[Histonet] Zeus (Michels) fixative and buffer wash

[Gayle Callis via Histonet]

We purchased from Poly Scientific Corp, both the transport media (fixative)
and the buffer wash.  It comes in several sizes.  

 

Another supplier is Newcomer Supply.  

 

Gayle Callis

GCallis Histology Service LLC 

Bozeman MT 

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[Histonet] Attention Gudrun Lang Re EDTA method for bone marrow biopsies

[Gayle Callis via Histonet]

Dear Gudrun,

 

EDTA will always be slower than a buffered formic acid and there are caveats
about using it for some enzyme methods for bone. 

 

This EDTA method which is better than most but is avoid pH that is too low.
This EDTA method is made with tetra-sodium EDTA (Fisher BP 121-500 which has
a very high molecular weight).  

The pH is adjusted DOWN to pH of your IHC buffers.   I have a review of
decalcification which explains how EDTA decalcifies bone as a function of
pH.   The correct EDTA with a high molecular weight information is also
attached.  The method was developed by a bone guru. Webb Gee,  many years
ago, and has remained a favorite of mine and others for many years.  You
might get by decalcifying in 24 hours with the higher concentration of EDTA
but there is no true way to speed it up.We used acetic acid to adjust
the pH down since this EDTA is very alkaline but is highly soluble.  HCL can
be used to adjust the pH.  I liked pH 7.6 which alone, speed up the rate a
bit, and this is the same pH as TRIS buffered saline.   I made up my EDTA in
Dulbecco's PBS, or any PBS will work.   You decalcify at RT.   EDTA is not
affected by temperature since it is a chelator.A trick to help you is
cut off the cortical bone plug at some point during decalcification since
pathologists should be interested in only the trabecular bone with marrow
components.  This will speed up decal too. 

 

METHOD: 14% EDTA Tetrasodium DECALCIFICATION (Webb Jee, Stain Technology) 

 

140 g EDTA, tetrasodium salt in 800 ml distilled water.  Dulbeccos PBS
(DPBS) is preferred over distilled water.  Starting pH is approx. 11, very
alkaline and could damage alkaline sensitive protein linkages.  Hence using
a pH close to your IHC buffers is desirable. 

 

Adjust with glacial acetic acid (or HCL) using pH meter, mechanical stirrer
and continuous readout on pH meter.  It takes approximately 18 ml glacial
acetic acid for every liter of 14% EDTA (tetrasodium salt).Since one
adds a large volume of glacial acetic acid at beginning, start out with
suggested gm of EDTA in less buffer,i.e. 800 ml, adjust the pH down.  When
pH 7.4- 7.6 reached, bring final volume to 1 liter using the solvent i.e.
water, DPBS or PBS.   This is basically a titration technique with
continuous readout on a pH meter.  

 

Suspend bone in decalcifying solution and stir or rock gently.  

 

This decalcifying method is NOT used for articular or other cartilage work
since EDTA extracts proteoglycans and will change the tinctorial quality of
cartilage stains i.e. Toluidine Blue, and Safranin O/Fast green so these
appear weaker.   It can extract proteoglycans needed for IHC.  Samples can
be left in EDTA solution over a weekend, but change frequently to replenish
active decalcifying agent EDTA.   Use a sufficient volume, 20:1 for adequate
decalcification.  

 

I have a colleague using this EDTA method for murine bones, and she loves it
even for doing decalcified bone frozen sections.  

 

Contact me privately and I will send a publication explaining how EDTA works
chemically as a function of pH, Webb Jee publication and the weight
loss/weight gain decalcification endpoint method.   

 

Take care, and stay in touch.  

 

Gayle 

 

 

You wrote: 

 

I would be happy about some input about decalcification protocols with EDTA
of trephine bone marrow biopsies.

 

recommended duration of fixation?  Complete fixation.  NBF totally fixes in
24 h but you may not be able to wait that long.  

 

recommended duration of decalcification? Maybe 24 h, less is debatable.
EDTA will always be slow since it is a chelation process.  There is a weight
loss/weight gain decalcification end point test, highly advisable, to know
when sample is decalcified.  This requires a balance which weighs in mg, or
you can use xray method.  A chemical method for EDTA is painfully tedious.  

 

strategies for speed-up of decal? Use higher concentration 14% EDTA,
tetrasodium at pH 7.6 with some gentle agitation.

 

recommended EDTA-solution formula?  Provided above it 14% Tetrasodium EDTA,
pH 7.6.  This is highly alkaline EDTA and the pH is adjusted down with
acetic or HCl.  

 

Hopefully some experienced histotechs can share their knowledge with me.

 

thanks in advance

 

Gudrun

 

 

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Re: [Histonet] Decal following ISH?

[Gayle Callis via Histonet]

Dear Nancy, 

Try using EDTA after ISH.   Small embryos with shells will probably take
very little time in 14% Tetra Sodium EDTA in PBS (we use Dulbeccos PBS)
adjust the pH down to 7.4 - 7.6 with glacial acetic acid although some use
hydrochloric acid.  We adjusted the pH using constant stirring on a magnetic
stirrer with single electrode immersed in solution allowing continuous
readout on the pH meter.   Basically, you are titrating the pH down with
acid using a plastic Pasteur pipette, very easy.Tetra Sodium EDTA is
very alkaline, hence adjusting pH down with one of these acids is necessary.
7.4 is normal pH for Dulbeccos PBS and 7.6 is a working pH for TRIS buffered
Saline (TBS) or a use a pH compatible with ISH protocol.

EDTA will not be bothered by samples immersed in alcohol and you may be able
to preserve the ISH work done.  It is worth a try.   In the future, I
suggest the researcher do fixation, then EDTA decalcification before ISH
particularly if the signal is ruined on snails with shells.  I bet it would
make his results better too - no shell to deter penetration of reagents.
Good luck and let us know if you have success.   

Happy Holidays!!!   

Gayle Callis
HTL/HT/MT(ASCP) 

-Original Message-
From: Thomas, Nancy via Histonet [mailto:histonet@lists.utsouthwestern.edu] 
Sent: Tuesday, December 20, 2016 6:10 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Decal following ISH?

Hello all,
I received whole mount samples in 70% ethanol for paraffin processing.  The
samples are snail embryos and the researcher already did in-situ on them.
Because they are of varying ages, some of the shells will section without
decal, but some will need it.  However, this step was not done before the
ISH staining.  Does anyone know if decalcification can follow an ISH
procedure?  I have searched a few protocols online, and each of them did the
decal before hybridization.  I have not yet found one where the decal
followed the procedure.   My plan is to start with one embryo and try it to
see what happens.  Maybe the signal will remain, or maybe not.  Has anyone
done it this way and know the answer before I try?
Thank you,

Nancy Thomas
Senior Lab Manager, Histology Core
Stowers Institute for Medical Research
1000 E. 50th Street
Kansas City, MO  64110

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[Histonet] Kawamoto protocol online from Rowe lab

[Gayle Callis via Histonet]

Go to this link:   http://bonebase.org/bonebase/

 

Then click on Skeletal Phenotyping, then go to Histomorphometry to see how
to use Kawamoto tape system with all the sample handling.  Photos and an
excellent video titled High-Throughput, mutliimage cryohistology of
mineralized tissues is invaluable, with Protocol and Materials.   You can
also access a pdf of protocol.  Video access is on home page of this
website.   You will see cryomicrotomy, snap freezing and other techniques to
help you. 

 

Good Luck

 

Gayle Callis

HTL/HT/MT(ASCP)

 

 

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[Histonet] Sandersons Rapid Bone stain chemistry and history

[Gayle Callis via Histonet]

You worte:  RBS is the same as Methylene Blue, I believe.

 

Jessica Riggleman | Research Associate

 

Globus Medical, Inc.

Valley Forge Business Center

2560 General Armistead Avenue | Audubon, PA 19403

Ph: (610) 930-1800 ext. 2583 | Fax:

 

 

-Original Message-

From: Alicia Marie Ortega [mailto:alicia.ortega at colorado.edu
<http://lists.utsouthwestern.edu/mailman/listinfo/histonet> ]

Sent: Thursday, September 22, 2016 4:50 PM

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

Subject: [Histonet] Question on Sanderson Rapid Bone Stain

 

Hello everyone,

 

I am attempting to stain 30-40 micron thick undecalcified bone sections
embedded in poly(methyl methacrylate) with Sanderson Rapid Bone Stain (RBS)
with a Van Geison counter stain.

My first attempt at this stain resulted in a very faint stain from the RBS
(I could see very faint blue/green staining of osteoid/soft tissues) and a
very intense dark pink stain from the counterstain.  I was wondering if
anyone knows of a way to intensify the staining of the RBS?  I heated the
stain to 55-60 degrees Celsius as recommended by the manufacturer prior to
staining (with a 10 minute stain duration).

Thank you in advance for your time and help.

Sincerely,

Alicia Ortega

Postdoctoral Research Associate

University of Colorado, Boulder

 

***

Sandersons Rapid Bone Stain is oxidized methylene blue.  It is actually a
modified  Stevenels blue, a  polychromatic stain made by oxidizing methylene
blue with potassium permanganate that produces the byproducts i.e. methylene
violet, azure A, azure B, thionin, toluidine blue, thionin and possibly some
other thiazine dyes .Stevenels blue was first used by Maniatopoulos et al to
stain PMMA embedded, un-etched bone sections using a 60°C water bath and
optional counterstaining with either Van Gieson or basic fuchsin.   Cathy
Sanderson (Mayton) found a better,  easier way to make up Stevenels blue for
large production.   We found it tends to be a weaker stain unless you etch
the bone with 0.5% formic acid using a sonicator for 1 minute, rinse with
hot tap water very briefly, blot and look at the surface stained bone.
Over aggressive water rinsing allows the stain to release from this mildly
acid etched calcified bone.  All the etching is doing is a gentle removal of
calcium from only a few micrometers of bone surface and allow the stain to
penetrate better.   Acid etching intensified RBS staining but there are
drawbacks but ways to deal with that.   You can stain etched bone sections,
view, photograph and then do a very brief (only a few quick dips), blot
after counterstain, view and photograph the now counterstained section.
Avoiding over rinsing keeps the RBS in etched bone section.   If things
don’t look right, merely polish the stain from surface and start over.   

 

You can counterstain Sanderson RBS stained (etched) bone section with the
two mentioned counterstains, but it has to be done very briefly or the
counterstain will differentiate RBS from the bone and be overly red.
RBS, as used according to Sanderson and product brochure instructions, is
for un-etched mineralize bone sections in PMMA or even EXAKT preparations.
We preferred to acid etch, do the original Stevenels Blue instead of RBS,
and get a much deeper stain on osteoid, and other bone components.
Counterstaining was not done as it masked acid etched bone components
excessively or removed them,  

The joy of using RBS is avoiding a long, tedious, and messy making up of
Stevenels blue.   One can also try to stain longer in RBS per brochure
instructions to see if that deepens the staining.   Old bone picks up the
stain differently than young bone in terms of age of the animal.  Different
species can stain differently too.  

 

It is important to know that repeated heating of RBS (or Stevenels)  allows
the portassium permanganate to continue oxidizing the methylene blue, and
also keeps raising the pH  to be more alkaline.  One should replenish or top
off  the stain, always filter it into a clean coplin jar before using or
reusing this stain.  Eventually the stain does not work very well and you
need to start with new stock.  

 

We preferred to use a MacNeals tetrachrome combined with toluidine blue per
Sterchi method for more brilliant staining of osteoid other bone components
including cartilage.   

 

Take care

 

Gayle Callis

HTL/HT/MT(ASCP)   

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[Histonet] Attention Daniel Blackburn about plastics embedding

[Gayle Callis via Histonet]

You wrote: 

 

My lab hopes to get into plastic sectioning.  We need to be able to process
tissue pieces as large and thick as possible, but see that the largest
embedding molds for JB4 are only 13x19mm by 5mm deep.   We have two
questions:  (1) Do any of the available media (plastic or resins) allow one
to embed and section large  pieces (for example eggs with dimensions of 2 cm
or larger)?   (2) Is a special microtome (such as a retracting microtome)
needed?   Our reason for considering plastic is that we must section yolk,
which splits out of standard paraffin during sectioning. Any advice is
appreciated. -- Daniel Blackburn, Trinity College 

 



Glycol methacrylate is not designed for samples as large at 2 cm.  However,
I have one publication from an old Stain Technology for large
chondro-osseous (sp?) sample.   I also have a protocol from a lady who used
water based processing with GMA since it is water miscible.   This was a
protocol lipids which are removed by organic solvents.   You might be able
to develop a protocol with your 2 cm samples.   Kits are expensive and I
have information on making up GMA in house.   Hope you have a good fume
hood!!  As you can see from another post today, GMA contains sensitizing
chemicals, and a carcinogen.  It will require double gloving with nitrile
gloves, and other personal safety gear.   The fume hood is an absolute must
have. 

 

You will need a powerful microtome, i.e. Leica 2250 or 2265 or equivalent,
with very sharp knives, maybe even tungsten carbide.  We used glass knives
on a JB-4 microtome but I have a colleague who used a 2250 and disposable
microtome blades, but you blocks are pretty large.   You may want to
consider using Peel away molds which come in several sizes.   With your size
sample, infiltration and polymerization will be tricky.  You have to seal
air away from the top of molds in order to get the blocks to polymerize.  I
think people have used plastic wrap over the top of molds to exclude air
although our metal blocks fit in the embedding molds snugly and we did an
old school method to exclude air.  Melted paraffin around outside of molds,
a messy but effective air block.   

 

Please contact me personally and I will send this this information to you.
I think you can do this with a reply to all after reading this message.   

 

Take care

Gayle Callis HTL, HT, MT(ASCP)

GCallis Histology Service LLC

Bozeman MT/USA  

 

 

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Re: [Histonet] glycol methylacrylate GMA enzyme and IHC

[Gayle Callis via Histonet]

y, I can help with references
and  protocols.  
 
In today’s world, if wanting IHC on plastic embedded tissue, I would be
using Neil Hands recommendations and Poly methylmethacrylate (PMMA) plastic
embedding mainly since this plastic is removable.   I also have close
contact with Neil if you need to visit with him.   He is a good plastics
guru and has extensive experience with IHC using PMMA.Using this plastic
also requires stringent safety precautions for handling the chemicals.
 
Sorry I leaned towards a negative view of GMA.  
 
Take care
Gayle Callis HTL/HT/MT(ASCP)
GCallis Histology Service, LLC
Bozeman Montana USA   

 

 

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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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Re: [Histonet] 2 um sections H staining

[Gayle Callis via Histonet]

You wrote:  

When I cut at 2μm my H and special stains look pale. How can I get my
stains to pop or am I stuck with pale looking stains when sectioning that
thin? 

I run manual specials and a manual regressive H For H I've tried
increasing my time in hematoxylin (beyond the manufacturer recommendation),
diluting my acid alcohol differentiation, and increased time in eosin but

the slides still lack the vibrancy that many of the postdocs desire.

 

I use Shandon instant hematoxylin and alcoholic eosin by Thermo. Everything
else I prepare in house from scratch. Any recommendations?

 

 

First is a question.   Why do they require a  2μm thick section in the first
place?I had a pathologist many years ago fall in love with these very
thin section for all tissues with the same complaint of pale staining.   It
was explained to him that this thickness was excellent for bone marrow and
renal biopsies but too thin for the majority of other tissues.  Simple, you
are slicing through cells much of the time and leaving only cell walls for
staining.  It there isn't enough thickness there, then hematoxylin doesn't
have enough tissue thickness to be "vibrant", and the same for th eosin.
The pathologist went back to the former routine 5μm thick sections.   Some
laboratories do use  4 μm routinely. 

 

Things to try: 

 

1.   If this thickness is required to see basement membranes or marrow
cells

a.  Do not use regressive hematoxylin and eosin, but rather progressive
hematoxylin i.e. Gill III type formulation, and do NOT use any
differentiation solution which can remove hematoxylin. 

b.  Increase the thickness of section by trying 3 μm and 4 μm but use
progressive H  If you don't need 2μm, then go to 4 or 5

c.   Treat sections with 1% periodic acid for 10 min, rinse and then
stain with progressive H  This technique is found in Sheehan and Hrapchak
book.   It might improve the staining with your sections. 

d.

 

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Re: [Histonet] Bouins for testicular biopsies

[Gayle Callis via Histonet]

Richard, 

 

You wrote:  What are people fixing testicular biopsies in to evaluate
infertility?  In the past, I believe fixatives such as Zenker's and Bouin's
were used for this purpose since they enhance nuclear detail.  Obviously,
those fixatives can no longer be used.  Thank you.
 
***
 
I don't think Bouin's is forbidden in laboratories and we certainly used it
routinely for the Masson Trichrome connective tissue stain.  The problem is
having stock Picric Acid in crystal form, now frowned upon by chemical
safety people and eliminated from shelves these days.   I had no problem
storing stock picric acid under a layer of water and keeping crystals from
outside edges of lid.Zenkers is obviously not used due to mercury
content.   
 
Bouin's is still used for Masson's Trichrome staining and can be purchased
ready-made from Sigma, Fisher and elsewhere.   The key would be to use it to
fix testicular biopsies, with no more than 72 hour fixation.   Be careful to
wipe any drips from around lids where picric acid crystals form, collect and
dispose of this fixative per your lab's regulations.   
 
There is a B-5 substitute, sold by BBC,  which may do the job just as well.
This B-5 substitute is known to work well for bone marrow biopsies where
good nuclear detail is important, and may be a good option.  If your techs
are using Bouins for Massons Trichrome connective tissue staining, you could
get a small container for the biopsy.   
 
 
Good luck
 
Gayle M. Callis 
HTL/HT/MT(ASCP)
GCallis Histology Service LLC  
 
 

 

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[Histonet] NSH Journal of Histotechnology reminder on call of manuscripts for troubleshooting stains due July 1st

[Gayle Callis via Histonet]

Dear Histonetters,  

 

This is a reminder for people to submit a manuscript on troubleshooting H
and special stains on paraffin embedded tissue sections.   You do not have
to be a member of NSH to submit manuscripts to the Journal of
Histotechnology.   JOH wants to know how you solve staining problems for
both manual and automated staining for this special topics issue.  This can
be a short research communication previously known as a technical note.
There is still time to meet the submission deadline, July 1, 2016.  For
instructions for authors and submission, go to
http://www.editorialmanager.com/his/default.aspx. 
Inquiries can be directed to me or at JOH through contacts listed at under
publications and Journal of Histotechnology at www.nsh.org
  website.  

 

Share you expertise with others in a JOH publication.  

 

Thank you

Gayle M. Callis 

HTL/HT/MT(ASCP)

Assistant Editor/Acting Editor, Special Issue on Troubleshooting Stains

NSH Journal of Histotechnology

 

 

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Re: [Histonet] Reprocessing

[Gayle Callis via Histonet]

A quick technique, published in Histologic Vol XXXVI, No. 1, May 2003
http://www.sakura-americas.com/Histologic/Histo-Tips/1008.htmlby   

Michael Johnson,  A technic for correcting poorly processed paraffin blocks.

Melt the paraffin, blot to remove excess melted paraffin, place in cassette.  
Put on processor with rest of tissues sitting in NBF for daily processing run. 

This way you don't have to go through the agony of trying going back and then 
forward through reagents and the tissues will be less damaged by more heating 
and exposure to drying reagents.  

Good luck

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



-Original Message-
From: Mca Werdler via Histonet [mailto:histonet@lists.utsouthwestern.edu] 
Sent: Thursday, April 07, 2016 9:02 AM
To: Charles Riley 
Cc: histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] Reprocessing

What i would do, melt the blocks you made.

1. Put them back in paraffin for one hour 2. Put them back in another paraffin 
or one hour 3. Put them in xylene for one hour 4. Put them in another xylene 
for one hour 5. Put them in another xylene for one hour 6. Put them back in 
paraffin for one hour 7. Put them back  in paraffin for one hour.

It takes some time. The reason why it was dificult to cut is because the 
alcohol and paraffin dont mix, there is a possibility that the parafin was not 
well enough impregnated in the tissue.
With the above solution, you can restore that.

Maarten

UNAM neurobiologia, histología
Mexico

2016-04-07 4:43 GMT-05:00 Charles Riley via Histonet <
histonet@lists.utsouthwestern.edu>:

> Hello all,
>
>  We had an issue with our processor the other night. Someone 
> accidentally put 100% alochol into the last xylene station. The tissue 
> sections were difficult to cut and or were unreadable. What is the 
> best way to reprocess them?  They are mostly small GI biopsies and 
> only two larger specimens
>
> --
>
> Charles Riley HT(ASCP)CM
>
> Histopathology Coordinator/ Mohs
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[Histonet] Journal of Histotechnology special issue call for manuscripts

[Gayle Callis via Histonet]

Dear Histonetters, 

 

The Journal of Histotechnology is calling for manuscripts to be published in
December, 2016 . The manuscript submission deadline is no later than July
1st to allow for a review process.  Any late submissions would be published
in a future issue.This is a special issue on troubleshooting routine
hematoxylin and eosin and other special stains on paraffin embedded tissue
sections.   Modes of staining can be either automated or manual.  With
advanced staining technology, there is particular interest in problem
solving for automated staining.  Immunohistochemical troubleshooting is not
included in this issue.  The manuscript can be a Rapid Research
Communication, a longer, more comprehensive document or a scholarly review. 

To submit a manuscript,  go to:  http://www.editorialmanager.com/his/  then
click on button to "register now".  There also a link to "Instructions for
Authors" on that cover page.

If help is needed with manuscript writing, the JOH Writing Partners Program
is available to guide author(s) through the scientific writing process.   If
there are any questions, you can contact the journal via the NSH.org
website.  The email contact for JOH is found under publications.

You do not have to be an NSH member to submit manuscripts to this journal.


Thank you

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

Acting Editor for Troubleshooting Staining Special Issue 

Journal of Histotechnology

National Society for Histotechnology

 

   



 

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[Histonet] glycine method to remove aldehyde induced autofluorescence

[Gayle Callis via Histonet]

Dear All, 

 

Some person kindly mentioned my name as a source for the glycine method to
remove aldehyde induced autofluorescence.  We liked the simplicity of this
method, plus gentle to tissue sections.  

 

This was the original information but we modified it.   I have seen
concentrations of glycine range from 100 mM to 700 mM . 

 

Original method:  
 
1. Rehydrated tissue sections:  A Tris-glycine mixture (adjust 0.1M glycine
to pH 7.2-7.4 with 1M Tris base will saturate free aldehyde groups. (15-30
minutes at room temp in Tris-glycine for FFPE sections. Wash well in PBS.
If the tissue is fragile though, only use the Tris-glycine method.
 
2..  For tissues coming out of formalin, soak the tissues for 30 min to 1
hour and rinse well. 

 

 

Callis Modified Method: 

 

We did not use TRIS-glycine, preferring the same buffer used in IF staining.
Make fresh for a day's use.   500 mM glycine in pH 7.2 - 7.4  Dulbeccos PBS
(Sigma).  We increased concentration to 500 mM glycine for 15 - 30 minutes
at RT after we found 100 mM reduced autofluorescence while the higher
concentration did a better more complete removal.I don't think it makes
much difference if you use TBS or DPBS so you can use whatever your lab
prefers for IHC/IF staining should work equally well. 

  

We found  two changes of glycine solution worked well since you are
refreshing the solution to sop up those free aldhydes.  Do 15 minutes
incubation for each change, don't  rinse the sections between changes, just
tip, drain slides, blotted edges of sections, add solution on sections with
slides laying flat on a manual stainer.   Some people might prefer glycine
solution in a coplin jar if they are going to an automated staining system.


 

If you fear drying, one method was 700mM glycine, 0.15% BSA (use pure IgG
and protease free), and 0.1% sodium azide in PBS with 15 to 30 min RT
incubation.   Sodium azide can be left out since it is there to prevent
bacterial growth, and deemed unnecessary since our glycine solution was made
fresh before a one day/one time use.  Glycine is cheap and goes into
solution readily.  

 

FYI, lysine has also been used to get rid of free aldehydes (Elias J.
Immunohistopathology book) 

 

Good luck

 

Gayle Callis

HTL/HT/MT(ASCP)  

Bozeman MT   

 

 

 

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Re: [Histonet] auto-fluorescence

[Gayle Callis via Histonet]

Dear Joost, 

Yes and to read up on this, go to this website for an excellent, well
referenced free pdf i.e. Autofluorescence, causes and cures.
https://www.google.com/?gws_rd=ssl#q=autofluorescence+causes+and+cures+wrigh
t+cell+imaging+facility   They also posted a pdf on mounting medias for
fluorescence microscopy work.  

The pdf will tell you what tissue and cellular components autofluoresce.
The authors also provided various methods to reduce or remove
autofluorescence of these components but also how to remove aldehyde induced
autofluorescence if you are fixing tissues with NBF or PFA.   

Remember there is no autofluroescence in the Near Infra Red region so one
can use an NIR fluorophore, i.e. Alexa 750 (red) or another fluorophore for
the NIR region,  and even use tissue autofluorescence as a "counterstain
fluorescence" However, you cannot see these with the human eye but these
photograph beautifully.  OR if you have a spectral imaging or confocal
capabilities,  you can rule out autofluorescence without having to treat the
tissue section.

If you have difficulty access this pdf, I will send via private email.  

Gayle Callis HTL/HT/MT(ASCP)  
Bozeman MT USA



-Original Message-
From: Bruijntjes, J.P. (Joost) via Histonet
[mailto:histonet@lists.utsouthwestern.edu] 
Sent: Friday, February 26, 2016 3:00 AM
To: Histonet@lists.utsouthwestern.edu
Subject: [Histonet] auto-fluorescence

Hi histonetters

I hope one of you can help me. I don't have any experience with
fluorescence.

We are searching for some components and we will use  an alexa488 conjugated
secondary. Will there be an auto-fluorescence on different organs
(brain/spleen/liver)?

Greetings
Joost



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Re: [Histonet] formalin fixed tonsil frozen sections

[Gayle Callis via Histonet]

Dear Erin 

 

You wrote: 

 

Good morning!  My pathologists would like us to cut formalin fixed (not yet
processed) tonsil tissue on a cryostat for DIF staining.  Has anyone done
this?  I did a quick search that seemed to indicate that it was possible but
that the architecture of the cells would be altered because of ice crystals
and that it would be difficult to get the sections to stick.  If you have
any advice I would greatly appreciate it!

 


*

Several things to think about: 

 

FF tonsil should be cryoprotected with 20 -30% sucrose before snap freezing
to prevent ice crystal formation.  Simply take the FF tonsil, cut into
smaller pieces and immerse into 30% sucrose over night at 4C.  When the
tissue sinks to the bottom of this somewhat thick syrupy solution as which
indicates the tissue is cryoprotected.  Blot off excess sucrose, embed in
OCT and snap freeze.  You can snap freeze many blocks to stockpile controls
and store these at -80C, shorter time at -27C.  

 

One thing you did not mention is:  are tissues you are trying to do DIF on
for diagnosis also formalin fixed?  I would think a tissue control should be
handled the same way as the test tissue i.e. fresh tissue frozen sections
fixed with cold acetone compared to FF tissue.  If your tissues are not FF,
then why not collect fresh tonsil and snap freeze that as a control?  You
can make several blocks and store a -80C or even -27C (for a short time).
The concentration of your antibodies may be different on a FF tonsil section
due to the cross linking as compared to the concentration on a fresh tissue
frozen section fixed with cold acetone.   Antibody dilution would have to be
tested.  Personally, I would want my antibody concentration for a control
and test tissue to be the same.   If you already run DIF on a FF tissue from
patient, then a FF control works fine.   

 

If you are using FF tonsil sections as controls, then FF tissue will have
aldehyde induced autofluorescence but this can be removed.  Autofluorescence
is annoying when trying to read the sections and could mask what you want to
see unless you use a contrasting color fluorophore.

 

To get sections to stick, mount frozen section on reliable plus charge
slides, and air dry the frozen section a bit but be gentle when rinsing the
sections.  Formalin fixed frozen sections always have the possibility of
coming off, but many have success retaining the section and discussed many
times on Histonet.There may even be newer plus charge slides touted for
FF fixed frozen sections these days.  

 

Good luck

 

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

 

 

 

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Re: [Histonet] Glass knife holder

[Gayle Callis via Histonet]

Hi Jen, 

 

If you can't find a glass knife holder for your microtome, try using a
disposable blade, low profile.   I know Linda Jenkins sectioned GMA embedded
tissue (Immunobed is a formulation of GMA)  with a disposable blade, but the
microtome has to be powerful enough to do the work.   If I remember
correctly, she used Sakura Finetek's AccuEdge low profile and very sharp,
but not the same microtome as yours.  It would certainly be worth a try with
whatever disposable blade you have in your lab before investing in the
expensive holder.   You will be picking each section off the blade one at a
time with a fine forceps, and then float it on RT distilled water.  A glass
staining dish has always worked for this.   

 

Good luck and hope you have some success.  

 

Gayle Callis

HTL/HT/MT(ASCP)

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Re: [Histonet] Hematoxylin Precipitate

[Gayle Callis via Histonet]

Sandy, 

After years of using Richard Allan's hematoxylin 2 with great success,   if
we didn't filter daily before use, we had stain precipitate on sections.
Some of this comes from the hematoxylin continuing to oxidize in open air,
bacteria and other "crud".   Tim is absolutely correct ignoring
manufacturers no filtering instructions.   Being old school, we were taught
to faithfully filter any hematoxylin, regardless of progressive or
regressive types.If we topped off hematoxylin 2 or used new stock,  the
stain was filtered into a CLEAN staining container/dish.  Keep an extra
container around if possible.   We used a medium fast filter paper, Whatman
54.   I realize this takes time but junk on a slide is NOT good thing,
especially after IHC staining and have a photo to show this - the result of
being lazy and not filtering the hematoxylin on that particular day.   

We used a distilled water rinse before hematoxylin2, but DI H2O will be
contaminated with cellular debris and last hydrating alcohol carryover.
Change DI water frequently if you have many runs in a day.   We used 1
minute running tap water rinses after hematoxylin, clearant and bluing.   If
you are using running water rinses, take a look at the blue ppt in the post
hematoxylin container as you don't want that sticking to sections.   Non
running water rinses should be changed after each H run in my opinion.
Adequate clean water rinses are important to not have carry over of clearant
into bluing reagents or bluing reagent into eosin in order to maintain
correct pH for staining.

Good luck

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

-Original Message-
From: Tim H via Histonet [mailto:histonet@lists.utsouthwestern.edu] 
Sent: Tuesday, September 22, 2015 11:25 AM
To: histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] Hematoxylin Precipitate

You should be filtering your Hematoxylin on a daily basis regardless of what
the manufactures says.  We use to filter twice a day since we did a
traditional overnight run and then again in the afternoon for specimens that
had been microwave processed.  So much tissue washes off in the solutions
they should be changed or filtered fairly regularly to try and prevent cross
contamination on the slides.
 
You can also try increasing your rinse times and see if that doesn't help as
well.  
 
Thanks,
 
Tim
> 
> Message: 1
> Date: Mon, 21 Sep 2015 15:14:39 -0500
> From: "Sandra Cheasty" 
> To: "Histonet (histonet@lists.utsouthwestern.edu)"
>   
> Subject: [Histonet] Hematoxylin Precipitate
> Message-ID: <4cda87133587e64c965ce6c356d18...@svm.vetmed.wisc.edu>
> Content-Type: text/plain; charset="us-ascii"
> 
> Hello all,
> 
> Has anyone using Richard Allen Hematoxylin-2 noticed an
odd artifact on the slides after using the Hematoxylin for more than a few
days on their stainer? We are seeing small spore or pollen-like blue dots
here and there on the slides. It is not coming from the water bath or our
water supply on the stainer. I used sterile gloves, opened a new case of
slides, dipped them in DI water, then in the RA Hematoxylin 2 on the
stainer, then in DI again, air-dried and coverslipped them, and the blue
dots were there. The only way we got rid of the blue artifact was to use new
RA Hematoxylin-2 every 2-3 days, which is a bit expensive.
> 
> Thanks for your input, and if you can recommend a
different, reasonably priced hematoxylin, that would be awesome.
> 
> Cheers,
> 
> Sandy
> 
>  
> 
> Sandra J. Cheasty, HT (ASCP)
> 
> Histology & Necropsy Supervisor
> 
> UW-Madison, School of Veterinary Medicine

  
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Re: [Histonet] Hematoxylin Precipitate and filtering Gill formulations

[Gayle Callis via Histonet]

Yes, I have used Gill 1, 2 and 3 even in the early days of buying these
formulations from a vendor, and always filtered them before using.  

 Old school habits never changed..   

Gayle Callis 

-Original Message-
From: Manfre, Philip via Histonet [mailto:histonet@lists.utsouthwestern.edu]

Sent: Tuesday, September 22, 2015 12:26 PM
To: Elizabeth Chlipala <l...@premierlab.com>; Tim H <thiggin...@msn.com>
Cc: 'histonet@lists.utsouthwestern.edu' (histonet@lists.utsouthwestern.edu)
<histonet@lists.utsouthwestern.edu>
Subject: Re: [Histonet] Hematoxylin Precipitate

Wow, I agree with Liz.  There should not routinely be "so much tissue
washing off".  There is a fundamental problem, if this is the case.

With regards to hematoxylin, have you tried Gill's Hematoxylin, 1, 2, or 3?
These do not need filtering and do not produce a precipitate.

Phil.

Philip Manfre, B.A., HT (ASCP)
Associate Principal Scientist
Merck Research Laboratories
WP45-251
PO Box 4
West Point, PA 19486

215-652-9750
215-993-0383 (fax)
philip_man...@merck.com






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Re: [Histonet] Long reply on problems with hydrogen peroxide blocking

[Gayle Callis via Histonet]

 I suspect background is coming from another source which 
can be determined with an immunostaining reagent background test.   I will be 
happy to provide the simple background test  too. 

 

Take care

 

Gayle Callis

HTL/HT/MT(ASCP) 

 

 

Amos and Patrick Wrote: 

 

Hi,

 Peroxidase can really be a pain. If you look in the archives though

(or ask her really nice) Gayle Callis submitted a recipe for a glucose

oxidase for peroxidase quenching that does not include hydrogen peroxide.

If you aren't really a fan of making these things up I would bet dimes to

doughnuts that the recipe is *really* similar to the product from Biocare

Medical called PeroxAbolish.

Here's a link... http://biocare.net/product/peroxabolish/

I have used it and it did work, but I don't really use it regularly so

can't really compare it well. Incidentally if I am wrong in my assumptions

about the similarity of this to the glucose oxidase, I trust someone here

(even from the company itself) will gently correct me.

 

Cheers,

Amos

 

On Sat, Sep 12, 2015 at 1:00 PM, http://lists.utsouthwestern.edu/mailman/listinfo/histonet> >

wrote:

 

> Message: 5

> Date: Fri, 11 Sep 2015 20:33:45 +

> From: "Lewis, Patrick"  <http://lists.utsouthwestern.edu/mailman/listinfo/histonet> >

> 

> Hi Everyone

> 

> Thanks for your responses.

> 

> I am looking at cell surface markers,

> Sorry I should have said.

> 

> Based on what I found out below:

> 

> Methanol is out, even though I agree that Methanol does enhance the effect

> of H202 blocking.

> (I suppose I could try it to see how much/if any epitope loss there is in

> relation to H202 quenching,  At least It would help identifying false

> positives that are actually H202 artifacts.)

> 

> Also it looks like increasing the concentration of H202 is out.

> 

> As to when though,

> 

> It looks like with cell surface markers I should block after the primary,

> or even after the 2ndary?

> Can I do that successfully with a HRP labeled 2ndary?

> 

> thoughts?

> 

> Patrick.

 

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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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[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] Trichrome troubleshooting

[Gayle Callis]

Suzanne,   

I don't understand what you wrote here  We are using Leica's kit with the
Weigerts iron with Gills.  In a long histo career, I never used Gills half
oxidized hematoxylin for a trichrome stain nor heard of the combined
staining you mentioned.   Could you explain what you are doing here?  Is
this part of the Leica staining protocol?This is a new one for me to
combine Gills hematoxylin  with Weigerts iron hematoxylin and why it would
even be necessary???   Both of Leica's trichromes kits use Weigerts Iron
hematoxylin and do NOT have Gills in the method.  Their kits look more like
classic Mass Tri methods.   Personally I would only use Weigerts Iron
hematoxylin in a classic Trichrome method.   It is important to know
that iron hematoxylin is not stable over a long period of time since the
ferric chloride continues to oxidize the hematoxylin so it becomes very weak
or doesn't stain.   We never used iron hematoxylin for more than one day
although some people may have success over a week.   We never took a chance
in having weak staining with that continued oxidation by ferric chloride,
and only used freshly mixed iron hematoxylin for a day.   

I would be happy to personally send you a modified Weigerts Iron hematoxylin
that does NOT differentiate out so the iron hematoxylin staining is always
excellent.   It was found in a J of Histotechnology publication many years
ago, and is made up in house - even using other trichrome reagents from the
above vendors I mentioned.   The modified Weigerts iron hematoxylin is more
concentrated compared to the original iron hematoxylin recipe.   

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



-Original Message-
From: Suzanne Martin [mailto:smar...@lcpath.com] 
Sent: Tuesday, June 30, 2015 12:37 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Trichrome troubleshooting


Hi all,

We are having trouble troubleshooting our trichrome. It is too blue. We are
using Leica's kit with the Weigerts iron with Gills. Most of the small bowel
controls have seen improvement but patient tissue is not... strange. 

We have tried lessening the time in Gills, adding time for the last acid
step, even lessening time and adding time in the Weigerts. 


Thoughts?

Thank you.

Suzanne HT




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Re: [Histonet] Trichrome troubleshooting

[Gayle Callis]

Years ago, I was taught by Jerry Fredenburg, a stain guru, never to
microwave the Bouins step but do this for 1 hour at 60C or overnight at RT.
The post-fixation/mordant is very important in order to acidify the
connective tissue fibers properly for a trichrome stain, and the short time
in MW will not do the job.   Liz is correct here in letting the sections
stand longer in Bouins after microwaving or simply just do the 1 hour at
60C. 

I have removed Gills type hematoxylins from nuclei  by over exposure to
acetic acids so remember that ALL Masson's Trichrome reagents are acidified
with acetic acid and will automatically do this, even on Weigert's Iron
hematoxylin. We also examined our sections microscopically during
staining to make sure the check the depth of red staining reagents on smooth
muscle is correct - once again, Liz is correct about this fact.   

Gayle Callis   

-Original Message-
From: Elizabeth Chlipala [mailto:l...@premierlab.com] 
Sent: Tuesday, June 30, 2015 2:30 PM
To: Suzanne Martin; histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] Trichrome troubleshooting

Suzanne

How many times have you used the kit and reagents, I did look up how the kit
works but the trichrome stain can be tricky.  First of all you need to make
sure that the mordant (bouins solution) is at 60C prior to placing your
slides in them.  We normally heat up our bouins for at least an hour prior
to placing the slides in the solution.  We leave in bouins for an hour and a
half rather than an hour.   I see that this is a microwave protocol I cannot
comment on that but I don't think that the hematoxylin is the issue, if you
leave longer in 1% acetic acid that may pull some of the blue stain out or I
would try dehydrating with lower alcohol percentage that can pull some of
the blue stain out.   I would also try leaving it a bit longer in the bouins
after you microwave it - that might help.

Trichrome staining works best with fresh reagents so if you have used these
reagents too much that could cause problems.  I'm also not a big fan of the
one step trichromes, they are quicker but sometimes not as good as the two
steps, just my opinion.

FYI - to evaluate your staining look for a smaller vessel, the smooth muscle
should be nice a red, if its greyish or blue you have not done the stain
properly.  Good Luck

Liz

Elizabeth A. Chlipala, BS, HTL(ASCP)QIHC Premier Laboratory, LLC PO Box
18592 Boulder, CO 80308
(303) 682-3949 office
(303) 682-9060 fax
(303) 881-0763 cell
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: Suzanne Martin [mailto:smar...@lcpath.com]
Sent: Tuesday, June 30, 2015 12:37 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Trichrome troubleshooting


Hi all,

We are having trouble troubleshooting our trichrome. It is too blue. We are
using Leica's kit with the Weigerts iron with Gills. Most of the small bowel
controls have seen improvement but patient tissue is not... strange. 

We have tried lessening the time in Gills, adding time for the last acid
step, even lessening time and adding time in the Weigerts. 


Thoughts?

Thank you.

Suzanne HT




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Re: [Histonet] Xylene substitutes in staining line

[Gayle Callis]

You wrote: 

 

Hello Histo Land,

Does anyone use xylene substitutes in their stain line only? I am especially
interested in d-limonene options. Thanks in advance!

 


*

 

We used xylene substitutes in the stain line for both deparaffinization and
dehydration but did not use limonene (citrus smelling).   We preferred the
Clearite 3 (Richard Allan) i.e. single aliphatic hydrocarbons  also
available from other vendors, or Propar (Anatech)  for this purpose.
Limonene was particularly offensive and personally made me turn green with
nausea!  It was banned from my lab due to intolerance to the smell which
also included banning limonene based household cleaners in my home.I
think people can become sensitized to this substitute so be careful about
handling and working under a hood.   We also used Clearite 3 for tissue
processing to eliminate xylene.   

 

There are caveats about using xylene substitutes.  They are sensitive to
residual water carry over and do not clear water as well as xylene.  We
added an extra stations to deparaffinize sections and at the end of
dehydration sequence ensure NO residual water carry over before mounting a
cover glass.  On some high humidity days, Clearite 3 would be cloudy meaning
the last stations with Clearite had to be changed to fresh solvent not
exposed to humid air.  To combat a water carryover problem commonly not seen
with xylene and its ability to handle water carryover better, we rotated the
substitutes frequently during a work week so the last station was fresh.
If you have cloudy looking sections after cover slipping from a xylene
substitute, then you have water carry over.  A hint is if the last alcohol
in the dehydration series is pink, then there is water carryover.   Due
diligence is necessary to avoid poor paraffin removal and also good
dehydration.   In a very busy laboratory, this means extra time and expense
to deal with a xylene substitute.

 

The test for paraffin carry over in  deparaffinization sequence, one can
pipette a few mls of last 95% alcohol (closest to water) into a glass beaker
of tap water.  It the aliquot turns the water cloudy, then you have paraffin
carry over into the alcohol.  If not seen in last 95%, the test the 95%
station by working backwards in the deparaffinizing sequence.  If any
alcohol test is cloudy, then that alcohol and all previous stations were
changed in our lab.  Some rotation of clear test was done into an earlier
station slot. This will test also works when using xylene.  


However, I really like the emerging practice of using hot soapy water for
deparaffinizing sections recently discussed in depth on Histonet.   There is
a wonderful publication by Tony Henwood et al,  J of Histotechnol 2013;
36(2):45-50.  Tresa Goins also entered in on this discussion about their
lab's success with this method.  Go to Histonet Archives and read the
commentary.   It certainly means less dependence on organic solvents, less
exposure to potentially toxic chemicals, and the expense of solvent
disposal.   I think the soap method is definitely worth trying. 


Hope this helps


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


 

 

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Re: [Histonet] Cryojane bone frozen sections and immunofluorescence

[Gayle Callis]

Everyone wrote: 

 

Hi Ana,

 

The OCT on the coated slides will not disappear like it does on regular
glass slides. It appears to polymerize along with the coating, but it
doesn't seem to interfere on the tissue section with the staining.

 

According to Gayle Callis' recommendation, it's best to try using the least
amount of coating rather than more. More doesn't always mean better
adhesion, and as Nancy Thomas reported usually ends up causing more imaging
interference and stain uptake making for a messier slide.

 

Teri Johnson

Manager, Clinical Trial Testing

Genoptix, Inc., a Novartis company

BioPharma

1811 Aston Avenue

Carlsbad, CA  92008

USA

 

Ana,

We once tried 4x coated slides thinking, like you, that it would deliver a
better quality section and would stay on the slide better.  Our researchers
did not like them at all because of the high level of background  staining
and fluorescence.  We only use 1x now.

 

Nancy

 

-Original Message-

From: Ana Maluenda [mailto:
http://lists.utsouthwestern.edu/mailman/listinfo/histonet amaluenda at
svi.edu.au] 

Subject: [Histonet] Cryojane tape-transfer and hydration of sections

 

Hello Histonetters! 

 

I was wondering if there is anyone out there who is currently using the
CryoJane Tape-transfer system for bone sections. I have been trying to get
good sections for mice tibia and femur and often find myself with problems. 

 

I spent a good time trying to improve the quality of sections (tried
different temperatures, thicknesses and speed as well as 1x vs 4x coated
slides). The sections improved (although not perfect), but I was planning to
move on for a staining trial. Currently, the sections are taken at 5um, in a
4x coated slide, at -26oC. However, now I get myself with a problem when
hydrating the sections. At the moment, once the sections are taken, they are
kept in -20oC (for short-storage). They are then taken out and left at RT
for 30 min before hydration with PBS for another 30 min. I have noticed that
it seems as if the OCT around the sections doesn't completely dissolve. I
have already tried PBS vs dH2O and played around with times (from 5 min to
overnight) with no differences. 

 

Has anyone had this happening before? Can it be because of the 4x coat? Is
there anything I can do to? And would this be a problem for
immunofluorescence? 

 

Any advice would be much appreciated! 

 

Thanks in advance, 

 

Ana Maluenda 

 


**

 

To Teri and all, 

 

Thanks Teri for reiterating my suggestions along with more information. 

 

We found the 4X coating to be unacceptable.  It is  more sticky but the
polymer is too thick and will cause more background when one needs to deal
with for immunofluorescence work.With murine turbinates, unfixed and
calcified, we did use the 1X and even 1/2X but it was necessary to flash the
UV three times, waiting 30 sec between flashes so the capacitor could build
up charge.   It takes patience.   We sectioned at 5 µm but the d profile
tungsten carbide knife was very sharp so do not cut on an edge you use for
trimming into the block.  Careful removal of the pink tape is required,
inside cryostat chamber, brace the corner of slide, then pull pink tape
diagonally across the section from one corner to opposite corner.  You have
to play with temperatures with everything the same temperature.   For
turbinates -30C worked well, but tape, slides, rollers, etc. blade and
samples were at that temperature, including the UV platform.  Sectioning
temperatures vary with different laboratories, the sections can be air dried
like any other frozen section destined for solvent fixation.  I would go
ahead and air dry a frozen section and store at -80C, not -20C since the
colder temperature is more suitable for retaining antigenicity.   We
preferred using fresh, unfixed tissues, snap frozen correctly (not in a
cryostat!) over NBF or PFA prefixed/cryoprotected snap frozen bone as we
found the fresh tissue frozen section stayed transferred to the slide
better.   Other may have a different experience.   

 

You cannot remove the OCT from the polymerized surface and don't need to do
that.  There are some things your researchers will have to live with.  1)
polymer exists even with 1X, but focusing on the plane of the section and
what is in the section will work.   2) make sure you work with the brightest
fluorophors i.e. Alexa dyes.  3) autofluorescence is caused by many things
and if your sections are prefixed with NBF or PFA, then aldehyde induced
auto-fluorescence will happen but can be treated.  Go on web and get
Autofluorescence causes and cures pdf.  4) if you work with Near infra-red
fluorophores, there is no auto-fluorescence in the NIR region but the eye
can't see it but the photos are spectacular.  5) Totally  fill in those
weird polymer gaps, be overly generous with antifade

Re: [Histonet] toluidine blue for cartilage with controls and mast cell staining

[Gayle Callis]

You wrote:  

 

We use a canine mast cell tumor as positive control - veterinary lab
naturally.

Probably looking for mast cells in the core.

 

Tresa

 

-Original Message-

From: Bernice Frederick [mailto:b-frederick at northwestern.edu
http://lists.utsouthwestern.edu/mailman/listinfo/histonet ] 

Sent: Tuesday, June 02, 2015 12:02 PM

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

Subject: [Histonet] Toluidine blue

 

Hello all,

I was taught to do Toluidine Blue O without a control. Is there actual one
and what would it be? I'm staining a bone core. Don's ask why, it's research
and what a researcher wants... Plus they have a protocol they are following
for this cartilaganous defect.

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

 

*

Bernice and Tresa, 

 

Having done a bone research study like this in the past, controls should be
and were carefully done.  You need to know normal cartilage from treated or
defect in cartilage.  The researcher certainly should have set their
experiment up accordingly but may have controls in place now???You did
not say if this is articular cartilage from exterior joint surfaces where
they took the core or deeper in the bone at the growth plate?  These two
cartilages will stain differently with T blue.  Normally and when studying
articular cartilage defects, it is wise to also do a safranin O/fast green
stain along with the T Blue. Controls are extremely important and need
to be carefully set up.   Hopefully, you have a contralateral bone normal
core from the same animal OR a core sample from an untreated, naive control
animal.It was never stated what the experimental animal model is being
used? I have done a study like this in the past.

 

When core is  decalcified with an acid or EDTA, then the control needs to be
decalcified exactly the same way and at the same time as experimental cores
with defect.  If you are decalcifying with EDTA, then you should have a
normal core that is not decalcified.  This is difficult with mouse but
possible with larger animals.  The reason is to see if the proteoglycans in
the articular or even the growth plate cartilage will be extracted by EDTA,
and not appreciably by buffered formic acid.  Articular cartilage where
proteoglycans have been removed by a decalcifying agent will have different
tinctorial quality (lighter) than cartilage never exposed to a decalcifying
agent.  EDTA is used by biochemists to extract proteoglycans for biochemical
studies, and will the same thing in a cartilage section.  Hence, there will
be less staining seen with the toluidine blue or the Safranin O/fast green
stain after EDTA.  Hence you should run two controls, 1)a decalcified
cartilage control and 2) an undecalcified control.   How you decalcify will
be important in order to retain proteoglycans in the cartilage.  I strongly
suggest using buffered formic acid, available commercially.   You will find
recipes for buffered formic acid in text books that contain sodium formate
or sodium citrate.  Look for these ingredients in product MSDS before you
buy the formic acid decalcifying solutions.   If there is any question about
EDTA versus buffered formic acid and other acid decalcifiers i.e HCl, Nitric
acid, etc.  for cartilage studies, I will be happy to send publications
concerning this topic privately.   

 

The Toluidine blue that we do for cartilage is designed to show cartilage
staining and not mast cells.  It could be the mast cells might be seen along
with the cartilage staining but that is not the point. 

 

The toluidine blue stain I do for mast cells is entirely different from the
toluidine blue cartilage staining protocol. 

 

I will be happy to send you a toluidine blue stain procedure for cartilage
and also the SafO/Fast green protocol.   I have a superb  T blue mast cell
stain from Churukian which allows mast cells to stand out without any blue
background in surrounding tissues that is often seen with other T blue
staining protocols.  

 

Hope this helps.  

 

Gayle M. Callis

HTL/HT/MT(ASCP) 

 



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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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[Histonet] Staining net dishes for brain sections

[Gayle Callis]

Out of curiosity and having received an email from Brain Research
Laboratories about this new item, has anyone tried the staining net dishes
for free floating brain or other tissue sections? 

 

I thought this a clever, useful device to keep the sections from physical
damage without having to lift the sections in some way.   

 

Gayle Callis

HTL/HT/MT(ASCP)

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[Histonet] Can't log into Histonet to do anything

[Gayle Callis]

Dear Histonettters, 

 

At the risk of being pesky, is Histonet having problems.   I generally go to
Histonet via Firefox/Google and haven't been able to get to the website for
two days.   I only hope someone out there can even get this message.  I have
tried finding Marvin Hanna's email address.  

 

Stymied and dead in the water.   I would love to unsubscribe, but not sure
anyone gets this message. 

 

Gayle Callis 

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[Histonet] Passing onto PAS discussion RE: GMS question

[Gayle Callis]

Bob, 

Culling said exactly the same thing about not needing precision weighing for
periodic acid. I discovered, by a huge error in PAS staining results,
that  pre-weighing 1 gm of PA into a clear sample vial and letting the
pre-weighed PA stand at RT should NOT be done.  The PA degraded to the point
of very poor staining.   I thought I was doing myself a favor to have faster
solution preparation, and the pre-weighing efficiency idea was abandoned.
We sometimes learn the hard way.   Daily freshness is the key and we liked
1% periodic acid  (or 0.5% per the original McManus and Mowry method).   

 Our lab used 1% PA for a standard PAS stain for 10 minutes (never used for
fungus) most of the time.  It is interesting to note how many variations of
the PAS stain exist in this world.It's important to check what PAS
method variation works best for the component you are looking for as one
variation may not be the best for the tissue component you want to see.
If over oxidation is suspected, reduce oxidation time to 5 min (original
McManus/Mowry method) and/or reduce concentration of PA to 0.5% periodic
acid instead of 1% PA OR do both. Good positive controls are mandatory
for any particular tissue component in mind.   

The only time we used a periodic acid/methenamine silver staining was for 2
micrometer renal biopsies aka Jones Basement membrane stain, and our
methenamine silver was kept fresh with a 6 month expiration date on in house
preparation.   Fresh methenamine silver stock solution is important too.


Chromic acid cannot be drain dumped and hopefully people will have hazmat
collection available.  If you have to collect DAB, silver solutions, then
chromic acid (chromium trioxide is the stock chemical) can be collected.
It isn't that difficult.   

Thanks for the supportive comment on GMS.  Your comments on Histoplasma sp.
staining were interesting as not many of us have the opportunity to work
with ancient tissues.   

Gayle Callis  

-Original Message-
From: Bob Richmond [mailto:rsrichm...@gmail.com] 
Sent: Saturday, May 02, 2015 7:30 AM
To: Histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] GMS question

Gayle Callis's discourse on GMS staining is must reading for anyone who does
the GMS stain. Freida Carson in 1999 showed the importance of using chromic
acid - I was distressed that she didn't cite this paper in the 4th edition
of her book.

The most rigorous test of the GMS stain is probably the dead histoplasma in
ancient fibrotic granulomas. I've gotten these things to turn up with GMS
more than once - a finding of potential clinical importance.

For control material, I don't think there's any substitute for infected
human (or mammalian) tissue, preferably though not necessarily with a
species identification of the fungus. I think either active histoplasmosis
or invasive aspergillosis might provide the best material. The big academic
centers that are still doing autopsies could be helpful in getting control
material for everyone.

Periodic acid should indeed be made up fresh with the dry chemical, but
precision weighing isn't required - I used to keep a small plastic measure
in the stock bottle so I could simply spoon out what I needed for the day.

I'm not sure what the rules are for disposing of chromic acid, but it's a
significant hazmat - toxic metal, strong acid, strong oxidant. I think you
can neutralize it and precipitate the chromium, but I'd have to look up the
method.

Bob Richmond
Samurai Pathologist
Maryville TN
**
Gayle Callis wrote:

The same site of infection for your control and patient tissues does not
mean the fungus species was the same.  Now for addtional commentary.

The classic GMS uses chromic acid as the oxidizer, stronger than periodic
acid.   This is something to beware of since you had a false negative result
with the kit. Periodic acid should be freshly made up from periodic acid
crystals just before use (Culling insisted on fresh periodic acid reagent)
then discarded after that day - something that is not going on with a kit
where all components are sent to you in ready to use liquid form.   I think
if you had used the classic GMS with chromic acid to start with, you would
have had the results seen with Periodic Acid/Schiffs.   Part of the problem
is the not all fungus species stain well either PAS or Periodic acid/GMS,
and even classic GMS.   All these factors can present a problem when trying
to diagnose fungal infections.Lee Luna in AFIP manual, pointed out it
has been found that the time of exposure to methenamine-silver nitrate
solution for complete development may vary according to type and/or strains
suspected.   He advised if Nocardia asteroides is suspected, then two
slides should be run:  one for 60 minutes and one for 90 min in the silver
solution.   Histotechs should be aware these possible problems. Fungi
are difficult at best even when doing fungus cultures and treat

[Histonet] long answer RE: GMS Question

[Gayle Callis]

Hi Paula, 

The same site of infection for your control and patient tissues does not
mean the fungus species was the same.  Now for addtional commentary. 

The classic GMS uses chromic acid as the oxidizer, stronger than periodic
acid.   This is something to beware of since you had a false negative result
with the kit. Periodic acid should be freshly made up from periodic acid
crystals just before use (Culling insisted on fresh periodic acid reagent)
then discarded after that day - something that is not going on with a kit
where all components are sent to you in ready to use liquid form.   I think
if you had used the classic GMS with chromic acid to start with, you would
have had the results seen with Periodic Acid/Schiffs.   Part of the problem
is the not all fungus species stain well either PAS or Periodic acid/GMS,
and even classic GMS.   All these factors can present a problem when trying
to diagnose fungal infections.Lee Luna in AFIP manual, pointed out it
has been found that the time of exposure to methenamine-silver nitrate
solution for complete development may vary according to type and/or strains
suspected.   He advised if Nocardia asteroides is suspected, then two
slides should be run:  one for 60 minutes and one for 90 min in the silver
solution.   Histotechs should be aware these possible problems. Fungi
are difficult at best even when doing fungus cultures and treat.
Fortunately, there are antibodies for some fungi i.e. Aspergillus, Candida)
but you would probably need the specific fungus as a control for the
antibody being used.   Fungus IHC experts can weigh in on the latter topic.
I don't know what fungus species was in your  fungus ball control block,
but our clinical lab fungus ball control contained Aspergillus sp. from a
post mortem human lung.  With the researcher, he only had pure Aspergillus
sp. infecting the tissues.It would be nice to know what species of
fungus is in your control tissue block 

The publication by Frieda Carson along with Jerry Fredenburgh and John
Maxwell   Inconsistent detection of Histoplasma capsulatum with periodic
acid in GMS fungus stain  , J Histotechnology, 1999 is a profound statement
about what you just experienced.   Their tissue control i.e. Aspergillus sp.
stained adequately with periodic acid/GMS but the H. capsulatum fungus in
tissue did not stain.  They studied several different times, temperatures,
periodic acid concentrations and fungus species.  They had additional
controls containing fungi other than Aspergillus sp. for better sampling of
PA/GMS on 
different  fungi.   Having different fungus species control blocks is a
luxury many labs do not enjoy.  The reason chromic acid is not in kits is
due to shipping, health and safety hazards, must be handled carefully and
collected for proper, safe disposal so it is easier to make up GMS kits
with periodic acid.If you have to collect your silver solutions for safe
chemical disposal, then chromic acid shouldn't be a big problem to do the
same. 

Also, since Periodic acid/Schiffs is popular and commonly used for fungus
staining,  PAS can also present false negative results or weak staining  due
to the weaker periodic acid oxidation, even when the periodic acid is made
in house, fresh for the day.   Chromic acid/Schiffs has been recommended by
people on Histonet to improve fungus staining over PAS.   PAS stain always
seem to work well with Candida sp. and Aspergillus sp. but classic GMS was
always better in my hands.   I only used classic GMS, prepared in house, and
controlled the development with a microscope to avoid under and over silver
staining of fungus.   

I will be happy to send the Carson et al publication and scan the AFIP
manual pages with photos on fungus staining by Luna.   I apologize for long
discourse as chromic acid/GMS stain is one of my favorite special stains to
perform along with a long standing interest in fungus staining. 

I hope this helps.

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

-Original Message-
From: Paula Lucas [mailto:plu...@biopath.org] 
Sent: Friday, May 01, 2015 8:17 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] GMS Question

Hello,

 

I think I already know the answer but I'm not sure why so if someone can
help me understand the theory behind it, I would greatly appreciate it.

 

Currently, we use the Richard Allen kit for the GMS stain and it uses
Periodic Acid as the 1st step.  

We use a control tissue from a case we had that was positive for fungus and
it's a fungus ball from the Rt Maxillary. 

We ran a test for fungus on a different and current case of the same tissue
(different patient): Rt Maxillary sinus.

 

The control tissue did work, but the patient's tissue did not, so the doctor
ordered a PAS for fungus and this clearly showed the fungal elements nicely.

 

My question is why would the control and patient tissue have different
results when they are both fungus balls from the 

Re: [Histonet] GMS Question

[Gayle Callis]

Hi Tresa, 

What staining parameters do you suggest for seeing  mature and/or immature
fungal cell walls?   I don't think you will know what level of maturity is
present before doing a fungus stain.  But wouldn't both levels of maturity
both be present if the fungus is actively growing in a tissue?  

What do you recommend, using both a PAS, or chromic acid/Schiffs along with
chromic acid GMS method?   

We used 4% chromic acid at RT for 1 hour but would you recommend duplicating
slides so you do pull one slide out of chromic acid at 30 minutes and one
for 60 minutes to stain either mature or immature fungal and/or both cell
wall structure.   

Gayle Callis
HTL/HT/MT (ASCP) 



-Original Message-
From: Goins, Tresa [mailto:tgo...@mt.gov] 
Sent: Friday, May 01, 2015 11:29 AM
To: Paula Lucas; histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] GMS Question

To get a positive PAS or GMS fungal stain, one must oxidize the carbohydrate
in the fungal cell wall.  
Chromic acid is a stronger oxidizer than periodic acid, so would work better
with mature fungal cell walls that are highly polymerized.
Treat an immature cell wall for too long, and you may get a false negative
because the carbohydrate structure no longer resembles a fungal cell wall.

Tresa

-Original Message-
From: Paula Lucas [mailto:plu...@biopath.org] 
Sent: Friday, May 01, 2015 8:17 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] GMS Question

Hello,

 

I think I already know the answer but I'm not sure why so if someone can
help me understand the theory behind it, I would greatly appreciate it.

 

Currently, we use the Richard Allen kit for the GMS stain and it uses
Periodic Acid as the 1st step.  

We use a control tissue from a case we had that was positive for fungus and
it's a fungus ball from the Rt Maxillary. 

We ran a test for fungus on a different and current case of the same tissue
(different patient): Rt Maxillary sinus.

 

The control tissue did work, but the patient's tissue did not, so the doctor
ordered a PAS for fungus and this clearly showed the fungal elements nicely.


 

My question is why would the control and patient tissue have different
results when they are both fungus balls from the same specimen source?

 

Thanks in advance,

Paula

Lab Manager

Bio-Path Medical Group

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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: 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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[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: 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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[Histonet] Staining dishes and slide racks for sale

[Gayle Callis]

Dear Histonetters, 

 

Janet Maass has these new and used staining dishes, covers and slide racks
for sale.   I will be happy to forward your email message to her since
Histonet messes with email addresses. 

 

22 glass staining dishes with covers (for 50 slides) 4 extra covers
available

 

8  glass staining dishes with covers (for 30 slides) 6 extra dishes
available

 

12 new glass staining dishes still in original boxes (for 30 slides)

  

10 large (50 slide capacity) metal staining racks with 16 handles

  

4 small (30 slide capacity)metal  staining racks with 3 handles  

 

Gayle M. Callis

HTL/HT/MT(ASCP) 

 

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[Histonet] Acetone fixing and tissue damage

[Gayle Callis]

You wrote:   

 

Hi Everyone.

 

When I fix my cryosections  in acetone,  I am using HPLC grade 99.9% for 10
minutes at -20C.

 

Would the Histology grade 99.5% be less damaging to them?

 

Higher H20 content, i.e. less than 99.5% apparently is also very bad.

 

With the HPLC grade I often get tissue damage, the tissue also floats off
the slide causing a stringy effect.

 

Fixing with 4% p-formaldehyde or 100% Methanol, prevented the antibody from
recognizing the Nuclear Antigens.

 

Looking for advice,

 

Patrick.

Patrick Lewis

Research Associate II Bench

Seattle Childrens Research Institute

206-884-1115

 


*

HPLC grade acetone is not necessary plus very expensive.  Use ACS Certified
Reagent 99.5% grade, not histology grade, which you can buy in gallon size.
Maybe what you are calling histology grade is the ACS Certified Reagent
grade but  Histology grade implies a practical grade of acetone which is
not a pure as the ACS certified Reagent grade.   Also, 4°C acetone works
just as well.If you are storing your acetone (in a staining jar) inside
the cryostat to maintain a 20°C temperature, don't!!!  If your staining
container tips over, you will ruin your cryostat!!   Hopefully you are using
high quality plus charge slides?

 

We had frozen sections come off a plus charge slide after single 4°C acetone
fixation on occasion.  You can prevent frozen section loss is a Double
Acetone fixation that also increases permeabilization. An IHC guru gave
me this hint years ago and was given to her by a company selling
immunostaining products.   A small fan will be your best friend for RT air
drying and/or evaporating acetone.  However we air dried all  FS were dried
at RT for 30 minutes minimum  or longer before fixation.  By air drying, you
get rid of the water.  You can put your FS in front of a small fan, or
inside a hood for faster drying, and never store just cut FS in the cryostat
where water condensation occurs when you take them out of cryostat
environment to RT.DRY  frozen sections were the rule in our lab before
any solvent fixation.  

 

1.   Air dry frozen section at RT for 30 min 

2.   Fix FS in  4°C acetone for 10 minutes

3.   Air dry FS for 15 minutes to evaporate acetone

4.  Return FS to 4°C acetone for 10 minutes

5.  Air dry to evaporate acetone 

6.  Proceed to immunostaining 

 

Gayle Callis

HTL/HT/MT(ASCP)

 

 

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[Histonet] RE: Masson Trichrome stain

[Gayle Callis]

I have been following the string of inquiries about using metal forceps with
Masson's Trichrome staining.   I was taught many years ago to avoid metal
forceps or the older metal tissue cassettes with Bouins.   I am scrambling
to find the actual reference.   The reason given was acids in Bouins corrode
metal.  This may be a lost bit of information since the overall majority of
labs now use plastic tissue cassettes.   Case in point:   using acidic
descaling solutions for household cleaning i.e. showers/tubs or coffee
machines.   These solutions come with warning to avoid metal fixtures and
stainless steel sinks.  Accidental contact of acids in a stainless sink
causes the metal  discolor, indicating corrosion - been there, done that to
a stainless steel sink.  I so use metal forceps to move slides between Mass
Tri staining solutions (and silver staining solutions) without problems per
John Kiernan's comment.   

 

Not using metal forceps with silver stains i.e. GMS, reticulin, is to avoid
metal ion contamination which is more likely due to with poorly washed
glassware.  In the past, we dipped metal forceps in melted paraffin, very
messy since paraffin comes off on slides and in hot staining solutions.
Disposable plastic forceps are cheap but break easily resulting in a dropped
slide.   Teflon forceps are pricey but it was a challenge to hold slides.
Hopefully there are teflon forceps that work better than the one we used?
We tried a teflon tipped metal forceps but not worth the price as teflon
wears off the tips to rexpose metal.Weigerts hematoxylin  is not
affected by metal forceps since there are no acid components to corrode the
metal although Weigerts can stain the forceps.   Simply wash the forceps
in dilute chlorine bleach then soap and water.   I agree with John Kiernan
and now use metal forceps to move slides between staining solutions in both
Massons trichrome (and silver methods)  without problems.  If people want to
use plastic or teflon forceps, I understand the reasons.  

 

As for not rinsing before going into Aniline Blue (or light green) in
Massons trichrome, there is a reason for this.Sheehan and Hrapchak state
verbatim  The phosphomolybdic acid and phosphotungstic acid thus acts as a
link connecting basic groups of the connective tissue fiber to the basic
groups of the dye i.e. aniline blue.   The PM/PT acid treatment has the
ultimate effect of making an amphoteric dye that would ordinarily act as an
acid dye to change and act as a basic dye.  These authors also say
Although the exact mechanism of how the stain works is unknown, some
theories are available. By rinsing away the PT/PM, the link may be
weaker hence one goes from PT/PM directly into aniline blue (sometimes light
green or fast green).Bierbrich Scarlet/acid fuchsin and aniline blue
(light green or fast green) solutions can be filtered back and reused many
times.  PT/PM and 1%  acetic acid solutions  should be discarded after use.


 

Instead of kits due to expense and some kit deviations from classic Massons
Trichrome method, I found I could buy excellent, reliable single staining
solutions i.e. Biebrich Scarlet/Acid Fuchsin and Aniline Blue from Newcomer
Supply or Poly Scientific to avoid exposure when weighing out carcinogenic
dyes.   Bouins is purchased from the vendor with the best price.  However,
PT/PM and acetic acid single use solutions were still made in house to save
costs.  

 

I strongly recommend reading John Kiernan's   Methods for Connective
Tissues  from his book , Histological and Histochemical Methods Theory and
Practice  for better explanation and understanding of Massons Trichrome
chemistry. Collagen and muscle staining methods in Sheehan and Hrapchaks
Theory and Practice of Histotechnology is not recent but a good start.


 

Whew, a long reply but hope helps...

 

Gayle Callis

HTL/HT/MT(ASCP)  

 


 

Written is:  

 

Justine,

 

I do not have any metal forceps in the special stains area, due to the
reaction that they can cause when staining with silver.  As a rule of thumb,
it is just easier to use plastic all the way around.  

The Carson text does not state the use of only plastic forceps, but I would
think that maybe they are concerned with a reaction between the Weigert's
and the metal.  That would be a stretch.

As for no water before aniline blue, I believe the concentration is very
weak and the water may dilute they dye even further.  This would affect the
staining results.

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: 4

Date: Tue, 10 Mar 2015 00:31:56 -0500

From: John Kiernan jkiernan
http://lists.utsouthwestern.edu/mailman/listinfo/histonet @t uwo.ca

Subject: Re: [Histonet] FW: Masson's trichrome stain

To: Linda Margraf lindamargraf
http

RE: [Histonet] Acetone fixing and tissue damage

[Gayle Callis]

Dear Bernice, 

This has always been my favorite solvent fixative for murine CD markers and
some other markers/antigens but it can't be used for human CD4 and CD8 IHC.
These human  markers do not tolerate the alcohol component  and won't stain.
The late Dr. Chris van der Loos imparted this bit of information after I
recommended he should try it.   I think most people do human CD4 and CD8 on
FFPE tissues now but other human markers might be similar to the CD4 and CD8
and not stain after acetone/absolute ethanol. 

Acetone/alcohol in this ratio has been by fixative of choice for the mouse
and rat CD markers but I used it at RT, not cold.  The alcohol should be
absolute ethanol. With this in mind,  a recommendation is do a  fixation
panel with different fixatives to optimize for any given antigen. 

Air drying the fresh tissue frozen section is a must though. 

Gayle Callis 
HTL/HT/MT(ASCP)  

  

-Original Message-
From: Bernice Frederick [mailto:b-freder...@northwestern.edu] 
Sent: Thursday, March 12, 2015 11:49 AM
To: gayle.cal...@bresnan.net; histonet@lists.utsouthwestern.edu
Subject: RE: [Histonet] Acetone fixing and tissue damage

We use a 3:1 ration of cold acetone/100% ETOH to fix frozens for IHC. Slides
are always air dried first to remove any moisture.
Usually 75 mls acetone,25 mls alcohol.

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 Gayle Callis
Sent: Thursday, March 12, 2015 12:02 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Acetone fixing and tissue damage

You wrote:   

 

Hi Everyone.

 

When I fix my cryosections  in acetone,  I am using HPLC grade 99.9% for 10
minutes at -20C.

 

Would the Histology grade 99.5% be less damaging to them?

 

Higher H20 content, i.e. less than 99.5% apparently is also very bad.

 

With the HPLC grade I often get tissue damage, the tissue also floats off
the slide causing a stringy effect.

 

Fixing with 4% p-formaldehyde or 100% Methanol, prevented the antibody from
recognizing the Nuclear Antigens.

 

Looking for advice,

 

Patrick.

Patrick Lewis

Research Associate II Bench

Seattle Childrens Research Institute

206-884-1115

 


*

HPLC grade acetone is not necessary plus very expensive.  Use ACS Certified
Reagent 99.5% grade, not histology grade, which you can buy in gallon size.
Maybe what you are calling histology grade is the ACS Certified Reagent
grade but  Histology grade implies a practical grade of acetone which is
not a pure as the ACS certified Reagent grade.   Also, 4°C acetone works
just as well.If you are storing your acetone (in a staining jar) inside
the cryostat to maintain a 20°C temperature, don't!!!  If your staining
container tips over, you will ruin your cryostat!!   Hopefully you are using
high quality plus charge slides?

 

We had frozen sections come off a plus charge slide after single 4°C acetone
fixation on occasion.  You can prevent frozen section loss is a Double
Acetone fixation that also increases permeabilization. An IHC guru gave
me this hint years ago and was given to her by a company selling
immunostaining products.   A small fan will be your best friend for RT air
drying and/or evaporating acetone.  However we air dried all  FS were dried
at RT for 30 minutes minimum  or longer before fixation.  By air drying, you
get rid of the water.  You can put your FS in front of a small fan, or
inside a hood for faster drying, and never store just cut FS in the cryostat
where water condensation occurs when you take them out of cryostat
environment to RT.DRY  frozen sections were the rule in our lab before
any solvent fixation.  

 

1.   Air dry frozen section at RT for 30 min 

2.   Fix FS in  4°C acetone for 10 minutes

3.   Air dry FS for 15 minutes to evaporate acetone

4.  Return FS to 4°C acetone for 10 minutes

5.  Air dry to evaporate acetone 

6.  Proceed to immunostaining 

 

Gayle Callis

HTL/HT/MT(ASCP)

 

 

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RE: [Histonet] Re: Embedding

[Gayle Callis]

A fabulous idea!   I suspect one could use a cheap travel iron although  one 
needs to devise a way to collect melted paraffin.   Even our fancy para trimmer 
didn't have catch pan for paraffin drippings.I suggest using a new or 
receycled aluminum baking pans available in supermarkets, discount stores or a 
recycled frozen food pan without separations.  These pans come in various sizes 
and depths.   The joy is toss pans when full of paraffin.   

Gayle Callis
HTL/HT/MT(ASCP)  

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Tony Auge
Sent: Thursday, January 22, 2015 9:58 AM
To: histot...@imagesbyhopper.com
Cc: histonet@lists.utsouthwestern.edu; Morken, Timothy; Goins, Tresa
Subject: Re: [Histonet] Re: Embedding

If you want a cheaper alternative you can use a ski wax iron. They cost about 
$40. I mounted one upside down in a bucket and it works just as well as the 
$500 para trimmers.



-Tony Auge HTL (ASCP) QIHC
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[Histonet] Re: squamous cell contamination on slides

[gayle callis]

After following this thread on a topic that has always been a pet peeve and
a problem at times, a few suggestions. This is a problem discussed many
times on Histonet. 

 

If people are using the water bath as a finger bowl, they need to learn to
not let fingers go snorkeling when picking up sections.  This is sloppy,
poor technique.  Anyone can learn to never touch the water surface with bare
skin. I have seen people immerse their fingers up to the first joint and
their stained sections were covered by a snowstorm of squamous cells.

 

Be aware that squamous cells are going to exfoliate from other than hands,
so not touching face and hair is good advice even if one wears gloves.



Hand lotion is helpful except for those slather on lotion or the lotion is
particularly heavy duty and then they still touch the water. This can cause
a double problem - an oil slick which is a terrible section adhesive along
with squamous cells from bare skin. 

 

Hold slide at top or on sides, as mentioned previously.

Wear gloves. Not always popular with a common argument one loses dexterity
handling slides.  If gloves do not flop around loosely but fit the hand
well, then dexterity is not lost.  

 

Good luck

 

Gayle Callis

HTL/HT/MT(ASCP) 

 


***

 

We have that problem mostly during the winter months when our hands get dry.
Use hand lotion, that usually helps a bit.

 

Thanks,

 

Michele Margiotta-Watz

Histology Supervisor

BMHMC

101 Hospital Rd.

Patchogue, NY 11772

631-654-7192

 

-Original Message-

From:  http://lists.utsouthwestern.edu/mailman/listinfo/histonet
histonet-bounces @t lists.utsouthwestern.edu [mailto:
http://lists.utsouthwestern.edu/mailman/listinfo/histonet histonet-bounces
@t lists.utsouthwestern.edu] On Behalf Of Amber McKenzie

Sent: Tuesday, October 28, 2014 3:55 PM

To:  http://lists.utsouthwestern.edu/mailman/listinfo/histonet histonet
@t lists.utsouthwestern.edu

Subject: [Histonet] Squamous cells staining on HE and IHC

 

 

Does anyone else have problems with what looks like squamous cells staining
on your HE's and IHC's?  I'm trying to figure out how to eliminate that
problem in our lab...wear gloves while cutting?  Change out water bath
several times during shifts? Any suggestions?  Thanks!

 

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[Histonet] Re: Decalcified bone eosin overstaining

[gayle callis]

What Hazel is saying is true but with acid decalcifiers i.e. formic, nitric
or HCl.  These acids damage nucleic acids aka acid hydrolysisand if the
decalcification is not controlled by endpoint testing, then the nuclei will
appear pale and in some cases, don't stain at all (over-decalcification).
However, with EDTA is not going to damage the nucleic acids, and you should
have normal hematoxylin and eosin staining.After EDTA decalcification,
we had good HE staining.  After acid decalcification, we had over-staining
with eosin and reduced staining time in eosin to 10 continouslu moving dips,
followed by three 95% and three 100% and clearing.  

 

I think your problem is going from acid/alcohol directly into the eosin, a
no no.   This does NOT allow for correct staining of Eosin at pH 4 to 4.5.
Because you now have such an acidic environment, this is probably the cause
of eosin over staining - often an ugly brick red color. You should have
running tap water rinsing after hematoxylin for at least 1 minute, also do a
1 minute water rinse after the acid/alcohol.  You obviously answered your
own question Perhaps a tap water rinse after the Acid Alcohol? and that is
a resounding YES!!!

 

Where is your bluing step?  You are leaving your differentiated hematoxylin
in a reddish/blue state.  

 

A correct sequence for Harris Hematoxylin staining:  

 

Deparaffinize: 

 

Xylene x 3 changes

100% x 2 changes

95% x 2 changes

70% x 1 change

Distilled water rinse - 1 minute to hydrate the section

Harris Hematoxylin - up to 10 minutes

Tap water rinse - 1 minute

0.5% Acid/alcohol -  1 dip.  Do this quickly and go right into running tap
water. 

Tap water rinse - 1 minute  Water should flow start under and go up over the
slides.  

Bluing solution - 1 minute  This is a very mild base.  Buy a commercial
solution specified for Harris hematoxylin.

Tap water rinse - 1 minute

70% alcohol - 1 minute

Eosin - time desired

95% alcohol X 3 changes

100% x 3 changes

Clearing

 

You will see the sections turn blue after a bluing solution which requires a
tap water rinse and also a 70% alcohol rinse make sure there are no bluing
solution cations carried over into the eosin.   Optimal pH for eosin
staining is 4 to 4.5 (which you have altered by going from acid/alcohol
directly into eosin).A 70% alcohol also equilibrates to the alcohol
percentage of the eosin solution.   If the eosin still is over staining,
reduce time to 30 sec.  If dipping, be sure you are move the slides up and
down continuously. After the eosin you should be able to just rinse in 95%
alcohols to differentiate out excess eosin.  If any of the post-eosin
alcohol rinses have too much eosin carryover, you will continue to stain the
bone.   Make sure alcohol rinses after eosin are fresh or at least rotated
so there is a fresh last 95% and fresh last 100%.   Any pink color in the
last 100% means you have water carryover which can cause 1) cloudiness 2)
eosin bleeding after cover slipping.

 

On the side and many years ago,  we used Harris Hematoxylin for acid
decalcified bone followed by running tap water rinse but eliminated the
acid/alcohol step entirely to avoid even weaker nuclear staining already
caused by acid hydrolysis from the acid decalcifier.  A bluing solution was
used followed by a water rinse before eosin staining sequence.  I learned
this from the AFIP bone lab and it worked with excellent results.   However,
you are working with EDTA decalcified bone and when you eliminated the
acid/alcohol, it worked great without eosin over staining.

 

Good Luck

 

Gayle M. Callis

HTL/HT/MT(ASCP)

 

 

 

You wrote:  

 

When the bones are over decaled there will be very little hematoxylin
staining and too much eosin.  You can use a buffer before the hematoxylin to
aid the hematoxylin in staining.  I think one/two dip(s) in eosin will be
sufficient.

 

Hazel Horn

Supervisor of Histology/Autopsy/Transcription

Anatomic Pathology

Arkansas Children's Hospital

1 Children's Way | Slot 820| Little Rock, AR 72202

501.364.4240 direct | 501.364.1241 fax

 http://lists.utsouthwestern.edu/mailman/listinfo/histonet hornhv @t
archildrens.org

archildrens.org

 

Sent: Thursday, October 02, 2014 1:11 PM

To:  http://lists.utsouthwestern.edu/mailman/listinfo/histonet histonet
@t lists.utsouthwestern.edu

Subject: [Histonet] Decalcified Bone Eosin Overstaining

 

Hey histonetters! I'm currently doing some HE Staining for EDTA Decalcified
Bone Tissue and it seems that the tissue is staining very intensely with
Eosin...perhaps too much. Was curious to see from some of the more
experienced histotechnicians out there which methods you guys have used in
the past to perhaps fix an issue like this. Here is my protocol that I have
used once I have sectioned the paraffin embedded tissue. I have omitted the
1 second Acid Alcohol Dip in a few of my trials and it seemed that this
method worked great to keep the tissue from overstaining with Eosin.

[Histonet] At long last, a Murine CD4 monoclonal antibody that works on FFPE tissues

[gayle callis]

Dear Histonetters, 

 

This antibody is going to be very welcome for those working with NBF fixed
paraffin embedded murine tissue.It is a rat antiMouse CD4  for FFPE
tissue.   For those interested go to Affymetrix/eBioscience, Rat antiMouse
CD4, clone 4SM95 (L3T4/Ly-4,  Catalog number 14-9766.   It also comes
conjugated to the company's eFluor 570, equivalent to Alexa 555 although one
could do their own conjugation to their favorite Alexa or DyLight dye.   

 

Now all we need is a mouse CD8 that works on FFPE for total happiness since
we often do these two antibodies in tandem as a double IF stain or on
adjacent sections for enzyme IHC.

 

Take care

 

Gayle M. Callis

HTL/HT/MT(ASCP)  

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[Histonet] Re: Processing whole rat heads

[gayle callis]

 in holder
evenly.If you try to orient the block so it is on vertical axis rather
than horizontal,  your sectioning will be more difficult., fewer sections in
ribbon for flotation.The top of skull is rounded so that should take
care of trying to embed the skull at an angle, or you can try more of an
angle if it works for you.The old fashioned way is to lay each ribbon on
a flat black construction paper, cut the section from ribbon, float and pick
up each section so as to NOT lose a serial section.  This way you can have
many ribbons laying out in sequence before floatation.  Tedious but
accurate.

 

A regular rotary style  paraffin microtome should work without problems.  5
to 7 um should be adequate.   Overly thick sections will curl up, making
them hard to flatten on water bath.  To section, you should mount the block
in holder so the edge of the blade passes through top of the skull first and
those tough, harder teeth last.  Teeth and inner ear bones will be the
hardest thing to section.   I strongly suggest using high profile disposable
blades for stability as these are just as sharp as low profile.   Doing
serial sections will not be a simple task but certainly possible, requires
practice and patience.   Mount sections on plus charge slides, drain water
off well and dry sections FLAT at 37°C to 40°C for several days.  Do NOT dry
in a hot oven.  Bones, teeth and brain should flatten and adhere well
without excessive drying heat.  Make sure the blade is always sharp, changed
frequently, and soaking not excessive.  You may want to soak the block face
with a gauze wet with cold water with icing the wet gauze frequently while
the block mounted remains in the block holder so you can pick up all
sections.  After a each soak, you will have to back up the block a tiny bit
in order to get the next section in the series.

 

For staining, avoid overly vigorous rinsing as the softer brain can still
dislodge from bone and/or fold over.

 

Good luck and send photos to histonet when you have results.  

 

Gayle Callis

HTL/HT/MT(ASCP)   

   

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[Histonet] Re: antibody suggestions CD8, FAP, Cd11b, CD11c

[gayle callis]

You Wrote:

 

I need your help to find good antibodies for IHC on mouse and human FFPE

tissues like normal tissues and  tumors. Please share any information you

think might be helpful.

 

CD8 for IHC on mouse FFPE spleen or other lymphoid organs.

FAP (Fibroblast Activated protein) for IHC on mouse and human FFPE tumors.

CD11b  for IHC on mouse FFPE spleen  or other lymphoid organs..

CD11c  for IHC on mouse FFPE spleen  or other lymphoid organs.

 

Thanks in advance for all your help.

 

Regards,

Mesru


___

 

I can answer some of the murine CD marker questions.  

 

CD8 on FFPE mouse tissues will never work. These markers are too strongly
cross linked by the aldehyde based fixation, and PFA is not an alternative.
You can try HIER and Enzyme digestions until you are blue in the face, and
the staining will NOT work. This will also be true of CD11c dendritic cell
Clone HL3, and CD11b Mac1.  

 

This means you will have to use fresh tissue frozen sections, acetone fixed
for any staining success.  We use an acetone/absolute ethanol mixture for
fixation of murine tissue for all our CD marker staining, especially when
you need to do all these CD markers on the same sample.   

 

As for positive tissue controls, 

 

CD8 Rat antiMouse monoclonal - normal spleen 

CD11c - Peyers patches from small intestine, lymph node and probably spleen
will work.

CD11b, Mac1 - we stimulated mouse to produce macrophages in Peyers patches
located on small intestine. Stimulation was oral inoculation with an
attenuated bacteria.  

FAP (Fibroblast Activated protein) for IHC on mouse and human FFPE tumors.
Go to ABCAM, and look at cross reactivity for this polyclonal antibody.  It
works on FFPE mouse, human and two other species.  

 

 

I can't provide an answer for FFPE human tissues with any of these CD
markers.  

 

I strongly suggest you go to BD Bioscience and do a search for the CD
markers, especially the mouse that will be either rat and Armenian Hamster
monoclonals.   Read the Technical data sheets and see what applications will
work or not in terms of IHC. We have used BD Bioscience, and eBioscience for
our murine CD marker with great success but you have to be aware that CD8
has more than one clone, some working better on solvent fixed murine tissues
than others.   This is also true of CD11c.  ABCAM has an excellent website
for these antibodies, but compare prices.   

 

There is another fixative that is formalin free, the Becksteads IHC zinc
fixative and also PLP i.e. paraformaldehyde-lysine-Periodate.  PLP may allow
staining of these aldehyde compromised CD markers since the lysine helps get
rid of free aldehydes, along with the periodate and is, however, a fixative
that is bit tedious to use since it has to be made up fresh every time you
use it. Beckstead originally developed IHC zinc fixative in order to do IHC
on human tumors, lymphomas if I recall.I have original and other
publications for both fixatives showing success with murine CD markers and
other species.   The IHC zinc fixative allows paraffin embedding and IHC
staining of these markers, (also a publication by Nitta et al)

 

Our lab used fresh tissue frozen sections, acetone/alcohol fixed exclusively
for all CD marker IHC and/or single, double and triple immunofluroescence
protocols. 

 

I will be happy to provide acetone/alcohol fixative method and alternative
acetone fixations that work with murine CD markers.  You cannot use
acetone/alcohol for human CD8 or CD4 as the alcohol ruins the antigen -
however, FFPE probably will work with human markers. I can also provide
publications, information about PLP, Becksteads IHC zinc fixative. PLP
recipes can be found on the internet but I have one that is excellent for
larger volumes of this fixative.  Biocare has information on this fixative
with their IHC products too.

 

Gayle M. Callis

HTL/HT/MT(ASCP) 

 

 

 

 

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RE: [Histonet] RE: paraffin

[gayle callis]

I want  these in doorways of my house.  We used tacky mats at entrance of
designated room for prion tissue work i.e. paraffin microtomy and
cryomicrotomy, etc.  Fabulous way to keep the floors clean.

We also had industrial rugs i.e. the kind seen at entrances to large
buildings in  lab where routine paraffin microtomy was done.  These were
vacuumed daily by the janitorial staff and could be shampooed if needed.
These rugs  were put in after a  visiting professor slipped on paraffin
slick,  painted concrete floor. He would have bashed back of his head
but managed to catch himself on way down, clutching a door knob.   I think
the rugs avoid potential injuries and lawsuits against our university.   

Go for the sticky mats!!!   I liked how Esther used them in all areas, and
certainly better than rugs which are eventually trashed and unsightly with
ground in paraffin residue.   I suggest in front of the processors too. 

Gayle Callis
HTL/HT/MT(ASCP) 




-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Esther C
Peters
Sent: Wednesday, August 13, 2014 7:18 PM
To: Walter Benton; Algeo, Lacie A; histonet@lists.utsouthwestern.edu
Subject: [Histonet] RE: paraffin

The nice thing about those is that sticky mats will also collect the dust
from shoes; I found that these stop a lot of dust coming in at the outside
door entrances, as well as having them to collect paraffin under the
embedding center, in front of the tissue processor and fume hoods, and under
the microtomes. It is amazing how much dust and dirt can come in with
people's  shoes!

Esther C. Peters, Ph.D.
Associate Professor
Environmental Science  Policy
George Mason University
4400 University Drive, MS 5F2
Fairfax, VA 22030-
Office: David King Hall, Room 3050
Phone: 703-993-3462
Fax: 703-993-1066
e-mail: epete...@gmu.edu
https://bluprd0511.outlook.com/owa/redir.aspx?C=ET8XhF-xC0ytBErXdaN3U3lGqWmZ
NdAI_N-4nsEb0IjgUpeIoQa7EcVMJMh2oePPPKrrDjhwOvk.URL=http%3a%2f%2fesp.gmu.ed
u



From: histonet-boun...@lists.utsouthwestern.edu
histonet-boun...@lists.utsouthwestern.edu on behalf of Walter Benton
wben...@cua.md
Sent: Wednesday, August 13, 2014 9:05 PM
To: Algeo, Lacie A; histonet@lists.utsouthwestern.edu
Subject: [Histonet] RE: paraffin

You may want to purchase the large tack mats for staff to walk over when
going in and out of your embedding area. These come in blue, white, or gray:
http://www.fishersci.com/ecomm/servlet/itemdetail?storeId=10652langId=-1ca
talogId=29104productId=5264212distype=0highlightProductsItemsFlag=YfromS
earch=1searchType=PRODhasPromo=0



Walter Benton HT(ASCP)QIHC
Histology Supervisor
Chesapeake Urology Associates
806 Landmark Drive, Suite 127
Glen Burnie, MD 21061
443-471-5850 (Direct)
410-768-5961 (Lab)
410-768-5965 (Fax)
Chesapeakeurology.com

Voted a Best Place to Work by
Baltimore and Modern Healthcare
Magazines.

From: histonet-boun...@lists.utsouthwestern.edu
[histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Algeo, Lacie A
[lacie.al...@providence.org]
Sent: Wednesday, August 13, 2014 6:24 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] paraffin

Hi all,
I am dealing with an issue with slippery floors outside of our Histology
department.  The concern is that it is from paraffin scraps tracked on the
bottoms of peoples shoes.  The floor is cleaned every night and an anti-slip
agent applied.  I have tested whether our paraffin makes the floors more
slippery and it actually makes it harder to slipalmost tacky.  I really
do not think it is our paraffin.  I think it is a thin layer of dust that
settles on the floors after they are cleaned.  I have seen this happen in
other places.  Is anyone dealing with this?  Any insight?
Thanks,
Lacie

Lacie Algeo, HTL (ASCP) MBCM
Histology Supervisor
Providence Sacred Heart Medical Center Laboratory
101 W 8th Avenue
L-2
Spokane, WA 99204
509-474-4418
FAX 509-474-2052
lacie.al...@providence.orgmailto:lacie.al...@providence.org


This message is intended for the sole use of the addressee, and may contain
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that you may not use, copy, disclose or distribute to anyone the message or
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in error, please immediately advise the sender by reply e-mail and delete
this message.




This message is intended for the sole use of the addressee, and may contain
information that is privileged, confidential and exempt from disclosure
under applicable law. If you are not the addressee you are hereby notified
that you may not use, copy, disclose, or distribute to anyone the message or
any information contained in the message. If you have received

RE: [Histonet] Weight Loss/Weight Gain Decal _Histonet Digest, Vol 129, Issue 18

[gayle callis]

 the lighter side... (Mayer,Toysha N)
  12. Re: RE:On the lighter side... (Barry Rittman)
  13. Re: Weight Loss/Weight Gain decalcification endpoint test
  (gayle callis)
  14. Consult for taking abs from RUO to IVD (Patsy Ruegg)
  15. RE: On the lighter side... (susan.wal...@hcahealthcare.com)
  16. RE: Cytospin validation (Jamal)
  17. Re: On the lighter side... (Michael Ann Jones)
  18. looking for TS replacement in Histology due to retirement
  (Webb, Dorothy L)


--

Message: 1
Date: Mon, 11 Aug 2014 13:03:38 -0400
From: Shirley A. Powell powell...@mercer.edu
Subject: RE: [Histonet] On the lighter side...
To: Ingles Claire cing...@uwhealth.org,
histonet@lists.utsouthwestern.edu
histonet@lists.utsouthwestern.edu
Message-ID:
 
9BF995BC0E47744E9673A41486E24EE25BFB25FD10@MERCERMAIL.MercerU.local
Content-Type: text/plain; charset=iso-8859-1

I agree. :)

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Ingles
Claire
Sent: Monday, August 11, 2014 12:00 PM
To: histonet@lists.utsouthwestern.edu
Subject: RE: [Histonet] On the lighter side...


Old histologists never die, they are just well fixed...
Claire

From: histonet-boun...@lists.utsouthwestern.edu
[histonet-boun...@lists.utsouthwestern.edu] on behalf of Michael Ann Jones
[mjo...@metropath.com]
Sent: Monday, August 11, 2014 9:07 AM
To: Edwards, Richard; histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] On the lighter side...

25 years, (what?s that in micron?s??) Bernice, you are too funny!!
(lots of tenure here . . .lotsa brain cells) Michael Ann Jones, HT (ASCP)
Histology Manager Metropath
7444 W. Alaska Dr. #250
Lakewood, CO 80226
303.634.2511
mjo...@metropath.com




On 8/11/14, 5:17 AM, Edwards, Richard r...@leicester.ac.uk wrote:

Sniffed my  first formalin and  saw  first post-mortem November 1965.

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--

Message: 2
Date: Mon, 11 Aug 2014 13:18:57 -0400
From: Heather Knight heather.l.knig...@gmail.com
Subject: [Histonet] Bcl-2
To: histonet@lists.utsouthwestern.edu
Message-ID:
CABEU4_xVVjqyPu9o8=ONJA__mLNOXAcRmt6pK2oLcSAKp=n...@mail.gmail.com
Content-Type: text/plain; charset=UTF-8

Hi everyone-

Just wondering if anyone has a working protocol for Bcl-2 in FFPE mouse
tissue?  If so, can you share both the protocol and the antibody
information?

We have tried numerous antibodies over the years with very limited success.
 Thank you for your help!!

Best,
Heather Knight


--

Message: 3
Date: Mon, 11 Aug 2014 10:18:53 -0700
From: Jb craiga...@gmail.com
Subject: [Histonet] IHC Steam Strips:
To: Histonet@lists.utsouthwestern.edu
Message-ID: 375f4ef9-47b6-46cd-b5b1-b511309ef...@gmail.com
Content-Type: text/plain;   charset=us-ascii

Does anyone use IHC steam strips in their decloaking chamber?  The question
is do you run a new steam strip each run and log each individual run w/it's
own steam strip?

Thank you,

Sent from my iPhone


--

Message: 4
Date: Mon, 11 Aug 2014 17:43:34 +
From: Michael LaFriniere michael.lafrini...@ccplab.com
Subject: RE: [Histonet] Are Paraffin Blocks Biohazard
To: Dawn Bugge drbu...@gmail.com,
histonet@lists.utsouthwestern.edu
histonet@lists.utsouthwestern.edu
Message-ID:
4a2a16b9707ce04e9cb6c82dc18c1d296a0...@ahcmsasexch00.my.ahc.local
Content-Type: text/plain; charset=us-ascii

Dawn,

The only study I know of is on CJD crutsfeldt-Jacobs Disease (know to
survive formalin fixation and routine processing protocols, the CDC web site
has additional information, In my laboratories I put all blocks in hazardous
waste for incineration disposal.  It is not that costly just to be on the
safe side.



Michael R. LaFriniere, HT (ASCP)
Executive Director


Capital Choice Pathology Laboratory
12041 Bournefield Way, Suite A * Silver Spring, MD 20904
P: 240.471.3427 * F: 240.471.3401 * Cell 410-940-8844
michael.lafrini...@ccplab.com


-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Dawn Bugge
Sent: Friday, August 08, 2014 2:41 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Are Paraffin Blocks Biohazard

Hello Histo World!

Our pathologist for our private GI lab would like me to find out if anyone
has done a study to determine if the paraffin blocks, once they have been
processed, are considered biohazard.  I have searched high and low and can
find many people stating that the blocks are not bioharzard, with the
exception of neurological tissue, but they don't state how they know this.
He would like me to reference an actual study to prove

[Histonet] Liquid cover glass for HE stained cell culture in 6 well plates; toluidine blue stain for cell cultures

[gayle callis]

What are people using as a liquid cover glass for stained cells in 6 well
culture plates to be view with inverted microscope with having that annoying
refractile, funky look to the cells.  

 

Cells are grown in 6 well culture plates and with previous cultures, been
stained with Hema 3 (hematological stain) and Giemsa of which neither has
been successful to see cell membranes and nuclei clearly.   Viewing is done
with inverted microscope and cells are counted.   

 

I know there are some stains that can be viewed left under water with
inverted scope.  HE obviously cannot be left under water due to eosin
washing out of cytoplasm.   

 

A possibility is  letting HE or T blue stained cells dry, then put a thin
layer of permanent mounting media to flow over the cells to fill in spaces,
let dry and view with inverted microscope.  I think one could do an old
bonehead trick we used on surface stained, thicker ground MMA embedded
undecalcified bone, flood with immersion oil and put a cover glass on top.
Very messy!!!   However one could let HE dry after last alcohol rinse,  put
a layer of immersion oil over the cells, and view with inverted microscope.
Anything to fill in the cellular spaces on a confluent sheet of cells but
let the oil be the cover glass.  Growing cells on a cover glass or on slide
with wells/gaskets is not an option.   

 

Any suggested stains with protocols would be appreciated too with thoughts
here about using Toluidine blue.  The recipe for making up the stain,
staining method and preferred fixative would be appreciated.   You can send
information privately if you wish.  

 

A challenge to be sure

 

Thanks 

 

Gayle Callis

HTL/HT/MT(ASCP)

 

   

 

 

 

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[Histonet] Re: Weight Loss/Weight Gain decalcification endpoint test

[gayle callis]

Trevor and Jennifer, 

 

This method originally came from Mawhinney et al.  Control of rapid nitric
acid decalcification J Clin Pathol 1984 37:1409-1415 and was cited by Cathy
Sanderson (Mayton)in a publication using EDTA, found in Biotechnic and
Histochemistry.  Mawhinney acutally did a chemical test on final acid change
to see that calcium was not present, but we never had to do that.   If you
end up with a bit of residual calcium in block, I would surface decalcify at
microtomy.   I used it for years when we downsized and gave away our
FAXITRON.   Radiography is still the most accurate, and if you had either a
micro CT or digital FAXITRON available, it would be a better test. I
used Cathy's method and will be happy to send the original publication for
anyone's files/future referencing.   A chemical test for EDTA is a pain to
do if a FAXITRON is not available.   

 

I have put the full method below, with a bit more detail.   

 

WEIGHT LOSS/WEIGHT GAIN ENDPOINT TEST

 

This is a method of choice for EDTA decalcification although it was
originally used by Mawhinney et al for testing nitric acid decalcification.

Many samples can be decalcified together in one container, i.e. 25 mouse
femurs in 1 liter of 10% formic acid.   If all the samples are the same
size, i.e. mouse femurs, tibia, paws, choose several as representative
samples and test only those to save time.   Always suspend bones in the
decalcifier. 

Requires a balance that reads in milligrams to 3 places for greatest
accuracy.  Specimen must be blotted free of fluid for accurate weighing each
time you weigh the sample.  We suspend bones in nylon specimen bags for easy
removal to weigh.  Bags can be marked with pencil too. 

Technique:

 

1.  Rinse NBF off bone, blot with paper towel, WEIGH BONE, RECORD BEGINNING
WEIGHT.  Suspend bone in acid or EDTA decalcifier.  During acid
decalcification CO2 bubbles are given off, so stir during decalcification to
release bubbles or small samples will float.   EDTA does not create CO2
bubbles, only acids.  Large bones can be started at end of day in acid
decalcifier and sit overnight with testing the next morning.   

 

2. After 4 to 5 hours in acid or overnight in EDTA, remove bone, rinse with
water, BLOT, weigh.  RECORD WEIGHT.  If bone shows loss of weight, change
acid decalcifier to refresh acid.  Return samples to resume decalcification,
and repeat as many times as necessary.  EDTA should be changed but not as
often as acid.   Always use a large volume of decalcifier i.e. 20:1 or more.
Remove bone from specimen bag, and place in weighing boat to protect balance
from acids/EDTA. 

 

3.  When bone begins to GAIN WEIGHT, the bone is decalcified.  Once calcium
is removed, water is taken on and the weight increases.  This water does not
affect the bone.  

 

4.  Rinse bone with running tap water for an hour or longer to remove these
decalcifiers. Either store in 70% alcohol or process.Store endpoint
tested decalcified bones in 70% alcohol while waiting for other samples to
finish decalcifying and mass processing run.  

 

Reminders:  For EDTA, one can suspend bones and check every day for accuracy
but bones can be left in the EDTA over a weekend or several days without
damage as long as the bones were well fixed. Acid decalcified bones cannot
be left over a weekend, remove from acid, put in NBF to stop
decalcification. Bones should be endpoint tested before stopping
decalcification so you can resume decalcification on the next working day.
Rinse off NBF briefly before resuming decalcification.   Do not overexpose
bones to acids or you will damage antigens and nuclear staining.   

 

Enjoy the method, as it truly is fast and easy.

 

Gayle M. Callis

HTL/HT/MT(ASCP) 

 

 

 

 

 


_

Ha Wow...that's almost too easy. Thank you for this!

 

 

Trevor Jordan Wait

University of Texas Health Science Center, San Antonio

Class of 2017 MD Candidate

Abilene Christian University Class of 2013 Graduate

B.S.  Biochemistry



From: Jennifer MacDonald 
http://lists.utsouthwestern.edu/mailman/listinfo/histonet JMacDonald @t
mtsac.edu

Sent: Monday, August 11, 2014 2:33 PM

To: Wait, Trevor Jordan

Cc:  http://lists.utsouthwestern.edu/mailman/listinfo/histonet histonet
@t lists.utsouthwestern.edu;
http://lists.utsouthwestern.edu/mailman/listinfo/histonet histonet-bounces
@t lists.utsouthwestern.edu

Subject: Re: [Histonet] Weight Loss/Weight Gain Decal

 

I believe this was originally from Patsy Ruegg

 

Decalcification End Point: Weight Loss, Weight Gain

 

 

1.Blot sample to remove excess fixative

2.Weigh bone in mg, record as beginning weight

3.Next day, rinse bone, blot and weigh bone daily, record weight.
Change decalcifying solution to refresh acid OR EDTA. Return bone to fresh
decalcifying solution.

4.When bone begins to GAIN weight, remove 

RE: [Histonet] Gayle Callis, You Made Our Monday Morning!!

[gayle callis]

You have me laughing at myself!!!  I'll  blame the computer for the typo
error and  spend the rest of my day laughing at myself.This isn't the
first time this has happened in one of my posts.  Whatever you do, do NOT
make the mistake of misspelling fuchsin or your good friends will come
back and haunt you with that f-bomb  mistake.   I laughed even harder then
but am more careful when typing out that word. 

 Busy?   I don't think I was referring to any feminine anatomical features
we all know so well.   I am quite sure some histo guys are laughing too.   

Still chuckling and glad I provided some unintentional  amusement in that
lengthy post.   
 
Gayle Callis ;)

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Jones, Laura
Sent: Monday, August 04, 2014 7:51 AM
To: Histonet@lists.utsouthwestern.edu
Subject: [Histonet] Gayle Callis, You Made Our Monday Morning!!

From Gayle's very knowledgable post about storage of Schiffs:

However, if I had to have immediate PAS staining in a busty lab, 

We would like a definition of busty, please!!  Thank you for the laugh!


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[Histonet] Re: Schiff's storage temperature - long reply

[gayle callis]

Dear Histonetters, 

 

At the risk this may be more information than you need.   My fascination
with PAS and Schiffs reagents prompted the following.  

 

Our Schiffs reagent,  prior to available commercial products,  was made in
house.  We made Lillies Cold Schiffs solution stored at 4C according to
protocol but no time limit was given.Another in house Barger and
DeLamater reagent said several months at 4C - a rather vague time.  Luna
said 3 months storage at 4C. I think the kicker here is to test any
Schiffs before use since it is unstable over time to remove any doubt.
Other differences is  basic fuchsin versus pararosaniline dyes and there are
publications discussing these dyes for Schiffs reagents.   Confusing - yes!!


 

Back in the dark ages when practical exams were still done for HTL,
curiosity led to testing both in house and a commercial Schiffs reagent.
All staining was done at RT in parallel fashion using fresh in house and
commercial Schiffs on adjacent sections/separate slides.   We noted no
visible staining difference i.e. intensity.This led exclusive use of
commercial Schiffs to avoid last minute, tedious and exposure to
carcinogenic basic fuchsin i.e. the weighing out dry,  and strong HCl fumes.
We didn't always have good fume hoods back in the dark ages of histology.
Old habits die hard to store Schiffs at 4C, so RT storage was ignored, and
4C prevailed with commercial Schiffs with staining at RT. 

 

However, if I had to have immediate PAS staining in a busty lab, I would
store Schiffs at RT to avoid waiting for equilibration to RT.   This brings
up a question.  Has the person asking at temperature storage tried staining
with cold versus warm as the staining times I saw i.e. up to 30 minutes
could very well make up the difference.  Cold Schiffs longer time versus
warm Schiffs shorter time?   1% versus 0.5% periodic acid will also make a
difference.   

 

The best discussion of Periodic acid Schiffs reaction staining mechanisms
and technical comments can be found in Sheehan and Hrapchak Theory and
Practice of Histotechnology, Carbohydrate Chapter 9, p9 164 166.  Second
Edition, 1980.   This book is a classic and still available in Paperback
form from NSH. This gives excellent hints on performing the stain i.e.
1) always used freshly prepared periodic acid (PA) something Culling
insisted on.  PA is cheap and goes into solution immediately.   I do not use
premade periodic acid solutions in kits as I don't consider these freshly
made, simply unused but made days, weeks, months before purchasing the kit.
If they are used, should be discarded and go to making PA fresh daily.
Another finding was a decision to preweigh  dry aliquouts of  PA into a
clear glass Erlenmeyer flasks to save time, seal the top of flask tightly,
store on shelf until distilled water was added.  This did NOT work as  the
preweighed PA went bad due to exposure to air, and probably light.  PAS
staining was poor, renal pathologist complained, and was embarrassing.
Freshly weighed, solution prep daily was reinstated.  

 

Some things we did to maintain good PAS staining. 

 

1.  Since shelf life varies with use and probably storage conditions,
expiration date is important but in house Schiffs could go bad before that.
This is probably true of commercial solutions too.   

1.  If Schiffs reagent turns pink, discard.  

3.  Never freeze Schiffs reagent.   Shipment to cold weather states i.e.
Montana in winter was avoided after Schiffs arrived frozen.   Plan ahead.   

4.  Never return used Schiffs to stock bottle.   Store in another bottle,
dated and tightly sealed.  Expendable plastic wrap makes a good tight seal
around and under lids on bottles or Coplin jars with used Schiffs. 

6. Test used and even new if the latter hasn't been used for a time after
opening stock bottle.  Sheehan and Hrapchak have test in chapter.  We just
used a few drops of Schiffs in NBF and watch the bad color...deep
blue-purple.   You want instant bright red with slight tinge of purple. If
the lab is formalin free, use histologic test by staining cross section of
human appendix, look for good staining of fine meshwork sarcolemma between
smooth muscle cells. 

7.  Reuse Schiffs for up to a week, unless it turns pink.  

8.  Do not dump Schiffs down the drain, discard using appropriate chemical
safety facility.

9.  Don't get Schiffs on your skin...nitrile gloves.   Even though some
vendors may have Schiffs removal reagents, once on the skin, you are already
exposed to basic fuchsin - the contamination/exposure horse is already out
of the barn. 

10.  Sulfurous acid rinses were discontinued after hearing Culling (in
person) lecture on PAS staining.  He said a 10 minute running tap water
rinse was sufficient to intensify the color and we never had a problem using
only water rinse.  

 

We have used Sigma's Schiffs, and also Fisher reagent grade Schiffs with
great success.   I think commercial solutions 

[Histonet] Request for copy of old Technicon operation manual

[gayle callis]

Dear All, 

 

A friend is donating her old Technicon to be used in another country, and
needs a copy of the operation manual.   If anyone still has this, could they
send a scanned pdf copy to me asap.I will forward it to person in need

 

Thank you in advance

 

Gayle Callis

HTL/HT/MT(ASCP) 

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[Histonet] Re: VIP 1000 charcoal filters

[gayle callis]

Years ago, we went to a pet store (Petco, Petsmart) or pet section in
Walmart, etc. for bags of bulk charcoal used in fish tank filters. I am sure
Amazon will have this too.   The charcoal pieces were very similar to what
was in prefilled VIP filters.   We emptied the  VIP charcoal containers
under a hood to avoid xylene fumes captured by charcoal then refilled with
the fish tank filter charcoal.  If you try this, note the level of charcoal
in the boxes for quantity needed.   It was actually cheaper than buying
prefilled boxes from the vendor's, and no shipping costs either.
Unfortunately, Sakura discontinued charcoal milk carton  packaging,
probably to avoid the fume problem during filter change - a safety factor.  

This is somewhat of a stop gap measure but certainly worked for us.  The
more recent charcoal boxes are cardboard and sealed, if I remember
correctly, but careful opening and then resealing with duct tape might work.
However, first VIP charcoal filter containers were made of the same plastic
as the solvent containers and not easily damaged during a change.  

 

Good luck

 

Gayle Callis

HTL/HT/MT(ASCP)

  

 

 

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RE: [Histonet] Staining FFPE with biotinylated SNA - how to block?

[gayle callis]

Dear Merissa, 

Ray is correct about using serums with lectins.   In fact you really are NOT 
doing immunostaining unless you are using an antibody made to recognize that 
lectin e.g. anti-lectin therefore a normal serum block is not needed.Serums 
in fact contain glycoproteins that bind to lectins, so BSA is a better protein 
carrier for buffers and diluents if you want to make up your own.  We used 
Jacksons immunoglobulin protease free BSA.   Ray gave good advice on blocking 
techniques.   However, if you do anti-Lectin, you would be doing IHC and then 
blocking is legal.   If you want a protocol for lectin IHC,  I will be happy to 
forward via private email.   With some lectins for direct staining,  you should 
use the Lectin buffer that has no phosphate ions, as explained by Vector 
technical services.   I also have a recipe for the Lectin buffer if you want 
it.   Avidin/biotin blocking is still needed with biotinylated lectin,  
particularly is the tissue is known to have endogenous biotin.  Be sure to find 
out if your lectin is sensitive to phosphate ions, or use TBS or the Lectin 
buffer.  
 
Also, if you are doing a non-IHC direct lectin-biotin staining, you must do the 
correct negative control.  For SNA, this is an elution with 500 mM lactose in 
buffered saline followed by 500 mM lactose in acetic acid to finish elution.
Buffer alone is NOT a negative control.   For the lectins we worked with, we 
diluted the lectin (working concentration) in the recommended mM inhibition 
sugar and let it sit in the refrigerator overnight, warmed to RT just before 
use as negative control.   This allows the lectin to bind to its specific 
sugar, and not to glycoproteins in the tissue, but keeps the biotin, and in our 
case, fluorophore in the negative control. 

There is an excellent, inexpensive book, Lectin Histochemistry, a Concise 
Practical Handbook by SA Brooks, AJC Leathem and U Schumacher that tells all 
about using many lectins, protocols for lectin IHC and lectin direct binding 
staining. An interesting side history is the founder of Vector is a lectin 
expert.

For those doing IHC with anti-lectin,  antigen retrieval may be needed per 
James Watson reply and is included in the Lectin IHC protocol I have.I am 
presuming he was doing true IHC for his lectin work.  

Gayle Callis
HTL/HT/MT(ASCP)

 



-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of 
koelli...@comcast.net
Sent: Wednesday, April 16, 2014 10:53 AM
To: M.O.
Cc: histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] Staining FFPE with biotinylated SNA - how to block?



Hi Merissa, 

don't know if you got any private idea responses so I'll throw in my opinion.  
I would always worry about some of the things you are mentioning and that are 
standard thoughts regarding biotin block, retrieval, etc in IHC. 

But I would think about your serum, which I steadfastly avoided with SNA or any 
lectin I used.   Lectins look at glyco components and serum (or serum 
substitutes) can be full of glycoproteins and the target then is the blocking 
serum for your lectin which can cause bad background.  I did and would use 
washes, diluents, etc that had NO serum or milk or anything like that in them.  
You can make your own, completely free of potentially having glycoproteins or 
Vector sells some.  For some lectins (look at a list of target sugars) you 
maybe can get by with serum or milk and such to block but many I've found you 
just can't. 


Ray (still in, whoever would have guessed, once again rainy Seattle) 

- Original Message -

From: M.O. modz9...@gmail.com
To: histonet@lists.utsouthwestern.edu
Sent: Tuesday, April 15, 2014 5:59:51 PM
Subject: [Histonet] Staining FFPE with biotinylated SNA - how to block? 

Hello Histonet! 

I ran a trial on FFPE mouse samples with a biotinylated lectin, SNA from 
vector.  The SNA is Biotinylated Sambucus Nigra Lectin (Elderberry).  I have 
never stained with anything like this, so I ran a test. 

I deparaffinized, blocked with NGS, incubated overnight at 4C with the diluted 
biotinylated SNA.  On the second day, I used Vector's ABC kit and alkaline 
phosphatase (red) kit. 

Once stained, I noticed a lot of background.  After looking into the blocking 
step, would a biotin/avidin blocking step be the correct step instead of a 
serum because I don't have a secondary?  How do I know what needs to be blocked 
- biotin, avidin or both?  Is there a way to do this without a kit and use 
solutions I may have in my lab? 

Thank you for your help! 

Sincerely,
Merissa
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RE: [Histonet] Treatment of calcified human arteries

[gayle callis]

Dear Galina, 

 Formic acid, buffered that contain either sodium citrate or sodium formate
that have an approximate 4.5% formic acid concentration (yes, I did the
calculation one time) and as made from 90% stock formic acid.   You can
buy these decalcifying solutions from a vendor, but look at the MSDS so you
know what is in the solution.  These are both well known e.g. classic
formic acid solutions.If you are into making up your own solutions, what
Liz Chliplala suggested is excellent although not buffered.Make sure
your arteries are totally fixed in order to protect the antigens from
effects of acid decalcification.  

I replied yesterday about Massons Trichrome and have a protocol with a
modified, more concentrated Weigerts Iron Hematoxylin that proved to be far
superior for decalcified tissues/bones.   The nuclei were never
differentiated out to the point of weak staining.I will be happy to send
if you want it.  

Good luck

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

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Galina
Deyneko
Sent: Friday, February 14, 2014 9:06 AM
To: histology; histonet@lists.utsouthwestern.edu
Subject: [Histonet] Treatment of calcified human arteries

Dear Colleagues
 Please share your expertise on treating heavily calcified human arteries.
It is impossible to cut them without deminiralization in my opinion. Should
I use decalcification solution before processing? If yes, what reagent do
you recommend taking in account that I will do IHC for macrophages and other
targets on these samples, as well as Sirius Red and Masson Trichrome
staining.
 Thank you in advance

Galina Deyneko
Novartis, Cambridge, MA
 
617-871-7613 w
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RE: [Histonet] Mouse Joints

[gayle callis]

Dear Becky, 

You didn't say what decalcifier you were using? It is a good idea to
test the decalcification endpoint on these bones before processing .   I
have a very easy weight loss/weight gain method to make sure the calcium is
removed from these joints or any bone for that matter.We used this for
murine knees, all long bones and paws and never had problems with residual
calcium.   

Two things: 

1.  Now that the bones are embedded and contain calcium, you can surface
decalcify by immersing the trimmed block face into any acid decalcifying
solution, particularly one from a vendor,  for a few minutes.  This will
remove the calcium from only a few um of the sample, so be careful to NOT
trim this minimally decalcified portion away when you resection the block.
Be sure to rinse the acid off the block or metal parts on the microtome will
corrode.  You can re-cool the block after surface decalcification.   This
will take time for each block as you would have to repeat this each time you
want to cut further into the sample.  

2.  If you have a tungsten carbide knife, you could try sectioning with the
TC knife,  and not surface decalcify.   

Good luck 

Gayle Callis
HTL/HT/MT(ASCP) 

 

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Berger,
Rebecca
Sent: Friday, February 14, 2014 9:07 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Mouse Joints

Hey Histonet!
 I have to troubleshoot some mouse knee joints that were not completely
decalcified before processing. I received them already embedded and they're
almost impossible to cut. Any tips to get them decalcified?
Thanks!
Becky
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[Histonet] Caveats Re: Bouins in microwave for Massons trichrome

[gayle callis]

Dear Histonetters, 

 

Microwaving Bouins can be an absolute disaster as experienced  by me.   It
went all over the interior of microwave (borrowed one at that!) and created
an unbelievable mess of picric acid that then dried out,  and formaldehyde
going into every nook and cranny of the microwave interior along with
terrible fumes.  I spent a good deal of timeAll I can say is DONT
MICROWAVE BOUINS!!  

 

Other caveats:  

 

#1 is already mentioned

 

#2 Microwaving sections in Bouins can lead to spurious staining results.
Too short a time can create poor staining with this protocol.   As stated by
Liz Chlipala,  use a heated oven or waterbath at 56°C - 60°C for one hour to
ensure the section is properly exposed Bouins acid components (picric and
acetic acids) is a must for staining the connective tissue fibers.
This is the classic way of doing Massons Trichrome and still works the best.
A microwave must be properly vented in a hood, or it is a toxic mess of
fumes, let alone having to clean up nasty picric acid that will dry out in
crevices/corners of MW.   You can use a one zip lock baggie,  left open at
one end of top to vent fumes, and collect any boil over, but staining
problems have to considered due to short exposure to Bouins.   This is
something I learned from Jerry Fredenburgh many years ago, and after bulk
staining several hundred decalcified bone sections, the classic method,
correct time in Bouins gave the best and most consistent results.  

 

#3  If you are doing Mass Tri the next day, deparaffinize the sections,
rehydrate and immerse in Room Temperature Bouins and let sections sit
overnight.   No heating means:   no toxic formalin fumes from either MW or
heated oven,  problem and decalcified bone sections stay on slide as these
can dislodge due to  mechanical/physical forces from MW heating and
sometimes with heating in an oven, and is safer for user.  This became our
favorite method, and no more spillage/toxic fume problems.   

 

If anyone wants an excellent Massons Trichrome method, the classic one I
have from AFIP lab is superb.  It also gives hints on achieving best results
with proper differentiation of connective tissue fibers - something we often
do not think about when doing this stain.   

 

Take care

 

Gayle M. Callis

HTL/HT/MT(ASCP)  

 

 

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[Histonet] Need help with a publication reference

[gayle callis]

Dear Histonetters, 

 

If anyone has an original copy of Woods and Ellis, Laboratory
Histopathology: A Complete Reference, 1994 Churchill Livingstone, I would
like to know the chapter and page numbers for Anthony S-Y Leong,  Fixation
and Fixatives chapter.   Title of book, publisher, date and ISBN # is fine.
Unfortunately, the book is out of print.  

 

What I found on the internet for Roy Ellis Histology Page doesn't not have
the page numbers for the Leong  Fixation excerpt.   And the email address
for Roy Ellis is no longer valid or up to date.   

 

Unfortunately, publishers need chapters and pages in references.   If anyone
has the book and has the time, a pdf copy of the table of contents would
very helpful.

 

Thank you for your help

 

Gayle M. Callis

HTL/HT/MT(ASCP)

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[Histonet] Re: Activated charcoal for VIP 3000?

[gayle callis]

Dear Mike, 

 

Our VIP K series had plastic containers to hold the charcoal and was
refilled from bulk, and hopefully your VIP 3000 (you said 300?) has the same
containers.  I used charcoal for fish aquarium filters from a local pet
store.It was cheaper than activated charcoal from other laboratory
charcoal suppliers and although the charcoal pieces were a bit larger, the
fume filtration still worked.   One reason for trying this was charcoal from
Sakura for the K series processors was being discontinued for older models,
and a bit of ingenuity with lack of funding drove me to try it.   Newer
aquarium charcoal can be smaller as I refill my kitchen over the stove top
filter at home since I can't find the replacement filters anymore.

 

It may take some price comparison at PetSmart and Petco versus what you have
found already.  Buying in bulk would be ideal.  Try Amazon and/or do a
Google search.   

 

Gayle Callis

HTL/HT/MT(ASCP)

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[Histonet] Re: Filtering stains

[gayle callis]

Filtering stains was a long standing habit in our lab.  It didn't matter if
the solutions were coming from a stock bottle or reused and filtered into a
clean container. 

 

However, the kind of filter paper is  important and something I learned
early on from Lillie and Fullmer's book, Histopathological Technic and
Practical Histochemistry.   It was recommended to NOT use a high retention
paper (slow filtration) and use a medium fast paper.   If the porous filter
paper is  too slow, dye molecules are retained on the paper plus it takes
forever to filter a solution when tends to be thicker or flocculent.
Nuclear Fast Red is one that takes forever to filter - very annoying when
there was a time crunch.  Plan ahead!   

If you use a fast paper e.g. more porous, then fine particles can go through
the pores,  creating some contamination on the section.   Some stains were
not filtered e.g. working silver solution for GMS. 

 

As for hematoxylin solutions, regardless of a new stock solution or reusing
the stain the next day, we always filtered to get rid of any oxidation
product and/or tissue particles.If I didn't filter hematoxylin solution
before IHC counterstaining, there was always an ugly precipitate on the
sections.  

 

Some stains in alcohol, e.g. Luxol fast blue and eosin were not filtered
although reused solutions do contain tissue particles, and then were
filtered. 

 

Rule in our lab:   Filter stains through medium fast paper, daily for HE,
but top off to keep replenished and at correct level for staining racks.   I
know that some vendors say not to filter the hematoxylin before routine use,
but I could never get out of the filtering habit.   

 

An addition bead of wisdom for people.Filtering is a messy procedure but
using the correct funnel is important to prevent splashing.   Years ago and
in a chemistry class, I learned to use a tapered end funnel for
filtering/introducing a fluid into a container.  Make sure the tapered end
touches the side of the container.   Use wide, open ended (non-tapered)
funnels are for introducing dry chemicals to a container as wide ends will
cause splashing. 

 

Gayle Callis

HTL/HT/MT(ASCP)  

 

 

 

   

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RE: [Histonet] CD4 and CD8 mouse tumor IHC, thanks..

[gayle callis]

Another solution, and one we did almost exclusively for murine T and B cell
IHC was use biotinylated Rat anti mouse CD4 and CD8.   The isotype can be
purchased biotinylated from BD Bioscience/Invitrogen or you can buy Jacksons
Rat IgG-biotin. 

This totally eliminates a secondary antibody hence anything rat will not be
detected.   A bonus is staining is much faster too.  We routinely did
Streptavidin/biotin block from Vector since Streptavidin has an affinity for
integrins in epithelial cells.Our normal serum block with biotinylated
rat antimouse primaries is 10% goat or donkey serum + 2.5% mouse.   The
block is also the diluent for the primary antibody and isotype/IgG negative
control.   Rabbit serum and antibodies were avoided since the bunny is
sticky, and can cause more background.   My immunologist frowned on anything
using a rabbit host unless we couldn't avoid it. 

Good luck
Gayle Callis
HTL/HT/MT(ASCP)  



-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Erickson,
Jamie E
Sent: Monday, December 09, 2013 12:06 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] CD4 and CD8 mouse tumor IHC, thanks..

Thank you to those that sent me suggestion on my CD4 problem. I have
recently discovered the problem.
The tumors are mouse tumors but they are injected in-vivo with a test
antibody that is Rat anti-mouse, specifically Isotype (Rat IgG2a). Hence
this is why my secondary Rabbit anti-rat stained so much..

Next I will try to block the Rat in-vivo injected antibody with a goat
anti-Rat secondary, as a  block prior to staining with CD4.
Or I could precomplex the CD4 with the Rabbit Anti-Rat and bind excess
Anti-Rat with Rat Serum.

Any thoughts or suggestion on this as always would be welcomed...

Thanks again..

Jamie

Abbvie Bioresearch Center
Pharmacology
100 Research Dr.
Worcester, Ma 01605
OFFICE+1 508-688-3134
FAX  +1 508-793-4895
EMAIL  jamie.erick...@abbvie.com
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[Histonet] Re: Questions about IHC on Frozen Sections

[gayle callis]

Dear Brett, Liz and Patrick,

 

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

 

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

 

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

 

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

 

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

 

PEROXIDASE BLOCK (0.03% hydrogen peroxide)

 

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

5 ul 30% hydrogen peroxide 

50 ul 10% sodium azide

 

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

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

 

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

 

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

 

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

 

Take care

 

Gayle Callis

HTL,HT/MT (ASCP)

 

 

 

 

 

 

-Original Message-

I agree with Liz,

 

We usually fix with acetone/ethanol 5-10 min then go right into buffer, but
occasionally use 2.0% NBF for some antibodies.  Our buffer contains 0.1%
Tween and our sections can be anywhere from 8-20um depending on the specific
project. I think the 30min in acetone is messing up your morphology.

 

Brett

 

 

Brett M. Connolly, Ph.D.

Principal Scientist, Imaging Dept.

Merck  Co., Inc.

PO Box 4, WP-44K

West Point, PA 19486

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

T- 215-652-2501

F- 215-993-6803

 

-Original Message-

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

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

To: Lewis, Patrick; '
http://lists.utsouthwestern.edu/mailman/listinfo/histonet Histonet @t
lists.utsouthwestern.edu'

Subject: [Histonet] RE: Questions about IHC in Frozen Sections

 

Patrick

 

Here is what we do for frozen IHC, this is based upon methods that I
received from Gayle Callis.

 

Cut frozen sections and let air dry

[Histonet] Re: ImmEdge Pap Pen problems

[gayle callis]

The problem is using the pen without redistributing the liquid and solid
contents in the pen.   Years ago, the Vector technical service rep said to
shake the daylights out of the pen.   We use a vortex mixer, at highest
speed and press hard, to make sure the contents were well mixed before use.
We then have no problems of the goo lifting off the slide when in buffer
or using a solvent fixative for frozen sections after applying the barrier.


 

We had pens that worked for years as long as we shook (or vortexed)  them
well before applying the barrier around sections.   We used the pens for
both deparaffinized slides (well dried around section) and air dried frozen
section (on dry slides).   

 

Good luck

 

Gayle Callis

HTL/HT/MT(ASCP)  

 

 

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[Histonet] RE: goat anti Rabbit Alexa 594 woes.

[gayle callis]

You wrote: 

 

I would like to get your keen ideas about two issues I am facing now in my
immunohistochem. procedure:

 

1- I am using goat anti rabbit Alexa 594 (red) as secondary antibody for
detecting C-Fos in rat nervous system. When I am looking at the slides, it
seems always I have some residues of my

Alexa remained on the surface all over the section. I've added one extra
quick wash in every washing steps (Using PBS+TX100). But still they are
exist. Is anybody has suggestion to improve the quality of the picture and
get ride of these residues?

 

 

Leila Etemadi

M.Sc., Ph.D Candidate

Neuronano Research Center (NRC)

Lund University, BMC F10

Sweden

 



This is only a reply to #1 inquiry about residues on tissue after staining
with antibody-fluorophore conjugate.   What you have is deposition of
fluorophore-protein aggregates from antibody that tends to break down over
time and this is unavoidable and quite normal.  

 

Solution is dilute your goat antiRabbit Alexa 594 to working concentration
in micro centrifuge tubes, and spin down diluted antibody in a desk top
micro centrifuge a couple of minutes or so just before applying to the
tissue section.   Pipette the spun antibody carefully, keeping pipette tip
out of bottom of micro centrifuge tube,  to avoid remixing the fluorescent
aggregates and apply to section.   Every time I did not spin my diluted
antibody to take a shortcut in time (the lazy approach),  I ended up with
what we called glowing garbage. More rinses or longer rinsing did not
solve this problem as the aggregates tend to sit on the sections and slide.


 

Over the years of working with fluorophore conjugated antibodies, there some
things I did faithfully to keep sections free of glowing garbage 

 

1. Rinse buffer contained detergent, with our preference 0.025% Tween 20 and
NO protein carriers e.g. no BSA or normal serums.   Triton X 100 is ok to
use.   This rinse buffer was used to make up antibody diluent and blocking
reagent.   

  

2.  The final rinse buffer had no additives, just pure buffer before I cover
slipped with Prolong Gold Antifade reagent from Molecular Probes. I did more
final rinses X5 to make sure all antibody-fluorophore is removed. 

 

Take care

 

Gayle M. Callis

HTL/HT/MT(ASCP) 

 

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[Histonet] Re: Holes in paraffin, remedies

[gayle callis]

Hugh wrote: 

 

Sophia,

 

Holes like you mentioned, are probably from the polymers or impurities in
your embedding wax separating.  Gayle Callus mentioned this several times
(see histosearch), and recommended Stirring the embedding paraffin every
so often (before every use).  It could also be your tissue processing.
Check to make sure it is properly processed.  Embed without letting the wax
solidify before doing so.  You may have a bad batch of reagents?

 

As for the stainer, Hazy nuclei, as long as you are sure your hemotoxylin is
fine, are indicative of incomplete deparaffinzation, which could mean your
OVEN or your reagents.  It could also be your acid/base solutions.

 

Note:  Have you started a new lot # of wax?  How about xylene
(xylene-substiture) or alcohols?  I have heard people saying X-brand
paraffin was having problems with impurities that could cause these
problems.  I also heard it was solved last year. 

 

Also, your stainer has very little leeway in deparaffinzation.  You might
have to do things by hand until this problem is resolved.  You know,
deparaffinize to water using the longer method, then finish the HE in the
stainer.  Does this help?

 

 

Good luck (whoa, you have your work cut out for you),

 


***

 

Hugh, 

 

Thank you for the kind words and your reply was right on the money.In
fact, we stirred the paraffin daily before embedding, since the settling of
polymers is not just occasional.Also, clean your embedding center
frequently, before adding more paraffin.   

 

There is a clever little test for paraffin carryover into your rehydration
alcohols when removing paraffin.   1)  Use a glass beaker, add a few ml of
used alcohol  starting with last 95% just before 70% in deparaffinization
setup.   2) Pipette a few mls of  tap water into this aliquot of 95% and
look for cloudiness.   If a white cloud occurs, you have paraffin carryover
all the way down your deparaffinization process.   3) If you don't see
cloudy in this last 95%, test the 95% before this one, and keep going
backwards towards the xylene or xylene substitute.  If the next 95% is
cloudy, there is paraffin carryover.  Change out that alcohol station and
all the ones before that including the xylene/xylene substitute. 

 

Also, you can do rotation where you move the second ( closest to water)
95% into first 95% spot, then replace the second 95% with fresh, replace all
100% and xylene/xyl sub  as these latter are probably heavily contaminated
with paraffin which is carrying over into your alcohols at some level.   We
used two or three xylene/xylene substitute changes (three preferred when
doing IHC at 5 min/change) , two (or three)  100%, two 95% and one 70%
alcohol changes before distilled water, 3 minutes per change, with hand
staining.Change distilled water frequently, fresh daily and more changes
when staining many slides as you don't need alcohol carry over into your
hematoxylin.   Careful monitoring of your solvents and distilled water
should allow better hematoxylin staining. 

 

We also changed our acidic solution after hematoxylin and bluing solutions
daily, these solutions are cheap plus always doing a 1 minute running water
rinse after hematoxylin, acidic solution, and bluing.If you don't have
running water rinsing, at least change your water rinses before each
staining run.  

 

This all sounds very picky, but our HE staining was very successful.  

 

Gayle Callis

HTL/HT/MT(ASCP)  

 

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[Histonet] Re: Oil Red O and Mayers

[gayle callis]

Stacey, 

 

You wrote:  You wrote:  

 

We are trying to stain frozen cut sections of aorta with Mayer's Hematoxylin
following Oil Red O staining. We cannot get hematoxylin staining to work. We
are mainly seeing blue background but not labeling of nuclei.

Tissue is fixed with 4% PFA prior to sectioning. After tissue is cut;
sections are stained with Oil Red O

. ddH2O

2'

. 60%

isopropanol 30s

. Oil

Red O 18'

. 60%

isopropanol 30s

. 2x

ddH2O 1'

 

Sections are then stained with Sigma Mayer's Hematoxylin 

.Rinse in deionized water

 

.   Stain in Mayer's Hematoxylin 1-5 min

 

.  Rinse in running tap water until nuclei are blue

.  Rinse in deionized water

 

We have tried staining in Hematoxylin for 3 min up to 15 mins

We have also tried rinsing 1 min up to 15 mins (checking at for staining at
various timepoints) and always see the same result -some blue staining but
no clear nuclear staining.

 


***

Possibilities here are:  

 

1.  Not getting rid of the isopropyl alcohol adequately.  Try a longer rinse
than 1 minute to rehydrate the section, and add an additional DI water rinse
to make sure the alcohol is removed

 

However there is a much better Oil Red O stain than the one you could be
using now (you didn't give method e.g. messy Oil Red O with propylene
glycol, and without isopropyl rinsing.I am forwarding Churukians Oil Red
O under separate cover which has never failed on our frozen sections.  It is
easy to make up and use.   We also prefer Gill 2 hematoxylin staining which
takes less time and requires a mild bluing solution or you can simply use
warm water to blue the section.   If Gill 2 seems too dark, use Gill.   I
have the original Journal of Histotechnology publication by Churukian if you
want it.  We always do Oil Red O stain before counterstaining with
hematoxylin.  

 

2.   How fresh is your  Mayers?   It could be outdated or needs changing?


 

Also, after final tap water rinse, we coverslip without an extra deionized
water rinse. 

 

Good luck

 

Gayle Callis

HTL/HT/MT(ASCP)  

 

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[Histonet] FW: CD4 CD8 Zinc Tris PE staining

[gayle callis]

Hello Patsy, 

 

You wrote:   Has anyone tried this:  zinc salt fixed mouse tissue PE stained
for CD4/cd8.  I found some papers that look impressive.

 



I presume you mean the Beckstead zinc TRIS fixative that is formalin free?
It should work with murine or rat tissues fixed in  ZnTRIS  since the
tissues can then be paraffin processed, but avoid any solvent that have NBF
carry over so these antigens are not compromised by that sniiff of aldehyde.
The fixative can be purchased from BD Bioscience/Invitrogen.   However, PE
(phycoerythrin) fades rapidly before your eyes while you are looking at the
stained sections.  Been there, and seen it happen resulting in a lot of
wasted time and effort.PE is for FACS not fluorescence microscopy.
Not only that, but doing direct immunofluorescence using CD4 or CD8
conjugated to a fluorophore will result in poor to no staining since the
fluorphore molecules in close approximation to each other will exchange
electrons and quench fluorescence.   It is better to do CD4 or CD8 then come
back with an excellent secondary e.g. Jacksons donkey anti mouse F(ab')2
frag of IgG conjugated to Alexa 594  OR use a biotinylated CD4 and CD8 and
come back with Streptavidin Alexa 594.   Double staining using this fixative
should be very possible, even triple IF.

 

I have several publications for this fixative including the original
Beckstead (for human) and Nitta et al (for murine) CD marker publications.
We tried ZnTris for chromogenic IHC with success but didn't continue to use
it since we did the work much faster with frozen sections.We found this
fixative did not work well on fresh tissue frozen sections though, and
preferred to use our favorite 75% acetone/25% absolute ethanol for 5 minutes
at RT  on air dried frozen sections then into buffer from this solvent
fixative (sections were not alllowed to air dry again after this fixaton).  

 

As for papers you have seen, can you give the references please?  

 

Gayle Callis

HTL/HT/MT(ASCP) 

 

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RE: [Histonet] eosin in the processor

[gayle callis]

The only immunostaining any residual eosin in tissue might affect is if you
do immunofluorescence.  Eosin does fluoresce.   Others can address any
effect on IHC.  

Gayle Callis
HTL/HT/MT(ASCP)
Bozeman  MT 

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Kim Merriam
Sent: Wednesday, November 28, 2012 5:55 AM
To: Histonet
Subject: [Histonet] eosin in the processor

Hi Everyone,

Years ago, my lab used to put eosin in the processor to lightly tint the
smaller mouse tissues.  I can't remember which station we put it in (I think
it was the 2nd 100% ethanol).  Also, back then my lab didn't do any IHC;
will the eosin affect any IHC that might be done (I am guessing no, but I
want to be sure).

Thanks,
Kim
 
Kim Merriam, MA, HT(ASCP)QIHC
Cambridge, MA
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RE: [Histonet] Looking for US Distributor for biological dyes

[gayle callis]

Amy,  

Cellpoint Scientific.   You have to call them for information on what you
want, or a catalog?  Telephone:   800-999-9734 or email:
i...@cellpointscientific.com

They also have brain matrix devices for rodents, etc., but no website other
than contact information.   

Gayle Callis
HTL/HT/MT(ASCP)
Bozeman MT 




-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Amy Porter
Sent: Tuesday, November 27, 2012 7:24 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Looking for US Distributor for biological dyes

Does anyone know of a U.S. distributor that carries Chroma-Gesellschaft
dyes???

 

Amy S. Porter, HT(ASCP) QIHC

Michigan State University

Investigative HistoPathology Laboratory

William S. Spielman, Ph.D. - Director

Patricia K. Senagore, M.D. - Consulting Pathologist

Department of Physiology / Human Pathology

Biomedical Physical Sciences Building 

567 Wilson Road - Room 2133

East Lansing, MI  48824-3320

Phone:  517-884-5026

Fax:  517-432-1368

port...@msu.edu

www.humanpathology.msu.edu

 

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[Histonet] RE: Wrinkle Out Water Bath

[gayle callis]

You wrote: 

Is anybody using this product, Wrinkle Out Water Bath H
http://lists.utsouthwestern.edu/mailman/listinfo/histonet @t O, in their
waterbaths and what precautions are you taking if so?
The label states that the product is highly toxic, a hepatoxin.
I received a sample of it and wasn't aware of its toxicity before I
requested the sample at the NSH S/C.


In reading the MSDS for this product, all it contains is 1% isopropyl, 18%
Ethanol, and 1% methanol citing possible damage to liver.  Why not avoid
spending the money and just use 20% ethanol in your water bath according to
how this product is used.It will work just as well as this.You could
even make up your own mixture with isopropyl and ethanol, forget the more
toxic and expensive methanol.   These alcohols are all used for tissue
processing e.g. denatured reagent alcohols, isopropyl, ethanol and/or
methanol alone, and just as toxic to the liver. 
 
Beware!  Too high of an alcohol concentration can cause your sections to
explode on a warm water bath.  We never used alcohol in warm water bath to
flatten tissue sections.  Instead, we used RT 10% ethyl alcohol in a glass
staining dish, floated a section on this, picked up section onto a slide
then went to a warm water bath, gently lowered the section but kept top
paraffin portion of section on the slide, and watch the section flatten in
the warm water.   This prevented a section from exploding wildly or losing
the section on the water bath. All the alcohol or a detergent does is
reduce the surface tension of the water so the section flattens.   Tween 20
has been used too. 
 
I don't think any of these damage IHC since the antigens are still protected
by the paraffin in the tissue section.   
 
Gayle Callis
HTL/HT/MT(ASCP)
Bozeman MT 
 
 

 

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[Histonet] Rabbit/Mouse Polymer Detection Kit Recommendation?

[gayle callis]

Biocare and Golden Bridge International have kits.I believe Vector can
be added to this group too.  

 

Gayle Callis

HTL/HT/MT(ASCP)

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RE: [Histonet] Re: Changing dynamics in histotechnology

[gayle callis]

Well, this honey has been on the same page since the EARLY 1960'S.  I
crossed over from the MT side into histology and never looked back.   It was
obvious very early on that histology was far more interesting than working
as an MT, poor pay or otherwise. Way back in the Dark Ages, our MT
training included histology and the ASCP MT registry exam tested us on
histology. Becoming an MT simply led to histology, and the MT training in
clinical chemistry, microbiology, parasitology, virology, hematology, etc.,
enhanced our knowledge for working in histology.   

Your (plural) discourses have been interesting, to the point and certainly
no offense is taken about being an MT!   

It is admirable when histotechnicians go above and beyond their jobs and
take the time pass on their expertise to present workshops, teleconferences,
presentations and writing articles with hopes the written word is actually
being read.   Don't stop!  Ignore the critics, the complacent!   Educate!  

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


-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Weems, Joyce
K.
Sent: Tuesday, September 18, 2012 8:16 AM
To: 'joelle weaver'; tjohn...@gnf.org; histonet@lists.utsouthwestern.edu
Subject: RE: [Histonet] Re: Changing dynamics in histotechnology

Honey! We have been trying to get this group on the same page since the 70s.
We're a bit closer but we're still singing different songs... fa la la la,
la la la la...

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



-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of joelle
weaver
Sent: Tuesday, September 18, 2012 1:58 AM
To: tjohn...@gnf.org; histonet@lists.utsouthwestern.edu
Subject: RE: [Histonet] Re: Changing dynamics in histotechnology


TeriI think you are right about the promotion of the status quo, and this
is a definate concern for me in staying in this field. There seems to be so
much change resistance.  Also, it is my understanding that many MT programs
used to require histology rotations in histology here as well, but it
seems many now do not. It seems to me that many MT programs are 2 +1 or 3
+1, which is fine by me, but I never thought this was the same as doing a
full undergrad curriculum,  and never understood why it offers MT
gradstrumping  for any advanced lab roles, over any other similarly
educated lab person with equal or greater education and training? I have
concluded that we are fighting a perception, and that is not going to be
easy. Personally,  I have no issue with an MT doing histology if they want
to learn it sincerely by whatever means, but some seem to think that since
they know clinical lab, that it does not take any additional learning,
formal or otherwise. I often wonder why it seems outrageous to the same, if
it were to be worked the other way? I believe that I would be ignored
completely or scoffed at,  if I tried to apply, or walked into a clinical
lab to work. Also,  I think some people in histology have put considerable
effort into dialogue about our field and its needs for well prepared staff
in the main-stream media, but I agree that it is far below the level of
communication that will be needed to change the aforementioned perceptions.
Interestingly, most histotechs I have encountered are unwilling to dedicate
much time, since it is rarely for any pay,  to any activities like these-
since it often involves a lot of work and preparation to construct/publish
an article or give a presentation out in the public arena. I know that over
time, I have donated probably hundreds of hours, and most of the time it is
a fight just to be allowed to do this ( such as take time off from work
with your own vacation to travel or attend). If anything in my current
environment, people roll their eyes at me for doing anything of this sort.
If you want to encourage people to participate, we will have to work to see
it supported within organizations and applauded within the group. So what
usually is a frustration/dissappointment for me is when  people will
complain, but most won't bother to take any action ( not directed at you or
anyone in particular, just expressing frustration with general lack of
initiative)...anyhow your points are well taken. If we are to move forward
as a group, we are going to have to get on the same page ourselves and put
forth some consistent and concentrated efforts.




Joelle Weaver MAOM, HTL (ASCP) QIHC
  From: tjohn...@gnf.org
 To: histonet@lists.utsouthwestern.edu
 Date: Tue, 18 Sep 2012 00:06:37 +
 Subject: [Histonet] Re: Changing dynamics in histotechnology

 Ok, my workplace blocks Facebook, so here is the article for those of 
 you who can't read it from the original link provided: 
 

[Histonet] RE: zinc fixative

[gayle callis]

Amy,

 

You wrote: 

 

A wonderful colleague recommended to aid in my response that I add the
components I am using for preparation to assist in obtaining a more specific

response, about shelf life, so here it is: 

 

0.1   M Tris/HCl Buffer pH 7.4 - to which is added:

 

Calcium Acetate - 0.5 gm

 

Zinc Acetate - 5.0 gm

 

Zinc Chloride - 5.0 gm

 

The resulting solution was pH 5.45

 

According to my protocol my pH should be between 6.5  7.0 which is working.
I am just starting to work on developing this protocol and my Zinc Chloride
looked pretty pathetic in the bottle and I will be ordering fresh.  I think
this may have affected the pH - uncertain however at this point.

 

**

Invitrogen/BD Biosciences sells this fixative aka IHC Zinc Fixative
(formalin free) and are only vendors in the USA as far as I know although it
is sold in Europe under another name.   I did not find a shelf life in
either MSDS or Technical Data sheet, but you might want to contact them
about this.  When we tried this, we made it fresh.  You may want to look at
cost of buying the ready made compared to buying new chemicals or taking the
time to make it up - whatever is cost effective.If you make it up, I
suggest using only fresh chemicals if any of yours  have been sitting around
on the shelf as we ran into the same problem with zinc chloride going bad.
The commercial IHC Zinc Fixative is a 10X solution (storage is RT) and
diluted with distilled water just before use - very handy.

 

If you are interested, I have many publications on file about this fixative
including the original Beckstead (for human CD markers) and Nitta (for
murine CD markers) articles and would be happy to send these to you
privately.   At least the TRIS buffer can be made up ahead of time, maybe
even as a 10X solution and dilute when needed to add dry chemicals just
before use.   One thing is that the pH is never adjusted after adding the
chemicals to the pH 7.4 TRIS buffer.   This was mentioned in the original
publications.   

 

Some things to think about when using this fixative.Make sure your
tissues are not overly large/thick by reducing the sample size to achieve
total fixation since the time of fixation is limited.   BD Bioscience says
up to 48 hours.  I think one could perfuse nicely with this fixative too or
at least inject it into lumens, fill lungs, hearts, etc.   If you have
incomplete fixation with ZnTRIS buffer (Beckstead's fixative), then alcohol
during processing will complete the fixation which is something you do not
want to happen.Nitta et al had a processing schedule in their
publication but we found we had to use shorter processing schedule for
murine tissues which became too dry and friable leading to poor microtomy
with overly long soaking to get a section.   Also the first NBF station on
the processor should be replaced with this fixative.  When working with
formalin sensitive CD markers which is the purpose of this fixative, I
wouldn't want a sniff of formalin from  NBF carry over into ANY of the
solvents.   Others may have more suggestions on this.  

 

I am not sure what you are using zinc fixative for, but presume it is for CD
markers.   

 

Good luck, 

 

Gayle Callis

HTL/HT/MT(ASCP)  

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[Histonet] Re: DAPI stained smeared nuclei

[gayle callis]

Carol wrote: 

 

Someone sent me a question regarding DAPI staining ( I generally  use
Hoechst )...but anyway, they are having smearing of nuclear contents. I
would guess that is from rupturing of the nuclear membraneover fixation?
Over air drying?  They are fixing 10 mins in cold acetone and 10 mins to air
dry. What do the experts out there say? This could be a learning moment for
me, as well.

 


**

Interesting problem, but I have never had problems with DAPI staining after
either 4C acetone or my beloved acetone/alcohol fixation for murine CD
markers and other antigens that like solvent fixatives.  No problems occur
after air drying either.   We always air dry our frozen sections very well
before fixation and/or storage at -80C.   I don't think one can over air dry
unless they leave the sections out for days at RT, then antigenicity could
be damaged. 

 

I suspect there is something else going on with either how they store frozen
sections after sectioning rather than air drying, fixation in cold acetone
and then air drying again.   Possibly doing some kind of damaging thaw,
freeze, thaw?   Storing frozens in cryostat immediately after sectioning is
not advisable since condensation forms on FS when one takes them out of cold
environment. How do they pick up the sections as I can smear a frozen
section quite thoroughly if I am not quick and tidy when mounting the FS on
a slide.   The tiniest movement will mess up tissue components, even nuclei
The very act of picking up a section on a slide can be referred to as flash
drying and is, in some ways a means of fixation although not very good.

 

More details would help on HOW they handle the sections before fixation?
What tissues?  Cutting temperatures?   What is actually happening to their
immunofluorescence staining?   Is that fine but the nuclei are smeared?  

 

The only problem we ever had with DAPI staining is using mounting media with
DAPI which gave a staining gradient with dim staining of the nuclei in
certain areas of the tissue section.   We solved this by never using
mounting media with DAPI and now buy  DAPI solution from Biogenex or Pierce,
then stain with DAPI solution AFTER all the immunofluorescence staining was
complete followed by Prolong Gold Antifade reagent.  What kind of mounting
media do they use?  How do they rinse after all the IF staining?   Do they
use an in house prepared DAPI solution and if so, is it made correctly?   

 

Mark Tarango brought up an interesting solution to the problem, but what
happens after you pull the sections from freezer and go back to RT?

 

Tossing out a lot of things here.

 

Take care

 

Gayle Callis

HTL/HT/MT(ASCP)

Bozeman MT   

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[Histonet] JB fixative

[gayle callis]

Hi All,

 

 

You wrote:  I use a Zinc fixative called JB fixative for tissues that I want
to stain for CD antigens(see attachment). Does anyone know if this fixative
kills all microbes and prions?

 

You did not say what species you are working with?   

 

The answer to your question is no for both microbes and prions.   JB
(acronym for Jay Beckstead) fixative is formalin/aldehyde free, and is for
CD markers.   Beckstead developed this fixative when studying CD markers in
human lymphomas.

 

Beckstead JH (1994) A simple technique for preservation of
fixation-sensitive antigens in paraffin-embedded tissues. J Histochem
Cytochem. 42(8):1127-34.  Free online

Beckstead JH (1995) A simple technique for preservation of
fixation-sensitive antigens in paraffin-embedded tissues: addendum. J
Histochem Cytochem. 43(3):345. Free online

Did your attachment contain the information below (via IHCworld)?   

 

Zinc Fixative (JB Fixative) Formalin Free

 

0.1M Tris Buffer, pH 7.4

  Tris Base  12.1 g (TRIZMA)

  1N HCL --- 81.5 ml

  Distilled water -- 900 ml

  Mix to dissolve. Adjust pH to 7.4

 

Zinc Fixative

  Calcium Acetate -- 0.5 g

  Zinc Acetate -- 5.0 g

  Zinc Chloride -- 5.0 g

  0.1M Tris Buffer made above -- 1000 ml

Mix to dissolve. The final pH will be approximately 6.5-7.0. Do not readjust
the pH, as this will cause the zinc to come out of solution.   Store Zinc
Fixative at room temperature.   Fix tissues for 24 to 48 hours. Fixation
longer than 48 hours may make the tissue brittle and difficult to cut.

Description: Tissues fixed in this solution followed by paraffin embedding
and sectioning results in antigen preservation comparable to that in frozen
sections with antibodies to these cell surface markers (CD1, CD3, CD4, CD7,
CD8, CD19, CD31). Morphological preservation was also comparable to
formalin-fixed sections.

 

As killing microbes, and I presume bacteria, virus or fungi, you should use
neutral buffered formalin.  Prions are a whole other story since the only
way to totally eradicate these particles is incineration.  CDC has
guidelines on handling prions in a laboratory.

 

Gayle M. Callis

HTL/HT/MT(ASCP)

 

 

 

 

 

 

 

 

 

 

 

1

 

 

 

 

 

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[Histonet] Re:Glucose Oxidase peroxidase block that works for eosinophils

[gayle callis]

Dear Kim

 

I attached the peroxidase block method via personal email which works for
either frozen or FFPE sections.   You must buy the correct, exact glucose
from ICN as Sigma discontinued this.  I included the original publication.
You can also find this blocking method on the Vector website.   This has
been discussed on Histonet before but possibly not in the context of the
original publication for eosinophils in frozen section in minimally fixed
e.g. acetone.If the sections are FFPE, you may have to extend the time
or stick to 60 minute incubation in the glucose oxidase/glucose mixture.
We nicknamed this GLUOX Peroxidase Block.   For frozens,   we used 60
minutes incubation but later found a publication using 30 min blocking of
endogenous peroxidase (ordinary and not eosinophils) in frozen sections.
Timing is something you might play with to shorten the staining time.
Theory is slow, continuous production of H2O2 removes/blocks peroxidase and
pseudoperoxidases in tissues/cells. 

 

I had feedback that it worked on paraffin sections with stubborn endogenous
peroxidase.

 

Good luck

 

Gayle Callis 

 

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[Histonet] Re: Unsubscribing problems for the amusement with a touch of seriousness for all

[gayle callis]

Dear Histonetters particularly those irritated with people who incorrectly
unsubscribe from this list,   

 

After participating on Histonet since its very beginning,   it is noted that
incorrect unsubscribing (and other infractions) from the list has been a
continuous problem from the start.   This has been the proverbial thorn in
peoples' sides and always eliciting a rash of How to unsubscribe correctly
messages.  This rash of How To Unsubscribe messages can be found in
Histonet Archives for every year this list has been in existence.This
includes to the amusement of some, the misspelling of UNSUBSCRIBE in
interesting, amusing ways.Another irritation is Re: Daily Digest instead
of a valid topic in subject line.   All these things use to get my lab coat
in a bunch, decision were made to calm ruffled feathers and deal with these
life's less than important crises.

 

So the following suggestions/comments might help.   

 

1.  Don't waste time informing those few people How To Unsubscribe.
This is a lost cause, a proven fact shown not to work during all the years
of Histonet.  

 

2.  If one is subscribed to Histonet and a subject line contains
Unsubcribe me; Remove me from list; Take me off Histonet, etc. and Re:
Daily Digest. Use delete key.  

 

3.  Suggestion. If one finds these irritations too difficult to deal
with, causing indigestion, clenched jaw syndrome, gnashing of teeth, tension
headache, sleep loss , computer keyboard abuse, bad language habits, and
Histonet  eRoad rage leading a message for all to read,  try cruising
Histonet Archives daily.   You pass up subject(s) not interesting to you,
and the OMG not again (!) incorrect Unsubscribe practice.When a
message/topic piques your interest enough to pen a reply, then hop back onto
(whoops! subscribe to) Histonet to share your histo-expertise.   

 

Well I am off to unsubscribe correctly and will look for your replies with
commentary in the Archives. 

 

Gayle M. Callis J

HTL/HT/MT (ASCP)  

 

 

 

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[Histonet] Immunofluorescence in the clinical laboratory, some questions

[gayle callis]

Dear Histonetters, 

 

I know that immunofluorescence has been done for decades on renal biopsies,
but am curious if laboratories are using IF more these days?   If so, I
would be very interested to talk to you one on one about this as I have more
questions on why you deviate from standard chromogenic enzyme
immunohistochemistry and perform IF.   Comments about renal biopsy
procedures are welcome too.  

 

  Also, do you do mostly single IF or double IF, and the reasons why?   Is
your IF done primarily on FFPE or frozen sections/acetone fixation?  

 

When you do immunofluorescence in your clinical laboratory are you using an
automated stainer or a manual protocol?   Or is your clinical laboratory
associated with medical research groups?   

 

Any comments/information is most welcome.  

 

Thanks..

 

Gayle M. Callis

HTL/HT/MT(ASCP)  

Bozeman MT

 

 

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RE: [Histonet] Xylene Substitute for Counterstain Clearing

[gayle callis]

You are also  likely to have spheres dissolve in toluene based mounting
media.   Aqueous media may be the answer unless the neutral red runs out of
the tissue in the aqueous environment.   Hard to get the best of both worlds
if dye runs in aqueous media and spheres dissolve in solvent based .   Good
luck! 

Gayle Callis 

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Rene J Buesa
Sent: Friday, May 11, 2012 12:23 PM
To: histonet@lists.utsouthwestern.edu; Andrew Coleman
Subject: Re: [Histonet] Xylene Substitute for Counterstain Clearing

After staining dry the sections in an oven at 60ºC for 10 minutes. When
completely dried, coverslip as usual. Beware of the mounting medium solvent
because it may contain xylene as well. Use one mounting medium without
xylene.
René J.

--- On Fri, 5/11/12, Andrew Coleman andrewcoleman...@gmail.com wrote:


From: Andrew Coleman andrewcoleman...@gmail.com
Subject: [Histonet] Xylene Substitute for Counterstain Clearing
To: histonet@lists.utsouthwestern.edu
Date: Friday, May 11, 2012, 12:25 PM


Hi all,

We are performing a neutral red counterstain on tissue sections containing
colored polystyrene microspheres. The spheres are inert to alcohol, but are
washed out when we clear with xylene to coverslip.
The spheres are also supposedly soluble in DMF, acetone, acetonitrile,
chloroform and methylene chloride for what its worth.

Is it reasonable to coverslip these slides in permanent mount without
clearing with xylene after dehydrating the tissue? Or does anyone know of a
substitute clearing agent with chemical properties dissimilar enough from
xylene that might be worth trying instead?

Thanks,

Andrew

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[Histonet] Re: Massons Trichrome on decalcified bone

[gayle callis]

Dear Sara,  

 

You wrote: 

I work at an Orthopaedic research lab and I have been having some trouble
getting our trichromes to work on our bone. I have tried a masson's, gomori,
and goldner's and in all cases the bone stained red.

 

Our bone is arrive to our lab frozen, we fix them in formalin, embed in
paraffin, and decal with EDTA. I also post fix with Bouins before staining.

 

Techs that worked in this lab before me were also getting the same red
staining of bone.

 

Is there something during processing that could cause this reaction? We fix
in formalin, dehydrate through graded alcohols, and clear in xylene.

 




I doubt your processing has any effect on Mass Tri staining.   However,
incomplete decalcification can but I suspect it may be the staining protocol
itself.   Using a simple weight loss/weight gain decalcification end point
test with EDTA is a good idea, and can be used for acid decalcification
endpoint testing IF you are not presently using an endpoint test. 

 

Years ago, I visited the AFIP bone lab, and acquired a Massons Trichrome
protocol which worked much better than the standard Massons Trichrome found
in most textbooks.  It never failed to work well with our decalcified bone
work although we did use acid decalcification.This is NOT a kit method.
All reagents are made up in house, and post mordant heating Bouins is NOT
done in a microwave.  We preferred to let the sections sit in Boiuns
overnight  at RT, or heat in a water bath.   We never used a dry heat,
incubator type oven where one gets uneven heating in the chamber.  

 

There is more to doing Massons Trichrome on decalcified bone other than just
following the recipe from a textbook. One of the best discussions on
understanding the chemistry/theory of trichrome staining is found in Sheehan
and Hrapchak, Theory and Practice of Histotechnology.

 

The AFIP method has some different staining times, plus how to remove the
biebrich scarlet/acid fuchsin from collagen BEFORE going into aniline blue,
a much more controlled staining method.It also has a modified Weigerts
Iron hematoxylin that is superior, a bit more concentrated since Iron
hematoxylin tends to be removed by the acidic staining solutions.   I will
be happy to send these methods to you privately.  

 

Gayle M. Callis 

HTL/HT/MT(ASCP)  

 

 

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[Histonet] RE: Nocardia sp in FFPE

[gayle callis]

Jeanine, 

 

I always recall what the third edition AFIP manual (green) had to say about
staining Nocardia successfully.   Unfortunately, I didn't see an antibody
for Nocardia.  

 

Several stains work and maybe have to do a combination of stains correlating
results using adjacent sections?   

 

First choice would be GMS but you have to do some extra times in silver
solution according to AFIP manual/Luna.   In order to get complete
development of filaments if different types/strains are suspected, run two
slides with one at 60 minutes and the other at 90 minutes in the silver
staining solution.   Be sure to use chromic acid as the oxidizer.
Churukian Shenk had a modified GMS that is supposed to work too, but nothing
was said about extended times in methenamine silver solution.   

 

Fite's 

 

Rojas-Darby-Nochomovitz Method for Norcardia, TB and Leprae.  This is
similar to Fites but uses a Ziehl Neelsen carbol fuchsin modified with
mineral oil  (from  Luna's Histopathological Methods and Color Atlas book)

 

Luna Parker Method for Nocardia Asteroides  (Luna book) with filaments
appearing red.  

 

Brown and Brenn should show the organism too. 

 

Good luck

 

Gayle M. Callis

HTL/HT/MT(ASCP)

 

 

 

You Wrote: 

 

I need the absolute best, most fool-proof special stain protocol for the
staining of Nocardia in FFPE tissue.

 

Thanks in advance!

 

Jeanine H. Bartlett, BS HT(ASCP), QIHC

Centers for Disease Control and Prevention

Infectious Diseases Pathology Branch

1600 Clifton Road, NE

MS/G-32

Atlanta, Ga 30333

404-639-3590

Jeanine.bartlett @t cdc.hhs.gov

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[Histonet] Re: undecalcified bone IHC

[gayle callis]

Jeff, 

 

It is most certainly possible to do IHC on undecalcifed bone sections
embedded in PMMA although not the easiest task.   Sectioning is done on a
microtome that is powerful enough to cut the plastic and using tungsten
carbide knives.   The key is total removal of the plastic from MMA embedded
bone sections to allow antibody/ immunoglobulins to access antigenic sites.
Neil Hand has done IHC successfully on PMMA embedded tissues including
undecalcified bone on 2 to 3 µm thick sections.  I think one could cut
thicker sections at 4 to 5 µm and still be successful.  I do not recall what
Troiano et al used.   

 

The following publications will help you and should include protocols,
although conventional protocols will work according to Hand.  

 

Blythe D. Hand N et al 1997 J Clin Path 50:45-49.The use of methyl
methacrylate resin for embedding bone marrow trephine biopsies. 

Hand NM et al 1996 Antigen unmasking using microwave heating on formalin
fixed tissue embedded in methyl methacrylate J Cellular Path 1:31-37

Jackson P et al.   1996  Amplification of immunocytochemical reactions by
the catalytic deposition of biotin on tissue sections.   J Path
170(suppl):23A.  This was about tyramide amplification when one gets a weak
signal from conventional methods.   

Hand NM, Church RJ 1998 Superheating using pressure cooking: its use and
application in unmasking antigens embedded in methyl methacrylate.  J
Histotechnology 2`:231-236

Hand NM et al 1989 Immunohistochemistry on resin embedded tissue for light
microscopy: a novel post embedding procedure.  Proceeding Royal
Microscopical Society 24(1):A54-55. 

Hand NM Plastic Embedding media and techniques, Ch.30, p 663-677.   Theory
and Practice of Histological Technique,  5th edition by Gamble and Bancroft.
The 6th edition is updated under same title.  

 

Use Google Scholar to find Troiano N et al from Yale on doing IHC on PMMA
embedded bone sections with publications in J Histotechnology.

 

 

Hand mentioned several HIER methods, using citrate buffer.   Optimizing
retrieval will depend on the antigen and you may end up doing this with some
form of HIER, including microwave or other heat producing methods and with
different buffers. Enzyme digestion is also a possibility.   

 

Hand removed MMA with xylene, warm my speed up the removal, also more than
one change for 10 - 20 minutes or longer.   When I talked to him personally,
he said he had used warm xylene although temperature was not mentioned in
his chapter.   After MMA removal, rehydrate section through alcohol gradient
as one does paraffin sections.He was emphatic about never allowing the
sections dry out. 

 

Hopefully Jack Ratliff and Damien Laudier will provide more insight on this
topic.  

 

Good luck

 

Gayle M. Callis

HTL/HT/MT(ASCP)

 

 

 

 

 

 

**

 

Hi Jeff,

 

If is it possible a few more specifics of how the tissue has been received,

processed and evaluated would help.  Undecalcified bone sectioning

procedures vary and also what specific markers are you looking to do is

important.

 

Vikki

On Mon, Mar 12, 2012 at 11:06 AM, Rene J Buesa rjbuesa @t yahoo.com
wrote:

 

 Undecalcified? How are you going to section it?

 If you can section it, just use any IHC protocol for regular sections.

 Good luck!

 René J.

 

 --- On Mon, 3/12/12, Jeffery Howery Jeffery.Howery @t jcl.com wrote:

 

 

 From: Jeffery Howery Jeffery.Howery @t jcl.com

 Subject: [Histonet] Undecalcified bone IHC

 To: histonet @t lists.utsouthwestern.edu

 Date: Monday, March 12, 2012, 10:59 AM

 

 

 Does anyone have a protocol for Undecalcified bone for IHC?

 

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RE: [Histonet] Gram Stain

[gayle callis]

Stacy, 

You can buy a complete Brown and Brenn staining kit from Newcomer Supply
which is very good, and not change your method.   The picric acid/acetone
mixture is included and you wouldn't have to store stock picric acid nor
ether in the lab but just this small amount of Picric acid/acetone that
comes in the kit.  Disposal should not be that much of a problem.   They
also have a Hucker Twort Gram stain that uses acetone for destaining and you
supply the acetone.  Poly Scientific has Gram Stain kit (Hucker
modification) which uses Grams decolorizer, a mixture of acetone and
alcohol.  They have excellent staining kits too.   

Gayle Callis
HTL/HT/MT(ASCP)


-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Giroux,
Stacy
Sent: Monday, March 12, 2012 11:36 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Gram Stain

Hi,

Our lab is currently transitioning from the Brown  Brenn gram stain due to
no longer wanting to store picric acid due to its potential hazards. Our
pathologists have requested a gram stain for paraffin embedded tissue that
looks similar but does not use picric acid or ether. Does anyone have any
suggestions on stains that could be used or gram stain kits that are
available for purchase that are good?

Thank you for your help,
Stacy




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[Histonet] Re: PAMS Jones Basement Membrane

[gayle callis]

A stain I have never had a problem with.  However I did have the good
fortune to listen to Culling many years ago on the use of 1% periodic acid
for PAS staining and he insisted periodic acid, no matter what the
concentration, should be made fresh every time you do these stains e.g.  PAS
and PAMS.Some people use 0.5% but we used 1% for 15 minutes, and then
did microwave staining for the methenamine silver.A 56C - 60C water bath
also works plus monitoring the color of the sections -  the color of dark
tea.  

 

This is NOT a stable oxidizer once made up, and never should be reused under
any circumstances.   On the chance you have not used this solution in over 1
1/2 years, you may not be getting proper oxidation of the basement membranes
since the periodic acid and the methanamine silver may be bad.   Also, the
Methenamine silver should be no more than 6 months old.  We also kept our
silver nitrate solution fresh when we made up the methenamine silver
solution, and do store our silver nitrate salts in the refrigerator since
this is hygroscopic.  Check on storage of this salt in MSDS  I have
never been one to use kits for either of these stains, particularly with
periodic acid as a ready to use solution.   This goes into solution very
rapidly and be sure to oxidize for 10 minutes before going to the
methenamine silver.  None of these solutions is difficult to make up in
house.

 

If you want, I can send my method and also a protocol pdf from HistoLogic by
Stanley Shapiro, where he used freshly made 0.5% periodic acid via private
email.   The nuclei will pick up some silver, but one should be able to
discern nuclei from basement membranes on 1 to 2 µm sections.  

 

Good luck

 

Gayle Callis

HTL/HT/MT(ASCP)

Bozeman MT  

 

  

 

 

 

You wrote: Just wanted to give a quick update on this.  I had a suggestion
for using thiosemicarbazide for 10 minutes after the periodic acid, and of
all the things we tried, this was the only thing that worked.  It eliminated
the nuclear staining and the capillaries are now picking up the silver (as
they should be!).  Unfortunately, I accidently deleted the email that
suggested this so I don't know who to thank, but it was a great suggestion!

Liz

 

From: Elizabeth Cameron

Sent: Thursday, February 16, 2012 2:17 PM

To: histonet @t lists.utsouthwestern.edu

Subject: Jones/PAMS

 

Hi,

I was wondering if anyone has any suggestions for a Jones/PAMS stain that is
not working properly. This is something we don't do often.  The last time we
did it was a year and a half ago, and it seemed fine at the time.

We have tried 3 or 4 protocols, including an ammoniacal silver, and it is
still not working properly.  In some protocols, our red cells are staining
but the capillaries in the glomeruli do not seem to be picking up the
silver.  In other protocols, there are nuclei of some cells that should not
be staining that are, but again, the capillaries are not.  We are working on
mouse tissue that is fixed in NBF.  The strange thing is the stain seems to
be working well on Bouins and Telly's fixed tissue.  I even tried mordanting
in Bouins!  We have tried multiple kidneys with the same results.  We are on
new bottles of silver and periodic acid, although our methenamine has been
around a while.  Any suggestions would be greatly appreciated.

Thanks!

Liz

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Posting photographs RE: [Histonet] microtomy

[gayle callis]

There is a way to post pictures on histonet but just  not in the email
messaging.   The instructions for doing this are found on the Histonet
website. 

Speed does affect the thickness of sections.   It pays to turn the flywheel
at a steady, slow pace and not NASCAR racing speeds I have sometime
observed. Harder paraffin helps for thin sections but one should be able
to section at 2 um without difficulty on a properly adjusted microtome, a
sharp disposable blade and using any modern day paraffin. 

Gayle Callis
HTL/HT/MT (ASCP)
Bozeman MT  

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Kim Donadio
Sent: Friday, March 02, 2012 2:54 PM
To: Webb, Dorothy L
Cc: histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] microtomy

Love your question. Hate to hear that you are having a issue. My two cents
follow: 
Yes. The amount of time for a faced block will effect the section thickness.
The cells become bloated if they   Sit to long.  feel free to have a Friday
laugh on this one. 
Anyway. If you havnt changed the stain in any way or the tech isn't rushing
and hard facing causing extrenuated cell artifact and you don't have bloated
cells? Could it be possible his eyesight got better or maybe he is mad and
wants to pick a fuss? 
I sometimes wish we could post pictures if our issues. I'm wishing you the
best.
Kim D
Sent from my iPhone

On Mar 2, 2012, at 1:06 PM, Webb, Dorothy L
dorothy.l.w...@healthpartners.com wrote:

 We have run into an interesting scenario and wondering what the experts
think!  We cut bone marrow bx's and lymph nodes for lymphoma @ 3 microns on
one particular microtome.  Within the past month, the hematopathologist has
felt the sections are thicker than the usual 3 microns.  I had our service
technician  measure the microns and the equipment was cutting as set.  I had
the blocks cut on a different microtome and we have seen variations there
also.  My question is, does the amount of time on ice make a minor
difference in the section thickness?  I know a lot of responses may be the
difference in the tech cutting inasmuch as how fast they turn the rotations,
etc., but,we have ruled out that variable by having more than one tech cut
at the microtome in question. I am stymied as to how to remedy this
fluctuation!  This is why we love histology, so many variables to create a
problem and why I love histonet, so many techs to help one through a
dilemma!!  Thank you!!
 
 Dorothy Webb, HT (ASCP)
 
 
 
  
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RE: [Histonet] Cutting on an interview

[gayle callis]

I find the wording cutting on an interview interesting.   It could mean
cutting out e.g. leaving and not doing the interview.  Cutting =
sectioning?! 

Sorry if these appear to be a cutting  remarks. 

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


-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of joelle
weaver
Sent: Friday, January 27, 2012 12:29 PM
To: trathbo...@somerset-healthcare.com; Cheryl R. Kerry; Histonet
Subject: RE: [Histonet] Cutting on an interview


Yes, agree- this is essentially what I was trying to say eariler in the
thread.

Joelle Weaver MAOM, (HTL) ASCP
 
http://www.linkedin.com/in/joelleweaver

 From: trathbo...@somerset-healthcare.com
To: tkngfl...@yahoo.com; histonet@lists.utsouthwestern.edu
Date: Fri, 27 Jan 2012 19:18:06 +
Subject: RE: [Histonet] Cutting on an interview
CC: 

We also have cytotech applicants look at some slides as part of an
interview. These are cases which have been signed out already (patient
information is protected), the applicant is given a quiet room with a scope,
and records  their diagnosis.
Histology and Cytology are not especially large fields, and if you're
interviewing someone who works fairly local, chances are that you've met
them, or know someone who knows them. But with the applicant who has been
out of the field for a while, or has relocated, it is helpful to have as
much information as possible.
Unless your job requirements are very specific, and say how many
blocks/slides should be produced in a certain period of time, it will be
difficult to terminate the mediocre tech, but who interviewed exceptionally
well. This is also a good time to check for accuracy in labeling. Give them
blocks with six- digit, non-sequential  numbers, and see how they do.
 
 
 
-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Cheryl
Sent: Friday, January 27, 2012 1:53 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Cutting on an interview
 
 
 
 A comment on why we have techs cut on an interview-
 
 
How do you pick the BEST fit in the 60 minute interview?  The more
information you gather, the more likely you'll hire the best fit.  Most of
human communication is non-verbal.  Watching a potential new hire gives you
SO MUCH non-verbal information in addition to validating that they know
their way around a microtome.
 
The cost of mis-hiring is ASTRONOMICAL.  40% of companies polled say a bad
hire costs over $25000.  One in four polled estimated the cost closer to
$5.  Would you really want your Aunt Minnie's GI biopsy cut by someone
who COULDN'T cut a few blocks under a little new-interview pressure?
 
An authoritative article on just this kind of interview can be read at
www.fullstaff.org (A Histology Blog).
 
It's from Career Builders and gives a lot of impirical data to the value of
gathering the most information before making that hiring decision...we'd
love some feedback on the post.
 
Cheryl 
 
Cheryl Kerry, HT(ASCP)
Full Staff Inc. 
Staffing the AP Lab by helping one GREAT Tech at a time.  
281.852.9457 Office
800.756.3309 Phone  Fax
ad...@fullstaff.org 
 
Visit the FREE Webblog:  www.fullstaff.org  regarding Histology, Careers,
Tricks of the Trade, New Equipment review, and much more for our industry.
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[Histonet] Nail Polish sealant

[gayle callis]

You wrote: 

We are currently starting up some IHC on frozen tissue sections.  After
staining with different fluorescent antibodies, we end with applying DAPI
w/Prolong gold and then coverslipping.  We would like to seal the coverslip
so that we can keep the slides longer.  Any suggestions on where and how
best to apply the nail polish for a permanent fix on the coverslips?

 

*

Prolong Gold antifade reagent is a hard seal to begin with. We only seal
ends but not the sides of cover glass.  Our coverslips go right to edge of
slides e.g. 25 X 30, 25 x 40, or 25 X 50.We don't like having nail
polish slop over the edges onto back of slides but one could seal all sides
of coverslip if careful. We buy the thinner top or base coat nail
polish, but in general, prefer to use thinned mounting media rather than
nail polish.   There can be some issues here.If you are trying to view
GFP or RFP (red fluorescence protein) labeled cells or tissue components,
you should not seal the coverslip with nail polish since the alcohol in nail
polish leaches under the coverslip and causes GFP/RFP to fade.  This fact
was published in Science.   We found that dumping out cheap clear nail
polish from bottle, rinsing away the residue with acetone, and then adding
permanent mounting media and thinning that with toluene to the consistency
of top coat nail polish works best.  Toluene or xylene based sealants cannot
leach under the cover glass since these solvents are NOT miscible with water
in the PBS.Thinned mounting media is better sealant for GFP purposes (no
fading) and also works for IF stained sections (perfect seal).  We love the
little brush in the nail polish bottle for application.  Thicker clear nail
polish (for non GFP studies) or IF stained sections is messy during
application so we buy the cheapest top coat polish we can find at Walmart.


 

DAPI in the Prolong Gold will cause an uneven staining gradient so that some
of the nuclei in the center of a section are not stained as brightly as the
nuclei on the outer edges of a stained section.  The cause is not getting
enough thicker Prolong Gold/DAPI over the section or not having just the
right amount of buffer on the section to permit a good flow of this
wonderful mounting media over the section.We now complete all IF
staining then stain with a DAPI solution before cover slipping with Prolong
Gold.   You can buy ready to use DAPI solutions from Pierce or Biogenex, or
make up the solution in house.   You can find the recipe at IHCworld website
or simply Google.  

 

We do NOT store our IF stained slides in the cold, but in a folder at RT in
a dark drawer before viewing on the day after staining to reduce any
movement/flow under the coverslip.   Fluorophores can and will eventually
fade.   I do not recall any studies saying storing IF stained slides in the
cold reduces fading but we never have space to do cold storage anyway and
store slides at RT.The new fluorophores (Alexas and Dylights) remain
stable over a longer time even for several weeks compared to fluorescein
derivatives e.g. FITC TRITC.  

 

Gayle M. Callis

HTL/HT/MT(ASCP)

Bozeman MT   

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RE: [Histonet] saffron vs. safran du gatinais

[gayle callis]

Beth, 

In the past, people replied to Histonet with the suggestion to buy saffron
aka safran du gatinais from a grocery store spice section.   Depending on
where you are located e.g. a bigger city, try to find a store that sells
spices from India may have the freshest saffron.   However, you won't suffer
the sticker shock of buying it from a chemical supplier.  It still tends to
be expensive but not like chemical company prices.Once we made the
alcoholic saffron solution for Movat's pentachrome, we stored this solution
in a container with a dessicant to maintain a water free environment.   

I was fascinated that Tom actually grew and harvested saffron from the
flowers.   That is true devotion, but I suspect it was for those delicious
sounding nubbies .   

Good luck

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

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Villarreal,
Beth
Sent: Thursday, January 05, 2012 1:02 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] saffron vs. safran du gatinais

Hello histonet,
I have a protocol that calls for safran du gatinais and am experiencing some
serious sticker shock.  Can I substitute saffron in my solution or am I
asking for trouble?

Many thanks,
Beth


Beth Villarreal
Scientist I
Novartis Institutes for BioMedical
Research, Inc.
300 Technology Square
Cambridge, MA 02139
USA


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RE: [Histonet] Re: Histonet Digest, Vol 97, Issue 21

[gayle callis]

This same clone for Rat CD31 still sold by BD 
Biosciences/Pharmingen/Invitrogen, also Serotec, ABCAM and a  company called 
Research Diagnostics found in the IHC World protocol.   The latter protocol 
used Proteinase K digestion.   

Gayle Callis 

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu 
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Madeleine Huey
Sent: Sunday, December 18, 2011 9:19 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Re: Histonet Digest, Vol 97, Issue 21

Colleen,

Many years ago I used Pharmingen's  anti-Rat CD31, clone TLD-3A12 (I think, you 
need to check it out if they still sell it).

This antibody is not recommend for formalin fixed paraffin tissues, but it will 
work.

Following steps are critical;
1) Do not use HIER, use enzymatic digestion (need optimization with Trypsin or 
Proteinase K @ 37c.  Lot to Lot variation from manufacture)
2) Overnight Incubation @ RT, not 4C (need warmer  long time)
3) Do not over dilute the 1st ab (ie. 1:10 - 1:25)
4) Try use a more sensitive detection system (Leica Refine DAB system) Good 
Luck!
Madeleine Huey BS, HTL (ASCP) QIHC
Supervisor-Pathology, IPOX  Histology
El Camino Hospital
madelein...@elcaminohospital.org


On Sun, Dec 18, 2011 at 10:00 AM,
histonet-requ...@lists.utsouthwestern.edu wrote:
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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. Re: Histonet Digest, Vol 97, Issue 20 (Madeleine Huey)
   2. RE: CD31 for rat tissue (Patsy Ruegg)
   3. e-cadherin (Patsy Ruegg)
   4. RE:   mouse antiRat CD31 (gayle callis)


 --

 Message: 1
 Date: Sat, 17 Dec 2011 13:28:52 -0800
 From: Madeleine Huey madeleineh...@gmail.com
 Subject: [Histonet] Re: Histonet Digest, Vol 97, Issue 20
 To: histonet@lists.utsouthwestern.edu
 Message-ID:

 CAF2e4C+Xe45xiKcGK0D_4HcEN2kMbNT=l67w2rekln_unks...@mail.gmail.com
 Content-Type: text/plain; charset=UTF-8

 Magan,

 Your problem is very simple.  First of all, you do not need Harris 
 Hematoxylin after IHC, because Harris Hematoxylin is a regressive 
 hematoxylin.  What you need is a progressive hematoxylin, like Mayer, 
 Gill (I, II, III)  etc.

 Try this simple procedure after your DAB chromogen;
 1) Counterstain in Gill I (Sigma) or Mayer (American MasterTech) for
 0.5 - 1 min (longer if want darker counterstain, or use Gill II/III.
 Personal preferences)
 2) Wash off excess Hematoxylin with tap water (no need distilled 
 water, your experiment is done)
 3) Blue the Nuclei with PBS or TBS buffer (common buffers used by IHC)
 4) Wash with water
 5) Dehydrate  cover slip with permanent mounting

 You can write or call me if you still need help or have any questions.

 Madeleine Huey BS, HTL (ASCP) QIHC
 Supervisor-Pathology
 El Camino Hospital
 Mountain View, CA
 madelein...@elcaminohospital.org

 On Sat, Dec 17, 2011 at 10:00 AM,
 histonet-requ...@lists.utsouthwestern.edu wrote:
 Send Histonet mailing list submissions to
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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. 2% low melting agarose gel (awatan...@tgen.org)
   2. Re: 2% low melting agarose gel (Emily Sours)
   3. AUTO: Ramona Nelson is out of the office. (returning
  01/03/2012) (ramona_nel...@bd.com)
   4. Formalin cost account (Morken, Timothy)
   5. Looking for a used Biocare Decloaker (Sowmya Kedarnath)
   6. Re: Looking for a used Biocare Decloaker (Akemi Allison)
   7. CD31 for rat tissue (Colleen Forster)
   8. Re: CD31 for rat tissue (Lucie Guernsey)
   9. Diane Tokugawa/CA/KAIPERM is out of the office.
  (diane.tokug...@kp.org)
  10. paraffin recycler (Gudrun Lang)
  11. Re: paraffin recycler (Rene J Buesa)
  12. Re: DAB haematoxylin counterstain;too purple, 
 overpowering
  IHC (Maxim Peshkov)
  13. Re: New Lab (mequita praet)


 -
 -

 Message: 1
 Date: Fri, 16 Dec 2011 18:12:01 +
 From: awatan...@tgen.org
 Subject: [Histonet] 2% low melting agarose gel

[Histonet] RE: mouse antiRat CD31

[gayle callis]

Several sources for mouse antiRat CD31 (PECAM) were found by doing a simple
Google search with keywords mouse antiRat CD31 immunohistochemistry.  You
can buy this monoclonal from Serotec, BD Biosciences and probably
eBiosciences plus other companies that specialize in rodent antibodies.   As
for protocol, IHC World had a mouse antiRat CD31 procedure that worked on
FFPE tissue with recommended retrieval - certainly worth a  try with your
reagents.Be sure to check application e.g.  IHC on FFPE or frozen
sections for any particular company's technical data sheet before buying the
antibody.   They sometimes only test on frozen sections or with the formalin
free Zinc Tris Buffer (Becksteads  ZSF fixative) fixed tissue for paraffin
sections ala BD Biosciences Pharmingen.  

 

As for working concentration, one should always do a dilution panel since
your laboratory conditions and reagents will never be the same as in someone
else's laboratory.   We start our dilution panel at 10 ug/ml for solvent
fixed fresh tissue frozen sections, and 20ug/ml for FFPE, and often fill in
a wide gap if doing a serial dilution.   The gap could be 1:500 then 1:1000,
so we toss in a 1:750, and sometimes a 1:1500.   

 

Gayle M. Callis

HTL/HT/MT(ASCP)

 

 

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[Histonet] Re: Woelckes Myelin Sheath method

[gayle callis]

You wrote:  

 

I hope everyone had a great Thanksgiving!  I was wondering if anyone has

any experience with the Woelcke Myelin Stain.  A researcher is trying to

use it to identify damaged neurons in brain sections.  I have found one

article with a rough procedure, but any other help anyone has would be

great!  Thanks!

 

Erin Sarricks



 

Erin,  

 

If you contact me, I will personally send you the procedure.   It was called
Lee Luna Woelcke Myelin Sheath method in a publication I found.  I will scan
the method and send to you in pdf format IF you email me.  Pretty simple to
do but hopefully John Kiernan, a myelin staining expert will make
suggestions too.   He wrote a fantastic review of myelin staining about 6
years ago that is classic tutorial on the subject.   This was published in
Journal of Histotechnology, and can be accessed online by NSH members free
of charge.  If you go to Google Scholar, you can pick up this publication
Cereb. Cortex (1997) 7 (2): 166-177. doi: 10.1093/cercor/7.2.166   Or merely
cut and paste the whole DOI link into Google to get free pdf/full text
article.   

 

 

Gayle Callis

HTL/HT/MT(ASCP)

Bozeman MT 

 

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[Histonet] RE: DAPI on thicker sections, problem with gradient

[gayle callis]

Carmen, 

 

You wrote: 

 

Good morning:

 

I am Carmen Garcia from the Faculty of Medicine of the University of 

Valencia, Spain. I usually use your vectashield mounting medium for 

fluorescence with DAPI (H-1200) and I never have any problem.

But now I am working with 50um frozen sections and the problem that I 

have is that the DAPI staining does not arrive to the bottom of the 

section. Do you have any sugesstion of what can I do? (I already have 

the sections permeabilized with triton x-100) I have thougth to 

incubate with the mounting medium some hours at 4ºC or put some 

mounting medium in the antibody diluent and incubate...

 

any suggestion would be really welcome.

Thank you so much,

Kind regards,

 

Carmen

 


***

The problem you are experiencing is what we refer to as a DAPI staining
gradient.  Uneven staining occurs with the DAPI is not reaching all the
cells due to viscosity of the mounting media and thickness of the tissue
section.  

 

You did not say whether your thick sections have been fixed before
sectioning and staining for a tissue component, and then the DAPI - please
explain.  I suggest you do NOT use a mounting media with DAPI, but rather
make a DAPI solution which you can then apply to the section and incubate
mounting the coverslip with Vectashield that does not contain DAPI.  Using a
DAPI solution may shorten the time you now use, e.g. some hours.You
can buy the DAPI, ready to use from Biogenex or other vendors, but making it
in house is simple. I would assume you could add Triton X to the DAPI
solution  to improve permeability and match what you have already done with
that detergent.  After DAPI staining, mount the coverslip using  VectaShield
without DAPI.  DAPI recipes can be found on the internet, just Google DAPI.
I found it on a Johns Hopkins webpage, but IHC world may have it too.
There are other sources of ready to use DAPI if you look.  

 

We have used Prolong Gold antifade reagent with DAPI,  and get a staining
gradient on 5 um sections.   This is very annoying, and probably caused by
the actual technique of cover slipping itself. We are now going to use
Biogenex ready to use DAPI before the Prolong Gold antifade reagent without
DAPUI.   Staining for a thin section takes 5 mintues at RT, so you should
try several 50 um  sections (non experimental) to determine your optimal
staining time.   It may take only 15 minutes although cold, 4C temperature
may slow down the penetration of the stain through the thicker section.
This way you can achieve even staining of your nuclei without long
incubations in the more viscious mounting media containing DAPI. 

 

Good luck, and let us know if you have success...

 

Gayle M. Callis

HTL/HT/MT(ASCP)

 

 

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RE: [Histonet] Tissue left in processor

[gayle callis]

Some prefer to take the cassettes off the processor and let them harden
until embedding can be done, rather than leave in molten, hot paraffin.
This does not damage the tissue, and placing the cooled cassettes into the
embedding center holding area allows the paraffin to re-melt in a shorter
time before embedding.  This has been discussed in the past on Histonet.
Some tissues may dry out even more WITH difficult sectioning after prolonged
heat exposure.  One result will be a parched earth effect seen in sections.
I see you are from a veterinary facility, and if you work with rodent
tissues e.g. spleen and liver, you should not let the tissue sit in paraffin
or you will have little hard rocks to section.If your sectioning suffers
(dry, friable, shattered, hard)  after allowing tissues to sit in hot
paraffin as compared to the times when you embed asap after processing is
finished, then over exposure to hot paraffin can contributing factor.   

I personally do not like to cook my tissue any longer than necessary and
heat labile antigens will also be at risk.  I schedule so I can be there to
embed when the processing is completed. 

Our standard is to embed when processing is finished and schedule
accordingly.  

Gayle Callis
HTL/HT/MT(ASCP)

   

-Original Message-
From: histonet-boun...@lists.utsouthwestern.edu
[mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Janice
Mahoney
Sent: Thursday, October 06, 2011 9:45 AM
To: rjbu...@yahoo.com; histo net; rchar...@pa.gov
Subject: RE: [Histonet] Tissue left in processor


I agree with Rene, as long as the temp is only a few degrees above the
melting point of the paraffin.Jan,Omaha

 Date: Thu, 6 Oct 2011 07:13:54 -0700
 From: rjbu...@yahoo.com
 To: histonet@lists.utsouthwestern.edu; rchar...@pa.gov
 Subject: Re: [Histonet] Tissue left in processor
 CC: 
 
 I do not think that a well fixed, well processed tissue left in molten
paraffin for 2 hours after the processor finished will have any adverse
outcome.
 René J.
 
 --- On Thu, 10/6/11, Charles, Roger rchar...@pa.gov wrote:
 
 
 From: Charles, Roger rchar...@pa.gov
 Subject: [Histonet] Tissue left in processor
 To: Histonet (histonet@lists.utsouthwestern.edu) 
 histonet@lists.utsouthwestern.edu
 Date: Thursday, October 6, 2011, 10:05 AM
 
 
 Hi All,
 Is there any standard on how long tissue cassettes can remain in the
processor after processing before the tissue is subjected to unwanted
outcomes?  And if so  what type of artifacts can one expect from tissue that
was in the processor in molten paraffin for 2 hours after the processing was
completed?
 Thanks so much.
 Roger
 
 Roger Charles| Microbiologist II
 Pennsylvania Veterinary Laboratory
 2305 North Cameron Street | Harrisburg, PA 17110
 Phone: 717.787.8808 | Fax: 717.772.3895 
 www.agriculture.state.pa.ushttp://www.agriculture.state.pa.us


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RE: [Histonet] So long, and thanks for all the help

[gayle callis]

Dear Adam, 

 

Thank you for the kind words.   It has always been a pleasure to pass histo
information onto others to help them with their projects.   As for dinner
and a drink, I would take you up on that - hopefully with Andrea there too.
In lieu of that, I raise my morning cup of coffee to you in congratulations
on your PhD, a huge achievement.   I would love to know the title of your
thesis and you can email that via private email.   I wish you great success
and hopefully a return to the Histonet network in the near future.  

 

Take care, invest in a good coffee maker to ward off sleep and enjoy your
medical studies.  

 

Gayle Callis   

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[Histonet] Charcoal filters for VIP1000 K series

[gayle callis]

There are two charcoal filters (#6160) available for a Sakura Finetek VIP
1000, K series tissue processor. These are free for the lab  who can use
them.  Contact Maria Jerome [mjerome.mt...@gmail.com] for shipping
arrangements.   She is not on Histonet.  

 

 

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[Histonet] Re: Knife for trimming paraffin from blocks

[gayle callis]

You Wrote: 

 

I am looking for a stainless steel knife that we use to scrape off the 

paraffin from the embedded blocks.  I could not find the 6 inches size 

knife in any catalog. 

 

Thanks in advance

 

Mala

 

Nirmala Srishan

Histology Supervisor

Holy Name Medical Center.

 

*

Dear Mala, 

 

In the past, we used an old style permanent edge scalpel blade but after a
disaster with one person seirously cutting himself while trimming paraffin
from a block, we purchased a Paratrimmer with a heated, slanted metal
surface.  This trimmer has made everyone happy by eliminating the potential
for serious injury and does an even better job of getting rid of excess
paraffin. 

 

There are two models sold, one from Thermo Scientific and the other one was
recently spotted on a vendor website. (Sorry, I didn't jot the name down).
The trimmers  are worth the investment to keep you and your employees safe
from nasty cuts, no matter what the knife/blade, etc could be used.  

 

The only drawback to the Para Trimmer is the messy paraffin drippings have
to be collected in some separate container that the manufacturer doesn't
supply.  This is more than a bit annoying.  It seems to me the manufacturer
of these devices would design the trimmer with a paraffin catch tray.  A
small aluminum baking pan (from grocery store) could be used and disposed of
(not a Green consideration).

 

Be safe rather than sorry.  

 

Gayle M. Callis 

HTL/HT/MT(ASCP) 

 

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[Histonet] Re: 14% EDTA

[gayle callis]

There is no vendor I know of who sells 14% EDTA.  It is important to
remember that the EDTA you have to use is Tetrasodium EDTA, MW 452.2.   This
particular EDTA is highly soluble in water compared to EDTA without any
sodiums, or disodium EDTA.  Also, once dissolved it has a very high pH of
approximately 9 or more and you must adjust the pH DOWN to pH 7.2 - 7.4,
although we use pH 7.6 without problems.  If you do not adjust the pH down,
you will damage alkaline sensitive protein linkages in your tissues/bone.
Do not use HCl to adjust the pH, but glacial acetic acid.   The publication
supporting this EDTA decalcification was written by Dr. Webb Jee many years
ago.  We prefer to use tetrasodium EDTA because we can achieve a high
concentration with the highly soluble EDTA. 

 

You can buy tetrasodium EDTA if from any vendor, ThermoScientific, VWR, etc.


 

Simple recipe is

 

14 g Tetrasodium EDTA mw 452.2 (this may vary) 

80 ml Dulbeccos PBS or water (the latter is fine IF your bones are totally
fixed)

Adjust pH DOWN to 7.4 with glacial acetic acid (it takes a fairly large
volume)

Bring final volume to 100 ml with either buffer or water, depending on what
you dissolved the salts in. 

 

We adjust pH using constant stirring and pH electrode in the stock solution
to watch the pH change.  Keep track of final volume of glacial acetic acid,
so the next time you make this up, you can add a large volume without having
to keep adding glacial acetic acid with a pipette (in sense, a titration to
correct pH). 

 

As with all EDTA decalcification, it takes longer and you must rinse the
bones well for a couple of hours or the EDTA will ppt out in alcohols,
creating some annoying sectioning problems.   

 

Suspend your bones in this solution and use endpoint testing to know when
calcium is removed.  We use a simple weight loss weight gain endpoint test
that works very well for EDTA unless you have an xray machine or microCT
scanner to detect Ca removal.  EDTA is not affected by temperature, and can
be performed in the cold although room temperature works perfectly.  

 

Gayle M. Callis

HTL/HT/MT(ASCP)

   

 

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[Histonet] RE: Bluing

[gayle callis]

Dear All, 
 
We use commercial bluing reagents for convenience e.g. Richard Allan Bluing
Reagent ready to use and have used Scotts Tap Water Substitute Concentrate
diluted before use.  We do not use ammonia water anymore as the high pH can
cause section lift off from slide.  Here is recipe for Scotts. 
 

SCOTT'S TAP WATER SUBSTITUTE (You can make this up as a 10X concentrate if
you wish.  Store in house concentrate at 4C or growth will occur) 

Sodium bicarbonate2  g

Magnesium sulfate (anhydrous)  20  g

Tap Water   1 L

 
Peggy presented a good explanation of bluing.  We do not use tap water
anymore due to incorrect pH.   One of the failures for complete bluing is
not changing the bluing reagent daily with large volumes of slides. Lower
number of slides, one might be able to get by with fewer changes of bluing.
Tap water rinses dilutes the reagent and can change the pH, which should be
pH 8. If there is a large volume of slides, anything over 75 or more, we
change the bluing reagent daily. It is important to remember that Clarifiers
e.g. acetic acid reagents used with progressive hematoxylins (Richard Allan
Hematoxylin 1, Gill 1,2,3) will suffer from tap water rinse dilution too. We
also change those, and if Clarifier looks rather yellow, then you have
probably exhausted the acetic acid and its effectiveness.  Jerry Fredenburgh
has given excellent workshops on hematoxylin staining in the past and passed
on many of these facts. Consequently, we have no clarifying or bluing
problems.  
 
Fredenburgh recommendations for Richard Allan Hematoxylin 1 staining
procedure that we use: 
Hematoxylin 1 - 1.5 min, longer if needed
Tap water rinse-1 minute
Clarifier - 1 minute
Tap water rinse - 1 minute
Bluing - 1 minute
Tap water rinse - 1 minute
 
Remember, static or non-running water rinse stations are contaminated with
carryover hematoxylin, clarifier and/or bluing reagent.  If you do NOT
remove the bluing reagent with a good rinse, then eosin staining is affected
by cations carried over from the bluing reagent.  We also have an alcohol
step (matching % of alcohol in eosin) before going into this stain.  This
helps remove the cations plus prepares the section for eosin staining.
Perform good rinses and change static water rinse stations after a run, or
frequently during a day of heavy staining.  The second and third racks of
slides on a stainer should have the same tap water rinse conditions as the
first rack.   Yes, extra work but you need to ensure consistent staining
results of HE of ALL sections- the universal workhorse stain we depend on
so heavily. 
 
Years ago, Richard Allan had an excellent manual for HE staining guidelines
on how to perform and adjust your HE staining.  Sadly the manual is no
longer published but the good news is ..  I saved my copy, and scanned
it to pdf if anyone wants it.  
 
Gayle M. Callis
HTL/HT/MT(ASCP)
Bozeman MT 
 
 

*
Peggy Wenk wrote: 
 
It might just need longer time in the bluing agent.
 
Hematoxylin solutions stain the nuclei reddish (look at your slide right 
after coming out of hematoxylin - tissue will look reddish). The alkaline 
solutions that are used for bluing remove a H+ group on the aluminum 
hematein (that's the staining chemical of the hematoxylin solution), and 
change it to a -OH group. This changes the aluminum hematein from a reddish 
color to a blue color.
 
The more alkaline (higher pH) the bluing agent, the faster this reaction and

color change, and the less time is needed in the bluing agent. The lower the

alkalinity (not as high pH but still in the alkaline range) the bluing 
agent, the slower the reaction and color change, and more time is needed in 
the bluing agent. Dilute ammonia water usually only takes a few seconds, as 
it's pH is usually high (pH 10). Tap water can have a pH of around 7, and 
may take 5-10 minutes to blue. If the tap water is more acidic (pH 5 or 
below), the slides may not blue.
 
Now, we have to get all the nuclei from the reddish color to the bluish 
color. If the slide is not in the bluing agent for enough time (for the 
type/pH of bluing agent), then some of the nuclei change to blue, while some

still remain reddish (or within a single nucleus, some of the DNA has 
changed to blue, some remains reddish), hence a more purple color.
 
So, the easiest thing to try right now is to stain 2 slides from the same 
block (serial sections) in your hematoxylin for the same time, put one in 
the bluing agent for the usual time, put the other in the alkaline solution 
for extended time, and see if the nuclei on the second slide are now more 
blue than the first slide.
 
Let us know.
 
Peggy A. Wenk, HTL(ASCP)SLS
Beaumont Hospital
Royal Oak, MI 48073
 
-Original Message- 
From: Webb, Dorothy L
Sent: Friday,