[Histonet] Leica Bond Rx and clipped corner slides

[Johnson, Teri via Histonet]

Hi Histonetters,

Leica is recommending not to use clipped corner slides on their Bond Rx 
instrument. I know there are labs out there who use this instrument and may 
also use clipped corner slides in their slide printer. For those labs, do you 
have any problem with covertiles not working properly as a result of using 
clipped corner slides?

Thanking you in advance,

Teri Johnson, HT(ASCP)QIHC
Manager, Histopathology and IHC

T +1 760 602 1402 (new phone number)
teri.john...@navigatebp.com

Navigate BioPharma Services, Inc.
A Novartis Subsidiary
1890 Rutherford Rd.,
Carlsbad, CA 92008
USA

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[Histonet] Unsubscribe details

[Johnson, Teri via Histonet]

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Teri Johnson, HT(ASCP)QIHC
Manager, Histopathology and IHC

T +1 760 602 1402 (new phone number)
teri.john...@navigatebp.com

Navigate BioPharma Services, Inc.
A Novartis Subsidiary
1890 Rutherford Rd.,
Carlsbad, CA 92008
USA

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[Histonet] Microtome maintenance

[Johnson, Teri via Histonet]

Hi fellow histonetters,

I am interested in finding out what those of you in CAP/CLIA labs are doing for 
routine maintenance of your microtomes, especially for smaller volume labs. Do 
you have service contracts set up? Or regular (annual or every other year) 
maintenance schedules? The cost of "periodic inspection" is pretty significant.

Thanks in advance and Happy Friday!

Teri Johnson, HT(ASCP)QIHC
Manager, Histopathology and IHC

T +1 760 602 1402 (new phone number)
teri.john...@navigatebp.com

Navigate BioPharma Services, Inc.
A Novartis Subsidiary
1890 Rutherford Rd.,
Carlsbad, CA 92008
USA

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[Histonet] Open position - Associate Scientist: Clinical Trial Testing

[Johnson, Teri via Histonet]

Dear histonetters,

We have a newly open position available for an Associate Scientist at our 
facility in Carlsbad, CA.
ASCP certification is not required, experience in IHC staining a must. Ventana, 
Intellipath, Leica IHC platform experience desired.
Primary responsibilities for this position is testing and maintenance of 
fluorescent IHC multiplex assays and image analysis on clinical trial and 
research project samples, documentation, maintenance duties, etc.  Chromogenic 
staining may also be performed.

If interested or know someone who may be, please refer to 
https://www.novartis.com/careers/career-search#keyword=232747BR for full 
listing details and to apply.

Teri Johnson, HT(ASCP)QIHC
Manager, Histopathology and IHC

T +1 760 602 1402 (new phone number)
teri.john...@navigatebp.com

Navigate BioPharma Services, Inc.
A Novartis Subsidiary
1890 Rutherford Rd.,
Carlsbad, CA 92008
USA

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Re: [Histonet] Histology Hacks - The book

[Johnson, Teri via Histonet]

Hi histonetters!

Some clarification is needed I think.

In this context, the term "hacks" are used these days to describe work arounds 
to make our lives easier. I have seen videos which show many household hacks 
such as using the hole in the flip top can to hold a straw, or using a rubber 
band in a stripped phillips screw head to get the screw out. I'm sure you can 
add more that made you figuratively smack your forehead with disbelief you 
didn't come up with them!

Additionally, I do agree with those who point out that the best option is 
having the work done correctly the first time. However, we all know that one 
pathologist, student, intern, etc. in your practice that doesn't take the 
proper time to make the sample reasonable prior to processing. Or in our case, 
we receive mostly paraffin blocks that are processed elsewhere globally, and 
you never know what you will get. So knowing some workarounds to help us get 
the best section and stain possible is quite useful.

And finally, much of this knowledge is tribal. I learned it from the seasoned 
techs before me, and I pass it along to the techs who follow. I see absolutely 
nothing wrong with this.

Teri Johnson, HT(ASCP)QIHC
T +760 602 1402 
teri.john...@navigatebp.com


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Re: [Histonet] Kappa & Lambda ISH

[Johnson, Teri via Histonet]

Hi Renee,

Are you using two different fixatives for bone marrow vs lymph node? And, if 
yes, have the assays been validated on both fixatives?


Teri Johnson, HT(ASCP)QIHC
Manager, Clinical Trial Testing

T +1 760 516 5954
teri.john...@navigatebp.com

Navigate BioPharma, Inc.
A Novartis Subsidiary
1890 Rutherford Rd.
Carlsbad, CA  92008
USA



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[Histonet] Tracking lot numbers

[Johnson, Teri via Histonet]

Hi Histonetters,

I need to find out what labs are doing for best practices to track reagent lot 
numbers in their tissue processor as they rotate reagents. We currently have a 
preparation log where we track the lot numbers, but by the time we do a 
rotation we may be using a different lot.

As always, thanks for your help and input.


Teri Johnson, HT(ASCP)QIHC
Manager, Clinical Trial Testing

T +1 760 516 5954
teri.john...@navigatebp.com

Navigate BioPharma, Inc.
A Novartis Subsidiary
1890 Rutherford Rd.
Carlsbad, CA  92008
USA



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Re: [Histonet] Proficiency Testing for small labs

[johnson, teri via Histonet]

Hi Gareth,

Good luck on your upcoming inspection.

Regarding Proficiency testing, labs that don't do the usual barrage of special 
or immunohistochemical stains need to have some sort of alternate plan for PT. 
Our lab has outlined alternative assessment in our Quality Assurance Program.  
It requires that you provide details in how you perform all your quality items 
for the lab, including the PT (number of samples, method of comparative 
testing, criteria for successful and unsuccessful results, investigation 
details, documentation process, etc.)  See the information from 
www.cap.org on this:

"Use alternative assessment methods, when necessary. Tests for which PT is not 
available or that do not require enrollment in a formal PT program must be 
assessed twice a year to confirm their reliability. Methods of checking 
accuracy include split sampling with another laboratory, the use of previously 
assayed specimens, pooled material, use of PT products usually educational in 
nature, or clinical correlation. Another example of alternative assessment is 
the CAP's Sample Exchange Registry, which helps facilitate cooperation around 
genetic testing. It is important to emphasize to everyone that just performing 
an alternative assessment is insufficient. As the laboratory medical director, 
your job is to establish evaluation criteria for the alternative assessment. If 
a result falls outside the established acceptable range, conduct an 
investigation in the same manner as you would with formal PT."

In short, we have a bank of paraffin blocks that we use specifically for a 
specific assay's PT (previously qualified with known expression levels). Every 
6 months we perform our particular IHC stain using samples that have been 
blinded to the operator and pathologist. The operator performs a routine run on 
those samples and submits them to the pathologist for scoring. Those scores are 
recorded on a specific document and compared against the previous 6 month 
score. Concordance must be established according to a pre-defined threshold. It 
passes or fails, and any failure must be investigated, explained, and re-tested 
if applicable. Do this for each of your IHC assays. Run more than one sample, 
and when possible do both positives and negatives ; we do either 3 (100% 
concordance required) or 5 (80% concordance required, but any failure is still 
investigated).

Best wishes,

Teri Johnson, HT(ASCP)QIHC
Manager, Clinical Trial Testing

T +1 760 516 5954
teri.john...@navigatebp.com

Navigate BioPharma, Inc.
A Novartis Company
1890 Rutherford Rd.
Carlsbad, CA  92008
USA








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

[Johnson, Teri]

Adam,

I will miss your contribution on the Histonet. You have always been an asset to 
the listserv. I agree with your sentiments exactly, I know what I know because 
of the information I learned on here and from those kind souls willing to 
share. I continue to try to pay it forward and not get caught up in the 
insanity that breaks out from time to time.

Best of luck on your career path. I think you will do well.

Sincerely,

Teri Johnson, HT(ASCP)QIHC
Head, Histology and Electron Microscopy
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Incredible job opportunity - Kansas City, MO

[Johnson, Teri]

I am reposting this in case it was missed earlier. The Stowers Institute is 
looking for someone with a solid background in Histology, including research 
histology and supervisory experience to head the Histology and Electron 
Microscopy Core Facility. I will post the actual ad below, but before I do just 
let me say that working at this facility is a once-in-a-lifetime experience. I 
urge you to visit the website, www.stowers.org, and read about the type of 
reseach they are doing here, and learn more about the Institute and their 
mission. If you have been considering a career move, take and look and see if 
this is something that interests you.

This is a challenging but incredibly rewarding position. You will learn 
something new every day and contribute to really great science. Yes, this is my 
position they are replacing. It has been a great 10 years here, and whomever 
takes over will be managing a staff of really talented histology and electron 
microscopy specialists. I am leaving due to personal reasons, it has nothing 
whatsoever to do with the management, staff, or any issue I have with the 
Institute. Trust me, it was very difficult to make the decision to leave. This 
is by far the best job I have ever had.

I will be moving onward and forward and will still be a regular contributor to 
the histonet after starting my new position next month at GNF (Genomics 
Institute for Novartis Foundation). I am looking forward to the challenge.

Teri Johnson, HT(ASCP)QIHC
Head, Histology and Electron Microscopy
Stowers Institute for Medical Research
Kansas City, MO


The Stowers Institute for Medical Research has an opening for a Head of 
Histology and Electron Microscopy to oversee expert delivery of the highest 
quality service for the detection of gene and protein expression in tissue 
samples; histochemical staining; sample fixation; routine histology; and 
ultra-structural analysis.

 Primary responsibilities include maintaining the current status of projects 
and resolving issues in the EM and Histology labs; actively promoting team 
interaction and participation; monitoring workload and turn-around time through 
the LIMS system; providing oversight and feedback on complex or non-routine 
projects; monitoring usage of services; responding appropriately to unexpected 
peaks in workload and service requests; maintaining effective communication 
with all members of the Institute; troubleshooting problems and communicating 
appropriately with Principal Investigators and the scientific staff, 
administration, and other core facility personnel as needed; making formal and 
informal presentations on Core Center services; ensuring continuing education 
of staff through workshops, webinars, lab meetings, and email communications; 
and reading professional journals and other sources to stay current in 
Histologic and EM techniques.

In addition to outstanding communication skills and the ability to effectively 
multi-task in a team-oriented environment, the successful candidate will have 
QIHC (ASCP) Qualification, and HT or HTL certification.  Previous management or 
supervisory experience is preferred.

Minimum requirements include an undergraduate degree in the sciences; excellent 
knowledge of histologic technique to include fixation, processing, 
histochemical staining, immunohistochemical staining, microtomy (to include 
cryostat, paraffin, and plastic), and ultramicrotomy; experience in a research 
environment with animal model histology; and the ability to apply good 
judgement to troubleshooting, problem solving, and staff management.  Five to 
ten years relevant Histology and/or Electron Microscopy experience in lieu of 
education may be considered.

To apply, send resume and coverletter to care...@stowers.org




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[Histonet] Re: Von Kossa vs Alizarin Red

[Johnson, Teri]

Hi Betsy,

How timely is your question! We are (well, actually Nancy Thomas is) currently 
evaluating the two stains on some chameleon heads and we have found something 
really interesting we never considered. The chameleon has quite a bit of 
melanin pigment in places. The fact the von kossa will end up staining the bone 
black, it doesn't contrast well with the areas of pigment throughout the 
connective tissue. Otherwise, in other species I usually prefer the von kossa.

The silver technique might also work better for combined stains or combined 
histochemical and immunohistochemical staining than using the Alizarin Red. 
Just my preference.

~Teri

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[Histonet] Head, Histology and Electron Microscopy - Stowers Institute in Kansas City, MO

[Johnson, Teri]

Dear colleagues,

I am bringing your attention to the following job opening at the Stowers 
Institute in Kansas City, MO. It is a wonderful place to work, great Principle 
Investigators and research staff, fantastic physical location and beautiful 
facility. You can read all about it at www.stowers.org. The following is the ad 
as listed on the website. If interested, please send resume and coverletter to 
care...@stowers.org.

Best wishes,
Teri Johnson



Head, Histology and Electron Microscopy

The Stowers Institute for Medical Research has an opening for a Head of 
Histology and Electron Microscopy to oversee expert delivery of the highest 
quality service for the detection of gene and protein expression in tissue 
samples; histochemical staining; sample fixation; routine histology; and 
ultra-structural analysis.

 Primary responsibilities include maintaining the current status of projects 
and resolving issues in the EM and Histology labs; actively promoting team 
interaction and participation; monitoring workload and turn-around time through 
the LIMS system; providing oversight and feedback on complex or non-routine 
projects; monitoring usage of services; responding appropriately to unexpected 
peaks in workload and service requests; maintaining effective communication 
with all members of the Institute; troubleshooting problems and communicating 
appropriately with Principal Investigators and the scientific staff, 
administration, and other core facility personnel as needed; making formal and 
informal presentations on Core Center services; ensuring continuing education 
of staff through workshops, webinars, lab meetings, and email communications; 
and reading professional journals and other sources to stay current in 
Histologic and EM techniques.

In addition to outstanding communication skills and the ability to effectively 
multi-task in a team-oriented environment, the successful candidate will have 
QIHC (ASCP) Qualification, and HT or HTL certification.  Previous management or 
supervisory experience is preferred.

Minimum requirements include an undergraduate degree in the sciences; excellent 
knowledge of histologic technique to include fixation, processing, 
histochemical staining, immunohistochemical staining, microtomy (to include 
cryostat, paraffin, and plastic), and ultramicrotomy; experience in a research 
environment with animal model histology; and the ability to apply good 
judgement to troubleshooting, problem solving, and staff management.  Five to 
ten years relevant Histology and/or Electron Microscopy experience in lieu of 
education may be considered.



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[Histonet] Re: TEM using difficult tissue

[Johnson, Teri]

Daphne,

While I can appreciate wanting to try something just to see if it might be 
helpful, I really fail to see any benefit from trying to look at ultrastructure 
of tissues that have been fixed and stored in this way. The ultrastructure will 
be compromised, and it will be difficult to get any useful information from 
fixation or storage artifact in the tissues. You would need to validate your 
findings on freshly, properly fixed materials anyway, so why not just start 
with the best possible chance for success and use the right stuff right off the 
bat?

Good luck no matter which way you decide to proceed.

Teri Johnson
Stowers Institute for Medical Research
Kansas City, MO

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[Histonet] Re: iliac artery attachment to slides?

[Johnson, Teri]

Hi Jim, sounds like you are having a time of it.

I figure Jack Ratliff will chime in as soon as he sees this. In the meantime I 
will give you the same advice he gave to me.

If you are having troubles with tissues adhering, try Haupt's adhesive. You can 
find recipes on the internet to make it yourself, or you can buy it 
commercially ready to use (www.dornandhart.com). I have heard from several 
people who use this consistently with their MMA and they swear by it. Have you 
tried stretching the sections using a few drops of 50% alcohol and a couple of 
soft brushes prior to covering in plastic and clamping? It's going to be tough 
getting circular tissues wrinkle free. I hope others with more experience than 
me will chime in on this.

Best wishes,

Teri Johnson, HT(ASCP)QIHC
Head, Histology and Electron Microscopy
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: Embedding process improvement...

[Johnson, Teri]

Back to the question at hand,

You will need to engage the pathologists to provide information as to the 
correct way to embed the skin specimens. If you have a dermatopathologist in 
your practice, that person will need to provide the information about how 
he/she dissects it, what he/she wants, and why. I have attended a continuing 
education lecture locally by a dermatopathologist and he showed HE slides of 
incomplete and improperly embedded skin samples. He could not render a proper 
diagnosis due to this histologist's inability to give him the correct view of 
the samples. How would you feel if that was your biopsy and someone embedded it 
with complete disregard?

I would like to think that mistakes happen due to a misunderstanding and 
nothing more sinister. There was a time back in the day that we each had a 
grossing room rotation and watched how the pathologists did their grossing. I 
suspect in these busy labs and busy times, that happens less and less.

Having the paper trail of the process and/or quality improvement can hopefully 
demonstrate competency. But the expectation alone can not provide that. The key 
is education. Teach us how it needs to be done correctly. Show us the results 
of our work. Have it evaluated and give constructive feedback. Everybody wins 
in this scenario.

Teri Johnson, HT(ASCP)QIHC
Head, Histology and Electron Microscopy
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: Embedding process improvement and competency assessment

[Johnson, Teri]

Dear Shelly,

I would work for you without reserve.

I have managed both cytometry and Electron Microscopy successfully, and I 
cannot do either technique. However, I understand enough about it to make sound 
decisions and empower my people enough that it works well. It is possible to do 
well if done properly. The hardest part is proving yourself to those who have 
preconceived notions as to your worth and suitability because you are not an HT.

Best wishes,

Teri Johnson, HT(ASCP)QIHC
Head, Histology and Electron Microscopy
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: Baboon Tissues

[Johnson, Teri]

Mike,

It sounds like a classic case of either slow freezing or thawing and slow 
re-freezing causing ice crystal formation. One other guess is that they were 
fixed in formalin but not cryoprotected, and then frozen. I have seen freezing 
artifact in this way as well. Unfortunately there is nothing you can do at this 
point to save the samples. They are architecturally ruined.

It is entirely possible to snap freeze unfixed tissues properly without a 
sucrose gradient. Matter of fact, we only use the sucrose cryoprotection step 
on samples that have been previously fixed. As for the proper collection of the 
new samples, that depends on what will need to happen to them after they are 
mounted on the slide. The best histology results from fixed and cryoprotected 
frozen section. You can usually get something that looks almost as good as a 
paraffin section. Arguably the best IHC is achieved on unfixed and snap frozen 
tissues. Gayle Callis is the master at doing this, she has worked in rodent 
spleen in cryo for many years and can give you tons of good advice on 
sectioning and staining them. You should do a histonet search looking for her 
information on sample handling if you indeed need unfixed frozen sections.

For fixed frozens, definitely put them through the sucrose gradient and then 
snap freeze. Make sure your sectioning temperature is not too cold (shattering 
of the tissue), or too warm (ooey gooey sticky mess). You are looking for the 
Goldilocks temp - just right.

Good luck!

Teri Johnson, HT(ASCP)QIHC
Head, Histology and Electron Microscopy
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: MMA

[Johnson, Teri]

Reuel asks:

Can undecalcified bone MMA embedded tissue be use for Electron Microscopy.The 
tissue was fixed in formalin dehydrated in grades of alcohol, clear in xylene, 
infiltrated and embedded in MMA (MMA, dibutyl phthalate,perakdox). If not,can 
anyone have a procedure how to prepare tissue that are embedded in MMA for EM. 
Is this the same procedure done on a paraffin embedded tissue where you melt 
the paraffin with xylene then hydrate, wash in distilled water then transfer in 
osmium ,wash in water, dehydrate, PO,resin.

I am with Jack Ratliff, I think it is certainly worth a try. But in the 
meantime I would be working on getting additional samples and processing them 
properly for EM. I would expect that no matter what results you end up with, 
you will need to validate them on other samples not previously MMA embedded. 
Artifacts happen with every fixation/infiltration process.

First - ultrastructure details will not be optimal given they were fixed only 
in formalin.
Second - the size of the block might be too large to section on an 
ultramicrotome.You would need to cut down your block face considerably.
Third - if you are able to successfully remove the resin, post-fix in osmium 
and reprocess in epon, write it up. Share your success. This would be a 
fantastic technical paper.

Best wishes,

Teri Johnson, HT(ASCP)QIHC
Head, Histology and Electron Microscopy
Stowers Institute for Medical Research
Kansas City, MO




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RE: [Histonet] LR White

[Johnson, Teri]

Thanks for your information Jack. I will look into the Acrylosin as an option 
for doing MMA samples.

We are using the LR White as an alternative to GMA. We have had lots of trouble 
with getting it to polymerize evenly and consistently. So now, of course, we 
are having issues with knife lines and wrinkling with this resin.

I would be interested in getting your images if you would please send them to 
me directly. I believe the list serve blocks them.

Best wishes,
Teri

From: Jack Ratliff [mailto:ratliffj...@hotmail.com]
Sent: Wednesday, July 27, 2011 4:56 PM
To: Johnson, Teri; Histonet
Subject: RE: [Histonet] LR White

Teri,

You should try using Haupt's Adhesive 
(www.dornandhart.comhttp://www.dornandhart.com/) to secure your resin 
sections. I have never (knocking twice on wood...LOL) lost a section during the 
staining of thin resin sections when using this product. In fact, I have used 
this solution for over 14 years and I even subject my methacrylate based resin 
(Acrylosin SOFT Embedding Solution @ 
www.dornandhart.comhttp://www.dornandhart.com/) to deplastification with 
warmed xylenes prior to staining and still do not lose a section. Also, I have 
found that wrinkle free is proportional to the softness of the resin block 
and the section collection, transfer and mounting method. I have attached a few 
images of specimens embedded with Acrylosin and stained HE, Goldner's 
Trichrome, and Von Kossa - MacNeal's Tetrachrome for your consideration. Feel 
free to contact me if you have any additional questions.

On a related note, I am giving a teleconference sponsored by the National 
Society for Histotechnology (NSH) next month (August 17th) as part of their VIR 
Summer Teleconference Series. During this teleconference I will be talking 
about the use of resin for undemineralized bone histology. Definitely check 
this out if you have interest in working with undemineralized bone!

Resin Histology: A Practical Approach for Demonstrating Undemineralized Bone
Presented by Jack Ratliff, BioMimetic Therapeutics, Inc.

As musculoskeletal research progresses with new technological advancements in 
the areas of biological repair and replacement, histological evaluation 
continues to play a crucial role in the determination of safety and efficacy 
for these new treatments. While most will employ traditional and acceptable 
methods of decalcification and paraffin embedding for the demonstration of 
these critical components of evaluation, these techniques can sometimes be very 
challenging and/or impossible when presented with a variety of implant 
materials or devices. For example, to evaluate safety and efficacy of a 
metallic device coated with a biological therapeutic at the bone interface, one 
will need to forego traditional methods of decalcification and seek an 
undisturbed representation of the specimen by utilizing an embedding media that 
is both as hard as the specimen and the implant material. Additionally, it may 
also be important to use a media that will not distort or dissolve the coating. 
This seminar will address the use of resin histology techniques for the 
demonstration of undemineralized bone. Topics will include tissue preparation, 
fixation, processing, infiltration, and embedding/polymerization with acrylic 
resins. We will also discuss two types of microtomy as related to small and 
large undemineralized bone specimens and the presence or absence of implant 
materials.



Best Regards,


Jack


Jack L Ratliff
Senior Histologist
BioMimetic Therapeutics, Inc.

Chairman, Hard Tissue Committee
National Society for Histotechnology





 From: t...@stowers.org
 To: histonet@lists.utsouthwestern.edu
 Date: Tue, 26 Jul 2011 10:29:06 -0500
 Subject: [Histonet] LR White

 Is anyone out there using LR White for routine resin embedding, sectioning, 
 and staining? I am interested in learning some tips for mounting sections on 
 to the slide as wrinkle free as possible. Also our HE stains are a little 
 bit pale.

 Thanks in advance!
 Teri

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[Histonet] Re: What is 10% Buffered Formalin Acetate

[Johnson, Teri]

Jennifer, the only reason I am aware of using acetate over phosphate buffering 
would be to minimize the precipitation that happens when phosphate buffered 
formalin contacts alcohol concentrations greater than 70%. It is why some 
automated tissue processors have a warm water flush step. The two fixes to this 
issue I have heard are:

1 - use 70% alcohol or lower in the step following the last fixative or
2 - use sodium acetate buffered formalin instead

That's my wild guess for the day. I bet if you ask the investigator, the answer 
is nowhere close to this. grin

Teri Johnson, HT(ASCP)QIHC
Head, Histology and Electron Microscopy
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] LR White

[Johnson, Teri]

Is anyone out there using LR White for routine resin embedding, sectioning, and 
staining? I am interested in learning some tips for mounting sections on to the 
slide as wrinkle free as possible. Also our HE stains are a little bit pale.

Thanks in advance!
Teri

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[Histonet] Re: Histobath revised

[Johnson, Teri]

Dr. Richmond recommended using non-flammable and non-explosive Novec (tm) 
Engineered Fluid HFE-7100. We are currently using this for our freezing and it 
works very well. The main thing we have noticed is the blocks float in it and 
do not sink as they would with other solvents.

It is a 3M product, and my contact there last year was ebin...@mmm.com (Erin 
Binder).

Teri Johnson, HT(ASCP)QIHC
Head, Histology and Electron Microscopy
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: 100 micron paraffin sections?

[Johnson, Teri]

Tim,

The thickest paraffin sections I have cut are 50 microns and they curled so 
tightly right off the blade I can't imagine what you might get with a 100 
micron section. Even if you do get this slab intact, I think the tissue would 
show cracking artifact, not to mention the big chunks taken out at the block 
face like one would get from too aggressive rough trimming (facing into the 
block). Sections of this thickness are almost always better if done with a 
vibratome and mounted out of buffer onto a glass slide.

Good luck!

Teri Johnson, HT(ASCP)QIHC
Head, Histology and Electron Microscopy
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: Formalin fixed frozen sections

[Johnson, Teri]

Peggy,

We fix ours prior to freezing and cryosectioning routinely. You can use 10% NBF 
(commercial prep) or you can make your own using paraformaldehyde, 4% in PBS 
buffer. After fixation, you will need to cryoprotect in sucrose/PBS. We fix, 
then rinse, and then place in 15% sucrose/PBS until the sample sinks. We then 
place it in 30% sucrose/PBS until the sample sinks. We will put it into a 
cryomold with OCT and let sit for some time, then a fresh change and snap 
freeze. The tissues section very well.

Good luck!

Teri

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[Histonet] Re: endogenous peroxidases

[Johnson, Teri]

Hi Emily,

As for when is better to block endogenous peroxidases, I have no direct 
experience but I have heard from some that it should be done post-AR because 
the retrieval can reactivate the endogenous peroxidase. So that's when we do it.

Some folks worry about what effect that might have on their primary antibody 
binding with their target and will do their block after application of the 
primary antibody, prior to the secondary (or whichever step has the HRP label). 
I try to keep it simple and just do my H2O2 block prior to the protein blocking 
step.

Also, we never use methanol as a solvent for diluting the H2O2. It is 
contraindicated in some CD marker staining, and it's cheaper and just as good 
to use aqueous 3% H2O2. Some make it up in buffer, and you can do that as well 
if you wish. For cryosection peroxidase quenching, we use 0.3% H2O2 for 30 
minutes. Otherwise it's a 10 minute step right after the AR in our routine 
paraffin section IHCs.

Hope this helps!

Teri

Teri Johnson, HT(ASCP)QIHC
Head, Histology and Electron Microscopy
Stowers Institute for Medical Research
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[Histonet] Re: Histonet Biopsy Program for Sakura VIP 5/6 Processor

[Johnson, Teri]

Wow, somehow I missed this email. Sorry for the delay.

Andria, take the heat off your alcohols. Keep it on the formalin and the 
paraffin, but do the rest of the processing at ambient temperature. Make sure 
you put only small soft tissues on this schedule though, tougher collagenous 
skin specimens might not process well enough with the quick dehydration times 
you have.

I am curious how you arrived at the timing for the steps. 14 minutes, 9 
minutes, 7 minutes, 23 minutes, etc?

Good luck and best wishes,

Teri

Teri Johnson, HT(ASCP)QIHC
Head, Histology and Electron Microscopy
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: IHC pretreatment with NaOH/H2O2

[Johnson, Teri]

Dear Neil,

I was able to find a protocol like you described published where they had done 
a regular IHC protocol on free-floating tissue sections for a-MSH antibody 
using 0.3% H2O2, normal serum block, etc. They go on to describe that for 
P-STAT3, the tissue needed to be pretreated with 1% NaOH and 1%H2O2 in water 
for 20 minutes, 0.3% glycine for 10 minutes, and 0.03% SDS for 10 minutes. 
After that it looks like a standard protocol. Here's the URL for that paper: 
http://endo.endojournals.org/cgi/reprint/144/5/2121?ijkey=23f435f1cf61629953465217ca752dd73df366e9

They don't say why this was necessary, but it might be worth trying to track 
down one of the authors and ask them.

It seems like the sodium hydroxide and hydrogen peroxide serves as an oxidizer 
somehow.  Since they are not doing IF, maybe the glycine is needed to bind with 
glycine receptors in the brain prior to staining? And I suspect the SDS is 
needed for permeabilization, even though they are using a sectioned sample. The 
right detergent can make all the difference.

I would really be interested in knowing what the rationale is, but it seems 
like they figured out what physiologic changes needed to happen for this 
antibody to work in brain. If you do have a conversation with the authors, 
please report back here because my curiosity is now at high roar.

Best wishes,

Teri Johnson, HT(ASCP)QIHC
Head, Histology and Electron Microscopy
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: Mouse CD45 staining of bone marrow

[Johnson, Teri]

Dear Adam,

Thank you for your detailed email regarding your problem with CD45 
immunostaining! That helps tremendously know where to start in a reply. I have 
cc'd the Frozen section CD marker guru on this, so expect an equally detailed 
response to this in the near future. She'll be able to tell you how to proceed 
here, but I suspect your problem is aldehyde fixation. The amount of time the 
cells are fixed for flow cytometry is very minimal. Quite different from the 
overnight treatment your protocol uses.

Gayle will likely tell you to use unfixed, undecalcified Cryojane sectioned 
bone sections that are post-sectioning fixed in Acetone/Ethanol 3:1.

First I need to know if the immunostaining protocol you are using is currently 
providing nice strong immunostaining for other markers. In other words, is the 
detection you are using a sound one at those conditions? If no, then optimize 
your detection system first. If yes, then read on.

If you are liking the morphology of the fixed and cryopreserved bone (and who 
wouldn't?) then you can perhaps try using Zinc Formalin as a fixative prior to 
sucrose cryoprotection and see if that works.

Some CD markers work only with Zinc Tris buffer (Beckstead), that contains no 
aldehyde. You can find tons of information on this by doing a histonet or 
google search. Fix the tissues with the Zinc Tris, decalcify in EDTA, section, 
and then do what you usually do.

For the samples you already have available, maybe try using some gentle HIER 
with citrate pH 6.0 or a higher pH, 60 degrees C for 10-20 minutes to see if 
you can recover some of the antigens. Yes, you can do AR on frozen sections on 
fixed tissues.

Ok, start there. This is just a mere appetizer compared to the full course meal 
you are about to get.

Enjoy, and good luck!

~Teri

Teri Johnson, HT(ASCP)QIHC
Head, Histology and Electron Microscopy
Stowers Institute for Medical Research
Kansas City, MO




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[Histonet] Chromogenic ISH with riboprobes service

[Johnson, Teri]

Can anyone recommend a laboratory or company that does ISH service using 
chromogenic detection with riboprobes we supply on slide mounted cryosections?

Thanks for your help, as always.

Teri Johnson, HT(ASCP)QIHC
Head, Histology and Electron Microscopy
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: HE Stain

[Johnson, Teri]

Allison Scott wrote:

Hello to all in histoland and Happy New Year.  We are having issues with
our HE stain.  The nuclei are staining very blue to purple and the
mucin is staining blue to purple-blue.  It is difficult to see the
nuclear detail.  The mucin is obscuring things.  We have not changed our
process for staining or processing.  The funny thing is that it is only
in the Biopsy cases, and it is every few slides.  The surgical  cases
are all right.  We checked the alcohol and xylene for water, and there
is not any.  My tech changed out the stain and we are staining a new
batch of slides.  If anyone has any idea what is wrong, any help would
be greatly appreciated.  I have gone over our processes and nothing has
changed.  The reagents are the same, the staining times are the same,
and the processing times are the same.  We are using the Shandon Gemini
stainer and VIP processor.

Allison Scott HT(ASCP)
Histology Supervisor
LBJ Hospital
Houston, Texas

When you say it is every few slides, is there variation within multiple slides 
of one case? If all the slides (levels) of a case look the same, I would 
suspect a problem with that particular case, not the processing and not the 
staining.
Are these all GI biopsies? Or do you see this in other needle biopsies, 
cervical biopsies and the like? GI biopsies tend to show hazy nuclei in 
epithelial cells commonly. There are several explanations and mostly are 
attributed to inadequate fixation.

It could be you a client who has changed how they collect and fix the 
specimens. Are they letting them dry on a gauze before putting them in 
fixative? It could be you have something out of place on the tissue processor. 
It could be there is water under the tissue on the slide and it is subject to 
high heat (microwave or oven) and is cooking the cells prior to staining. As 
pointed out previously, it could be a problem with your hematoxylin pH.

Good luck, and please let us know what the fix for this is when you get it 
figured out!

Best wishes,
Teri Johnson, HT(ASCP)QIHC
Head, Histology and Electron Microscopy
Stowers Institute for Medical Research
Kansas City, MO




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[Histonet] Re: custom-made polyclonal antibody doesn't stain

[Johnson, Teri]

Barbara,

I see your antibody was purified by the manufacturer already so I don't think 
it would be useful to purify it again. It sounds as though it's not a good 
antibody yet. How did the company test it, by elisa? Since your western and IP 
shows no specific staining, I'm not all that encouraged this antibody will be 
useful for IHC studies.

Maybe someone with more experience with this than me will chime in here. We did 
some testing of custom antibodies for one of our PIs and never got anything 
useful for her experiments. Others here have had great luck with theirs.

~Teri

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[Histonet] Re: long term storage of tissue

[Johnson, Teri]

Melissa,

I definitely would store them in 70% ethanol rather than leaving them in 
fixative for that time period. They should be fine for BrdU immunostaining 
after paraffin processing.

Good luck with your studies!

Teri

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[Histonet] Question from one of our researchers

[Johnson, Teri]

Can someone give me ideas to pass along to one of our researchers? Of course 
adhesion molecule antibodies are the first thought, but not for heterogeneous 
cell populations. So I was wondering maybe an antibody cocktail? Are there any 
lectins that might show this? Thanks!

I am wondering what kind of cell membrane marker is recommended to stain 
tissue section for imaging (going to process the image with 3D Imaris software).
Cells are mouse tissue section in paraffin and very heterogeneous. To see 
cell-cell boundary clearly.

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[Histonet] Re: Microwave processing effect on DNA/RNA

[Johnson, Teri]

Matt,

You asked an interesting question and it got me to googling. Since microwave 
radiation is non-ionizing it should not adversely affect things like DNA and 
RNA. I found this summary of a group of publications on this and apparently in 
1995, Kakita demonstrated that microwaves were capable of  fragmenting viral 
DNA, and they were able to conclude it was not due to the heating effect. 
http://www.rfsafe.com/research/rf_hazards/dna_damage/microwave_effect.htm

Most of the data I can find seems to correspond to the type of electromagnetic 
radiation produced by cell phones, because that's the biggie going around these 
days. Here's a more recent paper on cell phone microwaves and their effect on 
human lymphocytes. It might be useful to check out the reference list, 
especially the ones cited in the introduction. 
http://www.hese-project.org/hese-uk/en/papers/sarimov_chromatin_heatshock_ieee04.pdf
  The thing I don't know is how the microwaves from cell phones equate, or if 
they even do, with microwave oven exposure, especially since we can control the 
samples from being overheated. Also, this study was done on cell cultures, 
which while it might give interesting and useful information, it does not mimic 
the conditions of a whole organism. In addition, the microwave effects were 
reported on unfixed cells, unlike the samples you will be processing.

What I would do if I were you is get a demo unit in, simulaneously process 
10-20 different patient samples with the microwave processor and with your 
routine processor after your routine fixation, and then send them off for 
analysis. That should tell you all you need to know right there.

If anyone has already done this, please speak up!

Best wishes,

Teri Johnson, HT(ASCP)QIHC
Managing Director, Histology Facility
Stowers Institute for Medical Research
Kansas City, MO




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[Histonet] Re: Turnaround time for Research Pathology

[Johnson, Teri]

Julie,

You asked about standard turnaround times for histology services in the 
Research environment. As you eluded to in your second paragraph, rarely do we 
have standard projects. Often we're asked to do serial sectioning of an entire 
organism and we have no idea how many slides we'll end up with, or how long 
it'll take is to do each one. Someone can submit one sample, or 30 samples. 
Paraffin processing and 1 HE can be turned over fairly quickly depending on 
the existing work queue. With some exceptions we take things as first in, first 
out.

Here's what I have on my intranet page:
Turn-around Time:
It is our goal to supply completed work to the researcher within 3 to 7 working 
days from submission. Fixation, decalcification, special handling, or other 
procedural steps may delay completion. Any special turn-around needs should be 
communicated to the histology staff and we will do our best to meet them.

There are times we can receive a sample, process it over night, section and 
stain it the next day and have it completed in 24 hours. The researchers are 
always delighted and surprised when we can do this. It's more usual that we can 
turn around fairly routine and fairly simple requests within 3 working days. If 
we have a long work queue, we try to let the researcher know there might be a 
delay in getting their samples back as soon as they submit the sample 
regardless of what the services are.

We always will move things up the queue when it's needed for publication 
submission reviews or other special circumstances. Communication is critical. 
Give them a reasonable timeline (you may have to determine this yourself based 
on usual workload and staffing). Give them timely information when there are 
circumstances which interfere with achieving that timeline, and update it when 
you need to. Finally, work together when exceptions need to be made.

It's worked out well for us so far. Good luck!

Teri Johnson, HT(ASCP)QIHC
Managing Director, Histology Facility
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: Autofluorescence in murine white adipose tissue cryosections

[Johnson, Teri]

Dear Frank,

Good luck with ridding your samples of autofluorescence. Fat is always very 
brightly fluorescent and I suspect it might even be so without any aldehyde 
fixation. First I wondered how successful you'd be using Sudan Black B, knowing 
it is a fat stain, and knowing that it's been published for this purpose. So 
you might try just that and see how that works. In addition you could try using 
a combination of UV irradiation and the Sudan Black B and see if the 
combination works for you. They use paraffin sections in this reference, but it 
might work as well with cryosections. 
http://www.microscopyu.com/references/pdfs/Viegas_etal_Eur_J_Histochem-51-59-2007.pdf

Good luck, and do report back if you find a solution to this dilemma!

Best wishes,
Teri Johnson, HT(ASCP)QIHC
Managing Director, Histology Facility
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: looking for Gillette super stainless Inoxydable Blades - Vibratome

[Johnson, Teri]

Hi Hadley,

We get our blades from EMS (Electron Microscopy Sciences): 
http://www.emsdiasum.com/microscopy/products/preparation/blades.aspx?mm=10#71990

On this page you'll find the injector blades. Further down they have the double 
edge blade you can break in half and use in the holder. Try those and see if 
they work for you.

Best wishes,

Teri Johnson, HT(ASCP)QIHC
Managing Director, Histology Facility
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: Cryostat vs decal sections of bone

[Johnson, Teri]

Hi Louise,

Remember me? It's been a while.

You asked if there were distinct advantages to using cryosections of bone over 
decalcified wax samples for IHC and ISH and the answer is yes, of course. Some 
antibodies and probes just won't work well in paraffin processed samples, 
either due to the decalcification process or the alcohol/solvent exposure. The 
ISH purists know that just paraffin processing can diminish signal greatly. 
Having said that, obtaining excellent sections of undecalcified, fresh frozen 
bone is tricky. Very tricky indeed, and what works one day doesn't work the 
next. You'll need to have at least one tungsten carbide knife for it, so make 
sure you also get the regular blade holder. I don't know if you still use steel 
knives on the cryostat, and if so you'll already have the correct blade holder, 
but we use disposables so we needed to order a separate one.

We've had pretty good luck getting beautiful sections of fixed and decalcified 
cryosections and the cryojane, and this would work well for samples that just 
flat don't like paraffin processing but the antibody staining does ok with 
formalin and formic acid treatment. Having the system opens up this possibility 
for your studies as well.

Best wishes always,

Teri Johnson, HT(ASCP)QIHC
Managing Director, Histology Facility
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: Special stain to demonstrate silver

[Johnson, Teri]

Maria,

I don't know of any particular procedure or stain to demonstrate silver, but I 
got to thinking wouldn't it reduce with formalin fixation to a visible state? 
Or maybe strong sunlight?

Kind regards,

Teri Johnson, HT(ASCP)QIHC
Managing Director, Histology Facility
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: Two antibodies from the same host with tyramide

[Johnson, Teri]

Adam,

Thanks for the detailed protocol. I can't explain it either but I'll throw some 
ideas out there and maybe it'll stimulate discussion. Hopefully Chris 
vanderLoos will chime in.

Have you tried doing the labeling in reverse, using the VE-Cadherin first and 
then blocking with the goat IgG, and then doing the other antibody/tyramide 
protocol? I'm not sure what effect, if any, the hydrogen peroxide might have on 
the anti-goat fluorescent 649 label, so you could certainly do that block prior 
to doing the first primary antibody incubation.

Also, you may have to double the concentration of the Cadherin to get it to 
label at the same intensity as you do in single staining. But still I might 
have expected to see a signal.

Do the two antibodies co-express? Could there be something with the covalent 
binding of the HRP-tyramide complex that shelters the antigenic sites for the 
second antibody? That's why I wondered what might happen if you reversed your 
protocol.

Let us know if you get it figured out and what fixed it for you.

Good luck,

Teri Johnson, HT(ASCP)QIHC
Managing Director, Histology Facility
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: Two antibodies from the same host with tyramide

[Johnson, Teri]

Adam,

I've been doing some searching and the tyramide shouldn't shelter subsequent 
IHC reactions, so I don't think that's it. I found evidence of co-localization 
of mRNA ISH signal with tyramide enhancement and IHC staining without 
amplification in a publication from J Histochem Cytochem.

I'm actually quite surprised you are getting single staining using the goat 
anti-mouse antibody from RD systems. The one I found online (AF1002) has only 
been tested in Elisa and western blot, and I'm not seeing any sort of antigen 
retrieval in your protocol. I would suspect that most endothelial markers (with 
the exception perhaps of Factor VIII) would require it. The fact this is 
actually working for you in formalin fixed paraffin embedded mouse bone should 
be cause for celebration.

I still would be interested in knowing if reversing the protocol would have any 
affect on the staining, especially if you can do the Cadherin first, verify 
staining briefly, then continue on with the other antibody and its detection 
and see what happens.

Best wishes,

Teri Johnson

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[Histonet] Re: paraformaldehyde

[Johnson, Teri]

Margaret, good luck getting 37 grams of powdered PFA to go into solution. Even 
making a 10% solution is difficult, usually requiring heat (60 degrees C) 
and/or sodium hydroxide to get it into solution. Once you do, it's going to 
want to repolymerize and you'll end up with a real mess. You'd do much better 
to use stock 37% formaldehyde you can buy commercially.

Teri Johnson, HT(ASCP)QIHC
Managing Director, Histology Facility
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: Acetone Fixation

[Johnson, Teri]

Atoska Gentry asked:
hello, if any of you have acetone fixation incorporated into you frozen
section H E staining protocol will you please advise me on adjustments
necessary for routine staining protocol. Also, out of curiosity please
enlighten me on the purpose for post sectioning acetone fixation on
tissue samples  initially fixed in   95% ETOH/ Acetic Acid? Your prompt
replies will be greatly appreciated. ~ Atoska

Several thoughts come to mind.

Acetone fixation is usually used only for preserving antigenicity on fresh 
frozen tissue samples. For slides needing HE I would use a formalin fixative 
prior to the HE procedure for fresh frozen tissues. Acetone fixed HE stained 
cryosections are uuugly.

You should not be doing frozen sections on tissues initially fixed in alcohol. 
Alcohol is an anti-freeze and you will not get good freezing/sectioning of the 
samples. You can section unfixed samples and then fix after mounting on glass 
slides in the Alcohol/acetic acid fix.

Teri Johnson, HT(ASCP)QIHC
Managing Director, Histology Facility
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] MO Society for Histotechnology 2010 Spring Symposium

[Johnson, Teri]

Hi all!

The Missouri Society for Histotechnology cordially invites you to our 2010 
Spring Symposium for continuing education and histopathology technique and earn 
up to 12 CEUs.

This year's conference will be held in: 
Saint Louis, Missouri
Thursday, May 20th to Saturday, May 22nd

We will stay at the Sheraton Westport Lakeside Chalet which is located on 
Westport Plaza, a centrally located 42 acres development offering an 
unparalleled combination of amenities and services, where you may enjoy over 20 
restaurant and entertainment venues. It is nestled in the heart of St Louis and 
surrounded by everything exciting that the city has to offer.

Room reservations can be made by calling 1-800-822-3535. Hotel reservation 
deadline for symposium rate is April 29, 2010. Reservations received after this 
date will be subject to room  rate availability. To secure your symposium rate 
please make your reservations early and indicate that you are attending the MSH 
symposium. 

Online room registration and meeting information is available at 
www.starwoodmeeting.com/Book/mohistotechnology. Visit our webpage for more 
information and links to the Symposium at www.missouri-histo.org/index.cfm For 
registration and more information about the program and work shop descritions, 
download the Brochure 2010 and Workshop Descriptions MHT 2010 in our 
downloads page. 

Obtain a discounted price on the symposium registration fee by becoming an 
active member of the society. Download a Membership/Renewal Application, fill 
it up and send it with your dues and your registration for the symposium.

For more information, contact Amanda Kelley, MSH President, at 
amanda.kel...@stlukes-stl.com or Sharon Walsh at userwa...@sbcglobal.net

Hope to see you there!

 



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[Histonet] Re: IHC Question

[Johnson, Teri]

Dear Ellen,

You wrote: I'm interested in performing IHC on a mouse ligament.Given 
their minuscule size,  in lieu of paraffin or cryostat embedding and 
sectioning, we would like to embed in plastic and section at 1 micrometer.   
Things we are interested in detecting include collagen type 1 and III, CD31, 
Arginase 1 and iNOS---to name a few.   Is there a plastic out there that is 
recommended for Immuno work?   I have performed a brief literature search and 
have come across Immunobed, JV-4 as well as LR White and Lowicryl.   Is anyone 
familiar with any of these and whether I would have any success?I am 
familiar with Epon-Araldite embedding and sectioning with glass knives, but 
have never used any of the above plastics.
Thanks for any info

I think you have a typo, it shoudl be JB-4 and not JV-4. JB-4 is not suitable 
for IHC.

Any Methyl methacrylate or polymethyl methacrylate formulas should work for 
you. In your list, Immunobed, LR White, and Lowicryl would all be suitable.

Sectioning and mounting these are different from usual mounting methods. You 
will need to wet the block face with alcohol for each section. You will put the 
cut section on a glass slide flooded with alcohol and stretch it, pulling out 
wrinkles. It will need to be covered with a plastic film, rolled with a roller, 
and put into a press of some kind and dried overnight. It helps to watch 
someone else do this first before you try doing it on your own.

We have limited experience with this. Perhaps one of the hard tissue folks will 
chime in here with some great resource to help you.

Good luck!

Teri Johnson, HT(ASCP)QIHC
Managing Director, Histology Facility
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] Re: Diff-Quik

[Johnson, Teri]

Dear Colleagues,

At the risk of beating a dead horse, I would like to thank Bryan Llewellyn for 
so eloquently and succinctly echoing my own thoughts on the matter that Dr. 
Richmond commented on which has drawn criticism. For those who may have missed 
it, I'll include it below my own meager response.

Teri Johnson, HT(ASCP)QIHC
Managing Director, Histology Facility
Stowers Institute for Medical Research
Kansas City, MO


Date: Wed, 27 Jan 2010 13:57:16 -0800
From: Bryan Llewellyn llewl...@shaw.ca
Subject: Re: [Histonet] Re: Diff-Quik
To: Histonet histonet@lists.utsouthwestern.edu
Message-ID: 26938660d7e848faa055bcb0a76df...@bryanpc
Content-Type: text/plain; format=flowed; charset=iso-8859-1;
reply-type=original

I have worked in several laboratories, both in Canada and England.  I
trained there and was taught by technologists, not pathologists, to check
the staining of all my slides, which I have always done.  I finds that even
when asked, most pathologists do not return poorly stained slides, yet how
can we know of deficiencies if they do not?

I worked in several histology labs over my career and slides were checked in
all but two.  I was able to introduce it in one of those, but not in the
other due to technologist resistance (we've always done it that way).  It
was the technologists who resisted it.

I don't see why Dr. Richmond needs to apologise.  He merely drew attention
to his work experiences in numerous histology labs and decried what is, in
his experience, a common practice.  In other words, he spoke about real,
actual events.  Why does being truthful require an apology?  Are our egos so
fragile that we can't take valid criticism as a profession nor as
individuals?  He certainly did not accuse those who got upset of failing to
check slides, so why take umbrage?  Noting and decrying poor histological
practices should be a concern for everyone.  Surely we are concerned about
service to patients not about our egos.

Bryan Llewellyn


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[Histonet] Re: Counterstain for fluorescent tissues

[Johnson, Teri]

John, would the neutral red technique work the same with unfixed tissues? The 
fluorophore Nick uses might not fluoresce after formalin fixation. I suspect he 
would also need to be sure it's stable after mounting with DPX or other 
resinous medium.

Teri Johnson, HT(ASCP)QIHC
Managing Director, Histology Facility
Stowers Institute for Medical Research
Kansas City, MO


Neutral red (CI 50040) is an excellent fluorescent Nissl stain: 0.002%, in 
water, for 5 minutes; dehydrate, clear, and mount in DPX or another 
non-fluorescent resinous medium. With excitation by either near-UV or blue 
light (range 325-500nm) the Nissl substance and nuclei fluoresce yellow-orange.

Reference: Allen, D. T.  Kiernan, J. A. 1994. Permeation of proteins from the 
blood into peripheral nerves and ganglia. Neuroscience 59(3):755-764.
   We used this very dilute neutral red as a fluorescent counterstain for 
paraffin and cryostat sections of  formaldehyde-fixed tissues from rats that 
had received iv injections of rhodamine-labelled albumin (green excitation, 
orange-red emission, localization mostly extracellular).
   Franz Nissl was a man, not a granule, substance or stain, so we should give 
his surname its capital N. Check out http://www.whonamedit.com.

John Kiernan
Anatomy, UWO
London, Canada
= = =


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[Histonet] Re: Bubbles on Fresh Frozen tissue after IHC

[Johnson, Teri]

Hi Nick,

Do your peroxidase blocking offline in a coplin jar or staining bucket, prior 
to loading them in the capillary gap holder.

Problem solved.

Teri Johnson, HT(ASCP)QIHC
Managing Director, Histology Facility
Stowers Institute for Medical Research
Kansas City, MO


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[Histonet] FW: OT fuchsia

[Johnson, Teri]

 
One of the researchers here summed it up nicely when he reported that most 
discoveries in the lab are not heralded with cries of Eureka!, but rather 
with hmmmnow that's interesting...

Teri Johnson, HT(ASCP)QIHC
Stowers Institute for Medical Research
Kansas City, MO
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[Histonet] Re: Compresstome

[Johnson, Teri]

Ed,

We have a 2 year old model of the Precisionary Instrument vibrating microtome 
that was made specifically for fixed tissues for histology. Back then it was 
the VF400 model. It is supposed to be able to cut 10 micron thick sections and 
we have not tested that it has this capability yet. It's design is simple and 
it is easy to use. It's not as sexy as some of the other Vibratome units, but 
it seems to work pretty well. One of our researchers has a model they use for 
live tissue work and they have had great success with it.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Surgipath Automated Slide Printer

[Johnson, Teri]

Is anybody in histoland using the Surgipath Automated Slide Printer, Item 
#04V5000? If so, could you please let me know how it's working for you?

Thanks!

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: Rabbit-on-Rodent HRP polymer

[Johnson, Teri]

Jennifer,

You will not need to use the avidin/biotin block in your protocol. The polymer 
technology does not use any sort of biotinylation or avidin system for 
staining. That should help knock some time off your protocol.

You will need to make sure you use a peroxidase block (3% hydrogen peroxide) in 
your staining protocol to block endogenous peroxidase activity (namely red 
blood cells/sera).

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: Isotype background

[Johnson, Teri]

Hi Adam,

We have had the exact same experience you have sometimes when we're using Goat 
isotype and Rabbit isotype negative controls. Sometimes you can fiddle with the 
protocol and minimize some of the background, and sometimes you can't. Rabbit 
and goat antibodies/Igs are notoriously sticky. We use a non-serum protein 
blocker in our stain protocol and still have this happen.

The most ideal negative control would be non-immune serum from the animals that 
were immunized to produce the antibody. Since most places do not make that 
available, we have to purchase just normal animal species isotype control and 
hope for the best. This might explain why your antibody is not showing the 
background staining, but your negative control is.

Are you using any detergent in your wash buffers? You can try adding some Tween 
20, 0.05% up to 0.1% or so and see if that cleans some of it up. Others might 
use Triton X-100, but we prefer to use Tween as it's a gentler detergent for 
slide-mounted samples.
How many rinses are you using? We rinse 3x between steps and that should be 
adequate.
You should be using a streptavidin block kit from Vector since you are using a 
conjugated streptavidin in your staining protocol.
Your use of donkey serum as a blocker and as an addition in your seconday 
antibody should block any Fc receptor staining (in theory).
Have you tried titering out the isotype control to the point of negativity? Do 
you ever get that?
What is your dilution of your secondary antibody? Streptavidin? Could those be 
titered out more to get better signal/noise ratio in your primary antibody 
treated sample?

Yes, you should try a null (no primary antibody - diluent only) control with 
your detection system on your sample and see if there is any background 
staining. But I would only use that to gauge the degree of background staining 
from your detection system in the absence of primary antibody. In most cases, 
unless there is some degree of endogenous staining, those slides are 
beautifully negative. Not ideal to use as a negative control other than 
demonstrating non-interference with the detection system.

I'm sorry I don't have any specific answers. Best I can do is sympathize and 
offer some possible things to look at.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: IHC on plasma cells

[Johnson, Teri]

Kim,

CD138 is a good marker for plasma cells. Contact Chris van der Loos for a 
protocol. He mentioned this at NSH in Denver 2007.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: ISH on decaled bone

[Johnson, Teri]

Gudrun is absolutely correct. The HCl destroyed your nucleic acids. The best 
decalcifier for RNA or DNA is EDTA.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: License

[Johnson, Teri]

Joyce,

You might have some luck calling the following places:

Physician's Reference Lab, Overland Park, KS - Donna Miller

Shawnee Mission Medical Center, Overland Park, KS

North Kansas City Hospital - N. KC, MO - Nancy Warren

University of Kansas Medical Center, Kansas City, KS - Timothy Chilton

Midwest Anatomic Pathology Lab, Overland Park, KS

Litton Laboratories, Blue Springs, MO

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: Harris hematoxylin weak staining

[Johnson, Teri]

Aazath,

There are several strategies you can use to improve your hematoxylin staining.

1 - use distilled or DI water before putting your slides into the stain. 
Sometimes the chemicals/minerals/metals in the tap water can weaken your stain.
2 - replace your stains when they start showing signs of weakening, or 
routinely in order to keep the quality even.
3 - try using more than one bucket/dish of hematoxylin and split the total 
staining time between them. Then you can rotate the last one up to the first 
position, and put a new change in the second station. This might help keep your 
staining more consistent.
4 - does your tap water seem to contain a high level of chlorine? If so, you 
can use a bluing agent to blue your stained slides, and then use DI or 
distilled water rinses.
5 - do not linger in the decolorizing acid alcohol. Usually one quick dip 
followed by an immediate plunge into water is all you need.
6 - some have reported that their eosin, if the pH drops too low, can leach out 
some of the hematoxylin. According to Carson, the ideal pH of eosin is between 
4.6 and 5.
7 - what thickness is your tissue? 3 microns thin tissue needs longer in the 
stain solution than 5 microns thin tissue.

There may be others that folks will chime in with, but these are the most 
obvious ones to me.

Good luck!

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: DIF tissue in GLUT

[Johnson, Teri]

Jim, unless they've changed methodologies since I left clinical, they process 
all DIF for renal biopsies for cryosectioning.

I'm very aware of the different methods for quenching autofluorescence, and I 
know that what works for some samples doesn't work entirely for all. 
Additionally, what may work for formalin induced AF may not even touch GLUT 
induced AF. Best thing to do is try various methods, and see what works. Even 
more important is to provide positive and negative controls (both treated with 
the chemical agents for AF) for this case so you can be reasonably certain that 
if you have diagnostic staining that's consistent with the controls, that it is 
real.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: DIF in GLUT

[Johnson, Teri]

P.S. Jim, thanks for the references, I'm on my way to dig them out as we speak.

Cheers!

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: uneven alternating sections on cryostat

[Johnson, Teri]

Nathan, you got great information already.  Be careful to make sure things are 
snug, but don't really clamp down on the holders. It's just not necessary. It 
just needs to be sturdy and not moveable.

If everything appears to be snug, then it's likely knife angle. Try making an 
OCT block and change the angle a little at time until you achieve complete, 
even, continuous sections. Then try it on your sample.

I'm not a neurohistotechnician but I'm sure there are some out there who can 
help with your LFB staining problem. I think a thickness of approx. 8-10 
microns are recommended for sections for this stain. You might be seeing 
variability due to the thin/thick issues you're having, not due to any staining 
problem.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: DIF tissue in GLUT

[Johnson, Teri]

Anne,

Tissue that has been fixed in glutaraldehyde has very, VERY bright 
autofluorescence. Unless there is some way to minimize this (none that I'm 
aware of), your immunofluorescence will be impossible to read over the 
background signal.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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RE: [Histonet] Re: DIF tissue in GLUT

[Johnson, Teri]

William, thanks for the reminder, I had totally forgotten about using a 
counterstain to counteract AF in aldehyde fixed samples. We have some 
glutaraldehyde fixed cryosections of kidney I think I'll try this with. I can 
assure you, the autofluorescence in these samples is magnificent! I worry about 
using sodium borohydride because they are cryosections and I've heard the 
treatment can be a bit harsh on tissue.

Here's some additional information since we're on the subject: 
http://www.uhnresearch.ca/facilities/wcif/PDF/Autofluorescence.pdf

Kind regards,
Teri
-Original Message-
From: WILLIAM DESALVO [mailto:wdesalvo@hotmail.com]
Sent: Thursday, August 27, 2009 12:51 PM
To: Johnson, Teri; histonet
Subject: RE: [Histonet] Re: DIF tissue in GLUT

I suggest you use a counterstain for your IF to reduce the autofluoresence. 
Evans Blue - the product can be used as a counterstain in immunohistochemistry 
when using FITC. After staining for immunofluorescence, dip sections in a 0.1% 
(w/v) in water solution of Evans Blue for 5-10 minutes. Rinse well in fresh PBS 
or water before coverslipping. Reference: Immunocytochemistry, Theory and 
Practice, p. 82 (1988). Purchase from Sigma-Aldrich.

William DeSalvo, B.S., HTL(ASCP)





 From: t...@stowers.org
 To: histonet@lists.utsouthwestern.edu
 Date: Thu, 27 Aug 2009 12:18:47 -0500
 Subject: [Histonet] Re: DIF tissue in GLUT

 Anne,

 Tissue that has been fixed in glutaraldehyde has very, VERY bright 
 autofluorescence. Unless there is some way to minimize this (none that I'm 
 aware of), your immunofluorescence will be impossible to read over the 
 background signal.

 Teri Johnson, HT(ASCP)QIHC
 Managing Director Histology Facility
 Stowers Institute for Medical Research
 1000 E. 50th St.
 Kansas City, MO 64110


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[Histonet] Re: Human samples in university research lab

[Johnson, Teri]

Denise,

You will need to contact your Institute Biosafety Committee (IBC) before you 
can start working on human samples in your lab. They should have everything 
you'll need to do in order to become compliant with federal regulations 
involving the use of human materials. It takes a ton of paperwork and 
preparation on the part of the collaborator and the university. In addition, 
anybody handling the material will need to take training so they are aware of 
the ethical and biohazardous issues involved.

Good luck!


Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110



Message: 5
Date: Mon, 10 Aug 2009 09:42:52 -0400
From: Denise Crowley dencr...@mit.edu
Subject: [Histonet] Human samples in university research lab
To: histonet@lists.utsouthwestern.edu
Message-ID: b5bfe03f-b706-4773-a776-e63d74ced...@mit.edu
Content-Type: text/plain;   charset=US-ASCII;   delsp=yes;  
format=flowed

Hi all,

We are a research core facility processing animal tissues for cancer
research.  We have been approached by a collaborator about bringing
human sample to us for processing, sectioning, and routine HE
staining, for use in research, not diagnosis.  In the past, we have
always encouraged these folks to have their slides cut at the
clinical facility which is supplying the tissue.  But we are getting
more and more of these requests and I need to think about the changes
we would need to make in our protocols, both in the processing
schedules and safety issues.  And I cannot even imagine the legal
issues involved in transporting patient samples and informed
consent.  Can anyone give me some guidance here?
Thanks,

Denise Crowley
Facility Manager Histology
David H. Koch Institute for Integrative Cancer Research Massachusetts Institute 
of Technology 40 Ames St. E17-427 Cambridge MA  02139 617-258-8183 
dencr...@mit.edu


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[Histonet] Re: double immunostaining with antigen retrieval and no antigen retrieval

[Johnson, Teri]

Cathy,

You absolutely can do this, but you have to use a permanent, stable end product 
on the first antibody staining. This is easy to do if you use DAB as the first 
staining technique chromogen. The antigen retrieval will probably remove all 
antibody-antigen bonds (elution) from your first IHC reaction, so having a 
permanent label on the first one is critical.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: IHC on frozens

[Johnson, Teri]

In response to this thread:

snip
Kimberly:
Your question has a two parts answer:
1- it cannot be done because the sections will peel off, but most importantly
2- it is not necessary since HIER was developed to undo the cross linkage 
produced by the NBF fixation, and the tissues used for?FS are not fixed. ren? 
J.?

--- On Thu, 7/2/09, Kimberly Tuttle ktut...@umm.edu wrote:


From: Kimberly Tuttle ktut...@umm.edu
Subject: [Histonet] IHC on frozens
To: histonet histonet@lists.utsouthwestern.edu
Date: Thursday, July 2, 2009, 12:55 PM


Can you do heat retrieval on frozens?

Kimberly C. Tuttle? HT (ASCP)
Pathology Biorepository and Research Core
University of Maryland
Room NBW58, UMMC
22 S. Greene St
Baltimore, MD 21201
(410) 328-5524
(410) 328-5508 fax
Please consider the environment before printing this e-mail.
snip

I disagree that you cannot do HIER on frozen sections. We do them all the time. 
All our samples are fixed in formalin, and then cryoprotected in sucrose prior 
to freezing them, so providing an antigen retrieval step usually produces good 
IHC results in frozen sections. We section and dry the slides, then use citrate 
buffer pH 6.0 in the microwave at 60 degrees C for 10 minutes, cool 10 minutes, 
then rinse and continue with IHC protocol. Use Plus slides and keep the 
solution under boiling temperature and you should be fine. For things like 
brain or bone, you might want to use lower temperatures for longer period of 
time instead of high temps for less time.

It can even work in fresh-frozen samples which are sectioned and then fixed 
prior to immunostaining. Some great information on this is included in this 
article: Yamashita and Okada, Application of Heat-induced Antigen Retrieval to 
Aldehyde-fixed Fresh Frozen Sections, JHC Vol 53(11): 1421-1432, 2005. A big 
thanks to Gayle Callis for the heads up on this paper!

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: Peroxidase block again....

[Johnson, Teri]

Gene,

Why would you want to eliminate peroxidase blocking as part of your 
immunostaining procedure? It's cheap, it's quick, it's easy, and if you think 
it interferes with the ability of the primary antibody to bind with the 
antigen, you add it as a step after application of the primary antibody. Simply 
use aqueous hydrogen peroxide instead of methanolic routinely, and you won't 
have to worry about the potential of methanol to interfere with the 
immunoreactivity of the tissues for CD markers.

You absolutely can leave it out and learn to ignore the positive staining of 
the RBCs and other tissues, and control what's labeled endogenous staining by 
including a negative control. Frankly, unless there is a scientific reason to 
have it included (i.e. looking for antigen positivity (demonstrated by a 
different enzyme) in cells/tissues which contain endogenous peroxidase), it's 
best to get rid of it.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] 2009 MO Society for Histotechnology Symposium

[Johnson, Teri]

Mark your calendar for this years MO Society for Histotechnology Symposium to 
be held at the beautiful Lake of the Ozarks, Port Arrowhead Resort on May 
29-30, with a vendor reception and MSH Quarterly Business Meeting on Thursday 
evening, May 28.

Our program is as follows:

Thursday, May 28:
6:00 pm - 8:00 pm - Registration and Exhibit Hall Open
8:00 pm - Enjoy the Portside Lounge with friends or attend the MSH Quarterly 
Business Meeting

Friday, May 29:
7:00 - 7:45 am - Registration
7:45 am - Welcome - Amanda Kelley, MSH President
8:00 - 9:30 - Brave New World: Introduction to Molecular Pathology, Part 1 - 
Dr. Thomas Haas, DO, FASCP
9:30 - 10:00 - Exhibit Hall Open
10:00 - 11:15 - Brave New World: Introduction to Molecular Pathology, Part 2 - 
Dr. Thomas Haas, DO, FASCP
11:30 - 12:30 - Green Histology - Laurence Patton BS, HT(ASCP)

12:30 - 1:15 - Lunch on your own

1:30 - 5:00
Workshop 1 - Muscle Biopsies: Gross Room to Reading Room - Konnie Zeitner, 
HT(ASCP)HTL SLS
Workshop 2 - The Histology Workout: Getting Lean in the Lab - Christine 
Charlie Dorner, HT(ASCP)QIHC
Workshop 3 - Methyl Methacrylate - Why and How? - Jack Ratliff, BA

6:00 - 8:00
Tropic Island Dinner Cruise (Cash Bar)

Saturday, May 30
7:00 - 7:45 am - Registration
7:45 - Welcome - JP Rey, MSH Vice-President
8:00 - 8:45 - Mohs at a Glance, The Benefits of Moh's Surgery - Gina Rodriguez, 
HT(ASCP)
8:45 - 10:00 - Where We are and Where we have been with Tissue Processing, Part 
1 - Christine Charlie Dorner, HT(ASCP)QIHC
10:00 - 10:30 - Exhibit Hall Open
10:30 - 12:00 - Where We are and Where we have been with Tissue Processing, 
Part 2 - Christine Charlie Dorner, HT(ASCP)QIHC

12:00 - 1:30 - MSH Awards lunch with Exhibitors

1:30 - 4:30
Workshop 4 - The Clinical, Technical, and Financial Benefits of Multi-antigen 
Immunostaining (MAIS) Procedures - Joseph D. Myers, MS, CT(ASCP)
Workshop 5 - Tissue Identification for the Histotechnologist - Dr. Thomas Haas, 
DO, FASCP
Workshop 6 - The What, When, Where, and How of Disaster Preparedness - Sylvia 
Jackson Casey, HT(ASCP), MA/MA

All workshops are CEU approved by NSH.

For complete information and a pdf copy of the brochure, visit 
www.missouri-histo.org or email Amanda Kelley kelleypa...@charter.net or JP Rey 
jp10...@hotmail.com

Hotel reservation deadline is April 27, 2009, and the toll free number is 
1-800-532-3575. Let them know you're attending the MSH symposium for a reduced 
room rate.

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[Histonet] Re: Strange Circles in IHC slides

[Johnson, Teri]

The circles could be either from air bubbles present when you mount the slides 
onto the coverplates, or they could have arisen after exposure to H2O2 as part 
of the staining procedure. Doing the H2O2 off instrument in a coplin jar and 
then rinsing and mounting them in the coverplates will help keep that from 
happening.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: IF

[Johnson, Teri]

Vanessa,

The answer of how long is suitable, the answer is it depends.

Routinely, many will air dry over night, even over a fan, in order to achieve 
the best adherence to the glass slide. We have found that we lose 
immunoreactivity with some antigens if we let it go that long. Our routine has 
now changed to 2 hour drying time. We tried 1 hour but the researchers had 
trouble with the tissues (especially neural tissue) trying to lift off during 
the staining procedure.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: wondering if anyone else has had this problem?

[Johnson, Teri]

Anita, I have seen this when using plastic embedding molds. The molds 
themselves hold an electric charge. It's annoying when your samples all migrate 
to the edges.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: processing v-e-r-y tiny samples

[Johnson, Teri]

Andi,

We process E7.0 mouse embryos and have problems sometimes because they are so 
very tiny and fragile. We've wrapped them and sometimes (but not all the time) 
had them break and flatten. Usually we use the histoscreen cassettes. They 
still will sometimes break apart using these but we have our best luck using 
them. You might find, though, that even with the mesh, the diameter of the 
holes may be big enough to let the sample pass through if the gut samples are 
as small as you say.

We tried the cell saver mesh inserts and they were a disaster. Lost the sample 
in them. I won't use them for our embryo work.

As for the histogel or agar, we've had difficulty using either in our paraffin 
processing. Recent histonet emails reveal other folks having a problem with the 
stuff randomly getting hard and brittle, like plastic and ruining the sample 
for sectioning. We may have 4 or 5 blocks, all go through the processor 
together, and one might have the problem of turning brittle. According to the 
other emails, Richard Allan folks hadn't any clue as to what was happening or 
how to fix it. And I'm clueless as well. Wish I knew how to consistently 
process things without this happening. Pre-embedding in a matrix like this 
would be so very helpful to allowing us to manipulate and properly orient tiny, 
fragile samples.

Good luck, and let us know what you come up with!

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: Bad sections

[Johnson, Teri]

Paula,

Try changing your microtome blade. Get as many samples as you can from the 
companies and try them all out. We see a difference in sectioning paraffin 
blocks just by changing blades. You'll find something that works well for your 
paraffin and room ambient conditions. If your room is warm, that may also 
contribute to compression. Ice down (or spray) the knife edge and see if the 
colder blade also helps.

Also, get rid of the Downy, you don't need it unless you need to soften some 
hard samples. Ice water should chill and rehydrate the blocks just fine. If you 
need more than that routinely, you are overprocessing your tissues.

Good luck!

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: GFP antibody

[Johnson, Teri]

We have had success staining murine paraffin sections using Rockland goat 
anti-GFP antibody.

You'll need to optimize it to your laboratory conditions trying with either 
antigen retrieval (citrate buffer pH 6.0) or Proteinase K. Make sure you have 
tissue with known positive GFP expression and a wild type (negative) animal for 
controls.

As for whether processing destroys the GFP in the sample, it will render the 
protein non-fluorescent. Fixation alone can do that. But the protein should 
still be there, even if it is not glowing.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: Ideal fixation of mouse prostate

[Johnson, Teri]

Vanessa,

Overnight fixation should not be a problem for these samples depending on what 
will be done to them at a later time. For samples needing studies for mRNA or 
some IHC markers, you want to avoid prolonged exposure to formalin fixation. 
You'll need to optimize your fixation based on those needs. If it is for HE or 
routine special stains only, overnight is fine.

Regarding leaving in 25% alcohol, that also shouldn't be a problem but if it 
isn't necessary, why do it? We routinely fix our samples overnight, and then 
dehydrate to 70% alcohol where they may have to sit until we can get them on 
the processor if it is already running another program. Our tissue processor 
starts in 70% alcohol. We have not experienced any issues with having tissue 
samples too brittle from sitting in this alcoholic solution.

For paraffin processing, you should only need 10-20 minutes per station on the 
VIP processor. You should avoid extended times in absolute alcohol and xylenes 
because your samples will get overly brittle. We have one xylene station on our 
processor, followed by two stations of Clear-Rite 3. We use the one station of 
xylene because it is more tolerant to water than the Clear-Rite 3. There is 
generally enough humidity around here that it may affect our processor 
solutions. The Clear-Rite 3 completes the clearing step and is a better agent 
(in my opinion) for mouse tissues than using 3 changes of xylene. Others on 
this list use xylene routinely with no trouble.


Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: Processing mouse seminal vesicles

[Johnson, Teri]

Kathleen,

Before changing your entire processing set-up, try decreasing the time in your 
alcohols and xylene to 10-15 minutes per station. Mouse seminal vesicles are 
very small and you don't need such rigorous dehydration and clearing to get 
proper processing.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: in utero mouse embryo

[Johnson, Teri]

Helen, you don't state the age of the embryo or what you plan to do with it 
afterwards.

I would recommend dissecting them out of the uterus and fixing them. 
Penetration/fixation will be greatly improved by doing this.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Histonet alias

[Johnson, Teri]

It is not uncommon for many/most online forums to have users create screennames 
that they use for their online moniker. You never really know who they are so 
everybody, unless they want to divulge their identity, stays anonymous. How can 
we really know all of the registered names on the histonet are real anyway? And 
why would it matter?

The value of the histonet is information. There are opinions for and against 
equipment and particular methods. I am not likely to discount good information 
just because it was given by a company rep, pathologist, med tech, hairdresser, 
or Mr. Anonymous. I argue that anonymity could be the best opportunity to give 
the most honest answer to a question posted to the board. It is up to each one 
of us to evaluate those opinions, take what is important, and leave the rest 
regardless of who posted it.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: LacZ Staining of Tumors

[Johnson, Teri]

Melissa wrote:

I am looking for a protocol to fix and stain in paraffin embedded 
subcutaneously grown mouse tumors for LacZ.  Can anyone help?  M.

Far as I know, one cannot stain paraffin embedded material with x-gal. We will 
do either whole mount X-gal staining, post-fix, and then paraffin process and 
section, or we will fix, sucrose cryoprotect, freeze and then cryosection to do 
the X-gal staining. Send me an email if you want to have our protocols for 
doing either.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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[Histonet] Re: PLS Help with IHC protocol

[Johnson, Teri]

Kimberly, you asked:

Step 10: Add antibody at 10ug/ml in PBS + 0.1%BSA + 1:10 volume CAS block

Can someone break this down for me in simple instructions?
And
Its 1.1mg/ml

Another question: they are asking that I add glycerol to the antibody to 
prevent freezing. Wont that change my concentration?

Some antibody companies request you add equal parts of glycerol to the antibody 
stock to prevent freeze/thaw cycles which is not good for antibody integrity. 
If you add equal parts of glycerol, your concentration would now be .55 mg/ml.  
In order to get ug/ml, you will need to convert .55 mg/ml to ug/ml. You do that 
by multiplying by 1000, therefore .55 mg/ml = 550 ug/ml. In order to get 10 
ug/ml concentration, you would divide 550 by 10, and you get 55. So you would 
use a 1:55 dilution of antibody/glycerol stock. The PBS/BSA/CAS block is a 
stock solution which you will then use as the diluent for the antibody. Some 
folks make it up using commercial antibody diluent. Some folks use whatever 
they use for the normal serum block (say 5-10% serum from the species source 
the secondary antibody is made in, i.e. normal goat serum if your secondary 
antibody is goat anti-rabbit). There is no one best diluent for antibodies. If 
you want to use the PBS/BSA/CAS block, go ahead. But you don't have to if you 
want to simplify things.

Teri Johnson, HT(ASCP)QIHC
Managing Director Histology Facility
Stowers Institute for Medical Research
1000 E. 50th St.
Kansas City, MO 64110


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