Re: [Histonet] cloudy cornea after Hartman's fix

[John Kiernan via Histonet]

The cornea consists of cells (nuclei with plenty of cytoplasmic protein) and 
parallel bundles (lamellae) of collagen fibres, also protein. It is wonderful 
that the normal cornea is transparent.

Doesn't any fixative make the cornea opaque?
Davidson's (=Hartmann's) fixative contains plenty of formaldehyde (to add to 
and slowly crosslink all protein molecules, including collagen fibrils) but 
perhaps not quite enough ethanol to rapidly coagulate soluble proteins in 
cytoplasm.
(Davidson's 22.2% v/v EtOH is slightly more than in sherry, madeira or 
vermouth; about half of that in whisky, gin etc.)  The acetic acid (about 10%) 
can be expected to make nuclear DNA molecules and their associated proteins 
shrink into the patterns we see as typical of different cell-types.

Most of the many formalin-acetic-alcohol fixatives contain enough ethanol to 
bring about protein coagulation before the more slowly reacting formaldehyde 
does its stuff. As Tony points out, more research is needed.

John Kiernan
https://www.schulich.uwo.ca/anatomy//people/faculty/emeriti/kiernan_john.html
= = =

From: Tony Henwood via Histonet 
Sent: June 27, 2024 7:17 PM
To: histonet@lists.utsouthwestern.edu ; 
Davoli, Katherine A 
Subject: Re: [Histonet] cloudy cornea after Hartman's fix

Hartman's (also known as Davidson's) fixative is sometimes used to reveal lymph 
nodes in resections where they appear opaque - white. It is not surprising that 
the tissues of the eye would react the same.
I assume that the alcohol causes the bleaching of the tissue (or is it the 
acetic acid?) - more research needed.

Regards,

Tony Henwood MSc, BAppSc, GradDipSysAnalys, CT(ASC), FFSc(RCPA) (Retired)
Principal Scientist, the Children’s Hospital at Westmead (Retired)
Adjunct Fellow, School of Medicine, University of Western Sydney.

From: Davoli, Katherine A via Histonet 
Sent: 28 June 2024 04:12
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] cloudy cornea after Hartman's fix

Anyone know why the cornea of my pig eye got white/cloudy on dropping it in to 
Hartman's fixative?  I'm used to working with mice where, if this happens I 
didn't notice.

Katherine Davoli, MDiv, HTL(ASCP)cm(they/them/theirs)
Lab Manager, Tissue Culture & Histology Cores, U. Pitt Dept of Ophthalmology
7.373 UPMC Mercy Pavilion1622 Locust St., Pittsburgh PA 15219
(412) 624-8508   this number cannot receive texts
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Re: [Histonet] Causes of false positive Congo Red

[John Kiernan via Histonet]

Dear Greg,

This is the same as on p.126 in my copy of Carson's 2nd edition (1997) and also 
in the 11th and last (2008) edn of Churukian's "Manual of the Special Stains 
Laboratory". The only stock solution that can be expected to change with time 
is the Stock Congo red solution, because solutions of dyes with large anions 
are unstable in the presence of inorganic salts. Churukian (p.195) said it 
could be kept for 2 months.

The correct staining you got with newly made solutions suggests that your 
earlier stock Congo red stock solution was too old. Evidently you solved the 
problem yourself!

In their "Troubleshooting Histology Stains"  book, Horobin & Bancroft (1998, 
p.45-47) stressed the need for a freshly made dye solution. They also suggested 
ignoring "pink background" and checking that it's not dichroic. They also 
listed various Congo-positive and dichroic materials that aren't amyloid. An 
unidentified yellow compound is often present in Congo red and it may sometimes 
cause generalized yellow background staining.

John Kiernan
https://www.schulich.uwo.ca/anatomy//people/faculty/emeriti/kiernan_john.html
= = =

From: Greg Dobbin 
Sent: June 20, 2024 8:44 AM
To: John Kiernan ; histonet@lists.utsouthwestern.edu 

Subject: Re: [Histonet] Causes of false positive Congo Red

You don't often get email from greg.dob...@gmail.com. Learn why this is 
important

Hi John,

I must apologize again. We used to use the method from Carson's book. We now 
make up the reagents as follows:


Stock alkaline salt solution

Sodium chloride. 2g

Distilled Water... 20mL

Stir until the salt is dissolved, then with continuous stirring on a magnetic 
stirrer add

80mL of 100% denatured ethanol.

Some salt may precipitate out after the ethanol is added.



Working alkaline salt solution

Stock alkaline stock solution...50 ml

1% Sodium Hydroxide0.5ml

Filter and use within 15 minutes


Stock Congo red solution

Congo red 0.1g

Stock alkaline salt solution 50mL

Stir well with the magnetic stirrer and let stand overnight or for a minimum of 
3 hours if the slides need to be ready the same day that the order was placed.



Working Congo red (Congo red)

Stock Congo red.. 50ml

Sodium hydroxide 1%... 0.5ml

Filter and use within 15 minutes.

On Thu, Jun 20, 2024 at 9:31 AM Greg Dobbin 
mailto:greg.dob...@gmail.com>> wrote:
Good day John,
Very nice to hear from you again! I have been consulting your textbook in my 
investigations!
Sorry about the brevity of the description of our method. I felt like my post 
was already too long
and it might scare off some would-be contributors! :-) And yes, I incorrectly 
referred to the dichroic green as "fluorescent"-thank you.

Our method follows the Puchtler method described on pages 132-3 in Frieda 
Carson's "Self-Instructional" textbook (1990) as does
the hospital that repeated our false-positive Congo Reds. Note, once we re-made 
our reagents, our results returned to accurate staining.
Greg

On Thu, Jun 20, 2024 at 2:49 AM John Kiernan 
mailto:jkier...@uwo.ca>> wrote:
Greg, your method is incompletely described in your Histonet post, but it looks 
quite different from the "traditional" Highman's procedure (Arch. Path. 
41:559-562). What method were they using "at another
lab" to get correct red amyloid that is green (dichroic, not fluorescent) with 
crossed polars?
John Kiernan
https://www.schulich.uwo.ca/anatomy//people/faculty/emeriti/kiernan_john.html
= = =

From: Greg Dobbin via Histonet 
mailto:histonet@lists.utsouthwestern.edu>>
Sent: June 19, 2024 8:53 AM
To: histonet@lists.utsouthwestern.edu 
mailto:histonet@lists.utsouthwestern.edu>>
Subject: [Histonet] Causes of false positive Congo Red

Hello experts,
*Some background:*
I know that Congo Red can bind nonspecifically to non-amyloid components
such as collagen and elastin under certain conditions (eg Carnoys fixative,
insufficient differentiation, insufficient alkalinity, etc). However,
everything I have been able to read on the topic suggests that
over-staining is "easily" differentiated from true amyloid staining by
using polarizing light microscopy. That is, true amyloid produces apple
green fluorescence while non-amyloid components produce silver/grey color.

*My question:*
I want to know if anyone has encountered false positive staining that *is
apple green* in color? We had a few bone marrow core biopsies that stained
bright green but were later found to be negative when stained at another
lab. We subsequently threw out all of our working solutions and made up
everything fresh and repeated the previous (false positive) specimens and
they were indeed negative in our lab as well.

*In order to prevent this from happening again, I need to attempt to
understand what may 

Re: [Histonet] Causes of false positive Congo Red

[John Kiernan via Histonet]

Greg, your method is incompletely described in your Histonet post, but it looks 
quite different from the "traditional" Highman's procedure (Arch. Path. 
41:559-562). What method were they using "at another
lab" to get correct red amyloid that is green (dichroic, not fluorescent) with 
crossed polars?
John Kiernan
https://www.schulich.uwo.ca/anatomy//people/faculty/emeriti/kiernan_john.html
= = =

From: Greg Dobbin via Histonet 
Sent: June 19, 2024 8:53 AM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Causes of false positive Congo Red

Hello experts,
*Some background:*
I know that Congo Red can bind nonspecifically to non-amyloid components
such as collagen and elastin under certain conditions (eg Carnoys fixative,
insufficient differentiation, insufficient alkalinity, etc). However,
everything I have been able to read on the topic suggests that
over-staining is "easily" differentiated from true amyloid staining by
using polarizing light microscopy. That is, true amyloid produces apple
green fluorescence while non-amyloid components produce silver/grey color.

*My question:*
I want to know if anyone has encountered false positive staining that *is
apple green* in color? We had a few bone marrow core biopsies that stained
bright green but were later found to be negative when stained at another
lab. We subsequently threw out all of our working solutions and made up
everything fresh and repeated the previous (false positive) specimens and
they were indeed negative in our lab as well.

*In order to prevent this from happening again, I need to attempt to
understand what may have caused this to happen in the first place. *

This is where the vast collective knowledge of this group comes in. :-)
Can anyone offer some insight as to possible causes?

*Our Congo Red method:*


Deparaffinize sections and bring them to water.

Stain in Hematoxylin for 1 minute

Add 0.5ml of 1% Sodium Hydroxide to 50 ml of stock alkaline salt solution.

Wash slides in running water

Place in *working* alkaline salt solution from step 2 for 20 minutes

Add 0.5 ml of 1% Sodium Hydroxide to stock Congo red solution.

Start to filter *working* Congo red solution when 15 mins are left in step 6

Place sections in the *working* Congo red from step #8 for 20 minutes.

Dehydrate the slides one at a time in 3 changes of absolute ethanol, 6 dips
each.

Dip the slide 10 times in a coplin of xylene.

Continue dehydrating the other slides.

Coverslip the slides.

*Greg Dobbin*
1205 Pleasant Grove Rd
Route 220
York,  PE  C0A 1P0
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Re: [Histonet] What is the best xylene substitute for histology?

[John Kiernan via Histonet]

I liked terpineol (mixed isomers). Smells nice and doesn't harden tissues, but 
I see that it has now become pretty expensive.
Mineral oil USP (also called liquid paraffin and mineral oil, heavy) is OK and 
cheaper than xylene, but it's not miscible with ethanol. You have to dehydrate 
specimens in  isopropanol and have the oil warmer than 44oC.
Tert-butanol is miscible with water, ethanol and paraffin; it's somewhat more 
expensive than xylene and must be used warm because it freezes at 26oC. It's 
often used for plant histology.

A recently proposed safe and economical alternative clearing agent for paraffin 
embedding is coconut oil, but it causes much more shrinkage than xylene and 
also induces ugly artifacts (in rats' prostates) if the clearing time is less 
than 4 hours. See OA Bright et al. 2024. J. Histochem. Cytochem. 72(4): 
233-243. 
https://www.researchgate.net/profile/Ebenezer-Senu/publication/379429541_Clearing_Properties_Between_Coconut_Oil_and_Xylene_in_Histological_Tissue_Processing/links/661013eb2034097c54f61dbd/Clearing-Properties-Between-Coconut-Oil-and-Xylene-in-Histological-Tissue-Processing.pdf

Just a few thoughts!  John Kiernan.
London, Canada

From: Kate Bummer via Histonet 
Sent: June 7, 2024 10:52 AM
To: histonet@lists.utsouthwestern.edu 
Cc: Kate Bummer 
Subject: [Histonet] What is the best xylene substitute for histology?

Hello Histonetters

I'm hoping I can get some recommendations for the best xylene substitute for 
histology that could be used in the following:

  *   Tissue processor
  *   Deparaffinization
  *   Coverslipper
  *   Mounting media that works with that particular xylene substitute

Thank you for your help!

Kate
SeqMatic
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Re: [Histonet] Inquiry on Tissue Softening for Microtomy

[John Kiernan via Histonet]

Jay, you are 100% right about "pseudo-scientific mumbo jumbo" for the 
mechanism. But have you (or anyone else) compared ammonia-water with ordinary 
pure water for hydrating/softening?  Water (with or without dissolved ammonia) 
won't go through solid wax. The embedded tissue has to be exposed by cutting 
into the block.
Whiffs of ammonia are unpleasant. If they upset some people in the lab, that's  
a good reason to try odorless water instead.
John Kiernan
Emeritus, UWO, London, Canada
https://publish.uwo.ca/~jkiernan/
= = =
From: Jay Lundgren 
Sent: May 11, 2024 4:40 PM
To: John Kiernan 
Cc: histonet@lists.utsouthwestern.edu ; 
IGNACIO GONZÁLEZ MASSONI 
Subject: Re: [Histonet] Inquiry on Tissue Softening for Microtomy

I wouldn't say softening, I would say hydrating.  Ammonia water accelerates 
hydration of FFPE blocks.  Nobody knows how it works, it's a mystery.  Or at 
least I've never heard a scientific explanation.  Only pseudo-scientific mumbo 
jumbo like "facilitates the removal of paraffin" which is false.  Go soak a 
solid block of paraffin in pure NH4OH for 24 hours, it won't do anything.

I was told as a student at AFIP, "It opens the pores of the tissue so water can 
get in."  In other words, pseudo-scientific mumbo jumbo.

 It works though.  Somebody needs to get a $150,000,000 NIH grant and do a 
research project on how ammonia water hydrates tissue.  I worked with some 
lovely Hmong people in California that called it "crying water".

Using it can cause a big interpersonal problem with certain people in the lab 
though.  Interestingly, hypersensitivity to smells is one of the symptoms of 
autism.  Hypersensitivity to smells is also highly correlated with bipolar 
disorder and heightened emotional reactivity.  Sooo...

Jay A. Lundgren, M.S., HTL (ASCP)

On Fri, May 10, 2024 at 11:14 PM John Kiernan via Histonet 
mailto:histonet@lists.utsouthwestern.edu>> 
wrote:
   If you apply the ammonia to the cut surface of the paraffin block, I suspect 
that it softens the tissue in the same way as applying water: by entering 
interstices of the tissue that are not occupied by paraffin molecules.
   I never tried ammonia for this purpose but in the 1960s to early '70s I 
occasionally used a proprietary product called Mollifex, which I see is still 
sold. In 1972 or '73 an elderly technician told me that water was just as good, 
and I soon found out that he was right. Indeed, water had the advantage of 
working in 15-30 minutes rather than taking several hours.
John Kiernan
Emeritus, UWO, London, Canada
https://publish.uwo.ca/~jkiernan/
= = =

From: IGNACIO GONZÁLEZ MASSONI via Histonet 
mailto:histonet@lists.utsouthwestern.edu>>
Sent: May 10, 2024 8:53 PM
To: histonet@lists.utsouthwestern.edu<mailto:histonet@lists.utsouthwestern.edu> 
mailto:histonet@lists.utsouthwestern.edu>>
Subject: [Histonet] Inquiry on Tissue Softening for Microtomy

Dear friends at Histonet,

I hope this message finds you well. I am reaching out to seek your expertise on 
a matter that has piqued my interest in the field of histology.

I am currently delving into the process of preparing FFPE (formalin-fixed, 
paraffin-embedded) tissues for microtomy. Specifically, I am curious about the 
role of ammonia in softening these tissues before sectioning. From my 
understanding, ammonia serves as an alkaline agent that helps neutralize 
formalin's effects and facilitates the removal of paraffin, thus aiding in the 
softening of the tissue.

However, I would greatly appreciate if you could provide a more detailed 
explanation of the chemical interactions at play here. How exactly does ammonia 
interact with the tissue components to achieve the desired softening effect? 
Moreover, are there any best practices or safety precautions that one should be 
aware of when using ammonia in this context?

Your insights on this topic would be invaluable to me and would greatly enhance 
my understanding of the intricacies involved in histological preparations.

Thank you for your time and assistance.

Warm regards from Santiago of Chile

Ignacio, Medical Technologist
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From: Jay Lundgren 
Sent: May 11, 2024 4:40 PM
To: John Kiernan 
Cc: histonet@lists.utsouthwestern.edu ; 
IGNACIO GONZÁLEZ MASSONI 
Subject: Re: [Histonet] Inquiry on Tissue Softening for Microtomy

I wouldn't say softening, I would say hydrating.  Ammonia water accelerates 
hydration of FFPE blocks.  Nobody knows how it works, it's a my

Re: [Histonet] Inquiry on Tissue Softening for Microtomy

[John Kiernan via Histonet]

   If you apply the ammonia to the cut surface of the paraffin block, I suspect 
that it softens the tissue in the same way as applying water: by entering 
interstices of the tissue that are not occupied by paraffin molecules.
   I never tried ammonia for this purpose but in the 1960s to early '70s I 
occasionally used a proprietary product called Mollifex, which I see is still 
sold. In 1972 or '73 an elderly technician told me that water was just as good, 
and I soon found out that he was right. Indeed, water had the advantage of 
working in 15-30 minutes rather than taking several hours.
John Kiernan
Emeritus, UWO, London, Canada
https://publish.uwo.ca/~jkiernan/
= = =

From: IGNACIO GONZÁLEZ MASSONI via Histonet 
Sent: May 10, 2024 8:53 PM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Inquiry on Tissue Softening for Microtomy

Dear friends at Histonet,

I hope this message finds you well. I am reaching out to seek your expertise on 
a matter that has piqued my interest in the field of histology.

I am currently delving into the process of preparing FFPE (formalin-fixed, 
paraffin-embedded) tissues for microtomy. Specifically, I am curious about the 
role of ammonia in softening these tissues before sectioning. From my 
understanding, ammonia serves as an alkaline agent that helps neutralize 
formalin's effects and facilitates the removal of paraffin, thus aiding in the 
softening of the tissue.

However, I would greatly appreciate if you could provide a more detailed 
explanation of the chemical interactions at play here. How exactly does ammonia 
interact with the tissue components to achieve the desired softening effect? 
Moreover, are there any best practices or safety precautions that one should be 
aware of when using ammonia in this context?

Your insights on this topic would be invaluable to me and would greatly enhance 
my understanding of the intricacies involved in histological preparations.

Thank you for your time and assistance.

Warm regards from Santiago of Chile

Ignacio, Medical Technologist
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Re: [Histonet] Golgi-cox staining

[John Kiernan via Histonet]

I suggest asking Dr Sami Zaqout, the corresponding author of the paper you 
cited. A German email address is given on the left side of the first page of 
the free PDF file for which you provided a web link: 
https://www.frontiersin.org/articles/10.3389/fnana.2016.00038/ful.
 He is easy to find with Google and he is now at Qatar University.
It's the most thorough description of Golgi-Cox staining that I've seen, and 
the pictures of results look superb. Vibratome sectioning evidently is better 
and quicker than embedding in nitrocellulose and cutting with a sliding  
microtome! The two authors' use of  the word "cryoprotectant" was unfortunate 
in the context of a technique that does not involve freezing.
John Kiernan
Emeritus, Anatomy, University of Western Ontario, Canada
https://www.schulich.uwo.ca/anatomy//people/faculty/emeriti/kiernan_john.html
= = =

From: Mariela Chertoff via Histonet 
Sent: April 10, 2024 10:28 AM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Golgi-cox staining

Hi all

We made the Golgi cox staining and  due to a problem with the vibratome, we
left the tissue several days embedded in  agarose and they get dryed, It is
possoble to recover the brains? it is better to repeat the agarose
embebbing o it is better to put the brains in crioprerervate solution to
rehidrated and after that put them in agarosa again? We are following the
Zaquot protocol
https://www.frontiersin.org/articles/10.3389/fnana.2016.00038/full

Thanks in advance for your reply

Mariela Chertoff, PhD
Laboratorio de Neuroepigenetica - QB75
 Departamento de Química Biológica
Facultad de Ciencias Exactas y Naturales - UBA
Ciudad Universitaria Pabellón II Piso 4
Ciudad Autónoma de Buenos Aires
C1428EGA - Argentina
Tel: 54 11 5285-8680/1/2
email:marielachert...@gmail.com
marielachert...@qb.fcen.uba.ar

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Re: [Histonet] Modified Davidson solution

[John Kiernan via Histonet]

The Davidson/Hartmann fixative is just another alcoholic formaldehyde mixture 
acidified with acetic acid. Like all others of that ilk it was intended for 
making up in the lab soon before using. Storage causes slow deterioration. The 
ethanol slowly gets esterified by the acetic acid, making ethyl acetate, which 
has no fixative properties. This change is evident from the fruity smell of the 
ester, which becomes evident after a week or two. All this has been in the 
textbooks for more than 50 years.
Why doesn't every lab spend perhaps $200 on a couple of books?  For obvious 
reasons I recommend my 5th edition (ISBN 9781907904325; $82.50,from the 
publisher, which is $30 less than Amazon's price), but there are other 
histotechnology books that are very good.  A book may cost much less than a big 
bottle of a deteriorating mixture of questionable value, and you and your staff 
could learn a lot by reading.
John Kiernan
= = =

From: Davoli, Katherine A via Histonet 
Sent: March 5, 2024 4:40 PM
To: Histonet ; Naira Margaryan 

Subject: Re: [Histonet] Modified Davidson solution

Hi Naira, I get Hartmann's from Electron Microscopy Sciences
Cat# 64133-10 (for 1L, but they have larger sizes I believe)


Katherine Davoli, HTL(ASCP)cm(they/them/theirs)

Lab Manager, Tissue Culture and Histology Core Facilities

U. Pitt Dept of Ophthalmology

7.373 UPMC Mercy Pavilion

1622 Locust St., Pittsburgh PA 15219

(412) 624-8508   this number cannot receive texts


From: Naira Margaryan via Histonet 
Sent: Tuesday, March 5, 2024 1:27 PM
To: Histonet 
Subject: [Histonet] Modified Davidson solution

Hello!

What is the best company to buy a ready to use Modified Davidson solution?

Thanks in advance,
Naira
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From: Davoli, Katherine A via Histonet 
Sent: March 5, 2024 4:40 PM
To: Histonet ; Naira Margaryan 

Subject: Re: [Histonet] Modified Davidson solution

Hi Naira, I get Hartmann's from Electron Microscopy Sciences
Cat# 64133-10 (for 1L, but they have larger sizes I believe)


Katherine Davoli, HTL(ASCP)cm(they/them/theirs)

Lab Manager, Tissue Culture and Histology Core Facilities

U. Pitt Dept of Ophthalmology

7.373 UPMC Mercy Pavilion

1622 Locust St., Pittsburgh PA 15219

(412) 624-8508   this number cannot receive texts


From: Naira Margaryan via Histonet 
Sent: Tuesday, March 5, 2024 1:27 PM
To: Histonet 
Subject: [Histonet] Modified Davidson solution

Hello!

What is the best company to buy a ready to use Modified Davidson solution?

Thanks in advance,
Naira
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Re: [Histonet] Modified Davidson

[John Kiernan via Histonet]

The "Davidson's" name is, as Tony says, applied to various mixtures. In these, 
formalin is acidified with acetic acid in solutions that also contain some 
alcohol (typically about 33% v/v) but probably not enough to contribute to 
fixation. The mixture quoted from Latendresse et al. 2002 has only 15% alcohol 
- about the same as in a good dry sherry.  (My gastric mucosa is not yet fixed.)

The Histonet archives must surely still contain Dr Bob Richmond's contributions 
about "Davidson's fixative" in the 1990s or 2000s.

Enough from me for now!
John Kiernan
London, Canada
= = =

From: Tony Henwood via Histonet 
Sent: February 8, 2024 5:31 PM
To: Naira Margaryan ; Histonet 

Subject: Re: [Histonet] Modified Davidson

There are several formulations (some are probably typos), but this seems to be 
one commonly cited.

Modified Davidson's Fixative:

(Latendresse, J. R., Warbrittion, A. R., Jonassen, H., & Creasy, D. M. (2002). 
Fixation of testes and eyes using a modified Davidson's fluid: comparison with 
Bouin's fluid and conventional Davidson's fluid. Toxicologic pathology, 30(4), 
524-533.)

37–40% formaldehyde30ml
Absolute ethanol 15ml
Glacial acetic acid5ml
Distilled Water 50ml

Regards,

Tony Henwood MSc, BAppSc, GradDipSysAnalys, CT(ASC), FFSc(RCPA) (Retired)
Principal Scientist, the Children’s Hospital at Westmead (Retired)
Adjunct Fellow, School of Medicine, University of Western Sydney.


From: Naira Margaryan via Histonet 
Sent: Friday, February 9, 2024 8:25:47 AM
To: Histonet 
Subject: [Histonet] Modified Davidson

Hello,

Could you please share your best recipe for the Modified Davidson?

Thanks in advance,
Naira
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Re: [Histonet] O.C.T. what MW PVA and PEG?

[John Kiernan via Histonet]

Your method makes no sense! It looks like something informally passed along 
among students and technicians who have never read a book.
Cryoprotection means preventing formation of ice crystals or, as it's usually 
done, minimizing their size. Sucrose is a cryoprotectant; the higher the 
concentration the better, but strong solutions penetrate the tissue slowly. For 
fixed specimens dissolve the sucrose in water.  (For unfixed, use an isotonic 
phosphate buffer, but that's only if a day or so at 4C is OK for your needs. 
Very fast freezing may be needed, especially for muscle.)
OCT is not a cryoprotectant; it's goop that surrounds the specimen and enters 
cracks and spaces but it does not penetrate. OCT serves to hold each section 
together during transfer from the cryostat's knife onto a glass slide or 
coverslip.
For some more information (much of it derived from Histonet) have a look at the 
Biological Stain Commission's FAQ, at 
https://biologicalstaincommission.org/faqlist.htm#CRYPRO.
John Kiernan
https://www.schulich.uwo.ca/anatomy//people/faculty/emeriti/kiernan_john.html
= = =

From: Tyrone Genade via Histonet 
Sent: November 30, 2023 9:46 PM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] O.C.T. what MW PVA and PEG?

Hello,

I'm using a cryoprotection protocol that involves 3-stage cryoprotection of 15% 
glucose O/N, then 30% + 50% OCT O/N and then finally O/N in OCT. Compared to 
previous protocols this works very well -- even when cutting through eyes and 
lenses (which had previously given a lot of grief).

My issue is that preparing the 30% + 50% OCT is a schlep. The OCT puts up a lot 
of resistance against mixing with the 60% sucrose. It would be much simpler if 
I could prepare 30% sucrose with powdered PVA and PEG. Does anyone know what MW 
polymers of PVA and PEG to use and what concentrations to approximate 
commercial (Scigen Tissue-Plus) OCT?

Thanks

Tyrone Genade Ph.D.
Quillen College of Medicine
ETSU
Johnson City, TN
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From: Tyrone Genade via Histonet 
Sent: November 30, 2023 9:46 PM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] O.C.T. what MW PVA and PEG?

Hello,

I'm using a cryoprotection protocol that involves 3-stage cryoprotection of 15% 
glucose O/N, then 30% + 50% OCT O/N and then finally O/N in OCT. Compared to 
previous protocols this works very well -- even when cutting through eyes and 
lenses (which had previously given a lot of grief).

My issue is that preparing the 30% + 50% OCT is a schlep. The OCT puts up a lot 
of resistance against mixing with the 60% sucrose. It would be much simpler if 
I could prepare 30% sucrose with powdered PVA and PEG. Does anyone know what MW 
polymers of PVA and PEG to use and what concentrations to approximate 
commercial (Scigen Tissue-Plus) OCT?

Thanks

Tyrone Genade Ph.D.
Quillen College of Medicine
ETSU
Johnson City, TN
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Re: [Histonet] Faded H tissue section

[John Kiernan via Histonet]

I found the Histo-Logic archive on one of Sakura's web pages. Lee Luna was the 
editor for several years. Nothing there before about 1980 periodic acid for 
re-working H The archive is huge and many contributions look good.  See
https://www.sakuraus.com/getattachment/103878b9-9854-469d-8203-9d9b1c45d850/761
but bear in mind that this is from a company. It's not a peer-reviewed 
scientific journal.
John Kiernan.
= = =

From: Tony Henwood via Histonet 
Sent: November 10, 2023 4:49 AM
To: Akemi Allison ; jayalakshmy p.s 

Cc: histonet@lists.utsouthwestern.edu 
Subject: Re: [Histonet] Faded H tissue section

I thought he may have reported this in Histologic.
This trick certainly works. You can notice the nuclear clarity on GIT and to a 
lesser extent skin biopsies stained with PAS.

Regards,

Tony Henwood MSc, BAppSc, GradDipSysAnalys, CT(ASC), FFSc(RCPA) (Retired)
Principal Scientist, the Children’s Hospital at Westmead (Retired)
Adjunct Fellow, School of Medicine, University of Western Sydney.

From: Akemi Allison
Sent: Friday, 10 November 2023 1:30 PM
To: jayalakshmy p.s
Cc: Tony Henwood; 
histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] Faded H tissue section

Hi Tony:
I actually gave that tip to Lee back in 1979 when he came to OHSU to give a 
seminar to the Oregon Histology Society.  I found out that when I was doing GMA 
and the pathologists didn’t like Gills Hematoxylin. They loved my PAS on GMA so 
I tried using 1% Periodic Acid before using Harris Hematoxylin for H’s on GMA 
and it turned out beautifully. Guess he shared my tip with the rest of our 
society. He and I became closer se friends and we shared several tips, 
including his Movat’s tips which he didn’t publish, but I shared them in 
Frieda’s 4th Ed.

Best,
Akemi Allison-Tacha, BS, HT/HTL (ASCP)

Sent from my iPhone

> On Nov 9, 2023, at 6:11 PM, jayalakshmy p.s via Histonet 
>  wrote:
>
> Thanks, I'll check it out.
>
>> On Fri, Nov 10, 2023, 6:58 AM Tony Henwood  wrote:
>>
>> You could treat the decoverslipped section in 1% periodic acid (same as
>> used in the PAS technique)  for 30 minutes prior to H staining. This
>> might improve the H staining.
>>
>>
>>
>> I believe Lee Luna suggested this but for the life of me, I can’t find the
>> reference!
>>
>>
>>
>> Regards,
>>
>>
>>
>> Tony Henwood MSc, BAppSc, GradDipSysAnalys, CT(ASC), FFSc(RCPA) (Retired)
>>
>> Principal Scientist, the Children’s Hospital at Westmead (Retired)
>>
>> Adjunct Fellow, School of Medicine, University of Western Sydney.
>>
>>
>>
>> *From: *jayalakshmy p.s via Histonet 
>> *Sent: *Friday, 10 November 2023 3:43 AM
>> *To: *histonet@lists.utsouthwestern.edu
>> *Subject: *[Histonet] Faded H tissue section
>>
>>
>>
>> Hello,
>> I would like to know how to effectively restain a faded H tissue section.
>> The color becomes dull when re stained. Somebody please advise.
>> Prof. Jayalakshmy. P S
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>>
>>
>>
> ___
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Re: [Histonet] Fixing and processing mouse eyeballs

[John Kiernan via Histonet]

The Journal of Histotechnology, Vol. 45 No. 4 (Dec 2022) is a special issue 
devoted to eyes and ocular tissues.  Two of the articles are specifically about 
fixing and processing eyes of mice: J. Pang et 9 al. pp172-181 and J. Li et 9 
al. pp161-171. Every member of the NSH should have received a paper copy of the 
journal. The cover has a splendid photomicrograph (from the Pang paper) of a 
whole mouse eye.

John Kiernan
University of Western Ontario
London, Canada
https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html

= = =

From: Charles Riley via Histonet 
Sent: October 11, 2023 5:00 PM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Fixing and processing mouse eyeballs

Does anyone have any helpful tips or protocols for fixing and processing
schedules for adult mouse eyes?
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Re: [Histonet] Long term museum specimen storage

[John Kiernan via Histonet]

I don't know anything about "Jore's fixative" or the rationale of using a very 
hypertonic unbuffered 4% formaldehyde with magnesium, sodium, chloride and 
sulphate ions. If brown stuff is now bleeding out of your museum specimens, 
Jore Juice evidently isn't a good preservative.

According to Chapter 26 in the late Charles Culling's excellent book (3rd edn 
1974; ISBN: 0407729011) the fixative/preservative for a museum specimen is 
optimized to preserve colour, which is the red or reddish-brown of haemoglobin 
and myoglobin. This usually is achieved with Kaiserling's fluid, which contains 
formalin, potassium acetate and also potassium nitrate (1.5% w/v) as an 
oxidant. Another approach involves treating specimens with carbon monoxide to 
convert all haemoglobin etc to a red carboxy derivative.

If your museum specimens have already lost all their meaningful colours, a 
neutral buffered aqueous formaldehyde may be the best that you can provide to 
preserve the sizes and shapes. 70% alcohol will cause some shrinkage, and it 
may not be as easy to seal this solvent into a museum container as a watery 
diluted formalin.

John Kiernan
= = =

From: Rhonda McCormick via Histonet 
Sent: September 20, 2023 11:39 AM
To: Histonet 
Subject: [Histonet] Long term museum specimen storage

Hi All,
I am looking to replace the fixative for veterinary specimens that have been 
preserved as "museum specimens". They are kept in jars in a glass case outside 
our lab, however, some of the fixative is starting to turn brown (and we've 
pulled a few jars that have some slight cracks in them).
The specimens are currently in Jore's Fixative: 100 mL Distilled water
10 mL 40% Formaldehyde2 g Magnesium Sulfate2 g Sodium Sulfate1 g Sodium Chloride
Preserving specimens is new to me. I've never heard of Jore's fixative before 
and I'm wondering if I could get some advice, please?  Do these specimens need 
to be replaced with the same solution? Could we rinse the specimen and replace 
the solution with 70% Alcohol? OR would 10% NBF be better to store the 
specimens in (or something al together different)? We have a varying display of 
specimens - anywhere from a small porcine optic nerve to a large equine 
granulosa cell tumor. Realizing it may be different based on the size of the 
specimen, approximately how often should the solution be changed ?
Thank you so much! Any help or insight is much appreciated.
Rhonda McCormickRhonda McCormick BS, HT (ASCP)cm
Histology Diagnostic Lab Supervisor

College of Veterinary Medicine
Texas A University

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Re: [Histonet] Formalin pH

[John Kiernan via Histonet]

Thank you Tony, for drawing attention to that excellent 1991 paper in the 
Journal of Histotechnology. It was published in the years when the NSH's 
journal was not included in Current Contents and was not taken by most 
libraries. Sadly, Norman Hew-Shue died in Toronto in 2022.

Papers in J. Histotechnol. (including all back-issues) are now easily available 
as PDF downloads from libraries that subscribe to the many journals published 
by Taylor and Francis. That's probably most academic and biomedical libraries.

Members of the NSH still get the paper journal and have access to the archive 
by way of an obscure members-only link on the society's web site; it is very 
difficult to find.  Why won't the NSH make its journal conspicuous and easily 
accessible to all by way of a single click from https://www.nsh.org/home?  Grrr!

John Kiernan.
= = =

From: Tony Henwood via Histonet 
Sent: September 13, 2023 7:01 PM
To: Bacon, Charles ; e...@pigs.ag 

Cc: HistoNet 
Subject: Re: [Histonet] Formalin pH

Hew-Shue (1991) has described a useful pH indicator for working neutral 
buffered formalin solutions. Bromocresol purple, when added to formalin 
solutions, serves as an indicator of pH as well as, from a safety aspect, 
labelling the solution as formalin making redundant the dangerous “smell” test. 
At an acidic pH (5.2) the intense indicator colour is yellow whereas at pH 6.8, 
the colour is purple. A saturated solution of the dye is prepared, and 2 to 4 
drops are added to 10 litres of neutral buffered formalin. Other advantages are 
that the fixative is more readily distinguished from other colourless solutions 
such as saline, thus preventing accidental misuse, and formalin spills are more 
easily recognised.

Hew-Shue N (1991) “Bromcresol Purple as a Colored Marker and pH Indicator for 
Ten Percent Neutral Buffered Forrnalin” J Histotechnol 14(4):257-260.


Regards,

Tony Henwood MSc, BAppSc, GradDipSysAnalys, CT(ASC), FFSc(RCPA) (Retired)
Principal Scientist, the Children’s Hospital at Westmead (Retired)
Adjunct Fellow, School of Medicine, University of Western Sydney.

From: Bacon, Charles via Histonet
Sent: Thursday, 14 September 2023 6:42 AM
To: e...@pigs.ag
Cc: HistoNet
Subject: Re: [Histonet] Formalin pH

We pull the COA from the vendors website. Some are better than others but the 
are usually listed by lot number. I keep them in a network shared folder so 
they can be pulled by anyone that is asked for the evidence during inspection.

Sent from my iPhone

> On Sep 13, 2023, at 8:22 AM, "e...@pigs.ag"  wrote:
>
> Sooner or later the people have got to rise up
> and tell those brain-dead paper-shuffling regulators
> to just stuff it.  Their mindless interference is
> just wasting time and interfering with people's
> ability to keep their own garden.
>
> -Original Message-
> From: Cooper, Brian via Histonet
> Reply-To: Cooper, Brian
> To: Paula Sicurello, HistoNet
> Subject: Re: [Histonet] Formalin pH
> Date: Today 10:19 AM
>
> Ask your vendor to send a certificate of analysis that includes the pH. 
> That's what we did and now they come with each lot. We do have to bug them 
> for them occasionally.
>
> I'm betting your vendor has at least heard about this from other customers so 
> it should be on their radar at the very least.
>
>
> Thanks,
>
>
>
> Brian D. Cooper, HT (ASCP)CMQIHCCM| Histology Supervisor
>
> Department of Pathology and Laboratory Medicine
>
> Children's Hospital Los Angeles
>
> 4650 Sunset Blvd MS#43- Los Angeles, CA 90027
>
> Ph: 323.361.3357
>
> bcoo...@chla.usc.edu
>
> 
> From: Paula Sicurello via Histonet 
> Sent: Tuesday, September 12, 2023 7:12:05 PM
> To: HistoNet 
> Subject: [Histonet] Formalin pH (EXTERNAL EMAIL)
>
> CAUTION: BE CAREFUL WITH THIS MESSAGE*
> This email came from outside CHLA. Do not open attachments, click on links, 
> or respond unless you expected this message and recognize the email address: 
> histonet-boun...@lists.utsouthwestern.edu.
>
> Hello Histoteckies,
> What are y'all doing regarding the CAP requirement to monitor the pH of 
> formalin?
> We buy tons and tons of the 5 gallon cubitainers and we are still debating 
> over how to check the pH.
>
> Looking forward to your replies.
> Toodles!
> Sincerely,
>
> Paula Sicurello
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Re: [Histonet] Formalin pH

[John Kiernan via Histonet]

You can check the pH with a pH meter. This is the most accurate way, but the 
meter's electrode must be calibrated against at least two standard (usually 
bought) buffer solutions, such as 4.0 and 7.0.  If you don't have a pH meter or 
the know-how to use one properly, you can use indicator papers, which are 
inexpensive but deteriorate with storage. With a few "books" of indicator 
papers covering the range 3 to 8 you should be able to get a value accurate to 
the nearest 0.2 pH units.

I'm assuming that by "formalin" you mean the concentrated solution that is 
nominally 37-40% by weight of formaldehyde gas in water. Its pH falls with time.
For more information about formalin, see 
https://www.cambridge.org/core/services/aop-cambridge-core/content/view/911F1870567E2487D56A51140EB8EA17/S1551929500057060a.pdf/div-class-title-formaldehyde-formalin-paraformaldehyde-and-glutaraldehyde-what-they-are-and-what-they-do-div.pdf.
 

  It's a free download, with >600 citations according to Google Scholar.

John Kiernan
Emeritus, UWO, London, Canada
= = =

From: Paula Sicurello via Histonet 
Sent: September 12, 2023 10:11 PM
To: HistoNet 
Subject: [Histonet] Formalin pH

Hello Histoteckies,
What are y'all doing regarding the CAP requirement to monitor the pH of 
formalin?
We buy tons and tons of the 5 gallon cubitainers and we are still debating over 
how to check the pH.

Looking forward to your replies.
Toodles!
Sincerely,

Paula Sicurello
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Re: [Histonet] Gram Stain

[John Kiernan via Histonet]

I don't know a method to obtain selective red coloration of Gram-negative 
organisms, with yellow for both Gram-positives and "background", and I cannot 
find one by looking in various books. It may not be possible because 
Gram-positivity relies on selective retention of an immobilized dye. 
Gram-negative bacteria stain only because the red dye is excluded by the 
organisms already filled with an insolubilized blue-purple dye in the 
Gram-positive cells.
The Brown-Hopps modification of Gram staining for paraffin sections clearly 
separates blue from red bacteria and gives a quite different yellowy-brown 
counterstain to the "tissue background".  This is explained and well  
illustrated in Freida Carson's textbook.  (I have only the 2nd edition, 1997; 
there is now a 5th, 2020, ISBN 9780891896760.)
John Kiernan  (London, Ontario)
= = =

From: Rhonda McCormick via Histonet 
Sent: August 30, 2023 2:42 PM
To: Histonet 
Subject: [Histonet] Gram Stain

Howdy,Does anyone have a recommendation for a Gram stain (or modification of a 
gram stain) that stains the background yellow with red gram-negative bacteria  
(no blue - gram positive staining).Thank you!
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Re: [Histonet] Sudan Black B

[John Kiernan via Histonet]

The Sudan black B method for lipofuscin is exactly the same as for frozen 
sections, but it is applied to hydrated paraffin sections. Remember that the 
dye solution needs to be fairly fresh (less than 4 weeks old) and must be 
filtered immediately before using. Use Sudan black B (CI 26150) powder that has 
been certified by the Biological Stain Commission.

Various controls and other stains are needed to be certain that the pigment is 
lipofuscin, not melanin or haemosiderin.

John Kiernan
Anatomy, UWO, London, Canada
= = =

From: Betsy Molinari via Histonet 
Sent: August 10, 2023 10:57 AM
To: Histonet 
Subject: [Histonet] Sudan Black B

Hi,
I have been asked to do a Sudan stain on a heart  biopsy for lipofuscin. The 
biopsy is in a paraffin block. They are looking  to better report and 
understand the IHC. I am totally unfamiliar with this stain. I did some reading 
but have been unable to find a protocol for paraffin sections. I found a 
reference to Sheehan & Hrapchak (1973) but unfortunately I don't have that 
edition.  Any ideas would be greatly appreciated .

Betsy Molinari HT (ASCP)
Texas Heart Institute
Cardiovascular Pathology
1101 Bates St.
Houston, Texas  77030
832-355-6524

Betsy Molinari, HT (ASCP)
Sr. Histology Research Technician
CV Pathology Research

The Texas Heart Institute (r)
6770 Bertner Avenue, MC 1-283
Houston, TX 77030

Office: 832-355-6524 | Fax: 832-355-6812
Email: bmolin...@texasheart.org
texasheart.org | 
texasheartmedical.org | 
facebook | 
twitter

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From: Betsy Molinari via Histonet 
Sent: August 10, 2023 10:57 AM
To: Histonet 
Subject: [Histonet] Sudan Black B

Hi,
I have been asked to do a Sudan stain on a heart  biopsy for lipofuscin. The 
biopsy is in a paraffin block. They are looking  to better report and 
understand the IHC. I am totally unfamiliar with this stain. I did some reading 
but have been unable to find a protocol for paraffin sections. I found a 
reference to Sheehan & Hrapchak (1973) but unfortunately I don't have that 
edition.  Any ideas would be greatly appreciated .

Betsy Molinari HT (ASCP)
Texas Heart Institute
Cardiovascular Pathology
1101 Bates St.
Houston, Texas  77030
832-355-6524

Betsy Molinari, HT (ASCP)
Sr. Histology Research Technician
CV Pathology Research

The Texas Heart Institute (r)
6770 Bertner Avenue, MC 1-283
Houston, TX 77030

Office: 832-355-6524 | Fax: 832-355-6812
Email: bmolin...@texasheart.org
texasheart.org | 
texasheartmedical.org | 
facebook | 
twitter

CONFIDENTIALITY NOTICE: This email and attachments contain information that may 
be confidential or privileged. If you are not the intended recipient, notify 
the sender at once and delete this message completely from your information 
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email is not authorized, and any such action should not be construed as a 
waiver of privilege or other confidentiality protections.
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Re: [Histonet] Von Kossa staining

[John Kiernan via Histonet]

Charles,

A handheld light of any kind isn't really suitable because you would have to 
hold it over the slides for 15 to 60 minutes, according to which variant of the 
von Kossa method you plan to use (see Lillie & Fullmer 1976 Histopathologic 
Technic ... 4th ed. pp 539-541).

An anglepoise lamp with an old-fashioned 100W bulb is OK, and so is a sunny 
windowsill. Silver salts absorb at the blue end of the spectrum, so a 
fluorescent light should be more efficient than an incandescent bulb. If no 
bright light source is available, it's possible to chemically reduce the silver 
phosphate and/or carbonate to black colloidal silver, with a traditional 
photographic developer. The method of Rungby et al.1993 may be better than 
other post-reduction methods 
(https://scholar.google.ca/scholar?hl=en_sdt=0%2C5=rungby+1993+calcium+deposits=rungby+1993).
 I never tried it, but Rungby's paper has collected 104 citations, which is 
very good for a paper in our field.

The von Kossa technique is simply explained in my Histological and 
Histochemical Methods textbook, 5th edn (2015). The book costs less than 1ml of 
any antibody.

Enough said!   John
John A. Kiernan
Emeritus, Anatomy & Cell Biology,
University of Western Ontario
London, Canada
https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html
Also  Secretary, Biological Stain Commission, Inc.
https://biologicalstaincommission.org
= = =








-Original Message-
From: Charles Riley via Histonet 
Sent: Wednesday, July 26, 2023 1:23 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Von Kossa staining


*** Externally sourced email message ***


Can anyone out there who performs Von Kossa staining provide me with any 
guidelines or suggestions for the light source to use for the Silver nitrate 
activation?

Is a standard handheld black light strong enough or does it need to be a UV 
sanitizing strength light if using UV versus incandescent bulb exposure?
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Re: [Histonet] Fast Green / Sirius Red - Unknown blue features

[John Kiernan via Histonet]

Your technique is the one first (I think) published by Lopez-De Leon A & 
Rojkind M (1985) A simple micromethod for collagen and total protein 
determination in formalin-fixed paraffin-embedded sections. J. Histochem. 
Cytochem. 33: 737-743. The photos in that paper show some of the collagen 
almost black - surely taking up both red and green dyes. More recent papers 
describe exactly the same method, and there are also some variants. Your 
technique, with an acid rinse after staining for an hour, then quick transition 
to rapid dehydration in 100% alcohol, is essential for any valid picro-sirius 
staining.

According to the entry for fast green FCF (CI 42053) in Conn's Biological 
Stains (10th ed, p.180-182), "chemically distinct blue-green dyes have been 
supplied under this name". Are you sure your fast green FCF is the real McCoy? 
Is it from a batch certified by the Biological Stain Commission? The jar of dye 
powder should have a small bluish label, with features that make forgery 
difficult. See 
https://biologicalstaincommission.org/certified-biological-dyesstains/ for 
pictures and other information. There are companies selling "certified stains" 
that have not been tested and certified by the Biological Stain Commission. 
Caveat emptor!

The Biological Stain Commission is a not-for-profit corporation that has been 
providing third-party quality control and other services for vendors and users 
of stains for 100 years.

Just a few thoughts; I could add more, but probably this letter already is too 
long for the Histonet censors.

John Kiernan
Professor Emeritus, Anatomy & Cell Biology
University of Western Ontario, London, Canada
https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html
Also  Secretary, Biological Stain Commission, Inc.
https://biologicalstaincommission.org
= = =

From: David Burk via Histonet 
Sent: June 21, 2023 5:48 PM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Fast Green / Sirius Red - Unknown blue features

We are working out an alternative method for collagen staining using Fast Green 
/ Sirius Red (Direct Red 80) rather than the standard picrosirius red method as 
I think it is prettier and easier to see the collagen on a green background.

What we’ve noticed, though, is that we are seeing intermediate blue staining in 
the tissue in particular regions or structures. I’ve not had any success in 
finding an explanation for this online and was hoping someone on the board may 
have an idea of what’s going on and what is being stained blue in our tissue 
sections.

We have examined a variety of murine or rat tissues including liver, heart, 
kidney, lung, skeletal muscle, spleen, brain, pancreas, and even decellularized 
human adipose tissue. There are, almost always, some structures/features that 
exhibit a denim blue to lighter blue-green color (at least to my eye) in 
addition to the expected red-colored structures that we would assume to be 
collagen, light green cytoplasm, and yellow-ish features stained with picric 
acid.

An interesting tidbit is that these blue-ish stained features are birefringent 
under polarized light so you would not know their color (with transmitted 
imaging) was atypical.

I don’t want to use a stain if I can’t let people know what a particular color 
represents and can also cause problems with the quantification of collagen 
using a color-based approach.

Our protocol is as follows:

  1.  Dewax
  2.  H2O rinse
  3.  Stain in a 0.1% Fast Green FCF (C.I. 42053) and 0.1% Direct Red 80 (C.I. 
35780) solution dissolved in saturated picric acid for 1 hour at room 
temperature
  4.  Dip 5x and then immerse in 0.5% acetic acid for 5 seconds
  5.  Repeat step 4
  6.  Dip 5x and then immerse in 100% Ethanol 30 seconds
  7.  Dehydrate in 100% Ethanol 1 min
  8.  Repeat step 7
  9.  3 x Xylene for 2 min each
  10. Coverslip

I’m uploading some images from mouse muscle and tumor tissue to the Histonet 
Image upload site. If that doesn’t work, here are links:

Mouse tumor:

https://drive.google.com/file/d/1VjOZFzvsQByQLuDtdGPfdwAwap_CVE58/view?usp=sharing

Mouse skeletal muscle:

https://drive.google.com/file/d/10vT_FKu3-Ad5uemM5gnZmqDCKb3fs2lV/view?usp=sharing


Thanks,

David Burk




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Re: [Histonet] Peroxidase stain on peripheral smears

[John Kiernan via Histonet]

What is the Kaplow method? I can't find it in textbooks. A quick Google search 
brings up only junk papers indicating that a Kaplov method may use carcinogenic 
benzidine (with wrong spelling) as the chromogen.
There are simple, safe and inexpensive methods for histochemical localization 
of sites of peroxidase activity.in tissue sections or smears of cells. Buy a 
textbook for about $50, read it, and save your lab a fortune in buying special 
kits for very simple routine histochemical jobs.

My ten cents-worth. June 2023
John Kiernan
J. A. Kiernan MB, ChB, PhD, DSc
Professor Emeritus, Anatomy & Cell Biology
University of Western Ontario, London, Canada
https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html
Also  Secretary, Biological Stain Commission, Inc.
https://biologicalstaincommission.org
= = =

From: Kendra Beechie ND-Bismarck via Histonet 

Sent: June 12, 2023 3:22 PM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Peroxidase stain on peripheral smears

Hello,

I am seeking some guidance in regards to a peroxidase stain. We have always
used the Kaplow Method to stain peripheral blood smears, and up until
recently, we have had no issues with getting it to work.  However, no
matter what we try in recent months, we have not been able to get it to
work. Several different techs have tried it and we have ordered new
reagents, but there are no granules to be seen

Does anyone have any experience with peroxidase kits? I can see that
Sigma-Aldrich has several available but I'm not sure what to go with

Any and all help would be greatly appreciated!

Thank you,

Kendra Beechie
MLS/HTL (ASCP), Technical Consultant
CHI St. Alexius Health
Bismarck, ND

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you have received this email in error, do not read any attachments. Instead, 
kindly reply to the sender stating that you have received the message in error. 
Then destroy it and any attachments. Thank you.
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From: Kendra Beechie ND-Bismarck via Histonet 

Sent: June 12, 2023 3:22 PM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Peroxidase stain on peripheral smears

Hello,

I am seeking some guidance in regards to a peroxidase stain. We have always
used the Kaplow Method to stain peripheral blood smears, and up until
recently, we have had no issues with getting it to work.  However, no
matter what we try in recent months, we have not been able to get it to
work. Several different techs have tried it and we have ordered new
reagents, but there are no granules to be seen

Does anyone have any experience with peroxidase kits? I can see that
Sigma-Aldrich has several available but I'm not sure what to go with

Any and all help would be greatly appreciated!

Thank you,

Kendra Beechie
MLS/HTL (ASCP), Technical Consultant
CHI St. Alexius Health
Bismarck, ND

Caution: This email is both proprietary and confidential, and not intended for 
transmission to (or receipt by) any unauthorized person(s). If you believe that 
you have received this email in error, do not read any attachments. Instead, 
kindly reply to the sender stating that you have received the message in error. 
Then destroy it and any attachments. Thank you.
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Re: [Histonet] Coffee at the desk

[John Kiernan via Histonet]

Bob, Why weren't they routinely buffering (or at least neutralizing) the 
formalin fixatives at Johns Hopkins as recently as 1970?

It had all been in the scholarly books (by Pearse, Lillie, etc) for >10 years, 
and was also in Lee Luna's 1968 Manual of Histologic Staining Methods, 
published by the USA's Armed Forces Institute of Pathology.

I was brung up on cold neutral buffered formalin for enzyme activity 
histochemistry when I was a medical student doing an interpolated research year 
in the early 1960s.


Cheers,  John

John Kiernan (London, Canada)

= = =


From: Bob Richmond via Histonet 
Sent: June 3, 2023 8:25 AM
To: Histonet@lists.utsouthwestern.edu 
Subject: Re: [Histonet] Coffee at the desk

This 84 year old pathologist recalls the histopathology laboratory at Johns
Hopkins Hospital around 1970, when I was a pathology resident there.

Histotechs, often laboratory clerks, sat in front of rows of 400 mL Stender
dishes, smoking cigarettes while they hand-stained slides, often carrying
out the entire procedure from xylene and descending alcohols, up to final
coverslipping.

It wasn't the xylene that worried me, so much as the dish full of a 20%
solution of picric acid in acetone, that removed most of the copious
formalin pigment (since buffering the formalin wasn't permitted).

I spoke to the chief histotechnologist about the issue. He responded by
stubbing out a lighted cigarette into one of the Stender dishes of xylene.
(I'm told you can also do that with gasoline, but not with acetone.)

He was never incinerated, but he died of smoking-related disease soon after
his retirement.

Bob Richmond
Maryville TN
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Re: [Histonet] Get the Short Term or Long Term Lab Staffing Coverage You Need

[John Kiernan via Histonet]

Hear, Hear! Let's see less advertising on Histonet.
John Kiernan
(London, Canada).
= = =

From: Jay Lundgren via Histonet 
Sent: June 2, 2023 1:15 PM
To: Melissa Owens 
Cc: Tom Walls 
Subject: Re: [Histonet] Get the Short Term or Long Term Lab Staffing Coverage 
You Need

As far as I know, Histonet doesn't really allow advertising for individuals
or agencies *looking* for jobs.  There are a very few, selective, agencies
that are allowed to post on Histonet  tastefully and infrequently, but they
are buying, not selling.  Or histotechs who let others know about open
positions in their organization.  That's cool.

I'm not a moderator, but this is supposed to be a collegial forum for
practical histopathology advice.  There are plenty of job search sites out
there already.

But if Histonet does start allowing this, then ***%%%$$$HEY SCRIPPS
INSTITUTE HIRE ME$$$***

***%%%$$$LONG OR SHORT CONTRACT!!!CHEAP$$$%%%***

 **%%%$$$BIG TIME IMUNNOHISTOCHEMISTRY$$$%%%***

 ***%%%$$$USAF HTL (ASCP) M.S. 30+ YRS!!!$$$%%%***

 Sincerely,

  Jay A. Lundgren, M.S., HTL (ASCP)

On Fri, Jun 2, 2023 at 10:50 AM 'Melissa Owens' via Histonet <
histonet@lists.utsouthwestern.edu> wrote:

> li {display:list-item;text-indent: -1em;}
> ul, ol{margin-left: 40px !important; padding-left: 0px
> !important;}
>
> When workloads spike, unexpected projects hit, employees are out
> for PTO, Sick or Parental Leave, we got your Lab Covered! Our
> Staffing Services for the lab can get the work done quickly. We
> have skilled laboratory talent at the ready and range of
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>
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> committing to hiring them for a permanent role)
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> laboratory staffing (Customized to your needs)
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>
> · All inclusive bill rates (Travel/Pay Rate/Employment
> Taxes, etc.)
>
> Candidates we have right now who can make an impact in your lab!
>
> Histotech Supervisor/ Manager (2 Available)
>
> * Certified Histotech (ASCP) with over 20 years of experience
> * Seeking Temporary or Permanent work
> * Location: Seeking Anywhere along the East coast from FL to ME,
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> * Available: Now
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> * Primarily looking for Permanent but would also consider
> Temp/Travel work
> * Georgia
> * Available: Starting in July 2023
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> * ASCP Certified and 5 years of experience
> * Looking for Travel/Temp Position but open for Temp-Permanent
> (North Carolina)
> * Locations of interest: Anywhere in the US, Des Moines, IA,
> South Charleston, SC, Birmingham, AL
> * Available: Now
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> Cytotechnologist-(1 Candidate Available)
>
> * Several years of experience with ASCP certification and Imaging
> Trained
> * Seeking primarily Temporary or Temporary to Permanent
> * Locations of interest: Anywhere in US (Not State Licensed)
> * Available: Now
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> * Several years’ experience as a Generalists
> * Seeking Temporary/Travel
> * Locations of Interest: Anywhere in US
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> * Has 2 years’ experience and MLT Certification
> * Seeking Permanent or Temporary Work
> * Location seeking: Houston, TX area
> * Available: Now
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> Molecular Technologist – (3 Candidates Available)
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> * ASCP Molecular Specialists with 2+ years of experience
> * Seeking Permanent, Temp to Permanent or Permanent
> * Location: Anywhere in US
> * Available: Now
>
> Phlebotomist- (2 Available Candidates)
>
> * Certified Phlebotomist (CPT) and ASCP Certified Phlebotomists
> and 5-10 years of experience
> * Seeking Temp/Travel Work Only
> * Location Seeking: Anywhere in US
> * Ability to Train other Phlebotomists: YES
> * Available: Now
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> Available)
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> * Has Medical Assistant Diploma with phlebotomy experience and
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> * Seeking Temporary or Permanent work
> * Location seeking: Houston, TX
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> *If you do not see a professional on this list that fits with
> what your talent needs are please inquire directly to discuss our
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>
> Ask me about our Laboratory Training Program, Government
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> https://app.loxo.co/agencies/13989/email_tracking/click?id=104910928=https%3A%2F%2Fcalendly.com%2Fmelissaowens
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>  

Re: [Histonet] history of H staining

[John Kiernan via Histonet]

Gudrun, your question got me looking through more than a dozen older books, 
several more recent ones and various articles, but with no clear answer!
H wasn't a routine combination in 1902. Pathology and normal human histology 
textbooks in the 1950s show pictures that are clearly H but with the stain 
getting scarcely a mention, and this is also true of the most recent (1999) 
path text on my shelf. Forty-five alum-haematein mixtures were published 
between 1868 and 1951. Of these, a majority (26) were in the period 1882-1916 
and these include the best-known ones: Delafield, Ehrlich, Mayer, Harris etc, 
but eosin alone was seldom the recommended counterstain before 1890. H has 
never been the "routine" stain outside the fields of human and veterinary 
histology and pathology.  Other staining combinations are preferred for 
invertebrates, protozoa, plants and bacteria.  My guess is that H gradually 
became "routine" for pathology in the period 1910-1930.  If someone has access 
to some non-technical textbooks from those decades they might be able to narrow 
down the dates.
I could go on and on, with references etc, but this reply may already be too 
long for Histonet.  John Kiernan.
= = =

From: Gudrun Lang via Histonet 
Sent: May 20, 2023 8:46 AM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] history of H staining

Hi all!

Does anybody know, when the H stain became that dominant routine-stain in
the pathology labs?

It was introduced by Wissowzky 1876, but I am curious when our usual
histoprocess became worldwide standard.



Regards

Gudrun Lang

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Re: [Histonet] Mice brain

[John Kiernan via Histonet]

Immersion in sucrose is for cryoprotection - to minimize damage from formation 
of ice crystals. It is needed only if you are cutting frozen sections. Specific 
demonstration of cholinergic neurons is with immunohistochemistry to detect 
choline acetyltransferase. Use frozen or paraffin sections, according to 
instructions that come with the primary antibody. For histochemical 
demonstration of acetylcholinesterase activity (seen also in some neurons that 
are not cholinergic), you need frozen sections of tissue fixed for 12h in 
neutral formaldehyde at 4oC. My favourite method for AChE activity was that of 
Karnovsky & Roots 1964 J. Histochem. Cytochem. 12:219-221.
John Kiernan (London, Canada).
= = =

From: Renee Fisher via Histonet 
Sent: April 19, 2023 1:34 PM
To: Histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Mice brain

Hi,
Does anyone know if immersing in  sucrose is essential for optimum 
visualization of cholinergic neurons in mouse brain.

Thank you
Sent from my iPhone
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Re: [Histonet] Alcian Green Dye for Attwood stain

[John Kiernan via Histonet]

Alcian green was a mixture of two cationic "Ingrain" dyes: one of ICI's alcian 
blues (a copper phthalocyanine) and their alcian yellow (CI 12840, a monoazo 
dye). ICI stopped making these dyes, which were a commercial flop, in the early 
1970s. It's extremely unlikely that anything now sold as "alcian green" will be 
the genuine article. The original "Alcian blue" dyes were unstable even as 
powders.

The staining procedure published by HD Attwood in J. Path. Bact. 76:211 (1958) 
has to be seen as an oddity from the days when carbohydrate histochemistry was 
still in its infancy. Any textbook published since the 1970s will provide very 
simple and rational staining techniques for showing cartilage matrix 
(basophilic) and keratin (acidophilic) in contrasting colours.

John Kiernan
London, Canada
= = =

To: 'histonet@lists.utsouthwestern.edu' 

Subject: [Histonet] Alcian Green Dye for Attwood stain

Hi All,

I could use some assistance.  I'm gotten a request to do an Attwood stain for 
squames and cartilage and I'm having a difficult time finding the Alcian Green 
2GX dye powder or even a kit if one exists.  Another question is the protocol 
that I've seen from Histonet archives states that you dilute the phloxine in 
cellosolve.  I would just like to ask if anyone has another version of the 
protocol with any updates to the protocol?

Thanks in advance.

Debbie Siena
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From: Debra Siena via Histonet 
Sent: April 5, 2023 2:47 PM
To: 'histonet@lists.utsouthwestern.edu' 
Subject: [Histonet] Alcian Green Dye for Attwood stain

Hi All,

I could use some assistance.  I'm gotten a request to do an Attwood stain for 
squames and cartilage and I'm having a difficult time finding the Alcian Green 
2GX dye powder or even a kit if one exists.  Another question is the protocol 
that I've seen from Histonet archives states that you dilute the phloxine in 
cellosolve.  I would just like to ask if anyone has another version of the 
protocol with any updates to the protocol?

Thanks in advance.

Debbie Siena





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Re: [Histonet] Sudanblack B on FFPET

[John Kiernan via Histonet]

It's true that Sudan black B won't stain ordinary lipids (fat, phospholipids 
etc) that are absent from paraffin sections. The lipid in lipofuscin is bound 
to protein strongly enough to resist extraction during passage through all the 
solvents used in preparing paraffin sections. Churukian's adaptation of the 
Lillie & Ashburn method (link to a free download methods book in my recent 
Histonet email) is OK. It is one of the Biological Stain Commission's tests for 
certifying Sudan black B.
John Kiernan
London, Ontario
= = =

From: AJ Cabral via Histonet 
Sent: March 28, 2023 4:12 PM
To: Tony Henwood 
Cc: Histonet@lists.utsouthwestern.edu ; Bob 
Richmond 
Subject: Re: [Histonet] Sudanblack B on FFPET

Sudan Black staining won’t work on FFPET. The alcohols and xylenes used in
the tissue processing dissolves the lipids in the tissue. However, you can
used formalin fixed tissue as an alternative if no frozen section is
available. Rinse the tissue in distilled water for several minutes, pat
dry, freeze the tissue on OCT, cut frozen sections and stain in Sudan
black.

Have you considered looking into acid phosphatase staining for lipofucshin?
It is non specific but it can be demonstrated in muscle biopsy.

Cheers,
Joanna



On Tue, Mar 28, 2023 at 12:22 PM Tony Henwood via Histonet <
histonet@lists.utsouthwestern.edu> wrote:

> I would also let the saturated solution stand for a few days. Like Oil Red
> O, it probably needs time to “mature”. I would also use a frozen section of
> skin as a control.
>
> Regards,
>
> Tony Henwood
> Sydney, Australia
>
> From: Bob Richmond via Histonet
> Sent: Wednesday, 29 March 2023 4:51 AM
> To: Histonet@lists.utsouthwestern.edu histonet@lists.utsouthwestern.edu>
> Subject: Re: [Histonet] Sudanblack B on FFPET
>
> >
> > Gudrun Lang in Austria asks:
> >
>
> >>Has anyone experience with Sudanblack B on paraffin slides for staining
> [lipofuscin]? A doctor wants the demonstration of the lipoid content of
> foamy cells or granulocytes in lung. I've found protocols that have
> incubation-times from 10 minutes to over-night. - I've made a saturated
> Sudan black B-solution in 70% ethanol and tried it with10 min on liver
> without real success.<<
>
> The main thing you need to do is demonstrate that it isn't hemosiderin with
> an iron stain (Perls prussian blue reaction), and perhaps also that it
> isn't acid-fast. Lipofuscin can be identified an H & E staining, except for
> these considerations.
>
> Bob Richmond
> Maryville, Tennessee
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Re: [Histonet] Sudanblack B on FFPET

[John Kiernan via Histonet]

You probably would get better results with a supersaturated solution of the dye 
in 60% isopropanol with added dextrin. For details see 
https://www.urmc.rochester.edu/urmc-labs/pathology/StainsManual/. Click on 
Table of Contents and follow the links to fined the method. Be sure to use a 
batch of Sudan black B certified by the Biological Stain Commission.
John Kiernan
London, Canada
= = =

From: Gudrun Lang via Histonet 
Sent: March 28, 2023 10:30 AM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Sudanblack B on FFPET

Hallo!

Has anyone experience with Sudanblack B on paraffin slides for staining
lipofuszin? A doctor wants the demonstration of the lipoid content of foamy
cells or granulocytes in lung.



I've found protocols that have incubation-times from 10 minutes to
over-night.

I've made a saturated Sudanblack B-solution in 70% ethanol and tried it with
10 min on liver without real success.



I would appreciate any input and help.

Thanks in advance



Gudrun Lang

Austria

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Re: [Histonet] IHC staining of tendons and cartilage

[John Kiernan via Histonet]

You might like to look at this 1999 article from Microscopy Today, about 
keeping sections on slides.
https://publish.uwo.ca/~jkiernan/adhesivs.htm
John Kiernan
London, Canada.
= = =

From: Shirley A. Powell via Histonet 
Sent: March 21, 2023 2:56 PM
To: Charles Riley 
Cc: Histonet@lists.utsouthwestern.edu 
Subject: Re: [Histonet] IHC staining of tendons and cartilage

Hi Charles,

Shirley Powell here in humid Georgia.  I ran an IHC reference lab here for many 
years.  I had a problem with using charged slides for a lot of the tissues I 
processed.  I used manual and automation methods.  My tissues were washing off 
a lot.  I changed to an adhesive for the water bath called Sta-On and I think 
Surgipath was the company that made it.  Surgipath was bought out by Leica but 
they still sold it.  Sta-On was the best adhesive I had found and that worked 
for me for many years.  Whenever I do IHC that is what I use, especially bone, 
cartilage, bloody specimens, autopsy tissues, they stay on better.  Some other 
companies may be selling it now, like VWR/Avantor.  Just Google it.

Shirley

Shirley Powell, HTL(ASCP)
Technical Director Histology Curricular Support Laboratory
Pathology Department
Mercer University School of Medicine
powell...@mercer.edu
Phone: 478-301-2374
https://medicine.mercer.edu/

-Original Message-
From: Charles Riley via Histonet 
Sent: Tuesday, March 21, 2023 2:31 PM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] IHC staining of tendons and cartilage

Hello all,

I am in a new position and it will potentially require doing a lot of IHC 
testing on cartilage and tendon samples. I have done some practice runs on my 
automated stainer and manually and am running into issues with the tissue 
sections falling off completely or folding over on itself during each process.

If anyone does staining like this routinely and has some pointers/tricks to try 
to get the samples to adhere to the slides better it would be greatly 
appreciated.

I have tried using charged slides from a variety of vendors and get similar 
results across the board.
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Re: [Histonet] Conferences 2023

[John Kiernan via Histonet]

The 2023 meeting of the Biological Stain Commission, marking the organization's 
100th anniversary, will be held in Rochester, NY, USA in June 2023. Probably on 
Friday June 9th. There will be invited presentations and poster sessions. 
Details are not yet available but should appear on the Commission's web site at 
some time in the next few weeks. Keep an eye on https://biostain.com.

On the BSC web site you can also see what was presented at about 20 earlier BSC 
meetings (at https://biologicalstaincommission.org/category/annual-meetings/).

The BSC web site also has many other links useful to histotechnologists, 
notably a Glossary and a variety of What's New? items, with conspicuous links 
on the Home page. The glossary comes up as the top item if you type stain 
glossary (or similar) into Google.

I can see, Lisa, that you won't be travelling from Australia to the USA for a 
one-day meeting!   Your Histonet query has provided an opportunity to advertise 
the BSC's forthcoming centennial celebration. The availability of BSC-Certified 
stains (about 60 dyes, as powders) underlies the validity of much medical and 
other biological research and of all routine staining done in pathology labs 
with bought pre-made dye solutions.

Beware of fake "certified" or similarly named stains sold perhaps from China 
and India and marketed in Europe and the Americas.  These do not carry the 
BSC's small and difficult-to-forge bottle label, which subtly indicates the 
history of every truly certified stain.

John Kiernan
https://biologicalstaincommission.org/category/news-about-dyes-and-stains/
= = =

From: Lisa whitham via Histonet 
Sent: December 8, 2022 3:06 PM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Conferences 2023

Hello

I am a grossing supervisor from Australia and am wishing to attend an 
international conference in 2023. Can anyone tell me what conferences are being 
held in 2023?

Kind Regards

Lisa Whitham
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From: Lisa whitham via Histonet 
Sent: December 8, 2022 3:06 PM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Conferences 2023

Hello

I am a grossing supervisor from Australia and am wishing to attend an 
international conference in 2023. Can anyone tell me what conferences are being 
held in 2023?

Kind Regards

Lisa Whitham
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Re: [Histonet] Immunofluorescense staining

[John Kiernan via Histonet]

If the antibody's supplier does not provide instructions for IHC you will need 
to test a wide range of concentrations of the primary antibody on sections 
known to contain the antigen in easily identifiable cell-types or other sites. 
Use a secondary antibody and detection system that you have already found 
reliable for primary antibodies raised in the same species as that in which the 
antibody for flow cytometry was raised. This would probably be a fluorescently 
labelled secondary antibody if "immunoflourescense" is specified.

The researcher probably needs to read a book about immunohistochemistry before 
expecting someone else to do it for her/him. An investment of $100 or so for a 
lab, along with a few hours of reading (by boss and workers), can save many 
hundreds of dollars that might otherwise be spent on antibodies and other 
reagents that don't work as expected. A good one to start with is Suvarna, S. 
K., Layton, C., Bancroft, J. D., eds. 2018. Bancroft's Theory and Practice of 
Histological Techniques 8th ed. London: Churchill Livingstone Elsevier. ISBN: 
978-0-7020-6864-5. (NO, I don't have any vested interest!). I'm sure other 
histonetters will come up with similar and perhaps better suggestions.

Beware of working from "protocols" informally passed around among technicians, 
grad students and research fellows without published references (papers, books) 
that you can check before investing time, effort and money in a technique 
that's new to you. Informal lab notes are often locally treated as if they 
"have Authority,  not as the Scribes". In practical science it's probably best 
to go first with the Scribes, because their writings have been severely 
reviewed (if in good journals). Good books have references to peer-reviewed 
papers.

I could go on and on, but that's enough of my grumpy advice for now.

John Kiernan
London, Canada
https://www.schulich.uwo.ca/anatomy/people/faculty/emeriti/kiernan_john.html
= = =

From: Charles Riley via Histonet 
Sent: August 24, 2022 2:55 PM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Immunoflouresence staining

I have been given an antibody that is used for flow cytometry yet the
researcher wants to use it for IF staining.  The protocol they provided is
for the flow cytometry staining. Will this produce the immunoflourescense
results they are looking for or do i need to use a different method?
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Re: [Histonet] The Passing Of Dr. James McCormick

[John Kiernan via Histonet]

Thank you, Tom Pella, for the web links.
Did James McCormick really invent the first cryostat?

I have always seen the early chapters of the late AGE Pearse's Histochemistry 
book as a good source for the history of the cryostat. In his 2nd (1960), 3rd 
(1968) and 4th (last, 1980 ISBN 0443019983) editions, Pearse cited many 
publications about the development of the cryostat in the late 1950s and early 
1960s but he did not mention McCormick.
I've not seen the 1st (1953) edition of Pearse's book. It may predate the 
cryostat.

As a student in the UK in 1962 I was taught how to use a "Pearse-Slee" cryostat.
It was quite similar to ones made since 2000 and used by exploited graduate 
students to this day. The temperature controls have steadily improved over the 
decades, and by the 1980s the microtome within the freezer had become something 
better than the Cambridge Rocker in the Slee cryostats of the 1960s.

The anti-roll plate to allow collecting a flat section was in place 60 years 
ago, and using it was never easy. It is necessary for doing some enzyme 
activity histochemistry methods (dehydrogenases, cytochrome oxidase etc) on 
nominally thin (~10um) sections of unfixed tissue.

The later invention of the vibrating microtome (Vibratome) has made it quite 
easy to collect and handle sections of unfixed specimens that have not been 
frozen, but the sections have to be quite thick (50um or more). That is often 
OK in research, especially for sections of brains. It is also comparable to 
traditional frozen sections for surgical pathology, which were cut from pieces 
of tissue briefly immersed in boiling hot 4% formaldehyde. The sections were 
cut with a traditional freezing microtome, collected from knife as they melted, 
and deftly moved onto slides with a paintbrush. This technology is not extinct.

John Kiernan
= = =

From: Tom Pella via Histonet 
Sent: July 23, 2022 3:44 PM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] The Passing Of Dr. James McCormick

I've skimmed over the posts to Histonet since late June and I haven't seen
any post on Histonet, where the passing of Dr. James McCormick was
mentioned. He died on June 26th, 2022. I saw this mentioned on Histology
Professionals on FB but not here.

Here are a few links to online obituaries, the second with some rolling
pictures:

https://www.legacy.com/us/obituaries/chicagotribune/name/james-mccormick-obi
tuary?id=35647547

https://www.swedishhospitalfoundation.org/news/in-memoriam-dr-james-mccormic
k

His contributions to the field of Histology can't be overstated. He invented
the first Cryostat, the first Histology Automated Tissue Processor, the
first Embedding Center, the first Tissue Cassette (and many other subsequent
cassettes), the first processes to use this instrumentation. His inventions
are in use every single day in most Histology Labs worldwide.

I only became acquainted with Dr. McCormick for a brief time later in his
life on a product collaboration. I found him to be the consummate gentleman;
a person in whom ideas were always bubbling to the surface; gracious and
intelligent and witty. His passing is a great loss for this community. His
public memorial service was held just today.

Tom Pella
President
Ted Pella, Inc.








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Re: [Histonet] Dr Freida Carson

[John Kiernan via Histonet]

Freida Carson was also well known to the Biological Stain Commission. She came 
to our meetings, and she will be missed by our members.
John Kiernan
Secretary, Biological Stain Commission Inc.
https://biologicalstaincommission.org
= = =

From: Tony Henwood (SCHN) via Histonet 
Sent: January 12, 2022 5:25 PM
To: Shirley A. Powell ; Willis, Donna G 

Cc: histonet@lists.utsouthwestern.edu 
Subject: Re: [Histonet] Dr Freida Carson

I agree,

She will be missed


Regards
Tony Henwood JP, MSc, BAppSc, GradDipSysAnalys, CT(ASC), FFSc(RCPA)
Principal Scientist, the Children's Hospital at Westmead
Adjunct Fellow, School of Medicine, University of Western Sydney
Tel: 612 9845 3306
Fax: 612 9845 3318
Pathology Department
the children's hospital at westmead
Cnr Hawkesbury Road and Hainsworth Street, Westmead
Locked Bag 4001, Westmead NSW 2145, AUSTRALIA


-Original Message-
From: Shirley A. Powell via Histonet [mailto:histonet@lists.utsouthwestern.edu]
Sent: Thursday, 13 January 2022 7:59 AM
To: Willis, Donna G 
Cc: Histonet (histonet@lists.utsouthwestern.edu) 

Subject: Re: [Histonet] Dr Freida Carson

Hi Donna,
So sorry to hear of this. I am proud to have known her and been a recipient of 
her wealth of knowledge.
She contributed so  much to the Histology Community.

Shirley

Shirley Powell, HTL(ASCP)
Technical Director Histology Curricular Support Laboratory Pathology Department 
Mercer University School of Medicine
1550 College St, Macon, GA  31207
O: 478-301-2374/F:478-301-5489
medicine.mercer.edu




-Original Message-
From: Willis, Donna G via Histonet 
Sent: Wednesday, January 12, 2022 3:19 PM
To: Histonet (histonet@lists.utsouthwestern.edu) 

Subject: [Histonet] Dr Freida Carson

For those of you that are not on Facebook you may not already know that 
yesterday Freida Carson, PhD became one of our Histology Angels.  I had the 
privilege of having her as my Histology Educator long before she wrote her 
first edition of Histotechnology.  But more precious to me than having her as 
my educator was that she was a friend.  She will be missed.  Thank you Freida 
for all that you have given to the Histology Profession and to me.  Rest In 
Peace singing in the Angels Choir.

Donna Willis
Anatomic Pathology Manager
Baylor Scott Health
Baylor University Medical Center
3500 Gaston Ave|Dallas, Texas 75246
214-820-2465 office|214-725-6184 mobile



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Re: [Histonet] Paraffin embedding following storage in 70% alcohol

[John Kiernan via Histonet]

Of course you are right!
This is yet another example of an error in a procedure informally handed on 
from person to person! Always work from a book. Even a very old one will be OK 
for paraffin embedding.
John Kiernan.
https://www.schulich.uwo.ca/anatomy/people/faculty/emeriti/kiernan_john.html
= = =

From: Charles Riley via Histonet 
Sent: November 12, 2021 11:03 AM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Paraffin embedding following storage in 70% alcohol

I am working with a grad student on a project dealing with equine articular
cartilage.

The protocol she sent me for embedding the tissue samples goes directly
from 70% alcohol to the embedding step in paraffin.


Correct me if I am wrong but shouldn't the tissue be dehydrated fully and
cleared before embedding the samples?
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Re: [Histonet] Reprocessing Protocol

[John Kiernan via Histonet]

It makes no sense to heat a paraffin-infiltrated specimen in saline. Sodium 
chloride isn't soluble in hot paraffin or in any of the organic solvents used 
in tissue processing.

Heating in water may melt and float out all the wax and rehydrate a specimen, 
but does it matter if some wax remains?

Reprocessing is an attempt to correct the effects of incomplete dehydration. 
This can be done by taking the specimen back into the clearing agent (xylene or 
similar) and then into 2 changes of 100% alcohol (methyl, ethyl or isopropyl). 
Why rehydrate?

For a rehydrated specimen, why go "from formalin, using a schedule that would 
have been of appropriate length as used initially"?  This will surely produce, 
again,  a block that is incompletely dehydrated.

John Kiernan
London, Canada
= = =

From: Etheridge, Sandra AFF:EX via Histonet 
Sent: October 12, 2021 5:19 PM
To: histonet@lists.utsouthwestern.edu 
Subject: Re: [Histonet] Reprocessing Protocol

Hi Curt,

We have used Taggart's Method quite successfully in the past for poorly 
processed tissues.  You can find it online.  The isotonic saline may help to 
rehydrate your tissues.

1.  Melt down the tissue block in the embedding centre block tray area and 
gently blot off the excess wax. Place the tissue in a newly labelled cassette.
2.  Place the cassette into a beaker of isotonic saline (0.9% sodium chloride) 
and place it in the 65 C incubator/oven for one hour. This will melt the 
residual wax which will rise to the surface of the saline.
3.  Remove the cassette from the saline, drain briefly and place in your 
processor from formalin, using a schedule that would have been of appropriate 
length as used initially.
4.  Embed and section as per usual.

Good luck!

Sandra Etheridge

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

Sent: October 12, 2021 10:00 AM
To: histonet@lists.utsouthwestern.edu
Subject: Histonet Digest, Vol 215, Issue 8

[EXTERNAL] This email came from an external source. Only open attachments or 
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Today's Topics:

   1. reprocessing tissue (Curt Tague)


--

Message: 1
Date: Tue, 12 Oct 2021 15:14:54 +
From: Curt Tague 
To: "histonet@lists.utsouthwestern.edu"

Subject: [Histonet] reprocessing tissue
Message-ID:



Content-Type: text/plain; charset="us-ascii"

I have a problem... some tissue got processed very poorly, there was water in 
the system somewhere and a few blocks just look burnt.. the nuclei are faint 
and cloudy, no detail at all. I've tried the process of rehydrating with the 
30% formaldehyde, glycerol and sodium acetate solution but they still process 
poorly, come out very brittle and just don't look good under the scope.

Does anyone have a magic bullet to salvage these specimens? I can send a pic 
directly if it helps.

Thanks,
Curt



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Re: [Histonet] frozen section problem

[John Kiernan via Histonet]

Yes, definitely ice crystal holes! If the tissues are unfixed you will have to 
freeze much more rapidly (isopentane cooled with liquid nitrogen.) If fixed in 
formaldehyde, cryoprotect by immersing the pieces in 20% sucrose, until they 
sink.

John Kiernan
Anatomy & Cell Biology, UWO
London, Canada
= = =

From: Bonello Dorianne M at Health-MDH via Histonet 

Sent: July 16, 2021 11:25 AM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] frozen section problem

Dear all,


We are experiencing freezing artifacts on our frozen sections. Basically, we 
are seeing cavity-like structures under the microscope, mostly elongated, 
especially when it's a frozen section on brain tissue. This is most probably 
happening due to ice crystal formation. We're not using cryospray, relying only 
on the cryobar boost function.


Does anyone has a solution to this problem please?


Regards,



Dorianne Bonello
Allied Health Practitioner (MLS)
Histology Laboratory - Pathology
Health-Mater Dei Hospital


[cid:image001.jpg@01D67184.63288530]


T +356 +356 25456434

E dorianne.m.bone...@gov.mt


Mater Dei Hospital, Triq id-Donaturi tad-Demm, l-Imsida, Malta MSD 2090 | Tel 
+356 2545  | 
https://deputyprimeminister.gov.mt/en/MDH/Pages/Home.aspx
 | https://www.facebook.com/materdeihospital/


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Re: [Histonet] Prussian Blue Reaction

[John Kiernan via Histonet]

Overstained? Doesn't that mean the tissue contains a lot of iron and you are 
seeing where it is - which was the reason for doing Prussian blue 
histochemistry. Gudrun Lang correctly says that mineral acids won't remove it. 
Oxalic acid is said to dissolve Prussian blue (? by chelation); I've never 
tried this. If it works, you will no longer see where the iron is. To see 
features other than the distribution of iron, why not just stain another 
section from the block with a general-purpose stain like Giemsa or H?
John Kiernan
London, Canada
= = =

From: Mac Donald, Jennifer via Histonet 
Sent: June 7, 2021 12:47 AM
To: Gudrun Lang 
Cc: histonet@lists.utsouthwestern.edu 
Subject: Re: [Histonet] Prussian Blue Reaction

The tissue was overstained and the blue was interfering with interpretation

-Original Message-
From: Gudrun Lang 
Sent: Sunday, June 6, 2021 2:18 AM
To: Mac Donald, Jennifer 
Cc: histonet@lists.utsouthwestern.edu
Subject: AW: [Histonet] Prussian Blue Reaction

  EXTERNAL SENDER- Exercise caution with requests, links, and attachments.

Hi Jennifer,
Why do you want to reduce the staining?

I ask, because the impact of hydrochloric acid on the tissue may influence the 
following results anyway.
I think, that the prussian blue pigment cannot be removed in an easy way. It is 
resistent to solvents and mineral acids.
https://nam12.safelinks.protection.outlook.com/?url=http%3A%2F%2Fwww.epsilonpigments.com%2Finorganic-pigment%2Fprussian-blue%2FPrussian-Bluedata=04%7C01%7Cjmacdonald%40mtsac.edu%7C0fbc82a2b13749a4222608d928cbfe52%7Ccc4d4bf20a9e4240aedea7d1d688f935%7C0%7C0%7C637585679205067185%7CUnknown%7CTWFpbGZsb3d8eyJWIjoiMC4wLjAwMDAiLCJQIjoiV2luMzIiLCJBTiI6Ik1haWwiLCJXVCI6Mn0%3D%7C2000sdata=KjvijcfrVPGZKGsAn6qX5rMKtulHpmsAzqHEkwz%2B96Y%3Dreserved=0
-for-Solvent-Based-Inks.html

On the other hand, if the blue colour doesn't interfere with your following 
staining, you can try to simple make a "double stain".

Regards
Gudrun

-Ursprüngliche Nachricht-
Von: Mac Donald, Jennifer via Histonet
[mailto:histonet@lists.utsouthwestern.edu]
Gesendet: Sonntag, 6. Juni 2021 06:34
An: histonet@lists.utsouthwestern.edu
Betreff: [Histonet] Prussian Blue Reaction


Does anyone know of a way to remove/reduce the Prussian blue reaction?
Thanks,
Jennifer



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Re: [Histonet] Protocol for DAPI staining on paraffin sections

[John Kiernan via Histonet]

Not everyone knows that the product of the Feulgen reaction is fluorescent. If 
you do the method on paraffin sections, DNA in nuclear chromatin is red with 
ordinary illumination. If you look with a fluorescence microscope (blue 
excitation) it shows as a brownish-red fluorescence in the chromatin. A similar 
fluorescent colour is seen in conventional PAS-stained sections, in mucus, 
basement membranes, etc. You need a non-fluorescent mounting medium: DPX or one 
of the various poly(methyl methacrylate)-based media such as Entellan.
John Kiernan
   Professor Emeritus, Dept of Anatomy & Cell Biology
   University of Western Ontario, London, Canada
  https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html
   Also  Secretary, Biological Stain Commission, Inc.
  
https://biologicalstaincommission.org
= = =





From: Alida Bailleul via Histonet 
Sent: June 3, 2021 7:53 AM
To: Histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Protocol for DAPI staining on paraffin sections

Dear Histonet list,

I have made some paraffin blocks of demineralized bone and cartilage. I
would like to stain paraffin slides with DAPI (or another DNA stain).
Does DAPI staining (or other fluorescent DNA stain) work well on paraffin
sections? Does anyone have a protocol they would like to share?

I am assuming that histological stains (like the Feulgen stain would work
better?). I could also try both.

Thank you in advance

All the best

Alida

--
Dr. Alida M. Bailleul
Associate Research Fellow/Associate Professor
Institute of Vertebrate Paleontology and Paleoanthropology, Chinese Academy
of Sciences
www.ivpp-avianevolution.com
& Research Associate of Paleontology, Museum of the Rockies, Montana State
University
Google Scholar
 -
ResearchGate

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Re: [Histonet] IF with permanent mounting media?

[John Kiernan via Histonet]

Espada J, Juarranz A, Galaz S, Canete M, Villanueva A, Pacheco M, Stockert JC 
(2005) Non-aqueous permanent mounting for immunofluorescence microscopy. 
Histochem. Cell Biol. 123: 329-334.  https://doi.org/10.1007/s00418-005-0769-2
A PDF can be downloaded from the Google Scholar entry for this article.

John Kiernan
London, Canada
= = =

From: Morken, Timothy via Histonet 
Sent: May 28, 2021 11:31 AM
To: Histonet 
Subject: [Histonet] IF with permanent mounting media?

Has anyone tried using xylene/permanent mounting media for immunofluorescence 
stains? I had a question from a pathologist who wondered if we could do this. I 
have never heard of anyone doing it.

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

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[Histonet] Luxol fast blue for staining myelin. Validity of sources.

[John Kiernan via Histonet]

This Histonet query is addressed to vendors of stains (powders and/or 
solutions), and to anyone who makes up luxol fast blue in the lab, avoiding the 
high cost of buying and transporting the flammable ready-made staining solution.

Most diagnostic labs buy ready-made staining solutions and follow a vendor's 
instructions, whereas research labs are likely to buy dye powders, make up 
their own solutions, and use them according to published books or papers. This 
means that the largest amounts of luxol fast blue (LFB) powder are sold to 
vendors of staining solutions. Am I right?

Are vendors having any problems with getting dye powders called luxol fast blue 
(LFB) that are consistently OK for staining myelin sheaths of axons in sections 
of brain, spinal cord etc?

The LFB name is informally attached to at least three dyes used as stains: LFB 
G (CI Solvent blue 34) and LFB ARN (CI Solvent blue37) are azo dyes, whereas 
LFB MBS (CI 74180, Solvent blue 38) is a copper phthalocyanine. All three have 
been used in similar (but not identical) published staining methods for myelin. 
LFB MBS is the dye most often used for this purpose, but not much is known 
about variation in the purity of commercial batches. It has many commercial 
uses and dozens of trade-names. See 
http://www.worlddyevariety.com/category/solvent-dyes. (It's free, unlike the 
Colour Index, but don't believe everything it says.)

Luxol fast blue (whichever one) may or may not be the best stain for showing 
myelinated axons and regions of demyelination, but it probably is the dye most 
used for doing this job. Is there a need for independent third-party 
identification, testing and certification of LFB powders? If you have 
encountered a bad jar of powder labeled luxol fast blue, please reply, 
including if possible the name on the label and the source.

The Biological Stain Commission (BSC) has never offered testing and 
certification for LFB as a stain for myelin, even though "luxol" dye have been 
in use for this job since 1953. Should third-party certification be made 
available for luxol fast blue? Would you submit samples for this service? For 
more information, see the link below.

John Kiernan
Secretary, Biological Stain Commission, Inc.
https://biologicalstaincommission.org/for-vendors/
For Vendors and Users of Stains | The Biological Stain 
Commission
Vendor FAQs: The Biological Stain Commission is a not-for-profit organization 
which is dedicated to the endorsement of histological materials. It is the 
mission of the BSC to ensure the quality of biological stains on the market 
that are sold by many US companies.
biologicalstaincommission.org
= = =







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Re: [Histonet] doing Sudan Black B

[John Kiernan via Histonet]

Hydrated paraffin sections of formaldehyde-fixed tissue are stained the same 
way as frozen sections of formaldehyde-fixed tissue, but in paraffin sections 
lipofuscin inclusions are just about the only things that stain. They are, of 
course, brown or yellow without any staining. If you need to identify 
lipofuscin inclusions with certainty you'll have to do a few other stains and 
histochemical tests to distinguish them from ceroid (a similar glycolipoprotein 
pigment), melanin and haemosiderin.

It is not a good idea to use a protocol informally passed on from someone else 
without checking its origins. Even a method printed in a book can contain 
errors, and for this reason just about all the methods books published since 
about 1950 have references to peer-reviewed papers that readers can and 
sometimes should check. Any book in our field is just a big collection of 
review articles that have not been reviewed as critically as more specialized 
reviews in a good journal.

The late RD Lillie made important original contributions to pigment 
histochemistry (two are cited below) and his techniques book (also cited) 
intelligently reviews the subject and provides technical instructions. So does 
Pearse's Histochemistry.

There are, of course, protocols for staining lipofuscin in all the ordinary 
histotechniques books. You don't need to have all the latest editions on your 
lab's bookshelf. The 3rd (1965) edition of Lillie is available for only $8 at 
https://www.abebooks.co.uk/book-search/title/histopathologic-technic-and-practical-histochemistry/author/r-d-lillie/
Histopathologic Technic and Practical Histochemistry by R D Lillie - 
AbeBooks
Histopathologic Technic and Practical Histochemistry by Lillie, R D and a great 
selection of related books, art and collectibles available now at 
AbeBooks.co.uk.
www.abebooks.co.uk
Snap it up! The 4th and last (1976) edition of Lillie is now $700 from Amazon, 
but the 1965 edition is still OK for most of what the book is about, and it's 
easier to read than his rather rambling 4th edition.

Lillie RD, Fullmer HM (1976) Histopathologic Technic and Practical 
Histochemistry. 4th ed. McGraw-Hill, New York. See comments above - the 
previous edition is cheaper and for you it may be just as good.

Pearse AGE, Stoward PJ (1980,1985,1991) Histochemistry, Theoretical and 
Applied, 4th edn. Vol. 1. Preparative and Optical Technology. Vol. 2. 
Analytical Technique. Vol. 3. Enzyme Histochemistry. v. 1,2,3. 
Churchill-Livingstone, Edinburgh.  Pigments are in Vol. 2. Third edn (1968. 2 
vols) probably just as good for most labs.

Lillie RD (1956) The mechanism of Nile blue staining of lipofuscins. J. 
Histochem. Cytochem. 4:377-381. I think all the papers in this journal are now 
free for anyone to download.

Lillie RD (1956) A Nile blue staining technic for the differentiation of 
melanin and lipofuscin. Stain Technol. 31:151-153. Sadly, not free unless you 
have access to a subscribing library or are a member of the Biological Stain 
Commission.

I hope this answer helps.
John Kiernan
= = = =

From: LEROY H BROWN via Histonet 
Sent: May 9, 2021 10:29 PM
To: histonet@lists.utsouthwestern.edu 
Subject: Re: [Histonet] doing Sudan Black B

Hi,  I am looking for a protocol for Sudan Black B staining on paraffin
embedded tissue.   Does anyone have a working stain for this?

Thanks

LeRoy Brown HT(ASCP) HTL
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Re: [Histonet] Movats

[John Kiernan via Histonet]

Dear Betsy,

Don't say you are sorry for putting a long post on Histonet! To get 
troubleshooting help you need to say exactly what you did. If you wrote only, 
"why are my sections brown after Movat staining", nobody would understand your 
problem.

Your procedure starts with an hour in hot Bouin. For many years this has been a 
routine prior to trichrome stains done on sections of specimens fixed in 
neutral formaldehyde. It isn't part of Movat's original method (Arch. Path. 
60:209-295, 1955), which probably was devised for sections optimally fixed for 
trichrome staining (in mixtures containing mercuric chloride).

Movat's pentachrome is a trichrome method preceded by alcian blue (for no 
obvious reason) and an iron-haematoxylin for nuclei and elastin. It differs 
from older trichromes in using a mixture of yellow polyene dyes  (saffron) to 
stain the collagen, instead of the blues or greens as in the Mallory and Masson 
methods.

Your method includes "5% sodium thiosulfate -1 min" after the iron-haematoxylin 
stain for black nuclei and elastic fibres. This also isn't part of Movat's 
pentachrome method, and I wonder why. Did you inherit an informal list of 
instructions passed on within the lab?  After a mercuric fixative, hydrated 
sections are dipped in iodine, followed by thiosulphate, before staining, to 
remove a black deposit (probably mercurous chloride) introduced by the 
fixative.I've been seeing similar informal passing of bad staining instructions 
in research labs for many years.  Are you a victim of this trend?

The thiosulphate step in your procedure obviously does no harm, because you got 
the right results with the dog tissues. There may be something different about 
your human specimens: perhaps inadequate fixation, or excessive acid treatment 
(if that's what Cal rite is) for decalcification.

If the sections of human arteries look OK with a microscope, it might not 
matter that grossly they are a different colour from the dog small intestine 
sections. They are, after all, different tissues.

A rather long, and not very helpful reply!

John Kiernan
London, Canada
= = =

From: Betsy Molinari via Histonet 
Sent: May 5, 2021 9:46 AM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Movats

Hi Histonetters,
I have received several human vessels for paraffin processing and to stain the 
sections for H and Movats. The H were fine. The human sections turned 
brownish yellow with the Movats.The control which is canine small intestine was 
perfect.
The protocol is standard
Bouins 1hr in 58C waterbath
Rinse till yellow disappears
Rinse in DH2O
1% Alcian Blue -20 min
Rinse in running tap H2O -5min
Alkaline alcohol-1hr
Rinse 10 min tap H2O
Rinse in DH2O
Verhoff's Hematoxylin -15 min
3 changes DH2O
Differentiate in 2% FeCl
Rinse in DH2O
5% sodium Thiosulfate -1min
Rinse in running tap-10 min
Rinse in DH2O
Woodstain scarlet/acid fuchsin-1.5 min
Rinse in DH2O
Rinse in 0.5% acetic acid water
5% aqueous phosphotungstic acid -2 changes 5 min each
Rinse in 5% acetic acid water
Rinse in 3 changes absolute ETOH
6% alcoholic  Safran solution
Absolute alcohol-xylene-coverslip
The human slides were fine until the Safran step. When I removed them from the 
stain into the 100% they were a yellowish brown .Under the scope the colors 
were there, blue, red, yellow and black. But on the slide the tissue was that 
brownish yellow. The researcher does not like to strong yellow color. Since my 
control was fine I question if something was going on with their tissue. I do 
not know how the tissue was handled before it came into the lab. They were very 
calcified and were decaled for 1-3 days in Cal Rite. I do know they were not 
rinsed after decal and were put straight back into 10% NBF before I got them 
for processing.
Should I have used a human control instead of canine?  These were very large 
pieces that were crammed into the cassette.
Thanks for the help. Sorry for the long post.
Betsy Molinari HT(ASCP)
Texas Heart Institute
Cardiovascular Pathology
1101 Bates St.
Houston,TX
832-355=6524 (lab)
832-355-6812 (fax)



Betsy Molinari, HT (ASCP)
Sr. Histology Research Technician
CV Pathology Research

Texas Heart Institute
6770 Bertner Avenue, MC 1-283
Houston, TX 77030

Office: 832-355-6524 | Fax: 832-355-6812
Email: bmolin...@texasheart.org
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CONFIDENTIALITY NOTICE: This email and attachments contain information that may 
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Re: [Histonet] Animal Histology

[John Kiernan via Histonet]

Jennifer MacDonald asked, "Can anyone recommend a good book for processing 
animal tissue?" Here's a list of 14 of the ones published since 1990, with very 
brief descriptive notes. There are, of course, plenty of good books that are 
older than these.

Lyon,H (1991): Theory and Strategy in Histochemistry. A Guide to the Selection 
and Understanding of Techniques. Springer-Verlag, Berlin. All on theory; no 
instructions. All aspects of preparation, staining etc are covered. 32 
chapters, by different people; one bibliography. Reprinted 2011. ISBN 
9783642737442.

Kok,LP; Boon,ME (1992): Microwave Cookbook for Microscopists. Art and Science 
of Visualization. 3rd ed.  Coulomb Press Leyden, Leiden. 432 pages. Plenty of 
microwave theory. Applications in fixation, infiltr, embedding, decalcif., many 
staining methods including conn.tiss., cytol., neuro. Enzyme, immuno- & 
hybridization histochemistry.

Sanderson,JB (1994): Biological Microtechnique. (Microscopy Handbooks, 28.) 
BIOS Scientific Publications & Royal Microscopical Society, Oxford. 224 pages.  
Detailed paractical instructions, with rationale, & titled bibliography for 
each of the 7 chapters. Collection, Fixation, Processing, Microtomy, Other 
prep. methods, Staining, Finishing (mounting etc).

Chayen,J; Bitensky,L (1991): Practical Histochemistry. 2nd ed. Wiley, 
Chichester. 321 pages. Special emphasis on use of unfixed cryostat sections.

Presnell,JK; Schreibman,MP (1997): Humason's Animal Tissue Techniques. 5th ed. 
Johns Hopkins University Press, Baltimore. 572 pages. Humason's Animal Tissue 
Techniques, 5th edition. Includes chapters on immunohistochem, safety, 
microwaves, lab hints, suppliers, etc.

Hayat,MA (1993): Stains and Cytochemical Methods. Plenum Press, New York. Book 
of techniques. Many are critically discussed. emphasis on EM, but plenty of LM 
too.

Carson,FL; Hladik,C (2009): Histotechnology. A Self-Instructional Text. 3rd ed. 
American Society of Clinical Pathologists, Chicago. 400 pages.  Freida Carson's 
textbook. 3rd ed, with Hladik, is 96 pages longer than 2nd (1997); with EM and 
cytopreparation; glossary. ISBN 9780891895817. (There is a 4th ed, 2015, which 
I haven't seen.)

Kiernan,JA (2015): Histological and Histochemical Methods: Theory and Practice. 
5th ed. Scion, Banbury, UK. 606 pages.

Kumar,GL; Kiernan,JA (Eds.) (2010): Education Guide: Special Stains and H & E. 
2nd ed. Dako North America, Carpinteria, CA. 300 pages.  33 chapters. Photos. 
PDF file is a free download from 
https://www.agilent.com/en/dako-pathology-education-guides/.

Wick,MR (Ed.) (2008): Diagnostic Histochemistry. Cambridge University Press, 
New York. 460 pages.  Multi-author book. Applications of techniques to path. 
diagnosis. Chapters by systems, organs, diseases etc. Emphasizes value, economy 
of pre-1970s (before immuno) histochemical methods. Only Ch1, pp.1-27 describes 
techniques. Many colour photos.

Orchard,G; Nation,B (Eds.) (2012): Histopathology. Oxford University Press, 
Oxford, UK. 396 pages. Multi-author textbook book. Applications of techniques 
to path. diagnosis. Chapters mainly by methods, with applications to systems, 
organs, diseases. One of an Inst. of Biomed. Science series, with web site at 
www.oxfordtextbooks.co.uk/orc/fbs/.

Mulisch,M; Welsch,U (Eds.) (2015): Romeis - Mikroskopische Technik. 19th ed. 
Springer-Verlag, Berlin. 603 pages. 25 chapters, 3 appendices. All types of 
microscopy, staining, histochem, etc. (in German) ISBN 9783642551901.

Exbrayat,JM (Ed.) (2013): Histochemical and Cytochemical Nethods of 
Visualization. (Series Ed: Morel,G. Methods in Visualization.) CRC Press, Boca 
Raton, FL. 335 pages. 13 chapters, 10 contributors. 1-7 are for LM; 8-12 for 
EM; 13 on image quantification. ISBN 978143987.

Suvarna,SK; Layton,C; Bancroft,JD (Eds.) (2018): Bancroft's Theory and Practice 
of Histological Techniques. 8th ed. Churchill Livingstone Elsevier, London. 672 
pages. In this 8th ed., histochem chapters for lipids, proteins, nucleic acids, 
enzyme activities are now compressed into Appx I (of VIII).

Hope this helps. For older books, look in AbeBooks, Amazon etc for ones by GG 
Brown, HC Cook, CFA Culling, RAB Drury, M Gabe, P Gray, RW Horobin, G Humason, 
JFA McManus, AGE Pearse.

John A. Kiernan
Dept of Anatomy & Cell Biology
University of Western Ontario
London, Canada
= = =

From: Mac Donald, Jennifer via Histonet 
Sent: May 3, 2021 1:07 AM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Animal Histology


Can anyone recommend a good book for processing animal tissue?
Thanks,
Jennifer


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Re: [Histonet] Oil Red O

[John Kiernan via Histonet]

It's in any textbook in the field of histotechnology published since about 
1950. This one from Amazon costs less than $8. Every lab should have a shelf of 
such books.

https://www.amazon.com/Introduction-Histotechnology-Geoffrey-G-Brown/dp/0838543405
[https://images-na.ssl-images-amazon.com/images/I/41xQTLdptPL.jpg1z]m]
An introduction to histotechnology: A manual for the student, practicing 
technologist, and resident-in-pathology: Brown, Geoffrey G: 9780838543405: 
Amazon.com: 
Books
An introduction to histotechnology: A manual for the student, practicing 
technologist, and resident-in-pathology [Brown, Geoffrey G] on Amazon.com. 
*FREE* shipping on qualifying offers. An introduction to histotechnology: A 
manual for the student, practicing technologist, and resident-in-pathology
www.amazon.com
John Kiernan
London, Canada
= = =

From: Niihori, Maki - (mniihori) via Histonet 

Sent: March 5, 2021 4:08 PM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Oil Red O

We would like to stain Right Ventricle (RV) and Lung (both from rat samples) 
with Oil Red O.
I appreciate if anybody can share a good protocol/kit information with me.

Thank you,
Maki


*

Maki Niihori, PhD

Life Sciences North Rm# 402,

Department of Medicine,

The University of Arizona

Phone: 520-626-6092

E-mail: mniih...@arizona.edu

*

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Re: [Histonet] Other Histonet-like listservers?

[John Kiernan via Histonet]

Try this.

https://biologicalstaincommission.org/bscglossary.html
Glossary of Staining Methods, Reagents, Immunostaining, Terminology and 
Eponyms. Currently Version 1.2.
Version 2.0 with another 300 or so items will go online quite soon.

Other items at http://biologicalstaincommission.org include QUESTIONS AND 
ANSWERS - Staining, histochemistry and histotechnology FAQ (Frequently Asked 
Questions),  Version  2.0 November 2019, at 
https://biologicalstaincommission.org/faqlist.htm, and information about 
BSC-certified stains.

There's also Bryan Llewellyn's StainsFile site:  
https://stainsfile.info/xindex.html, Last updated January 2019.

John Kiernan
   J. A. Kiernan MB, ChB, PhD, DSc
   Professor Emeritus, Dept of Anatomy & Cell Biology
   University of Western Ontario, London, Canada
  https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html
   Also Secretary, Biological Stain Commission, Inc.
  https://biologicalstaincommission.org
= = =

From: Morken, Timothy via Histonet 
Sent: February 11, 2021 11:39 AM
To: Histonet 
Subject: [Histonet] Other Histonet-like listservers?

Hi all, I'm going to give a presentation on online histology help sites like 
Histonet and the NSH Block. Does anyone know of other listservers that can be 
of help to histotechnologists in all fields? I checked the MSA EM listserver 
but it is down for some technical reason.

Thanks in advance for any help!

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

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Re: [Histonet] Alcian Blue staining

[John Kiernan via Histonet]

Pink with alcian blue?  Are you using an alcian blue pH2.5-PAS sequence?  If so 
you should get PAS-positive mucus (not in goblet cells) in the stomach and 
AB-positive mucus (greenish blue) in the intestine. Intestinal mucus, 
especially in the duodenum and jejunum, is also PAS +ve, so the cells appear 
purplish.  In the colon the mucus is mostly sulphated rather than sialylated, 
so it is only AB +ve, even if the stain is done at pH 1.

If a PAS stain is done before the AB, PAS +ve mucus also stains quite strongly 
with AB. For more about this, see Johannes ML & Klessen C (1984) Alcianblue/PAS 
or PAS/alcianblue. Remarks on a classical technic used in carbohydrate 
histochemistry. Histochemistry 80: 129-132. (Unfortunately the paper has only 
black & white photos.)

Various dyes are currently sold as "alcian blue" and not all are suitable for 
all applications. The Biological Stain Commission now certifies alcian blue 
dyes as either "alcian blue 8G or equivalent" or as "alcian blue variant".  See 
the recently updated (2020) entry for Alcian blue 8G (CI 74240) and other 
alcian blue dyes at   https://biologicalstaincommission.org/new-dyes/.
Current issues of interest to vendors and users of dyes and biological stains. 
| The Biological Stain 
Commission
Dyes are becoming more expensive! January 2018. BASF, a major dyestuff 
manufacturing company, recently announced that it has increased its prices for 
many pigments and dyes by up to 15% worldwide.
biologicalstaincommission.org
John Kiernan
Secretary. Biological Stain Commission
= = =

From: Charles Riley via Histonet 
Sent: October 9, 2020 10:14 AM
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Alcian Blue staining

We are having an issue where the goblet cells in our control tissue are not 
staining pink even though the patient tissue is staining beautifully on the 
same slide.

The stain is done manually.  What can be some causes for the issues in staining 
on the control section?
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Re: [Histonet] human shoulder joint fixation

[John Kiernan via Histonet]

Thanks for the compliments, Bob, but I've no experience of trying to section 
anything as big and bony as a human shoulder joint. Car Hobbs and Izak 
Dimenstein probably will be able to give better advice to Merissa.

Incidentally, I wasn't ever a pathologist. I moved from medicine to 
neuroanatomy (9-5 job!) at an early age. Some of my research could have been 
called experimental pathology. I also collaborated with some real 
neuropathologists, especially in the 1990s looking at the cerebral cortex and 
spinal cord in ALS and other motor neuron diseases.

Cheers,  John Kiernan.
https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html
= = =

From: Bob Richmond via Histonet 
Sent: 21 August 2020 17:25
To: Histonet@lists.utsouthwestern.edu 
Subject: Re: [Histonet] human shoulder joint fixation

>
> Merissa (where?) asks; >>I am doing some planning for a new project and
> wanted to get opinions on fixation of large pieces of tissue. We will have
> human shoulders, where we want to preserve the rotator cuff/joint. Cutting
> the tissue with a saw will damage the soft tissue, so we were thinking that
> post-fixation would be best for cutting slabs. - Does anyone have
> experience with fixing such large pieces of tissue? We typically use zinc
> buffered formalin for fixation. Would a vacuum work or a vacuum sealer?<<
>

This is a non-trivial problem, and I hope you're working closely with the
pathologist or other investigator who's going to be looking at the slides.

Formalin (forget the zinc) penetrates tissue slowly enough that you're not
going to get very good fixation if you put the whole specimen in fixative
and forget about it for a week or two. Some preliminary dissection is
needed to aid fixation, and you're going to need some serious help with
that.

A valuable resource person on Histonet is John Kiernan, a pathologist
turned research anatomist, and an expert on histologic technique. I hope he
responds, otherwise try to find him.

Bob Richmond
Samurai Pathologist
Maryville TN
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Re: [Histonet] Apple Green Birefringence in Amyliod slides

[John Kiernan via Histonet]

For another source of polarizing filters, go to a 3D movie, take home the 
glasses they provide, and poke out the lenses. They work very nicely as 
polarizer and analyzer with an ordinary microscope.
John Kiernan
Anatomy & Cell Biology
University of Western Ontario
London, Canada
= = =

From: Morken, Timothy via Histonet 
Sent: 28 July 2020 13:57
To: Ken M 
Cc: Histonet 
Subject: Re: [Histonet] Apple Green Birefringence in Amyliod slides

Ken, Yes, polarized light and apple green birefringence is diagnostic for 
amyloid with congo red and is the best practice. If you have a problem with 
known control slides  there are two possibilities: 1) make up fresh solution. 
The pH has to be right. Or 2) try other control slides. Maybe you cut through 
the amyloid area.

Because we have hundreds of microscopes in our department most just use 
polarized film as the polarizer (put over the light source) and another put 
over the top of the slide as the analyzer. Turn one of the polarizing slides 
and you will see the birefringence appear.

Source:
"Polarizing film, 2"" x 2"" , PK/10 (BEST For use as a microscope polarizer)"   
Cat# S07372 Thermo Fisher Sci Health$36.75  PK/10   "2" x 2"

These are polarized film mounted in 2" film holders (like the old Kodachrome 
slides).

Cheap and effective. (and avoids consternation from people losing expensive 
microscope polarizers)

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


-Original Message-
From: Ken M via Histonet 
Sent: Tuesday, July 28, 2020 11:43 AM
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] Apple Green Birefringence in Amyliod slides

Hi everyone.  I was wondering if anyone out there has any experience with 
diagnosing Amyloid tissue using Congo Red stained Kidney using polarized 
lenses.  Is it common to use polarized light to detect Amyloid deposits?  Does 
the absence of the "apple green birefringence" indicate a problem with the 
control tissue or the control slides?  Should this green bifringence always 
appear to confirm the diagnosis?  I know that the tissue should be cut thicker 
than normal (we usually cut at 5), but in the future maybe we will cut at 7 or 
8?
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Re: [Histonet] Fixed frozen non-paraffin mouse brain

[John Kiernan via Histonet]

And a very good pennyworth it is, Carl!

You wrote,  "... someone must've originally thought: 'Hang on, if we fix in 
commercially bought 40% Formalin, it's got 10% methanol added (to slow rate of 
formaldehyde repolymerisation) ...that will compete with formaldehyde fixation. 
So, we get coagulative and additive fixation. That is not good, folkslet's 
get pure and use depolymerised paraformaldehyde: pure methylene glycol 
polymer'".

That's almost how it came about: let's get pure.  A fixative made from PFA 
should have the same composition every time it is freshly prepared. The 10% of 
methanol (MeOH) in formalin (40% HCHO) isn't enough to coagulate proteins, and 
neither is the 1% MeOH in 10% formalin (with 4% HCHO). You need 60-70% alcohol 
to coagulate proteins, viruses etc. Formalin also contains some formic acid; 
the amount increases with age, from oxidation of the aldehyde by air. Dilution 
with water always gives an acidic solution. Marble chips can be added bring the 
pH up to neutrality. Buffering also takes care of the formic acid and can 
provide a neutral (pH7) or a "physiological" (pH7.4) fixative solution. The 
usual phosphate buffer also makes the fixative solution approximately 
iso-osmotic with mammalian extracellular fluid. Before the 1960s, dilution of 
formalin with with saline (0.9% NaCl) provided "formal saline", which had some 
advantages over 4% aqueous formaldehyde. See books by J. R. Baker, which are 
available as free downloads from http://archive.com.

Polymerization also increases with age. That's why you see a white precipitate 
in bottles of formalin stored for a long time. The precipitate is 
paraformaldehyde (PFA); its presence reduces the amount of formaldehyde that 
can be easily released by simple dilution of  the formalin with water. 
According to R. Cares (1945: A note on stored formaldehyde and its easy 
reconditioning. J. Tech. Methods & Bull. Int. Ass. Med. Museums 25, 67-70), 
milky formalin can be cleared by autoclaving, for 30 m in Kilner jars. I wonder 
if anyone else has done this?

John Kiernan
Anatomy & Cell Biology
UWO, London, Canada
= = =

From: Hobbs, Carl via Histonet 
Sent: 05 July 2020 14:25
To: histonet@lists.utsouthwestern.edu 
Subject: Re: [Histonet] Fixed frozen non-paraffin mouse brain

Prof. Kiernan, as usual, provides us all with such a depth/breadth of 
particular information/advice.
His Histological and Histochemical methods BIBLE is still my favourite read.

Respect
Most researchers fix in depolymerised Paraformaldehyde because someone must've 
originally thought:
" Hang on, if we fix in commercially bought 40% Formalin, it's got 10% methanol 
added ( to slow rate of formaldehyde repolymerisation) ...that will compete 
with Formaldehyde fixation.
So, we get coagulative and additive fixation. That is not good, folkslet's 
get pure and use depolymerised Paraformaldehyde: pure methylene glycol polymer"
I am sure Professor Kiernan can correct my inaccuracies!
Anyway..I've never noticed any difference: I've worked in diagnostic labs ( 
unfixed frozen muscle/renal/rectal bx) and also research labs ( unfixed/ fixed 
frozen tissues) using both fixing solutions
I have not noticed any IHC/IF difference in reactivity.
Many primary abs do NOT work even with fixed/unfixed  frozensome of them 
WILL need HIER ( at 90C rather than M/W or pressure cooker AR but, only fixed 
frozen of course), imho.
Part of the problem is whether  the antigen is linear or 3D...sorry for 
simplicity.
I can successfully snap-freeze fixed/unfixed rat/ms brain hemispheres without 
using sucrose ( success measured by lack of holes at the LM level).
This is because I was trained in a diagnostic lab to freeze fast but, 
effectively.
It is a technique that requires experience for consistency of 
successsometimes I fail!

The reason most use 20/30% sucrose is to give poor a snap-freezing technique a 
chance to avoid ice-crystal artefact, as stated by Kiernan).
Sucrose is no panacea.technique is everything.
My pennyworth-illy
Carl



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


020 7848 6813
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Re: [Histonet] Fixed frozen non-paraffin mouse brain

[John Kiernan via Histonet]

Dear Ed,

Adequate fixation is important. Formaldehyde penetrates quickly but reacts 
slowly with proteins. 4% formaldehyde made by depolymerizing paraformaldehyde 
(an insoluble high polymer) is the same as formaldehyde made by 10X dilution of 
formalin (a mixture of soluble low polymers).

For cryoprotection the sucrose must thoroughly penetrate the fixed specimen, 
which must sink in the concentrated solution.

Your Swiss cheese artifact is due to slow freezing. In badly frozen brains the 
ice crystal holes are as big as large neurons and the tissue architecture is 
destroyed.

Unless specimens are really tiny and frozen super-fast (special methods for 
electron microscopy), ice crystals always form and show later as holes in the 
sections. You get bigger holes with bigger specimens and slower freezing.
With very fast freezing, a skilled technician can get good cryostat sections 
even of small unfixed specimens such as muscle biopsies. These are needed for 
enzyme activity histochemistry methods used in diagnostic pathology and in 
research.

Cryoprotection of fixed specimens slows the growth of ice crystals. With luck, 
the holes are too small to interfere with light microscope studies of sections. 
In neuroscience research, quite thick frozen sections of samll animals' brains 
have been the norm for more than 50 years.

About 20 years ago I wrote a chapter that gave some quite detailed instructions 
and explanations, with references. (Don't do anything important just because a 
chapter or a review says so; check at least some of the refs!)   Here is the 
reference.

Kiernan, J. A. 2002. Freezing and fixation. Chapter 8 in Microscopy and 
Histology for Molecular Biologists. A User's Guide, ed. Kiernan, J. A. & Mason, 
I. G. pp. 103-143.  London: Portland Press.  ISBN 1855781417.

Your university's library in Urbana might have the book. It's out-of-print with 
its publisher. There are used copies on the web for much less than the original 
price.

John Kiernan
Emeritus neuroanatomist and histochemist
London, Canada
https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html
= = =

From: Roy, Edward J via Histonet 
Sent: 04 July 2020 20:08
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Fixed frozen non-paraffin mouse brain

As a research lab, we sometimes would like to use paraformaldehyde-fixed but 
non-paraffin embedded tissues; paraffin embedding alters antigens and 
necessitates antigen retrieval, but simple fixation does not. We have done the 
traditional 30% sucrose before OCT and freezing, with cryostat sectioning, but 
results are inconsistent, sometimes producing Swiss-cheese brains. Does anybody 
have an alternative to 30% sucrose that is more reliable?  I didn’t see 
anything in the Archives after a search for “30% sucrose”.
Thanks very much,
Ed Roy

Edward J. Roy, PhD
Professor Emeritus
Department of Molecular and Integrative Physiology
University of Illinois at Urbana-Champaign
Urbana, IL 61801
217 333-3375


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Re: [Histonet] Brain and spinal cord

[John Kiernan via Histonet]

10% neutral buffered formaldehyde is good, but a whole human brain needs to be 
immersed for at least 2 weeks, suspended by a string under the basilar artery 
to prevent squashing against the bottom of the container. The soft tissue has 
to be properly hardened before the brain can be sliced and small pieces 
selected for embedding and sectioning.

Fixatives other than formaldehyde can be OK for brains of small animals used in 
research, but for human neuropathology you need formaldehyde to get the 
expected results with regularly used stains and with immunohistochemistry.

See Adams & Murray 1982 Atlas of post-mortem techniques in neuropathology. ISBN 
9780521105682.
 Secondhand copies cost about US $30. Every lab should have plenty of books. 
They cost a lot less than making mistakes.

John Kiernan
Anatomy & Cell Biology
University of Western Ontario
London, Canada
= = =

From: Yahoo via Histonet 
Sent: 15 May 2020 15:42
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Brain and spinal cord

Hi All!
I’m looking for some suggestions please on fixation for brain tissue and spinal 
cord submissions from necropsies. We are currently using 10% NBF and ask our 
pathologists to leave the samples overnight (but that doesn’t always happen!!). 
Does anyone use alcohol-based fixatives? And if so, how long? Does it affect 
IHC or any other staining? Do you still process with other routine biopsies (14 
hour program)?
Thanks!

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Re: [Histonet] On-line references

[John Kiernan via Histonet]

Hello, Tom.

Some old classics are there for free, most notably JR Baker's "Principles of 
Biological Microtechnique" (1958), but almost anything more recent has to be 
bought.

There are plenty of cheap older editions of histotechnology books on sites like 
AbeBooks. Check it out for the last edition of  Pearse's Histochemistry!  I was 
amazed.

Even the latest editions of books in our field cost only about $100 from the 
publisher and most are good for several years.  Compare this with the price of 
a few drops of an antibody or (more realistically) a staining machine in which 
you must only use the liquids sold by its vendor.

John Kiernan
= = =

From: Tom Wells via Histonet 
Sent: 25 March 2020 14:34
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] On-line references

Given that our Institute's library is closed due to the pandemic, is anyone 
aware of on-line versions of Histotechnology/ Histochemistry textbooks? Thanks. 
Tom

Tom Wells BSc, MEd, MLT, ART
Faculty
Medical Laboratory Science
School of Health Sciences
SW03-3088
(604) 412-7594
BCIT

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Re: [Histonet] Glycogen detection; also Spring Forward

[John Kiernan via Histonet]

Glycogen (MW about 1,000,000) is soluble in water but insoluble in alcohol 
(Merck Index 12th ed.,1996, p.766). For this reason, non-aqueous coagulant 
fixatives may have advantages, especially for small specimens or thin layers of 
cultured cells.

Fixation immobilizes cytoplasmic proteins, which then entangle the big long 
polysacccharide molecules of glycogen, keeping them  approximately in their 
intracellular positions. Formaldehyde penetrates specimens rapidly, but its 
chemical reactions with proteins, especially the cross-linking that stabilizes 
structure, are slow (meaning 12-48 hours). During this time, the glycogen in 
liver cells dissolves and is carried in solution along the direction of  the 
fixative diffusing into the specimen. In each hepatocyte, this intracellular 
diffusion of glygogen is stopped by each hepatocyte's cell membrane, which has 
a lipid layers that are unchanged by an aqueous formalin solution. As a result, 
the stainable glygogen piles up in the side of each hepatocyte furthest from 
the surface of the specimen. This artifact is often called "polarix=zarion". 
With processing into paraffin, which removes lipids and coagulates any proteins 
not yet made insoluble by formaldehyde, glycogen is anchored into place by 
fixed cytoplasmic proteins, but it can still be attacked and removed by 
amylase/diastase/spittle.

All this has been known for at least 60 years. It's in the textbooks, as Bob 
Richmond pointed out yesterday. (Or was it the day before?)  It's now time for 
us all to advance our clocks by an hour, go to bed and wake up in time for 
Church on Sunday!

John Kiernan
= = =

From: Bob Richmond via Histonet 
Sent: 07 March 2020 13:47
To: Histonet@lists.utsouthwestern.edu 
Subject: Re: [Histonet] Glycogen detection

Galina Deyneko (where? asks: >>Does anybody have experience how fix the
tissues for successful glycogen ? detection in murine and humane
cardiomyocytes. I am wondering maybe the trace of methanol in 10% formalin
will dissolve glycogen?? - What would be better process for paraffin
embedding or use OCT embedding without fixation? Of course I prefer FFPE
blocks, since OCT blocks give bad morphology.<<

Ordinary neutral buffered formalin and paraffin embedding should be
adequate. R.D. Lillie (3rd ed.) notes good results with Carnoy's fixative,
alcoholic formalin, and acetic alcoholic formalin also.

The traditional stain for glycogen is periodic acid Schiff (PAS). You
verify the presence of glycogen by doing the stain with and without amylase
("diastase") predigestion. (A crude but adequate source of amylase is to
just spit on the slide.)

Bob Richmond
Samurai Pathologist
Maryville, Tennessee
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Re: [Histonet] How to Reduce Tissue Autofluorescence

[John Kiernan via Histonet]

There's a very brief article (downloadable PDF) from 2002 about suppressing 
autofluorescence, with a few references, at 
https://www.researchgate.net/publication/10971457_Suppressing_autofluorescence
[https://i1.rgstatic.net/publication/10971457_Suppressing_autofluorescence/links/00463520275ec6a5f100/largepreview.png]
(PDF) Suppressing autofluorescence - 
ResearchGate
A 'read' is counted each time someone views a publication summary (such as the 
title, abstract, and list of authors), clicks on a figure, or views or 
downloads the full-text.
www.researchgate.net
This PDF file also has short peer-reviewed histotechnical tips on 5 other 
topics. Fun for all there.

For something more recent on autofluorescence, try:
Davis AS, Richter A, Becker S, Moyer JE, Sandouk A, Skinner J, Taubenberger JK 
(2014) Characterizing and diminishing autofluorescence in formalin-fixed 
paraffin-embedded human respiratory tissue. J. Histochem. Cytochem. 62: 405-423.
They compared 9 procedures and favoured 3: sodium borohydride, Sudan black B 
and another dye called eriochrome black T. The last-named dye is CI 14645, 
Mordant black 11, a monoazo  dye very briefly described on page 108 in Conn's 
9th edn (1977) with the preferred name chromogen black ETOO; it's not in Conn's 
10th edn (2002). Sodium borohydride reacts with aldehydes and probably reduces 
fixative-induced fluorescence of proteins and the native fluorescence of 
lipofuscins. The black dyes may work by absorbing more weakly emitted light. 
Sudan black B can stain lipofuscin black even in in paraffin sections.

Using “home brew” reagents is always the best way to go, because you can avoid 
buying simple products sold at high prices with fancy names. Avoid trying 
anyone's unexplained "working protocol" because annotated pieces of paper get 
passed along in labs and can induce well educated people to do things that are 
obviously wrong.

It is necessary to know the reason for each step in a lab procedure. You 
identify as a research assistant, so you must have a boss. Probably your boss 
should be online along with you, asking histonetters for advice about reducing 
autofluorescence.

That's quite enough from me, on 9 Feb 2020.
John Kiernan  (Anatomy, UWO, London, Canada)
= = =

From: Arun Jyothi S.P via Histonet 
Sent: 06 February 2020 10:18
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] How to Reduce Tissue Autofluorescence

Dear All,

Kindly share your working protocol using  “home brew” reagents to reduce
tissue auto-fluorescence.

Thank you
Arun Jyothi S.P.
Research Assistant
Cancer Research
RGCB
Trivandrum
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Re: [Histonet] Question about gelatin embedding

[John Kiernan via Histonet]

Gelatin embedding is easy. You infiltrate the specimen and then fix it again in 
formaldehyde to cross-link the gelatin molecules and make the whole mass 
isoluble in water. You can than cut frozen sections of any kind: cryostat, or 
with an old-fashioned freezing microtome collecting thawed sections from the 
knife with a brush.  The formaldehyde-fixed gelatin holds everything together.  
With a Nissl stain it remains inconspicuous. If you do an H the gelatin will 
stain red.
John Kiernan
= = =

From: Alonso Martínez Canabal via Histonet 
Sent: 17 January 2020 16:29
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Question about gelatin embedding

Hello,
  I am here again. I am wondering if someone has good experience
embedding in gelatin-albumin for cryostat or vibratome sectioning.
Specifically we use brain tissue and is common in free floating techniques
non-attached parts of the same section float around and later that
generates all sorts of problems.
   Thank you very much.

--
Dr. Alonso Martínez Canabal PhD
Profesor Asociado "C"
Departamento de Biología Celular, Facultad de Ciencias, UNAM
Investigador Nacional "I"
56224833
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Re: [Histonet] Pap stains

[John Kiernan via Histonet]

As a student in the 1960s I was told by my elders and betters to filter all 
staining solutions before using. It was very good advice. Filtration does not 
prolong the life of a stain, but it does remove crud, a material that can be 
composed of polymerized dyes and bits of previously stained cells or tissues.

Fifty years on, I can identify a few staining solutions that never deteriorate 
into insoluble materials that fall out of solution even after 10+ years. Even 
for these, filtration before use is good laboratory practice. Not filtering any 
stain may therefore be bad laboratory practice.

Happy New Year,  John Kiernan
= = =

From: Haas, Elizabeth via Histonet 
Sent: 31 December 2019 11:46
To: S hay 
Cc: histonet@lists.utsouthwestern.edu 
Subject: Re: [Histonet] Pap stains


I believe filtering stains daily is a CAP requirement


Sent from my iPhone

> On Dec 31, 2019, at 9:23 AM, S hay via Histonet 
>  wrote:
>
> 1. Does everyone filter their pap stains daily?
> 2. Are you chaining all other reagents daily?
>
> Thanks in advance.
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Re: [Histonet] Diastase

[John Kiernan via Histonet]

Why buy?  Just think of squeezing a chunk of lemon over your helping of haddock 
and drool into a small beaker. Remove bubbles by wiping the surface of the 
collected liquid with the edge of a piece of filter paper. Incubate for 30 
minutes at 37C, just the same as for 0.1% malt diastase.

Saliva also contains RNase, but this doesn't matter because RNA is not stained 
by PAS or by other methods for staining glycogen. For a cheap source of RNase 
you can heat the collected saliva (80C for 10 minutes) to inactivate diastase 
and any other enzymes. Cool and centrifuge. Use the supernatant at 37C for 1 
hour.

Some references.

Bradbury S (1956) Human saliva as a convenient source of ribonuclease. Quart. 
J. Microsc. Sci. 97: 323-327.  (Free PDF available at 
https://jcs.biologists.org/content/s3-97/39/323.short).

Brown, GG (1978) An Introduction to Histotechnology. New York: 
Appleton-Century-Crofts. pp. 302 (diastase) & 292 (ribonuclease).

Drury RAB, Wallington EA (1967) Carleton's Histological Technique. 4th ed. 
Oxford University Press, Oxford. pp.163-164 (ribonuclease) & 208 (diastase).

John Kiernan
Dept of Anatomy & Cell Biology
University of Western Ontario, London, Canada
= = =


From: Paula via Histonet 
Sent: 26 November 2019 12:16
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Diastase

Hello,



We've been using STATLabs Diastase for our PAS with diastase digestion but
they have a backorder until January. Can anyone recommend an alternative
from other vendors?



Thank you in advance,

Paula

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Re: [Histonet] Metals

[John Kiernan via Histonet]

Timm's sulphide-silver method is very sensitive, and modifications (mostly by 
Danscher) even more so. Sulphide-silver methods detect only those metals that 
have insoluble sulphides (copper and zinc but not aluminium, in Jeanine's 
list). It is necessary to fix in a special solution containing hydrogen 
sulphide - stink and also serious safety precautions!

Tony's mention of Mallory & Parker's fresh hematoxylin stain prompted me to 
look it up. The 1939 paper is a free PDF download (Google Scholar: Mallory 
Parker Hematoxylin Stain Metals). Mallory FB, Parker F (1939) Fixing and 
staining methods for lead and copper in tissues. Am. J. Pathol. 15: 517-522 and 
Plates 83-85.
The authors noted the importance of fixation (neutral formalin was OK for 
copper but no good for lead, which needed 95% or 100% alcohol). Like F.B. 
Mallory's other papers about staining methods, it's rather vague on technical 
details and has no references.

The late Ralph D. Lillie reported more thorough investigations of staining for 
metals with haematoxylin in his classic book Histopathologic Technic and 
Practical Histochemistry (4th and last edn 1976, ISBN 0070378622), giving 
colours of the complexes with 30 metals introduced into tissues. I have tried 
Lillie's method on paraffin sections of rat tissues containing a few of these, 
and it works.  ISBN 9781907904325 (p.333-334)  may be more accessible than 
Lillie's book, which has become an expensive classic.

For the more specific stains mentioned in Tony's message you need to do some 
critical reading. The best place to start may be Frieda Carson's 
Histotechnology textbook. ISBN  978-0891896401.

Enough about metals for now!

John Kiernan
London, Canada
= = =

From: Tony Henwood (SCHN) via Histonet 
Sent: 30 September 2019 07:22
To: Sanders, Jeanine (CDC/DDID/NCEZID/DHCPP) 
Cc: Histonet 
Subject: Re: [Histonet] Metals

Two good screening stains are Mallory and Parker’s Fresh Hematoxylin Stain for 
Metals and  Timm’s Silver Sulphide Method for Metals. Malloy's results:
Aluminium   Blue-black
Copper  Greenish-blue
IronBlue-black
LeadBlue
ZincBlue

For more specific staining:
Aluminon Stain for Aluminium Hydroxide
Walton’s Stain for Aluminium  (Phloxine binds the aluminium)
Bedrick et al (1986) method for Zinc
Rubeanic Acid Technique for Copper
Rhodanine Technique for Copper

These methods are quite sensitive but  there are some specificity issues. I can 
provide further details and references if required. Here are some:

Ohtsuki, Y., Yamaguchi, T., Sonobe, H., Takahashi, K., Hayashi, K., Takenaka, 
A., ... & Terao, N. (1989). Stain Technology: A Simplified Aluminum Stain in 
Paraffin Sections of Bone from Hemodialysis Patients. Stain technology, 64(2), 
55-59.

Walton, J. R., Diamond, T. H., Kumar, S., & Murrell, G. A. C. (2007). A 
sensitive stain for aluminum in undecalcified cancellous bone. Journal of 
inorganic biochemistry, 101(9), 1285-1290.

Bedrick, A. E., Ramaswamy, G., & Tchertkoff, V. (1986). Histochemical 
determination of copper, zinc, and iron in some benign and malignant tissues. 
American journal of clinical pathology, 86(5), 637-640.

Regards
Tony Henwood JP, MSc, BAppSc, GradDipSysAnalys, CT(ASC), FFSc(RCPA)
Principal Scientist, the Children’s Hospital at Westmead
Adjunct Fellow, School of Medicine, University of Western Sydney
Tel: 612 9845 3306
Fax: 612 9845 3318
Pathology Department
the children's hospital at westmead
Cnr Hawkesbury Road and Hainsworth Street, Westmead
Locked Bag 4001, Westmead NSW 2145, AUSTRALIA


From: Sanders, Jeanine (CDC/DDID/NCEZID/DHCPP) via Histonet 

Sent: Monday, 30 September 2019 20:49
To: 'histonet@lists.utsouthwestern.edu'
Subject: [Histonet] Metals

Morning all!

I need some advice re: protocols to demonstrate metals in FFPE tissues. Metals 
such as copper, aluminum and zinc.

Thanks much!

Jeanine Sanders, BS, HT(ASCP), QIHC(ASCP)
Centers for Diseases Control and Prevention
1600 Clifton Road NE
MS H18-SB
Bldg. 18, Rm SB-114
Atlanta, GA 30329
404-639-3590

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Re: [Histonet] Permanent mountant for Oil Red O

[John Kiernan via Histonet]

Fructose syrup gives a hard set, but it's a bit acidic (I don't know why) and 
is therefore incompatible with simple basic dyes, but it's OK for Oil red O and 
the Sudans.
   Fructose (also called laevulose or levulose)  15 g
   Distilled water  5 ml
Leave at ~60C (in paraffin oven) for 2 or 3 days for all the fructose to 
dissolve. Don't let the water evaporate!  The transparent syrup keeps well for 
2-3 months at room temp.
With long storage fructose crystallizes under the coverslip; this can be 
retarded by sealing the adges with a resinous mounting medium such as DPX. (You 
can buy fructose powder for cooking - about $3 a pound on the the internet. Cf  
$25-80 from chemical supply houses.)

Another good one is polyvinylpyrrolidone:
 PVP (m.w. 10,000)   25g
 Water (or a phosphate buffer pH 7.4) 25ml
  When dissolved (several hours, magnetic stirring), add 1ml glycerol and a 
small crystal of thymol.
Keeps for up to 3 years. Discard if it becomes cloudy. Use the buffered variety 
if the preparation has been stained or counterstained with a basic dye like 
toluidine blue or neutral red.
This is less viscous than fructose syrup, and also has a lower refractive 
index, but with evaporation at the edges of the coverslip it gradually (weeks) 
becomes harder, and its refractive index increases almost to that of a resinous 
mountant.  Probably you won't want to wait before shipping the slides 
elsewhere.  PVP costs more than fructose (about $150 per pound for PVP10).

This short article about aqueous mounting media is rather old (1997) but 
probably still OK:
http://publish.uwo.ca/~jkiernan/aqmount.htm

John Kiernan
London, Canada
= = =


From: Hagon, Christopher (Health) via Histonet 

Sent: 15 September 2019 22:43
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Permanent mountant for Oil Red O

UNCLASSIFIED

Hello Histonetters,

We do Oil Red O stains on frozen section post mortem tissue, and need to find a 
permanent aqueous mounting medium. We used to use the Thermo-Fisher  
Perm-mount, but can't seem to get it anymore. As they are looking for fat 
deposits, we can't use any solvent based solutions, and the only aqueous ones 
we've found aren't permanent. We have to send the slides off site after 
staining, so the chances of the coverslip moving in transport is fairly high.

Does anyone else have this issue and what did you end up using?

Thanks in advance,

Chris Hagon | Senior Scientist, Anatomical Pathology
ACT Pathology | health.act.gov.au
Phone (02) 5124 2874





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Re: [Histonet] Mallory-Azan stain

[John Kiernan via Histonet]

Mallory's (easy) and AZAN staining (difficult) are different methods!

Frank B. Mallory's trichrome stain  (Journal of Medical Research 13: 113-136, 
1905) is the earliest and one of the simplest of its kind: acid fuchsine 
followed by a solution containing orange G, aniline blue and phosphotungstic 
acid (PTA).  Martin Heidenhain's trichrome is usually called AZAN (from 
Azokarmin and Anilinblau, the German names of two of the dyes used. I've not 
read his original 1916 publiication, but a very thorough account was given by 
Manfred Gabe in his 1976 Histological Techniques book (ISBN 3540901620), pp. 
219-223. I used this quite a bit in the 1990s mostly on paraffin sections of 
Bouin-fixed decalcified rats' heads. It is a 15-step procedure taking >2 hours 
and it includes two critical differentiations requiring careful microscopic 
control. Instructions based on my experiences can be found in Histological and 
Histochemical Methods (5th ed., 2015, pp.198-200).

AZAN gives a wider range of colours than Mallory's or Masson's trichrome or the 
various one-step trichromes (Cason, Gomori, Gabe). The related Romeis 
"cresazan" procedure was used to identify at least 6 anterior pituitary 
cell-types until the 1950s when more rational histochemically based stains were 
introduced by Adams, Herlant, Pearse and others. Nowadays, immunostainng 
accurately shows the hormones in pituitary cells, but much more expensively.

All trichromes give poor results after simple fixation in neutral formaldehyde. 
Bouin or (better) a mercuric chloride-containing fixative is needed. 
Zinc-formalin is probably also OK. (I haven't tried it myself for this 
purpose). If material fixed in NBF must be used, immerse hydrated paraffin 
sections in saturated aqueous picric acid either for 2h at 56-60C or overnight 
at room temperature, then wash well in water before staining. (Bouin's fluid is 
often used, but its ingredients other than picric acid are unnecessary.) 
Experiments are needed to learn the mechanism of this "rescue" of  staining 
properties of sections formaldehyde-fixed tissue, which is sometimes wrongly 
called "mordanting". My guess is that it's comparable to antigen retrieval. It 
has been claimed that citrate buffer is just as good, though the photos are 
unconvincing (J. Histotechnol. 26, 133).

It should be possible to identify Purkinje fibres with any staining method that 
shows nuclei and myofibrils, such as H or a trichrome method simpler than 
AZAN. A glycogen stain such as PAS might show this substance in the otherwise 
pale areas around the central nuclei of Purkinje fibres. I suggest persuading 
your researcher to let you try something simpler before attempting Heidenhain's 
AZAN. Wheater's Functional Histology has a nice photomicrograph of a section 
stained with H and for endocardial elastin (looks like orcein).

Enough rambling!
John Kiernan
Anatomy & Cell Biology
University of Western Ontario
London, Canada
= = =


From: Betsy Molinari via Histonet 
Sent: 09 September 2019 10:53
To: 'Histonet@lists.utsouthwestern.edu' 
Subject: [Histonet] Mallory-Azan stain

Hi histonetters,
I have a researcher that wants to stain Purkinje fibers and has requested a 
Mallory-Azan stain.
I have no experience with this stain. I have looked online for information but 
am reaching out to you for personal advice.
Thanks.
Betsy Molinari HT,ASCP
Texas Heart Institute
6770 Bertner Ave.
Houston, TX  77030
832-355-6524 (lab)
832-355-6812 (fax)

Betsy Molinari
Sr. Histology Research Technician
CV Pathology Research

Texas Heart Institute
6770 Bertner Avenue, MC 1-283
Houston, TX 77030

Office: 832-355-6524 | Fax: 832-355-6812
Email: bmolin...@texasheart.org
texasheart.org | 
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twitter
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Re: [Histonet] Recent issues with picro sirius red staining (entire liver section become red, no yellow background)

[John Kiernan via Histonet]

I've never seen the kind of staining you describe, abi jag, with "the complete 
section become stained as red" but I've never used the method on sections of 
liver.

You should get red collagen and yellow hepatocytes with blue nuclei. The 
strongly acidic picrosirius stain, applied for an hour, always greatly weakens 
(differentiates) a prior nuclear stain with Weigert's or Lillie's 
iron-haematoxylin.  It's also quite easy to lose some or all the yellow in the 
washing and dehydration of the stained sections. Most users of this method are 
interested only in collagen fibres and do not mind if the nuclear and 
cytoplasmic colours get lost. The iron-haematoxylin nuclear stain is often 
omitted.


It is necessary to have the right dye. It must be sirius red F3B (= CI 35780 = 
Direct red 80). This is still used as a textile dye, with several suppliers and 
trade-names. See 
http://www.worlddyevariety.com/direct-dyes/direct-red-80.html#respond

Direct Red 80 - 
worlddyevariety.com
www.worlddyevariety.com
List of Suppliers: Direct Red F3,Direct Fast Red BA,Direct Fast Red F3B. 
Tianjin Yadong Group . ACDI Red 8 BLN(Aakash Chemicals & Dyestuffs 
Inc)Alacodirect Red 2BL(Classic Dyestuffs Inc)AmbidirectRed 3BL( Thai Ambica 
Chemicals Co Ltd) Anadurm Red D-BA( Albion Colours Ltd) Arid Red 8 BLN ( 
Aashiana Dyestuffs Inc) Best Direct Supra Red F3B( Oriental Giant Dyes and 
Chemical Ind Corp)
Other dyes with "sirius red" in their trivial names probably are not suitable 
if they are not the product recognized in the Colour Index as  CI 35780,  
Direct red 80. An example of a different dye is sirius red 4B (= CI 28160 = 
Direct red 81), which has been prescribed for use in some staining techniques 
as a dye with properties similar to those of eosin Y and acid fuchsine.


The Biological Stain Commission has standards for sirius red F3B as a collagen 
stain. Dye from a batch that meets their standards will be OK for your method.


My Histonet post from which you are using this method must date from the 1990s, 
when I knew that picro-sirius red solutions were good for 5-6 years. (I would 
have written 3 years to be cautious.)  With more experience with stored and 
newly made solutions, I feel confident in saying they keep for more than 20 
years.


It might get contaminated from too much iron-haematoxylin extracted from 
previously stained  slides. I don't know what this would do.


The most obvious cause of red cytoplasmic staining by picrosirus is not enough 
picric acid (yellow powder in the bottom of the bottle) in the staining 
solution.


It's unfortunate that items found with HistoSearch are undated. It doesn't 
matter in this case, but many Histonet items become outdated after only a year 
or two; antibodies and automated staining are examples of fields in which you 
need to know the age.

Keep in touch about your sirius red problem.

John Kiernan
John A. Kiernan MB, ChB, PhD, DSc
Professor Emeritus, Anatomy & Cell Biology
University of Western Ontario, London, Canada
https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html
Also  Secretary,  Biological Stain Commission, Inc.
https://biologicalstaincommission.org
= = =

From: abi jag via Histonet 
Sent: 09 August 2019 11:29
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Recent issues with picro sirius red staining (entire liver 
section become red, no yellow background)

Hello Histonetters,I am writing this to seek your help regarding a very recent 
problem that I am currently facing with Picro Sirius red staining of lab animal 
(mouse and rat) liver samples. I follow the procedure that was provided by John 
Kiernan in the histonet archives (please see below), which was working very 
well. Quite recently, the complete section become stained as red. Usually, 
collagen in the sections get stained as red with a yellow back ground. Please 
note that there was no change in the procedure/reagents etc, It will be of 
great help if you help me in troubleshooting this issue.With my best 
regards,Abijag
Sirius red collagen procedure

|
|
|  |
Sirius red collagen procedure


 |

 |

 |




Solution A. Picro-sirius red

  Sirius red F3B (C.I. 35782): 0.5 g
  Saturated aqueous solution
of picric acid:500 ml
  Add a little solid picric acid to ensure saturation
(This is important).

  (Keeps for at least 3 years and can be used many times.)

Solution B. Acidified water

  Add 5 ml acetic acid (glacial) to 1 litre of
  water (tap or distilled).

Procedure

Fixation is not critical, The method is most frequently used on
paraffin sections of objects fixed adequately (at least 24 hours
but ideally 1 or 2 weeks) in a neutral buffered formaldehyde
solution.

1. De-wax and hydrate paraffin sections.
2. (Optional, and not usually done) Stain nuclei with
   Weigert's haematoxylin (as for the van Gieson method,
   but more 

Re: [Histonet] Troubleshooting Gomori's Trichrome Stain (Blue Collagen, Richard-Allen) staining

[John Kiernan via Histonet]

One-step trichrome methods (such as Gomori's) are OK when they work, but 
there's little you can do when the colours come out wrong. A simple thing to 
try would be a shorter time in the staining mixture, to reduce diffusion of the 
more slowly penetrating dye (aniline blue) into cells. Trichrome methods work 
better after coagulant fixation than after formaldehyde. (The Bouin 
pre-treament is to offset the undesirable effects of fixation in neutral 
formaldehyde; picric acid alone works just as well. See also Yu & Chapman 2003 
J. histotechnol. 26(2): 131-134.)  If you can fix your hearts in Bouin or 
Carnoy you will get a better result with any trichrome technique.


If the one-step method still won't work, use a multi-step trichrome where you 
can have some control over the actions of the different components. Masson's 
(which has several variants) is popular; Mallory's has fewer steps. Good luck!


John Kiernan

Anatomy, University of Western Ontario

London, Canada

= = =


From: abi jag via Histonet 
Sent: 25 July 2019 08:32
To: histonet@lists.utsouthwestern.edu 
Subject: [Histonet] Troubleshooting Gomori's Trichrome Stain (Blue Collagen, 
Richard-Allen) staining

Dear histo experts,Please provide me with your valuable suggestions for the 
problem described below.Objective: Staining rat hearts (fibrosis) with Gomori's 
one step Trichrome Stain (Blue Collagen, Richard-Allen) to demonstrate the 
collagenProcedure: Paraffin sections of 4 micron thickness; adequately fixed in 
10 % NBF,  Bouin’s Fluid treatment at 56°C for 1 hour before staining. Follow 
the procedure exactly recommended by kit insert (please see below)Problem: Non 
specific diffuse bluish discoloration of cardio myocytes in the normal hearts, 
which looked completely odd. The staining of same section with picro sirius red 
came beautiful.Any insights on the potential reasons of this and ways to 
resolve?Thanks a lot in advance for your time and your vision to make a wealth 
of knowledge available in histonet.
Best regards,Abi

Standard Staining Protocol

1. Deparaffinize and hydrate sections to deionized water.

2. Place sections in Bouin’s Fluid at 56°C for 1 hour.

3. Rinse sections in running tap water for 3-5 minutes untilyellow color is 
removed.

4. Place sections in Working Weigert’s Iron Hematoxylin Stainfor 10 minutes.

5. Rinse sections in running tap water for 5-10 minutes.

6. Stain sections in Trichrome Stain for 15 minutes.

7. Place sections in 1% Acetic Acid Solution for 1 minute.

8. Rinse sections in deionized water for 30 seconds.

9. Dehydrate sections in 95% alcohol for 1 minute.

10. Dehydrate sections in two changes of anhydrous alcohol for 1minute each.

11. Clear sections in three changes of clearing reagent for 1minute each and 
mount.

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Histonet -- For the exchange of information pertaining to histotechnology and 
related fields About Histonet

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Re: [Histonet] time for penetration of methacarn fixative

[John Kiernan via Histonet]

Peter Noyce, you did not say what tissue(s) you are planning to fix, or how big 
the specimens will be.


Carnoy (1886) and methacarn (1970) were developed for animal tissues, cleverly 
balancing the actions of acetic acid and an alcohol in the presence of  a 
hydrophobic solvent (chloroform) that diluted both these ingredients and also 
enhanced permeation by dissolving lipids. Carnoy's and other acid-alcohol 
fixatives are also used for plant specimens, which respond differently. If you 
are working with plant specimens you probably have a book by Steven E. Ruzin 
(1999)  ISBN 0195089561.


For the actions of alcohols and acetic acid in fixation of animal tissues, 
consult any text book of histotechnology or histochemistry published since 
about 1950.

For selfish reasons, I recommend ISBN 9781907904325 (published in 2015) as an 
item to buy for your lab's bookshelf. Answers to your question are discussed in 
Chapter 5.


With methacarn and other alcohol-acetic fixatives, no slow chemical reactions 
are involved (an important difference from formaldehyde-containing mixtures). 
Complete penetration accomplishes the fixation. The volume of fixative and 
procedure for subsequent processing into paraffin are very important. You need 
to read the paper by Puchtler et al (1970) and follow the instructions exactly. 
Probably you should also read Puchtler et al (1968) to use this type of 
non-aqueous fixative intelligently. PhD students are intelligent.


I have added a couple of more recent papers that relate to methacarn. Read 
Puchtler or a textbook first.


Not all modern investigators (especially molecular biologists) understand what 
the ingredients of fixative mixtures do to the different components of cells 
and extracellular materials. Current papers with micrographs full of ghastly 
artifacts abound, even in journals with very high citation indices. There are 
published mixtures with names like "modified methacarn" that may be OK for 
extracting RNA but do not have ingredients in correct proportions for 
minimizing distortion.


Here's your list of recommended readings.

- - - - -

Puchtler H, Waldrop FS, Meloan SN, Terry MS, Connor HM (1970) Methacarn 
(methanol-Carnoy) fixation. Practical and theoretical considerations. 
Histochemie 21: 97-116.

Puchtler H, Waldrop FS, Conner HM, Terry MS (1968) Carnoy fixation: practical 
and theoretical considerations. Histochemie 16: 361-371.

Uneyama C, Shibutani M, Masutomi N, Takagi H, Hirose M (2002) Methacarn 
fixation for genomic DNA analysis in microdissected paraffin-embedded tissue 
specimens. J. Histochem. Cytochem. 50: 1237-1245.

Buesa RJ (2008) Histology without formalin? Ann. Diagn. Path. 12: 387-396.
- - - - -

I wish you well with your research, and hope you will get your PhD while you 
are still young.

John Kiernan
London, Canada
https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html
http://biostain.com
= = =


From: peter noyce via Histonet 
Sent: 03 May 2019 19:47
To: histonet@lists.utsouthwestern.edu
Subject: [Histonet] time for penetration of methacarn fixative

Does any one have data for the time it takes for methacarn fixative (60%
methanol, 30% chloroform, 10%  glacial acetic acid) to penetrate and then
fully fix tissue?

PW Noyce -PhD student
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Re: [Histonet] PAP stains

[John Kiernan via Histonet]

Charles,  What do you mean by "dark nuclei"? Are you asking about the normal 
colour for the method, or about something you have not seen before that looks 
wrong?  Also, what is "PAP stain"?


If "PAP stain" means Papanicoloau, the nuclear stain is Mayer's haemalum. This 
is a progressive stain; you may need to shorten the time.


If "PAP stain" means peroxidase-antiperoxidase (unlabelled antibody-enzyme 
complex for amplifying detection of HRP-tagged secondary antibodies in 
immunohistochemistry), nuclei will be dark (usually brown) if they contain the 
antigen sought by the primary antibody. Nuclear staining might also be a 
false-positive artifact; if so, it would be present in the simplest of the 
routine controls (omission of primary antibody).  Dark nuclei might also be due 
to a nuclear counterstain that is too strong. The counterstain should be done 
in a contrasting colour. Usually it is a progressive haemalum (blue) to 
contrast with the oxidation product of DAB (brown).


Is this a Papanicoloau question or an immunohistochemistry question?  Does "PAP 
stain" have a third meaning?


John Kiernan

= = =


From: Charles Riley via Histonet 
Sent: 19 March 2019 14:17
To: Histo List
Subject: [Histonet] PAP stains

What causes dark nuclei in the PAP stain.

--

Charles Riley BS  HT, HTL(ASCP)CM

Histopathology Coordinator/ Mohs
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[Histonet] Words, phrases and names in histotechnology. A free glossary.

[John Kiernan via Histonet]

Hello histonetters. I'm a HistoNet old-timer, back again after a few years 
away. It's good to see that a few names are still around from the 1990s.


Here is something new that may interest all of us. I send it as a news item; a 
change from the usual initial question that initiates a Histonet topic.


A freely accessible online glossary of words, phrases and eponyms used in 
histotechnology, histochemistry and immunostaining was published by the 
Biological Stain Commission (BSC) at the end of December 2018.  It includes 
about 600 entries; it is to be revised and expanded from time to time. (A minor 
revision was uploaded on 19th January 2019.)


Notable features are extensive crosslinks between the entries, and explanations 
of terminology related to chemical and physical mechanisms involved in the 
methods. There are also definitions and explanations relating to all the stains 
(dyes) certified by the BSC.


The BSC glossary is readable on screens of all sizes (including mobile phones), 
and navigation among links is extremely rapid.


Check it out directly at 
https://biologicalstaincommission.org/bscglossary.html. Alternatively, see it 
in the broader context of the BSC:  http://biostain.com.


John Kiernan

= = =

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Re: [Histonet] axolotl lymphatics

[John Kiernan via Histonet]

Instead of an antibody, you might consider enzyme activity histochemistry, 
which is much less expensive. 
Demonstration of 5-nucleotidase activity in the presence of levamisole detects 
lymphatic endothelium. Sections can also be stained for alkaline phosphatase 
activity in the endothelium of blood vessels. Here are a few references.

Kato, S., Yasunaga, A. and Uchida, U. (1991). Enzyme-histochemical method for 
identification of lymphatic capillaries. Lymphology 24:125-129.
Ohkuma, M. (1994). Simultaneous double staining for the blood and lymphatic 
capillary. Lymphology 27, Suppl:106-107.
Okada, E. (1994). An improved enzyme-histochemical method for identification of 
lymphatic capillaries on paraffin sections. Lymphology 27, Suppl:732-735.
Ji, R.C. and Kato, S. (2003). Lymphatic network and lymphangiogenesis in the 
gastric wall. Journal of Histochemistry and Cytochemistry 51:331-338.

Needless to say, none of these relate to amphibian tissues!

John Kiernan
Anatomy, UWO, London, Canada
= = =
On 26/07/15, Jason Palmer via Histonet  histonet@lists.utsouthwestern.edu 
wrote:
 
 Hi all, 
 
 I need to find an antibody that will label lymphatic endothelial cells in the 
 axolotl. Does anybody have any experience or ideas? I have tried a couple of 
 our anti-mouse and anti-human Abs for podoplanin and LYVE-1 but no 
 cross-reactivity so far. I have no experience with staining of non-mammalian 
 tissues - maybe an anti-frog Ab would cross react? Does anyone have 
 experience with other amphibians? 
 
 Thanks, 
 
 Jason 
 
 -- 
 
 Jason Palmer 
 Histology Laboratory Coordinator 
 O'Brien Institute / St Vincent's Institute 
 42 Fitzroy St, Fitzroy Victoria 3065 
 Australia 
 tel +61 3 9288 4045 
 fax +61 3 9416 0926 
 email: jpal...@svi.edu.au 
 
 ___
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 Histonet@lists.utsouthwestern.edu
 http://lists.utsouthwestern.edu/mailman/listinfo/histonet
 
 
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Re: [Histonet] Toluidine blue stain for MMA

[John Kiernan via Histonet]

Yes. Probably hundreds of  Histonetters stain plastic sections. Let us all hope 
they don't all bombard the Histonet listserver with replies to your question. 
 
Instructions for staining plastic sections with toluidine blue are in every 
library that contains books with paper pages, and also (albeit with less 
authority) in great abundance on the Web.  
 
Try typing  SEMITHIN STAIN into Google. I just did, and an excellent web site 
came up on top of the heap. 
 
John Kiernan
= = =
 
On 27/07/15, Kai Hong via Histonet histonet@lists.utsouthwestern.edu wrote: 
  
 Hi,
 
 is there anyone have an experience with MMA toluidine staining?
 Im using T7200, T9100, Osteo-bed resin in lab now.
 
 Thanks,
 Kai
 Research Histotechnologist
 
 
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Re: [Histonet] National Academy of Sciences Confirms That Formaldehyde Can Cause Cancer in a Finding That Has Implications for Anatomic Pathology and Histology Laboratories

[John Kiernan via Histonet]

Dear Banjo,

A reference to the article would be helpful; there must be more to it than one 
sentence! 

Formaldehyde has been known for decades to be hazardous, and there are safety 
regulations in places where it is used. Plenty of old-timers are still alive 
and well after woking with formaldehyde in the days when there were few or no 
regulations. I'm one of them.
 
From about 1895 until about 1995 (and perhaps still, in some universities), 
every medical student spent most of the working day for at least a year with 
his or her nose and bare hands in a cadaver that had been embalmed in a 
cocktail containing phenol and formaldehyde. The predominant smell was the 
phenol, except when dissecting brains, which were fixed and stored in 4% 
formaldehyde. 

About 35 years ago, the American Association of Anatomists investigated effects 
of exposure to embalming chemicals on teachers of anatomy, who are in the 
dissecting room year after year. The only significant finding was eczema on the 
hands of some people, long known to be avoidable by wearing rubber gloves. Yes, 
I too should be able to provide a reference, but this was in the days of paper, 
which gets thrown out to make room for more paper ... There might be something 
deep in the archives at http://www.anatomy.org/

Other chemicals used in anatomy, pathology and histology labs also have their 
dangers; we avoid drinking them, rubbing them into our skin and inhaling their 
vapours, and we do our best to observe the safety regulations when it comes to 
getting rid of them. 

There is no substitute fixative functionally identical to formaldehyde. There 
are other fixatives, some less hazardous, but they have different effects on 
staining properties etc. The late Holde Puchtler published papers urging 
pathologists to use non-aqueous coagulant fixatives for routine fixation of 
small specimens, with her Carnoy variant methacarn (methanol 60, acetic acid 
10, chloroform 30) as the probable best, also good for some modern molecular 
methods. For this I can provide a few references:

Puchtler, H., Waldrop, F.S., Meloan, S.N., Terry, M.S. and Connor, H.M. (1970). 
Methacarn (methanol-Carnoy) fixation. Practical and theoretical considerations. 
Histochemie 21:97-116.

Cox, M.L., Schray, C.L., Luster, C.N., Stewart, Z.S., Korytko, P.J., Khan, 
K.N.M., Paulauskis, J.D. and Dunstan, R.W. (2006). Assessment of fixatives, 
fixation and tissue processing on morphology and RNA integrity. Experimental 
and Molecular Pathology 80:183-191.

Buesa, R.J. (2008). Histology without formalin? Annals of Diagnostic Pathology 
12:387-396.

Uneyama, C., Shibutani, M., Masutomi, N., Takagi, H. and Hirose, M. (2002). 
Methacarn fixation for genomic DNA analysis in microdissected paraffin-embedded 
tissue specimens. Journal of Histochemistry and Cytochemistry 50:1237-1245.

Milcheva, R., Janega, P., Celec, P., Russev, R. and Babal, P. (2013). Alcohol 
based fixatives provide excellent tissue morphology, protein immunoreactivity 
and RNA integrity in paraffin embedded tissue specimens. Brain Research 
Protocols 115:279-289.

Greer, C.E., Peterson, S.L., Kiviat, N.B. and Manos, M.M. (1991). PCR 
amplification from paraffin-embedded tissues. American Journal of Clinical 
Pathology 95:117-124.

Tissue processing is extremely simple after non-aqueous coagulant fixation, and 
most of the stages of a processing machine are not needed. Nuclear chromatin 
details are much sharper than after formaldehyde. This may not be seen as a 
blessing by young and middle-aged pathologists. In bygone days the routine 
fixatives contained mercuric chloride, which gives crisp chromatin and 
cytoplasmic details. The heterochromatin details probably are artifacts of 
fixation, but they are useful for identifying cells.

John Kiernan
Old neuroanatomist and histochemist
UWO, London, Canada
http://publish.uwo.ca/~jkiernan/
Also Secretary, Biological Stain Commission
http://biostain.com
= = =
On 13/07/15, Adesupo, Adesuyi (Banjo)  abades...@nrh-ok.com wrote:
  Hi,
  I read this article (National Academy of Sciences Confirms That Formaldehyde 
 Can Cause Cancer in a Finding That Has Implications for Anatomic Pathology 
 and Histology Laboratories) this morning.
  I wanted to know whether some of you guys out there are using Formaldehyde 
 substitute.
 
 
  Best regards,
 
  Banjo Adesuyi, BMLS, HT (ASCP) HTL, QIHC, QLS
  Histology Supervisor
  Norman Regional Health System,
  Norman, OK 73071.
  Tel: 405- 307- 1145
  abades...@nrh-ok.commailto:abades...@nrh-ok.com abades...@nrh-ok.com
 
 ==
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