevent growth. 

 

DO NOT USE PLUS CHARGE SLIDES WITH SUBBING SOLUTION.   GELATIN COATS OVER A
PLUS CHARGE COATING AND NEGATES THE POSITIVE CHARGE.  

 

For presubbing glass slides, wash these by dipping in acetone, air dry before 
using the pre-subbing protocol to get rid of any greasy/oily residues
on glass surface.   If you put the subbing solution in a water bath,
uncoated,  glass slides will work fine without further washing.

 

You can do either of the following: 

 

1.Add 10 ml subbing solution

[Histonet] Re: Pam Marcum colleague losing bone sections from slides

[Gayle Callis]

From Pam:  I am currently trying to stain L6 vertebrae from rabbits. They
have been decalcified and paraffin processed properly. I've tried cutting at
both 5 and 10 microns and my tissue is still not sticking to my slides. I
know my sectioning is fine because I'm successful with every other tissue
I've ever sectioned and stained. For some reason the bone I'm using won't
stick to any slides. I was using charged slides and I even tried
poly-L-lysine slides, but the bone keeps coming up even before I attempted
to stain them. I've even tried leaving them in the incubator for more than
the usual 48-72 hours. I know it's possible to do other stains beside HE on
bone, but I think my main issue is just getting good contact between the
tissue and slide. If you have any advice or thoughts, I would love to hear
them. 

  

I will get the messages to him ASAP. 

  

Pam 

*

 

What was meant by incubator and at what temperature?   It helps to dry
sections FLAT, at 37 to 40C for several days.  Do NOT dry at 60C.   

 

If the sections are not staying on plus charge or poly L lysine coated
slides,  then use chrome gelatin subbing solution in a water bath OR by
pre-subbing clean microscope slides.

 

This is the Chrome gelatin protocol that worked for our huge decalcified
bone sections and or problem bone sections. 

 

Chrome Gelatin Subbing Solution:  Section/Slide Coating Adhesive

 

0.1 g Chromium Potassium Sulfate (this is toxic.  Collect for proper
disposal, not down the drain is you pre-sub the slides). 

1.0 g Gelatin:  100 bloom, Sigma.  For large bone sections, use 200 or 300
bloom gelatin, Sigma).   200 and 300 bloom gelatins are very large gelatin
molecules made from pig collagen.  100 bloom is a much smaller molecule than
200 bloom.   Do NOT use household (cooking)  gelatin used for
cooking. Buy the pure gelatins only. 

1 liter Distilled Water

 

Dissolve chromium potassium sulfate and gelatin in hot but
not boiling water.  Cool subbing solution before use, and store in
refrigerator.  If gelatin gets growth, discard, make new.  A few crystals of
Thymol in stock subbing solution can help prevent growth. 

 

DO NOT USE PLUS CHARGE SLIDES WITH SUBBING SOLUTION.   GELATIN COATS OVER A
PLUS CHARGE COATING AND NEGATES THE POSITIVE CHARGE.  

 

For presubbing glass slides, wash these by dipping in acetone, air dry
before using the pre-subbing protocol to get rid of any greasy/oily residues
on glass surface.   If you put the subbing solution in a water bath,
uncoated,  glass slides will work fine without further washing.

 

You can do either of the following: 

 

1.Add 10 ml subbing solution to a warm water bath for paraffin
sections. Then mount sections onto the cleaned glass slide, drain, and air
dry, store in a cool, dry place. 

2.Dip acetone washed, dry slides into subbing solution, air dry,
and store in a dust free area.  Box subbed slides and store until needed.  

 

If you get background staining with hematoxylin (hematoxylin stains gelatin)
then dip  pre-subbed slides in NBF ~10 times, rinse with distilled water,
air dry and store slides.  The aldehyde fixative cross links the gelatin to
some degree, but still allows section to adhere without annoying background
staining.  

 

Pick up sections from water bath drain and lay flat to dry at 40C for
several days.   You will not need extra subbing solution in the water bath
if using presubbed slides. 

 

 

IF all else fails, try Sterchi tape transfer method with packaging tape.   I
have the method with photos and publication, and will send privately.   

 

Good luck

 

Gayle Callis

HTL/HT/MT(ASCP)

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[Histonet] Re: Histonet Digest, Vol 137, Issue 7

[Wanda Jones]

Wanda Platt Jones

 On Apr 5, 2015, at 12:12 PM, histonet-requ...@lists.utsouthwestern.edu wrote:
 
 Send Histonet mailing list submissions to
histonet@lists.utsouthwestern.edu
 
 To subscribe or unsubscribe via the World Wide Web, visit
http://lists.utsouthwestern.edu/mailman/listinfo/histonet
 or, via email, send a message with subject or body 'help' to
histonet-requ...@lists.utsouthwestern.edu
 
 You can reach the person managing the list at
histonet-ow...@lists.utsouthwestern.edu
 
 When replying, please edit your Subject line so it is more specific
 than Re: Contents of Histonet digest...
 
 
 Today's Topics:
 
   1. Re: Kidney Stone Histology? (Mike Andrews)
   2. kidney stone stain etc (mad...@verizon.net)
 
 
 --
 
 Message: 1
 Date: Sat, 4 Apr 2015 22:54:22 -0500
 From: Mike Andrews udsd...@gmail.com
 Subject: Re: [Histonet] Kidney Stone Histology?
 To: Histonet@lists.utsouthwestern.edu
histonet@lists.utsouthwestern.edu
 Message-ID:
CACPTdfKWwVfhHCJeK7MJM5RkdRR9iJ2i8Bi8U8Y-dez=4yx...@mail.gmail.com
 Content-Type: text/plain; charset=UTF-8
 
 What Everyone Else Wrote.
 
 They're stone (apatite; the kind I myself get) or hard crystals (uric
 acid). They're very unfriendly to a microtome blade. Getting a thin section
 of an apatite stone would require petrological thin-section techniques,
 which won't be in the repertoire of the typical soft-tissue histo lab.
 Think epoxy, abrasives, and time. Under a stereo scope, they can be
 amazing: covered with sharp corners and edges, and all the corners and
 edges have more corners and edges on them. For a big staghorn stone, a
 diamond rock saw might be required. Uric acid crystals aren't particularly
 better. But histo techniques pretty much don't apply.
 
 On Fri, Apr 3, 2015 at 1:42 PM, Norton, Sally 
 sally.nor...@seattlechildrens.org wrote:
 
 The only thing we do with stones is send them to the Mayo clinic for
 chemical analysis.
 
 Sally Norton
 Seattle Children's
 
 
 
 -Original Message-
 From: histonet-boun...@lists.utsouthwestern.edu [mailto:
 histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Jerry Ricks
 Sent: Friday, April 03, 2015 11:38 AM
 To: Histonet@lists.utsouthwestern.edu
 Subject: [Histonet] Kidney Stone Histology?
 
 How the heck do I process and section kidney stones?  And what kind of
 stain do you like for them?
 
 
 Thanks
 
 Jerry Ricks
 Research Scientist
 University of Washington
 Department of Pathology
 
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 for the sole use of the intended recipient(s) and may contain confidential
 and privileged information protected by law. Any unauthorized review, use,
 disclosure or distribution is prohibited. If you are not the intended
 recipient, please contact the sender by reply e-mail and destroy all copies
 of the original message.
 
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 Histonet@lists.utsouthwestern.edu
 http://lists.utsouthwestern.edu/mailman/listinfo/histonet
 
 
 
 -- 
 73 de
 
 Mike Andrews W5EGO
 
 
 --
 
 Message: 2
 Date: Sun, 05 Apr 2015 08:05:34 -0500 (CDT)
 From: mad...@verizon.net
 Subject: [Histonet] kidney stone stain etc
 To: histonet@lists.utsouthwestern.edu
 Message-ID:
28629564.780357.1428239134134.javamail.r...@vznit170116.mailsrvcs.net

 Content-Type: text/plain; charset=UTF-8
 
 
   I a   is  too  mu   kossa or Dahls might
   nbs
   Nick(Rocky) Madary, HT/HTL(ASCP)QIHC
   Jo
   On 04/04/15, histonet-request@lists.utsouthw
   Send Histonet mailing list submissions to   
 [1]histonet@lists.utsouthwestern.edu
   To subscribe or unsubscribe via the World Wide Web, visit
   [2]http://lists.utsouthwestern.edu/mailman/listinfo/   or, via email, 
 send a message with subject or body 'help'   
 [3]histonet-request@lists.utsouthwestern   You can reach the person managing 
 the list at
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 target=_blankhistonet-ow...@lists.utsouthwestern.edu
   When replying, please edit your Subject line so it is more specific
   than Re: Contents of Histonet digest...
   Today   1. Kidney Stone Histology? (Jerry Ricks)
   2   3. Re: Kidney Stone H   ---   
 ---
   Message: 1
   Date: Fri, 3   From: Jerry Ricks [4]rosenfeldtek@h   Subject: [Histonet] 
 Kidney Stone Histology?
   To  Mes   Content-Type: text/plain; charset=iso-8859-1
   How  the heck do I process and section kidney stones? And what kind of
  Thanks
   Jerry Ricks   University of Washington
   Department of P   
   ---   Message: 2
   Date: Fri, 3 Apr 2015 18:42:09 +
   Fr   

[Histonet] Re: Kidney Stone Histology?

[Bob Richmond]

I've been in pathology over 50 years, and I never heard of anybody trying
to cut a section of a kidney stone. You sure this wasn't an April Fool grab?

Stone analysis - once done chemically, now done mostly by physical methods
- is of course clinically quite useful and is very often ordered. There
aren't too many labs that do it - Louis Herring (see herringlab.com) is
perhaps the oldest and best known.

Bob Richmond
Samurai Pathologist
Maryville TN
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[Histonet] RE: cleaning glassware

[Tony Henwood (SCHN)]

Also check the gelatine - is it a new batch?

From: histonet-boun...@lists.utsouthwestern.edu 
[histonet-boun...@lists.utsouthwestern.edu] on behalf of Anne Murvosh 
[amurv...@advancederm.net]
Sent: Friday, 3 April 2015 6:01 AM
To: Jeff Halstead; histonet@lists.utsouthwestern.edu
Subject: [Histonet] RE: cleaning glassware

I always used to rinse the containers I used with alcohol and let them dry 
before doing the stain.  Some people use an acid alcohol rinse.  Anne

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Jeff Halstead
Sent: Thursday, April 02, 2015 11:48 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] cleaning glassware

Hi, All-I am currently experiencing problems with our warthin-stary silver 
stain. The finished product has silver particulates all over the empty slide 
and deposited on the tissue making the stain very difficult to interpet. Any 
ideas would be very helpful.  thanx
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[Histonet] RE: cleaning glassware

[Roberta Horner]

I work in a small veterinary lab and when I do the Warthin Starry stain I mix 
all the reagents in never used before disposable plastic beakers and I stain 
the slides in an un-used slide mailer.
Roberta Horner
Animal Diagnostic Lab
Penn State University

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Jeff Halstead
Sent: Thursday, April 02, 2015 2:48 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] cleaning glassware

Hi, All-I am currently experiencing problems with our warthin-stary silver 
stain. The finished product has silver particulates all over the empty slide 
and deposited on the tissue making the stain very difficult to interpet. Any 
ideas would be very helpful.  thanx 
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[Histonet] RE: cleaning glassware

[Bernice Frederick]

We always acid clean before doing any silver stain. No metal 

Bernice Frederick HTL (ASCP)
Senior Research Tech
Pathology Core Facility
Robert. H. Lurie Cancer Center
Northwestern University
710 N Fairbanks Court
Olson 8-421
Chicago,IL 60611
312-503-3723
b-freder...@northwestern.edu


-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Roberta Horner
Sent: Thursday, April 02, 2015 2:36 PM
To: Jeff Halstead; histonet@lists.utsouthwestern.edu
Subject: [Histonet] RE: cleaning glassware

I work in a small veterinary lab and when I do the Warthin Starry stain I mix 
all the reagents in never used before disposable plastic beakers and I stain 
the slides in an un-used slide mailer.
Roberta Horner
Animal Diagnostic Lab
Penn State University

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Jeff Halstead
Sent: Thursday, April 02, 2015 2:48 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] cleaning glassware

Hi, All-I am currently experiencing problems with our warthin-stary silver 
stain. The finished product has silver particulates all over the empty slide 
and deposited on the tissue making the stain very difficult to interpet. Any 
ideas would be very helpful.  thanx 
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[Histonet] Re: TRAP staining on formic acid decalcified bone reference

[Gayle Callis]

The reference within a reference from Ray is A Chimeric Form of
Osteoprotegerin Inhibits Hypercalcemia and Bone Resorption Induced by IL-1β,
TNF-α, PTH, PTHrP, and 1,25(OH)2D3 .   Sean Morony et al .  J Bone Mineral
Res V 14, pp 1478-1485.  

 

However, the formic acid decalcification method is not described in detail
and merely says formic acid but whether this is buffered formic acid or
just dilute formic acid in water only is not stated. Ray might elaborate
on what specific formic acid recipe he used as many in research don't always
use buffered formic acid decalcifiying solutions.  

 

I would assume Morony et all used a buffered formic acid with either sodium
formate or sodium citrate and controlled so as to not overexpose TRAP to
acids longer than necessary.   One publication,  i.e.,  Eggert and Germain.
Stable Acid Phosphatase I. Demonstration and Distribution.  Histochem 66, pp
301-317, 1980) discussed in detail the  rapid demineralization in acidic
buffers i.e. buffered formic acid for staining of stable forms of acid
phosphatase.  

 

I have both of these publications on file and will forward privately.   

 

I would err on the side of using a buffered formic acid with either sodium
formate or sodium citrate for doing this and use decalcification endpoint
testing to avoid over exposure to acid i.e. over decalcification.

 

Take care

 

Gayle M. Callis 

HTL/HT/MT(ASCP) 

 

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Re: [Histonet] Re: TRAP staining on formic acid decalcified bone reference

[koellingr]

Hi all, as usual Gayle was right on.  Use a buffered (more gentle) formic acid; 
not just formic acid per se of any water diluted concentration.  For end point 
testing, critical, we used a radiograph machine instead of chemical endpoints 
which is also fine;we just had access to a lot of equipment. 
Ray 
Washington 

- Original Message -

From: Gayle Callis gayle.cal...@bresnan.net 
To: Histonet histonet@lists.utsouthwestern.edu 
Sent: Thursday, April 2, 2015 3:12:55 PM 
Subject: [Histonet] Re: TRAP staining on formic acid decalcified bone
reference 

The reference within a reference from Ray is A Chimeric Form of 
Osteoprotegerin Inhibits Hypercalcemia and Bone Resorption Induced by IL-1β, 
TNF-α, PTH, PTHrP, and 1,25(OH)2D3 .   Sean Morony et al .  J Bone Mineral 
Res V 14, pp 1478-1485.   

  

However, the formic acid decalcification method is not described in detail 
and merely says formic acid but whether this is buffered formic acid or 
just dilute formic acid in water only is not stated.     Ray might elaborate 
on what specific formic acid recipe he used as many in research don't always 
use buffered formic acid decalcifiying solutions.   

  

I would assume Morony et all used a buffered formic acid with either sodium 
formate or sodium citrate and controlled so as to not overexpose TRAP to 
acids longer than necessary.   One publication,  i.e.,  Eggert and Germain. 
Stable Acid Phosphatase I. Demonstration and Distribution.  Histochem 66, pp 
301-317, 1980) discussed in detail the  rapid demineralization in acidic 
buffers i.e. buffered formic acid for staining of stable forms of acid 
phosphatase.   

  

I have both of these publications on file and will forward privately.   

  

I would err on the side of using a buffered formic acid with either sodium 
formate or sodium citrate for doing this and use decalcification endpoint 
testing to avoid over exposure to acid i.e. over decalcification.     

  

Take care 

  

Gayle M. Callis 

HTL/HT/MT(ASCP) 

  

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[Histonet] RE: cleaning glassware

[Anne Murvosh]

I always used to rinse the containers I used with alcohol and let them dry 
before doing the stain.  Some people use an acid alcohol rinse.  Anne

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Jeff Halstead
Sent: Thursday, April 02, 2015 11:48 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] cleaning glassware

Hi, All-I am currently experiencing problems with our warthin-stary silver 
stain. The finished product has silver particulates all over the empty slide 
and deposited on the tissue making the stain very difficult to interpet. Any 
ideas would be very helpful.  thanx
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[Histonet] RE: Question about Formic acid decal and TRAP stain

[Elizabeth Chlipala]

Debbie

In our hands it has not worked on formic acid decaled samples, EDTA decal only.

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
l...@premierlab.com
www.premierlab.com

March 10, 2014 is Histotechnology Professionals Day

Ship to Address:

Premier Laboratory, LLC
1567 Skyway Drive, Unit E
Longmont, CO 80504

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Debra Siena
Sent: Thursday, April 02, 2015 10:00 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Question about Formic acid decal and TRAP stain

Hi Histonetters,

I have a question to ask if you don't mind.  Can TRAP Histochemical staining be 
performed after decalcifying with formic acid? Any tricks of the trade, etc?  
If anyone has any experience or references that they could point me to, I would 
greatly appreciate it.  Thanks in advance for your help.




Debbie Siena
dsi...@statlab.com mailto:bbro...@statlab.com%7C | 
www.statlab.comhttp://www.statlab.com/

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[Histonet] RE: inconsistent HE staining

[Kienitz, Kari]

Hi Julie,
Try increasing your deparaffinization times, sometimes there is just not enough 
time in xylene so subsequent staining can be very inconsistent like described.


Kari Kienitz HT, (ASCP)
Histology Laboratory
Gastroenterology-EAST
The Oregon Clinic
 NE 99th Ave
Portland, OR  97220
503.935.8311
kkien...@orclinic.com




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From: histonet-boun...@lists.utsouthwestern.edu 
[histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Julie Cohen 
[juc2...@med.cornell.edu]
Sent: Wednesday, April 01, 2015 9:06 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] inconsistent HE staining

Hi,

I made slides of paraffin-embedded mouse small intestine (Swiss rolls), and 
stained them with Hematoxylin and Eosin.  Parts of the tissue on the same slide 
are stained dark with good structure.  Other areas look washed out with poor 
structure.  We realize that some of this could be caused by the 
orientation/structures captured, but similar tissue type looks paler as well.

Has anyone had a similar experience, and could suggest an explanation for me to 
give to our client?  At first we thought it might be due to poor fixation, 
since the centers of tightly-wound rolls were affected, but we also observed 
this in the outer parts of loosely wound rolls.  I soak the blocks before 
sectioning; could non-uniform swelling result in variations of the section 
thickness?  (These are 7 microns thick.)

Apologies if this information is available somewhere else; I tried 
unsuccessfully searching the archives.

Thank you,

Julie Cohen

Research Lab Tech
EM Core Facility
Weill Cornell Medical College
1300 York Avenue, Room A-105
New York, NY 10021
lab: 212-746-6146
email: juc2...@med.cornell.edumailto:juc2...@med.cornell.edu

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[Histonet] Re: inconsistent HE staining

[Teri Johnson]

Julie,

Are the inconsistencies consistent among different sections? In other words, 
with additional sections, are you seeing the paler areas in the same places or 
different ones? If you section it today, are the pale areas in the same place 
as when you sectioned it previously?

If it is the same area across multiple sections it is likely your tissues and 
not inadequate deparaffinization. Drying artifact is often seen in epithelial 
tissue structures and can happen when the tissue is out of fixative or have 
been placed in alcohol (which dehydrates much more quickly) and then made into 
a swiss roll. It can be a huge problem with GI biopsies if they are placed on 
dry gauze or paper towel prior to being placed in fixative at the time of 
biopsy. In my experience, there is not much you can do to fix it. Just be aware 
of keeping your tissues wet when creating the next swiss roll sample.

Teri Johnson, HT(ASCP)QIHC
Manager Clinical Trial Testing
Genoptix, Inc.
SAN5, Rm. 2005
760.516.5954 (office)
760.516.6201 (fax)




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[Histonet] RE: Histonet Digest, Vol 136, Issue 13

[Cynthia Hutchinson]

I can have them ready tonight or tomorrow

Cynthia Tily Hutchinson
Rsch Asst IV
Pathobiology
166 Greene Hall
Coll of Vet Med
Auburn Univ 36849
(334)844 7020

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of 
histonet-requ...@lists.utsouthwestern.edu
Sent: Wednesday, March 11, 2015 11:58 AM
To: histonet@lists.utsouthwestern.edu
Subject: Histonet Digest, Vol 136, Issue 13

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Today's Topics:

   1. anti-BRAF V600E (VE1) Ventana (Susan Foreman)
   2. RE: RE: Old slides. (suetp918)
   3. PRN (Kristy Castillo)
   4. Re: anti-BRAF V600E (VE1) Ventana (Patrick Laurie)
   5. Open histology positions: San Francisco (Cheryl)
   6. Direct Hire Histotech South of Tucson,AZ-New Grads Welcome
  to Apply! (Melissa Owens)
   7. Job opportunity (Hunter, Theresa)
   8. TDP43, PTG poly (victor_tob...@comcast.net)
   9. RE: Re: Bird head stored in 70% alcohol and possible
  decalcification (Rui TAHARA)
  10. BIG day today! (David Kemler)
  11. PA (Mike)
  12. Re: PA (Jennifer MacDonald)
  13. Diff Quik troubleshooting (Nancy Schmitt)
  14. THICK AND THIN SECTIONS ? (Klaus Dern)
  15. RE: Old slides (Mayer,Toysha N)
  16. 5-methylcytosine IHC in tissue (Mariela Chertoff)
  17. RE: Masson Trichrome stain (Mayer,Toysha N)
  18. RE: Old slides (Marcum, Pamela A)
  19. RE: Histonet Digest, Vol 136, Issue 12 (Solis, Bryan)


--

Message: 1
Date: Tue, 10 Mar 2015 13:15:05 -0400
From: Susan Foreman sfore...@labpath.com
Subject: [Histonet] anti-BRAF V600E (VE1) Ventana
To: histonet@lists.utsouthwestern.edu
Message-ID: 00bd01d05b55$c09b85f0$41d291d0$@labpath.com
Content-Type: text/plain;   charset=us-ascii

What vendor are you guys using for antibody anti-BRAF V600E (VE1)?  Spring
Bioscience or Ventana?  What dilution are you using?  Are you utilizing
amplification?   Expensive



--

Message: 2
Date: Tue, 10 Mar 2015 13:31:58 -0400
From: suetp918 suetp...@comcast.net
Subject: RE: [Histonet] RE: Old slides.
To: Gowan,Christie C christiecgo...@dermatology.med.ufl.edu,
'BerniceFrederick' b-freder...@northwestern.edu,
histonet@lists.utsouthwestern.edu
histonet@lists.utsouthwestern.edu
Message-ID: jqx50dw1pn5k224ex1acg53i.1426008646...@email.android.com
Content-Type: text/plain; charset=utf-8

So we actually cut the film around the section and mount to another slide and 
do pretty much what wascabove mentioned placing upside downband placing on 
paper towel. ??Actually works pretty good.
TJUH
Sue Paturzo


Sent from my Verizon Wireless 4G LTE smartphone


 Original message 
From: Gowan,Christie C christiecgo...@dermatology.med.ufl.edu 
Date:03/09/2015  4:01 PM  (GMT-05:00) 
To: 'Bernice Frederick' b-freder...@northwestern.edu, 
histonet@lists.utsouthwestern.edu 
Subject: [Histonet] RE: Old slides. 

Hi Bernice,
I have found that if you flood the slide with mounting media (don't use xylene) 
flip the slide over onto an absorbent lab wipe and put a heavy weight with even 
pressure and leave for a few hours. If the slide sticks to the wipe just put a 
few drops of xylene to clean up the slide. You may still have some tiny bubbles 
but it is much better than the alternative. Good luck.
Christie Gowan

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Bernice 
Frederick
Sent: Monday, March 09, 2015 3:42 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Old slides.

Hi all,
We received some old slides (1997-1998) that were coverslipped with film. 
Sakura I would imagine. The issue here is that the coverslips have come up from 
the slide and the tissue is adhered to the back of the coverslip. They need to 
be recovered so they can be evaluated. What do you all recommend? We use the 
CV5030 for coverslipping. I tried one with xylene and mounting media but there 
were still a couple of air bubbles in there.
Thanks,
Bernice

Bernice Frederick HTL (ASCP)
Senior Research Tech
Pathology Core Facility
Robert. H. Lurie Cancer Center
Northwestern University
710 N Fairbanks Court
Olson 8-421
Chicago,IL 60611
312-503-3723
b-freder...@northwestern.edumailto:b-freder...@northwestern.edu

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RE: [Histonet] Re: Post fixing IHC slides

[Obregon, Cecilia]

We bake our slide in the oven for 30 minutes at 65 degrees. 

Then placed them in a staining bucket 'covered' at room temperature for 30 
minutes. This small bucket contains a piece of gauze soaked in formalin at the 
bottom. 

We do not 'immerse' the slides in formalin, we just sort of use the contained 
fumes to post fix the slides. No rinsing after this step, just load them in the 
Benchmark Ultra, and coverslip as usual. (We do manually rinse all of our 
slides in Di soapy water before coverslipping..whether they have been post 
fixed or not to remove excess LCS)

Hope this helps,

I can be reached at 954-265-5317 if you have any further questions.

Thanks-Cecilia


From: histonet-boun...@lists.utsouthwestern.edu 
[histonet-boun...@lists.utsouthwestern.edu] on behalf of abt...@gmail.com 
[abt...@gmail.com]
Sent: Monday, March 30, 2015 8:30 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Re: Post fixing IHC slides

Hi Cecilia,
Glad to know post fixation will reduce section falling off from slide. We are 
bone research lab and always have problem of paraffin bone section falling off 
after antigen retrieval. There are many ways we have tried, but not improve too 
much. I know bone section will be easy to fall if it was incompletely fixed. I 
want to try your post fixation to secure the bone section on the slide. Would 
you tell me the exact way you are doing? You bake the slides in oven and 
deparaffin first? Then post fix the slides 30 minutes? How do you do vapor 
fixation? Then go on for IHC after rinse when you done post fixation?
Your answer is very much appreciated.
Dorothy

 1. RE: Post fixing IHC slides in formalin (Obregon, Cecilia)

 From: Obregon, Cecilia cobre...@mhs.net
 Subject: RE: [Histonet] Post fixing IHC slides in formalin


 Our facility implemented the post fixing of IHC slides years ago only for 
 tissues like breast, cell blocks, or anything that tends to fall off. We 
 baked the slides in the oven for 30 minutes, and then post fixed them for an 
 additional 30 min in formalin 'fumes'. It works great on breast resections 
 specimens, and it doesn't interfere with staining.
 Thank you,

 Cecilia M. Obregon
 Memorial Regional Hospital
 3501 Johsnon Street
 Hollywood, FL 33021

 
 From: histonet-boun...@lists.utsouthwestern.edu 
 [histonet-boun...@lists.utsouthwestern.edu] on behalf of Barry Rittman 
 [barryritt...@gmail.com]
 Sent: Saturday, March 21, 2015 2:20 PM
 Cc: histonet@lists.utsouthwestern.edu
 Subject: Re: [Histonet] Post fixing IHC slides in formalin

 There are always discussions about fixation but I have never seen comments
 about using vapor fixation for post fixing or for fixing fresh frozen
 specimens.
 Vapor fixation is simple to use and does not require any solvent as it uses
 the tissue fluids in the case of fresh frozen sections or the solution
 remaining after IHC.
 Especially useful for formaldehyde, alcohol,  acetone, osmium tetroxide etc.
 Barry

 On Sat, Mar 21, 2015 at 12:14 PM, Ann Specian thisis...@aol.com wrote:

 Does anyone post fix their IHC slides in formalin in an effort to try to
 reduce tissue loss?  If so, does anyone have a protocol for this that they
 have used and have seen good results?


 If you have any other suggestions which can help to reduce tissue loss
 during IHC staining, I would love to hear from you.
 Thanks,
 Ann
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