Re: [Histonet] Fixative in diff-quick

[Bryan Llewellyn via Histonet]

Diff Quick appears to be a modified Field's stain. The blue dye in the 
methanol is one of the modifications. Field's stain gives much the same 
staining, so using plain methanol should be of no concern. The simplest 
way to find out is surely to try it and see.


http://stainsfile.info/stain/micro/field.htm

Bryan Llewellyn


Corbin, Clay via Histonet wrote:

Hey folks,
I am shopping for a diff-quick kit.  However, all I really need is the 
fixative.  Generally, there is a blue stain (triarylmethane) added to the 
methanol in the fixative solution.  I have a giant jug of lab grade methanol.  
What would I lose by using methanol alone compared to the fixative solution 
included in a diff-quick kit?
Thanks!
Clay

Clay Corbin, PhD
Professor of Biology
Bloomsburg University
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Re: [Histonet] Jones' Methenamine Silver Stain for Basement Membranes of Kidney - Issues and Questions

[Bryan Llewellyn via Histonet]

Hi,
Try the method given in StainsFile at:
http://stainsfile.info/stain/metallic/jones.htm

Bryan Llewellyn


Hood, Jordan via Histonet wrote:

Hello,

I'm new to histology (and new to histonet), and I work in a small histology lab 
specializing in animal tissues that receives requests/submissions from 
researchers. I tried (and failed) to perform a Jones' Methenamine Silver stain 
on a client's submission of pig kidneys (formalin-fixed, paraffin-embedded, cut 
at 2.5 microns), and I need some help troubleshooting this stain since my 
co-workers are stumped, too.  I used the following procedure from Rowley 
Biochemical:


~
"Fixation: 10% Buffered Neutral Formalin (F-113) or Bouin's Solution (F-40) or 
Zenker's (F-155)

Sections: Paraffin, 2 microns

Procedure: Acid washed glassware must be used
1. Deparaffinize and hydrate to distilled water.
2. Oxidize in Periodic Acid 0.5% (F-396-1) 11 minutes. Wash in chloride-free 
water.
3. Prepare Methenamine Silver solution by mixing: 42.5 ml Methenamine 3% 
(F-396-2), 2.5 ml Silver Nitrate, 5% (F-396-3) and 12.0 ml Borate Buffer, pH 
8.2 (F-396-4).
4. Place slides in the solution and the entire jar in a water bath at 70°C for 
approx. 60-75 minutes. Check under microscope when slides appear medium brown 
microscopically. Every 10 minutes, once the medium brown color has been 
established, rinse a slide in 70°C, chloride free water and check under a 
microscope. Rinse again in hot water and return to the hot staining solution. 
As the staining time approaches the end point, check the slides, as above, 
every 1-2 minutes. The entire procedure must be performed quickly to prevent an 
uneven staining of the tissues. The slides should exhibit a brownish- yellow 
background, intense black reticulum fibers, and black basement membranes. If 
the slides become oversaturated, i.e. too black, destain in a dilute Potassium 
Ferricyanide Solution (F-396-11) for one or two dips.
5. Rinse well in distilled water. Tone in Gold Chloride 0.2% (F-396-5), 1 
minute. If sections are overtoned place in Sodium Metabisulfite, 3% (F-396-12) 
for 1-3 minutes. Rinse well in distilled water.
6. Sodium Thiosulfate 3% (F-396-9), 1-2 miutes. Wash in running tap water, 10 
minutes. Rinse well in distilled water.
7. Stain in Harris' Hematoxylin (F-396-6) containing 2-4ml of Glacial Acetic 
Acid per 100 ml for 5-15 minutes. Wash in water.
8. Differentiate in Acid Alcohol 1% (F-396-13) until the sections turn red.
9. Blue section in Ammonia Water, 0.3% (F-396-14). Wash thoroughly.
10. Counterstain in Eosin Y, 1%, Alcoholic Solution (F-396-7).
11. Dehydrate in 95% alcohol, absolute alcohol and clear in xylene 3 changes 
each. Mount.

Stain Results:
Basement membranes, reticulum fibers: Black
Nuclei: Blue
Cytoplasm, collagen, connective tissue: Pink-orange

References: Jones, D.B., Amer.J.Path. 27:99 (1951). AFIP Manual of Histolocical 
Staining Methods, 3rd ed., Ed. L. Luna: NY: McGraw-Hill Publications, c. 1968, p. 
97."
~


It became apparent that something went wrong during Step 4 when the slides were 
in the glass container (not a coplin jar - we have ten slides that we need to 
stain so we're using a rectangular glass container that holds ten slides on 
their sides - it does require a metal handle to move, but the handle is 
flexible and easy to remove after the glass slide rack has been transferred 
between containers) of silver solution in the water bath because there was lots 
of precipitate on the slides and floating on the surface of the silver solution.

In my first test, I used five test slides (extra slides that we cut from the same blocks 
that were submitted to us). I deparaffinized them in coplin jars (moving them with 
plastic forceps) and hydrated them to deionized water. I transferred the slides to a 
glass slide rack that holds ten slides on their sides, added five blank slides that were 
rinsed in deionized water (so that the displacement of reagents would be equivalent to 
when we stain our ten "real" slides after testing is complete), and completed 
Step 2. I don't recall exactly how long the glass container of silver solution and the 
glass container of deionized water had been heating up in the water bath, but I would 
estimate ~15-30 minutes. The thermometer said that the water in the bath (not inside the 
containers) reached ~60-65 degrees Celsius. The silver solution was clear and colorless 
when I made it up, but by the time I put the slides into the warm silver solution, the 
solution was beginning to turn a light brown color (though it was still clear and I did 
not see any precipitate floating around). I removed the metal handle of the glass slide 
rack after the rack was transferred into the silver solution, but the metal handle did 
dip into the silver solution briefly. At some point, I noticed precipitate floating 
around of the surface of the silver solution. After ~80 minutes, I used plastic forceps 
to remove one test slide from the warm silver 

Re: [Histonet] Bone decalcification

[Bryan Llewellyn via Histonet]

Use surface decalcification:

Surface the block, then remove from the microtome. Do not use the 
microtome for another block, so as to avoid adjusting the position of 
the block face to the knife.


Place the block face down in 4% nitric acid for 2 hours or longer.

Rinse the block and cool down.

Place back into the microtome. Use a fresh blade to ensure sharpness. 
You should be able to get 3 or 4 sections at 5 microns before hitting 
the calcium again.


Bryan Llewellyn


Patricia Latham via Histonet wrote:

To Histopeeps,
Does anyone know if there is a method to decalcify bone once it is FFPE?
Thank you,
Pat L
George Washington University
Washington, DC
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Re: [Histonet] On-line references

[Bryan Llewellyn via Histonet]

StainsFile is still available at http://stainsfile.info

In the downloads section there is a scanned, corrected and reformatted 
copy of the Microtomist's Vade Mecum, 7th edition. This text covers 
other areas as well as medical and is in the public domain in the US and 
Canada, and likely elsewhere. The problem with putting textbooks on line 
is that they must have their copyright expired and be in the public 
domain, which means they will invariably have been published in the 
first half of the 20th century, i.e. 50 or 60 years, or more, ago.


Bryan Llewellyn


Tom Wells via Histonet wrote:

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] contaminating patient requesition forms

[Bryan Llewellyn via Histonet]

We used to have a form with two sheets. The front sheet was a "magic 
carbon" so the patient information was automatically transferred. During 
booking in of the specimen, we separated the sheets. The back sheet 
accompanied the specimen for the gross description and the top sheet 
went directly to the office. The top sheets were retained as they had 
the original signature.


Bryan Llewellyn


Paula via Histonet wrote:

Hello..good morning,

  


Can anyone recommend a creative approach to avoid blood from being
transferred from disposable gloves onto the patient requisition form during
the grossing phase of histology?

  


I've seen a number of smudge marks and have asked the grossing doctor to
avoid touching the requisition form but she says that is unavoidable.  I've
asked her to change out gloves, but she says that is not practical.

  


In the meantime, if billing or lab personnel sees this, they will put on
gloves and insert the paperwork inside a protective sleeve cover.

  


But, what are the types of things the grossing doctor can do to avoid this
transfer besides changing out gloves or avoiding touching the paperwork?

  


Thank you in advance,

Paula

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Re: [Histonet] New lab set up

[Bryan Llewellyn via Histonet]

I have sectioned both standing and sitting, and much prefer sitting. 
However, due to back problems I found it necessary to have a stand built 
for the microtome so that when I was sctioning I reached slightly 
upwards rather than crouched over. This all but eliminated the strain on 
my back. The stand was basically a five sided box with an open front 
made from heavy plywood and sized to fit my requirements. The front was 
open and very useful for slides, pens, tissues and so on. I also used a 
heavy draughting chair which was higher than a normal chair. The 
combination worked very well for daily sectioning over a 20 year period.


Bryan Llewellyn


Vanessa Avalos via Histonet wrote:

We will be setting up our lab on the 2nd floor in the next few months and since 
it's been a while since I have set up a new lab I am looking for any ideas you all 
have. Our lab is a small one that does derm/H only for our 3 locations.
I have always stood to section, but am thinking of getting a higher chair and 
sitting. What do you all think of sitting vs standing, along with counter 
height suggestions? All input whether it pertains to our lab or not will be 
appreciated because I am sure there will be something I will forget.

Thanks!
V. Avalos
AD HISTO


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Re: [Histonet] 5% Chromic acid disposal

[Bryan Llewellyn via Histonet]

I suspect disposal might vary depending on the State. I live in British 
Columbia and we had permission from out city (Prince George) to use a 
dribble tank with lots of water and flush them into the local river (The 
Fraser), but I opted to collect all the toxic chemicals and ship them 
periodically to a facility in our neighboring Province of Alberta for 
recovery and reuse. There must be facilities of that kind somewhere in 
the United States, I would think. That is probably the best option. We 
used the same procedure for mercury salts, osmium tetroxide, etc.


Bryan Llewellyn


Bob Richmond via Histonet wrote:

Sharon at Celligent Diagnostics in Spartanburg SC asks:


We are changing our GMS stain over from a Periodic acid kit to a 5%

chromic acid kit. What do the labs that use chromic acid in special
staining do with the waste/ used chromic acid?<<

I hope someone can give an authoritative answer to this question, with
references. Chromic acid (chromium trioxide, CrO3) is a strong oxidant, and
chromium is toxic and an environmental hazard. I'd be comfortable with
pouring it down the drain with a LOT of water, but what do the authorities
want us to do?

Bob Richmond
Samurai Pathologist
Maryville TN
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Re: [Histonet] Muscle Enzyme Histochemistry

[Bryan Llewellyn via Histonet]

Hi,
I presume you are talking about light microscopy. Glutaraldehyde is 
often used for electron microscopy. If that is the case, ignore this.


Glutataraldehyde fixes similarly to formalin, so the morphology should 
not be much different. However, it leaves the tissue with free aldehyde 
groups and these can react with Schiff's reagent or silver reductions. 
That means if doing a PAS or methenamine silver you should first (before 
oxidation) do an aldehyde block. I recommend aniline-acetic acid for 30 
minutes, although others work as well. If you don't you will have a dark 
background. It is unlikely that enzyme histochemistry will be 
successful, but you never know. Esterase may still be demonstrable, 
possibly, but the ATPase iso-enzymes are unlikely to survive.


Go to http://stainsfile.info and read the pages on glutaraldehyde for 
more information about it.


Bryan Llewellyn


MONICA D. LOCKHART via Histonet wrote:

Hello Histology World!!!

Can muscle placed in Gluteraldehyde be processed for structure?  In this 
situation, all tissue was placed in Glut and we need to perform Enzyme 
Histochemistry but we have no fresh tissue.

Thanks for your help.

Monica D. Lockhart, BBA, HT (ASCP) PBT
Supervisor Clinical Labs Histology
Loyola University Medical Center
2160 S. First Ave, Bldg 110 Rm 2290
Maywood, IL  60153
(o) 708.327.2608
(c) 708.692.8361
monica.lockh...@luhs.org


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Re: [Histonet] Frozen sections and cold acetone...

[Bryan Llewellyn via Histonet]

Cold acetone, and cold ethanol, were used to fix tissues because they 
left enzymes unaffected and still demonstrable. This was in the early 
days of enzyme histochemistry. Pearse' Histochemistry: Theoretical and 
applied,3rd edition, volume 1, page 85 discusses it. I could send a scan 
if you wanted.


Bryan Llewellyn



Morken, Timothy via Histonet wrote:

Can anyone give me a rational for using cold (refrig or freezer-temp) acetone 
to fix frozen sections? Or a rational for using RT acetone.

This is for kidney or muscle bx frozens for immmunofluroescence or 
immunoperoxidase staining.

Normally they air dry for at least 15 minutes (just waiting for frozen 
sectioning to be completed) before going into acetone. Just wondering if we can 
reduce complexity...

I haven't seen anything saying why cold acetone is used, just instructions to 
do so. I always wonder about such things...

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

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Re: [Histonet] (no subject)

[Bryan Llewellyn via Histonet]

All of the Picro-Mallory variants are trichrome stains. An explanation 
of how they work is here:


http://stainsfile.info/StainsFile/theory/tri_gen.htm
http://stainsfile.info/StainsFile/stain/fibrin/fibrin.htm

Follow the links as well for added information.

Bryan Llewellyn



Пешков Максим via Histonet wrote:


Dear Histonetters!
I need in your proffessional help.
Can you explain for me a the chemical mechanism of Picro-Mallory V stain by 
chemcial language?
I will appreciate references about this issue except original article of 
Lendrum A.C. et al (1962)  
https://www.ncbi.nlm.nih.gov/pmc/articles/PMC480427/pdf/jclinpath00070-0009.pdf
Before asking I was read some histotechnological books, but can not find it. They 
are: Bancroft (from 5 to 8th ed), F. Carson A Self-instructional text 3 
ed, C.F.A. Culling (3rd ed), AFIP manuals (3-4 ed), Woods and Ellis (Histology lab: 
A complete reference), Lilli RD, 1962, Romeis 2014 (18 auflage) and some others 
book.
-- Russia,
Taganrog,
Maxim Peshkov.
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Re: [Histonet] Iron Stain

[Bryan Llewellyn via Histonet]

1% neutral red in 0.01% acetic acid (1 mL of 1% per 100 mL stain) works, 
as does 1% safranin. Both are stable, although should be filtered from 
time to time. Stain 1 minute, water rinse, ethanol, xylene and mount 
should work. I had one pathologist who preferred a very light 
progressive H She said she was interested in increased amounts of 
iron and normal amounts not being glaringly obvious didn't bother her.


Bryan Llewellyn


Bob Richmond via Histonet wrote:

Tasha Campbell, B.S.,HTL(ASCP), at Frederick Gastroenterology Associates in
Frederick, Maryland asks:


Is nuclear fast red the only counter stain for the Prussian blue stain? I

have a Masson's trichrome kit and was wondering if [Biebrich] scarlet could
be [used as] a counterstain. I won't be doing the iron stain very often at
all so I am trying to keep dry reagents on hand to make up as needed so
they do not expire so quickly.<<

This old pathologist never found Biebrich scarlet or the various fuchsins
to be satisfactory substitutes for nuclear fast red - they're too dense,
and they don't stain nuclei specifically.

Nuclear fast red (Kernechtrot, C.I. 60760) is a dye with problems - poor
shelf life both dry and in solution, and it will probably go out of
manufacture eventually. It's used as an aluminum lake for staining nuclei.

I'm not sure what would serve as an alternative. I'd like to know more
about brazilin (particularly about Anatech's Brazilliant), a catechol dye
similar to hematoxylin, with good specificity for nuclei.

Bob Richmond
Samurai Pathologist
Maryville TN
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Re: [Histonet] Wright-Giemsa for sections?

[Bryan Llewellyn via Histonet]

Go to http://stainsfile.info/StainsFile/stain/oversight/romanowsky.htm

Bryan Llewellyn


Tyrone Genade via Histonet wrote:

Hello,

Can the Wright-Giemsa stain be used on fixed, paraffin embedded sections?
Does anyone have a protocol?

I want to examine hematopoietic tissue of fish, i.e. the head kidney. No
smears or imprint possible. I would like to use Wrights so I can use the
same stain for blood smears.

Thanks



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Re: [Histonet] Metal embedding molds-large

[Bryan Llewellyn via Histonet]

Sorry!

That should be TSP - trisodium phosphate - not TCP, which might make it 
worse.


Bryan


Bryan Llewellyn wrote:

This used to be a common problem years ago. It is due to crud buildup on
the metal. Boil them with TCP for half an hour, then thoroughly wash
them in cold water. Coat them with a VERY light smear of glycerol before
you use them, preferably each time. That should help.

Bryan Llewellyn.

Diane Satterfield via Histonet wrote:

We are using large metal molds to embed mouse brains.  We are having a
hard time getting to block out of the molds, the paraffin blocks are
sticking.  Sometimes they are coming out cracked.  Sometimes the
cassette comes off the paraffin block.  Any idea why this is
happening? Any advice on how to fix this problem?


Diane L. Satterfield, BS
Manager Brain Tumor BioRepository
Research Program Leader
Duke University Medical Center
Brain Tumor Center Biorepository and Database

diane.satterfi...@duke.edu
office  919-684-4642
pager  919-970-7328
fax  919-684-4975

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Re: [Histonet] Metal embedding molds-large

[Bryan Llewellyn via Histonet]

This used to be a common problem years ago. It is due to crud buildup on 
the metal. Boil them with TCP for half an hour, then thoroughly wash 
them in cold water. Coat them with a VERY light smear of glycerol before 
you use them, preferably each time. That should help.


Bryan Llewellyn.

Diane Satterfield via Histonet wrote:

We are using large metal molds to embed mouse brains.  We are having a hard 
time getting to block out of the molds, the paraffin blocks are sticking.  
Sometimes they are coming out cracked.  Sometimes the cassette comes off the 
paraffin block.  Any idea why this is happening? Any advice on how to fix this 
problem?


Diane L. Satterfield, BS
Manager Brain Tumor BioRepository
Research Program Leader
Duke University Medical Center
Brain Tumor Center Biorepository and Database

diane.satterfi...@duke.edu
office  919-684-4642
pager  919-970-7328
fax  919-684-4975

CONFIDENTIALITY NOTICE:  The information contained in this electronic mail is 
sensitive, protected information intended only for the addressee(s).  Any other 
person, including anyone who believes he/she might have received it due to an 
addressing error, is requested to notify the sender immediately by return 
electronic mail, and to delete it without further reading or retention.  The 
information is not to be forwarded to or shared unless in compliance with Duke 
Medicine policies on confidentiality and/or with the approval of the sender.


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Re: [Histonet] Elastic stain

[Bryan Llewellyn via Histonet]

Please read the article at 
:http://stainsfile.info/StainsFile/stain/elastic/elastic.htm
There are numerous alternatives discussed there. I particularly like the 
Humberstone variant of iron resorcin fuchsin.


Bryan Llewellyn



Nirmala Srishan via Histonet wrote:

Histonetters,

Is there someone who can recommend a Elastic stain other than Verhoeff?

Thanks
Mala









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Re: [Histonet] Metabisulfite rinse for PAS reaction (potassium vs sodium)

[Bryan Llewellyn via Histonet]

You can use either the sodium or potassium salt. Both can also be used 
to make Schiff's reagent as well. In fact, many histotechs leave out the 
sulphite rinse step and simply rinse off in water and wash well with tap 
water. It seems to work just as well.


Bryan Llewellyn


Angela Lamberth via Histonet wrote:

In Carson’s 3rd edition, the reducing rinse following Schiff for PAS is
0.55% potassium metabisulfite (pg 137-138). For PAS/AB the reducing rinse
is given as sodium metabisulfite (Carson pg 149).



My other texts (Sheehan/Hrapchak as well as Vacca) list the reducing rinse
as sodium metabisulfite which makes sense to me since the Schiff is made
with sodium metabisulfite.



Is this a printing error in the Carson book? I appreciate any light anybody
can shed on this.




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Re: [Histonet] aniline oil/Holzer stain

[Bryan Llewellyn via Histonet]

Aniline oil is more commonly known just as aniline. Sigma list it for 
sale at: 
http://www.sigmaaldrich.com/catalog/search?term=aniline=All=0=match%20partialmax=en=CA=product


Bryan Llewellyn



Susan Bachus via Histonet wrote:

I'm trying to locate aniline oil for the Holzer glial fiber stain.  I
purchased what was listed as aniline oil from ENG, but the first hint
that something was wrong was that it was incompatible with the
chloroform also included in the differentiating solution.  Looking at
the fine print on the bottle, I saw that it was actually "aniline oil,
2.5% aqueous", which I would have interpreted as only containing 2.5%
water, except that it seemed to be completely aqueous (2.5% aniline?),
inasmuch as it wouldn't mix with the chloroform.  I can't find any other
source online.  I see that it is also called aminobenzine or
phenylamine, but am also having trouble locating sources under those
names.  I realize aniline oil is relatively toxic, but am using it under
a fume hood.  I tried dipping slides separately in the (apparently
aqueous) aniline and then chloroform, but still didn't get good
differentiation.  Can anyone suggest a source or a substitute?  Many
thanks for your help!   Susan

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Re: [Histonet] Medical/health related post

[Bryan Llewellyn via Histonet]

I think it is rather unfair to accuse Dr. Raff of harassment or misusing 
this forum. I recall that when he first posted references to his blog he 
asked for the members' permission to do so, and he was given that 
permission. Rather than abusing the privilege of posting to Histonet, I 
think he showed distinct courtesy for asking before he did so, and 
remarkable restraint in not responding to the, sometimes rude, 
responses. If members have now changed their minds about giving 
permission to him to past references to his blog, then let that 
permission be withdrawn with the same courtesy as he showed when first 
asking for it.


Incidentally, it takes me about 0.5 seconds to click delete on his 
posts. I do not read them since I am a Canadian and have no real 
interest in US health care experiences or problems.


Bryan Llewellyn



Tyra Connor via Histonet wrote:

I have been on this list for some time now, and I have seen Dr. Raff contribute 
very useful information to many posed questions. Recently this situation has 
shifted, with more posts concerning his blog than actual helpful responses. 
This is not the second or even third time this situation has been addressed in 
this forum. It's sad because I would not want to lose his voice in these 
discussions, but these days whenever I see any post from him at all, I 
immediately delete it. I feel at this point that his refusal to stop is a small 
form of harassment, like a telemarketer that refuses to stop calling even 
though you request that they do so.

Be full of wonder, be wonderful!!


On Apr 28, 2016, at 10:48 AM, WILLIAM DESALVO via Histonet 
 wrote:

Again, why the non-Histo post. Take this to another source. I do not understand 
why Dr. Raff has not been removed from this list serve. This is a valuable site 
for histotechnolgy related issues, please let us keep it that way.

Sent from my Windows Phone

From: Lester Raff MD via Histonet
Sent: ‎4/‎28/‎2016 6:59 AM
To: 
'histonet@lists.utsouthwestern.edu'
Subject: [Histonet] Medical/health related post

http://www.chicagonow.com/downsize-maybe/2016/04/running-mates-and-other-mates-fighting-trump-fighting-cancer/

Lester J. Raff, MD MBA
UroPartners
Medical Director Of Laboratory
2225 Enterprise Dr. Suite 2511
Westchester, Il 60154
Tel: 708-486-0076
Fax: 708-492-0203

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