[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
 ]

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

To: histonet at lists.utsouthwestern.edu
 

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]

You wrote: 

Hi all,

 

I am going to be exploring some tissue embedding using Glycolmethacrylate

(GMA), as I read that it can preserve enzyme function in some cases and

potentially can be used for IHC. However, information is a bit scant and

can be contradictory at times, to say the least.

 

If anyone has any experience doing any histological or IHC staining on GMA

embedded tissue, or knows of anywhere I could get protocols from or any

papers that are a bit more recent (I'm mostly finding 90s, early 00s),

pleas let me know, I would really appreciate any resources I can find!

 

-- 

Casey Berridge

 


**

GMA references from the 80’s and 90’s pretty much remain current for GMA.
Be sure to do a literature search in J Histochem Cytochem (JHC.org) to see
if there are more recent publications.  Google Scholar was not turning up
much for recent years.

 

We worked with GMA for many years but never for IHC.   There are many
considerations when using GMA both good and bad with emphasis on the
negative side of things from my point of view.   We had success when
studying some single cell protozoa, including Cryptosporidia in a research
setting.   

 

There is one publication in the old Stain Technology, now Biotechnic &
Histochemistry using GMA for successful enzyme staining.  Namba M et al.
Improvement in histochemical demonstration of esterase in glycol
methacrylate tissue sections by cold temperature embedding in glycol
methacrylate. 1983 58(4):207.

 

Several things about GMA. 

 

Requires a fume hood in order to work with toxic and carcinogenic chemicals.
Glycol methacrylate is sensitizing and several colleagues are so allergic to
fumes after working with this plastic over several years, they can’t be in
the same room where GMA is being worked with.  Double gloving is advisable,
and wearing safety glasses is a must since the sections are small and can
fly into an eye (know of this happening) which is not a good situation.
There should be no skin contact with the plastic, nor breathing the n, n, di
methylaniline, a carcinogen.Controlling polymerization can be a problem
unless you place embedding molds on top if ice, and cooling the embedding
mixture with ice water.   The polymerization is exothermic, and actually as
blocks polymerize gets uncomfortably hot which may be damaging to enzymes
and sensitive antigens although the heat can be dispersed.   

 

Samples cannot be any thicker than 2 mm, with 1 mm X 1 mm is recommended.
This plastic was first used for liver needle biopsies.Polymerization for
larger samples is hard to control as is the infiltration by this plastic
hence smaller, thinner samples.   Sectioning is commonly done with glass
knives although tungsten carbide knives work, and I know of one group using
disposable blades with a Leica 2255 model microtome.More powerful
microtomes i.e. Leica 2650 or equivalent works best.  We had a JB-4
microtome with a special block holder to accommodate the metal
“chucks”/block holders sold by Polysciences.   The metal block holders were
a better heat sink to disperse heat of polymerization. Sections are
generally no thicker than 1 to 3 µm, and were wonderful when studying single
cell protozoa.  We never used GMA for more routine tissue sections although
it was popular for bone biopsies in clinical labs over the many years.   I
personally found it labor intensive, and expensive for our projects although
the staining results for H, PAS-H and some other special stains very nice.


 

Routine stains can be used, including PAS-H, H, Massons trichrome with a
modified method, and others.   IHC will not work well, even with JB-4
Immunobed.  GMA, once polymerized, cannot be removed from the section.
Immunobed is probably just a looser matrix than JB-4 and some people have
success.   GMA  plastic is less hydrophobic but still will not allow  large
immunoglobulins to reach antigenic sites.   There has been some success with
IHC but in general, GMA is not the ideal embedding media for immunostaining.
Neil Hand worked with Poly methylmethacrylate for IHC since the plastic can
be completely removed from a thin section, followed by stringent HIER using
a pressure cooker.   PMMA is another world for processing,  sectioning and
staining.  

 

When doing H, the staining protocol is different from paraffin section
staining.   If you do an extensive, time intensive search on Histonet, there
are many discussions about GMA staining both for routine and IHC.  

 

You can buy kits, JB-4 and Technovits.   The JB-4 discolors over the years
to a dark tea/brown color making it more difficult to see the tissue while
Technovits remains clear.   When you cut sections, you work with one section
at a time, not a ribbon.   

 

I have a huge file on GMA collected from the early 70’s all the way to
current years.  If you reply to me personally, I 

[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 ; Tim H 
Cc: '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]

Hey Amos, 

 

I will be nice!   It is true peroxidase blocking can be a pain.  However, after 
looking at the PeroxAbolish SDS from Biocare, I did not SEE any of the 
chemicals involved in this block  so I am skeptical this is the Glucose oxidase 
method.  Big Ho Hum!   PeroxAbolish could be similar to a KPL peroxidase block 
from years past but not sure if  KPL still sells it.  KPL block was not an 
enzyme mixture and had to be used carefully on frozen sections.   Also, Biocare 
said nothing about blocking pseudoperoxidases but neither did Vector in their 
method although the latter did provide the reference which I encourage people 
to read.   The glucose oxidase method was developed for delicate, minimally 
(acetone) fixed frozen sections and really does work without H2O2 chewing a 
delicate FS off the slide or damaging morphology.The bonus:  glucose 
oxidase method works on FFPE tissues particularly when an  in house H2O2 method 
fails to remove persistent background caused by peroxidase or pseudoperoxidses. 
  It could be that Patrick is actually getting rid of peroxidase but NOT 
pseudoperoxidases with his H2O2 method?   Glucose oxidase is a good old fashion 
enzyme/substrate chemical reaction:  glucose oxidase +   β D (+) glucose = 
production of slow steady rate of hydrogen peroxide to remove 
peroxidase/pseudoperoxidases in tissues and cells.   It  gave us the cleanest 
background ever for murine CDmarker/HRP- IHC/solvent fixed frozen sections 
using a DAB enhancer.  We never could  get rid of endogenous peroxidase 
background with a commercial  low concentration H2O2 peroxidase block.   This 
is  another long story.  

 

There is no mystery about the protocol and Vector has this on their website 
(see below)   

 

CAVEAT:   the β D (+) Glucose, 97% pure can be hard to find but here is the 
source.   MP Biomedicals , beta-D-Glucose Catalog Number: 100953.  Sigma 
discontinued this glucose some years ago (heavy sigh!).   I will be happy to 
provide the original publication for this method under separate email.   



Glucose Oxidase Peroxidase Block (GLUOX),  *Jasani et al: 

 

Working Solution

β D (+) Glucose, 97% pure (MP Biomedicals Cat. No. 100953)
0.180 g  (180 mg)  

Glucose oxidase (Sigma G 6641)  
  0.005 g  (5 mg) 

Sodium azide
   0.0065g  (6.5 mg)

Dulbeccos PBS   
  50 ml

Do not preheat working solution.  

 

Protocol:

1. Immerse sections in working glucose oxidase solution 30 min - 1 hr 
(per cited reference).  Use a 37°C water bath for even heating.  

 2.After incubation, rinse sections 3X in DPBS, 5 min per change then 
proceed to IHC.



Results:  Clean background without endogenous peroxidase or pseudoperoxidases 
in minimally fixed (acetone) FS or FFPE. 

 

NOTES:  A.   Glucose oxidase can be weighed into screw top micro-centrifuge 
tube and stored in a freezer.  Pay attention to the expiration date and storage 
conditions.  This way, one can make up a  ready to use stock buffer in large 
quantity i.e. 2 liters or more.  We called this "GLUOX Buffer" due to sodium
azide content.  Weighing out chemicals each time was painful to say the 
least.   A stock buffer permitted quick preparation for a desired working 
solution volume  e.g. 50 ml or more.  To prepare desired quantity of working 
soluton:  pipette 500 µl GLUOX buffer into micro-centrifuge tube containing 
  pre-weighed glucose oxidase, vortex mix to dissolve,  add mixture remaining 
volume of buffer, stir, pour into staining container/coplin jar.   

 B.  Use a 37°C water bath for even heating, NOT AN INCUBATOR 
with air currents causing uneven heating.  

Reference:*Andrew SM, Jasani B.  An improved method for the inhibition of 
endogenous peroxidase non-deleterious to lymphocyte surface markers.  
Application to immunoperoxidase studies on eosinophil-rich tissue preparations. 
 Histochem J 19:426-430, 1987

 

FYI:  Vector has this method in a free downloadable pdf brochure;  
http://vectorlabs.com/brochures/   Multiple Antigen Labeling Guide 
p.20/Appendix 2/Method 3  

I took the liberty of copying this from Vector who used mg 
terminology. 

Method 3. 0.180 g b-D(+) glucose, 5 mg glucose oxidase, 6.5 mg 
sodium azide in 50 ml PBS.

Incubate sections for 1 hour at 37 ºC. Rinse in PBS 3 x 5 minutes. 
This reaction slowly and steadily produces very low concentrations of H2O2 by 
enzymatic reaction. This method consistently and completely inhibits peroxidase 
activity.

(Andrew S.M., Jasani, B.; Histochem J. 19, 426-430,1987.) 

 

Final comment.   If Patrick still has background staining after using Glucose 
oxidase method, then I suspect 

[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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