[Histonet] Histonet Digest, Vol 214, Issue 14

Tue Sep 28 06:51:15 CDT 2021

LABORATORY SUPERVISOR POSITION AT DUKE UNIVERSITY MEDICAL CENTER
DURHAM NC

We have an opening for a Laboratory Supervisor position in the Division of Anatomic Pathology and Digital Analytics.  The position requires a BS degree in biological science, certification as HTL or HT, and 5 years of Histology experience.  Experience preferred but not required includes supervisory, IHC, FISH, or digital pathology.  Contact Kathryn.perkinson at duke.edu.

Thank you
Kathy

Kathryn R. Perkinson, BS, HTL(ASCP), CSSGB
Manager, Division of Anatomic Pathology and Digital Analytics
Box 3712
Room 4710 Duke South Clinic Building, Yellow Zone
Duke University Health System
Durham, NC 27710
Phone: 919-684-5822

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-----Original Message-----
From: histonet-request at lists.utsouthwestern.edu <histonet-request at lists.utsouthwestern.edu> 
Sent: Friday, September 24, 2021 1:00 PM
To: histonet at lists.utsouthwestern.edu
Subject: Histonet Digest, Vol 214, Issue 14

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Today's Topics:

   1. Re: Multiple H&E stainer maintenance (Bacon, Charles)
   2. Jones' Methenamine Silver Stain for Basement Membranes of
      Kidney - Issues and Questions (Hood, Jordan)
   3. Bone marrow clot IHC tissue sections washing
      (Martha Ward-Pathology)
   4. Re: Bone marrow clot IHC tissue sections washing (Regan Fulton)
   5. Re: Jones' Methenamine Silver Stain for Basement Membranes of
      Kidney - Issues and Questions (Bryan Llewellyn)
   6. Re: Jones' Methenamine Silver Stain for Basement Membranes of
      Kidney - Issues and Questions (Tony Henwood (SCHN))
   7. Re: Jones' Methenamine Silver Stain for Basement Membranes of
      Kidney - Issues and Questions (Colleen Forster)

----------------------------------------------------------------------

Message: 1
Date: Thu, 23 Sep 2021 17:17:42 +0000
From: "Bacon, Charles" <Charles.Bacon at baystatehealth.org>
To: "Hanson, Leslie I :LLS Lab" <LIHANSON at lhs.org>,
	"'histonet at lists.utsouthwestern.edu'"
	<histonet at lists.utsouthwestern.edu>
Subject: Re: [Histonet] Multiple H&E stainer maintenance
Message-ID: <2211e9fd3b204d3790182cdbf0026398 at ZXSWEXCHMXPR06.bhs.org>
Content-Type: text/plain; charset="iso-8859-1"

Hi Leslie,

I have attached our current log. You can augment it based off the solutions you are using and how many shifts you have. We have two stainers and each has their own log. Our stainers are connected to running water thus the drain lines that need cleaning. If you have any questions reach out anytime. Good luck!

Best,

Chuck Bacon, HTL(ASCP)CM
Supervisor Histology
Baystate Medical Center
361 Whitney Ave., Holyoke, MA 01040
Telephone: 413-322-4786? Fax: 413-322-4790 Charles.Bacon at baystatehealth.org

-----Original Message-----
From: Hanson, Leslie I :LLS Lab [mailto:LIHANSON at lhs.org]
Sent: Wednesday, September 22, 2021 2:29 PM
To: 'histonet at lists.utsouthwestern.edu'
Subject: [Histonet] Multiple H&E stainer maintenance

Hi all,

We recently acquired a second H&E stainer and are working up a maintenance log. Looking for input from others who have already tackled this issue!

  *   Do you have a log for EACH instrument or one combined log?
  *   Do you have a space for listing reagent lots?
  *   What is your rotation schedule for changing reagents?
  *   Other tips and tricks are always appreciated!

Thanks!

Leslie Hanson, HT(ASCP)
Tech Specialist
IHC / Pathology
Phone: (503)944-7923
lihanson at lhs.org<mailto:lihanson at lhs.org>

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------------------------------

Message: 2
Date: Thu, 23 Sep 2021 19:49:52 +0000
From: "Hood, Jordan" <jordhood at med.umich.edu>
To: "'histonet at lists.utsouthwestern.edu'"
	<histonet at lists.utsouthwestern.edu>
Subject: [Histonet] Jones' Methenamine Silver Stain for Basement
	Membranes of Kidney - Issues and Questions
Message-ID: <48bd31f00b074079baca10d963e1bc2f at med.umich.edu>
Content-Type: text/plain; charset="iso-8859-1"

Hello,

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

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

Sections: Paraffin, 2 microns

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

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

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

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

In my first test, I used five test slides (extra slides that we cut from the same blocks that were submitted to us). I deparaffinized them in coplin jars (moving them with plastic forceps) and hydrated them to deionized water. I transferred the slides to a glass slide rack that holds ten slides on their sides, added five blank slides that were rinsed in deionized water (so that the displacement of reagents would be equivalent to when we stain our ten "real" slides after testing is complete), and completed Step 2. I don't recall exactly how long the glass container of silver solution and the glass container of deionized water had been heating up in the water bath, but I would estimate ~15-30 minutes. The thermometer said that the water in the bath (not inside the containers) reached ~60-65 degrees Celsius. The silver solution was clear and colorless when I made it up, but by the time I put the slides into the warm silver solution, the solution was beginning to turn a light brown color (though it was still clear and I did not see any precipitate floating around). I removed the metal handle of the glass slide rack after the rack was transferred into the silver solution, but the metal handle did dip into the silver solution briefly. At some point, I noticed precipitate floating around of the surface of the silver solution. After ~80 minutes, I used plastic forceps to remove one test slide from the warm silver solution, dipped it several times into the warm deionized water to rinse it, and wiped off the back of the slide with gauze. The amount of precipitate was so extreme that the gauze did nearly nothing. I showed the slide to one of our pathologists and they could hardly see beyond the precipitate, but said that they couldn't see any staining of the structure that they were looking for (I forget exactly what it was, but I know it's supposed to turn black).

In my second test (to see if the metal holder was the problem) that I performed immediately after the first test, I used one test slide. I deparaffinized it in the same coplin jars as before (moving it with plastic forceps) and hydrated it to deionized water. I used new glass containers for the periodic acid and deionized water rinse in Step 2, for making the silver solution in Step 3, and for the warm silver solution and warm deionized water in Step 4. I used plastic forceps to move the slide into the periodic acid, and propped it up in the container so that no glass rack or metal handle was used at all. I used plastic forceps to transfer the slide to the deionized water rinse, and dunked it several times and swished the slide around a bit. I used plastic forceps to transfer the slide into the warm(-ish) silver solution and propped it up against the side again. After approximately 20 minutes, I saw precipitate floating around, and I used plastic forceps to remove the slide from the silver solution. I dipped the slide into the warm(-ish) deionized water several times, and saw that the precipitate was again covering the slide and the tissue so I stopped there for the day.

We purchased all of the reagents listed in the above procedure from Rowley Biochemical (except for the Glacial Acetic Acid mentioned in Step 7, but I didn't even get that far).

Questions:

1. Could this indicate that the acid-washing was not done correctly? I made up a ~1% Hydrochloric Acid solution (with deionized water) and filled a plastic bin with the solution (I rinsed the bin with deionized water first). I then submerged all glassware (in several batches) for at least five minutes, then rinsed well with deionized water (not by filling a bin - I just used the hose of deionized water in our lab sink and poured it over the glassware) and left them to air-dry overnight.

2. Are using acid-washed glassware and avoiding metal even necessary precautions after the sodium thiosulphate in Step 6? I read that sodium thiosulphate "stops the reaction," and the procedure stops specifically saying to use deionized water after Step 6 and starts saying to use just "water" or "tap water." My lab refers to our waters as either "tap" or "deionized," so I'm assuming that using my deionized water is fine when the procedure calls for "distilled" or "dechlorinated."

I don't even know enough to ask more questions, but I'm sure many more will arise after I test the stain again next week, so I welcome any and all advice about silver stains, acid-cleaning glassware, and literally anything else...

Thank you!!!

Jordan H.
University of Michigan
Ann Arbor, MI
**********************************************************
Electronic Mail is not secure, may not be read every day, and should not be used for urgent or sensitive issues 

------------------------------

Message: 3
Date: Thu, 23 Sep 2021 19:55:42 +0000
From: Martha Ward-Pathology <mward at wakehealth.edu>
To: "histonet at lists.utsouthwestern.edu"
	<histonet at lists.utsouthwestern.edu>
Subject: [Histonet] Bone marrow clot IHC tissue sections washing
Message-ID:
	<B2CECB1B6665A4479056478F6DE3C4AB01ED75B093 at exchdb7.medctr.ad.wfubmc.edu>

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

It was brought to my attention that we had significant washing on  3 of 8 bone marrow clot sections the other day; this is not the first time so we would like to get to the bottom of this.   We use positively charged slides and all 8 cases were cut and run the same morning but allowed to air dry and then bake at 60C for 20 minutes before being run on our Bond 3 stainer.   Has anyone out there experienced this type of problem and if so, what were your solutions?    The repeat of the 3 cases today showed similar washing of tissue.

This hasn't just started but has occurred periodically but the pathologists have tried to live with it and usually we can finally get enough tissue to stay on after 1-2 attempts.   Suggestions include cutting and drying the slides overnight and/or going to a gelatinated  slide versus a sialylated slide.   We have been using this particular brand of positive charged slide with good results for several years and rarely have issues with other tissue types unless they are particularly bloody.

Thoughts or suggestions are greatly appreciated.

Martha Ward MT(ASCP) QIHC
Atrium Health Wake Forest Baptist

------------------------------

Message: 4
Date: Thu, 23 Sep 2021 13:16:27 -0700
From: Regan Fulton <regan.fulton at gmail.com>
To: Martha Ward-Pathology <mward at wakehealth.edu>,
	Histonet at lists.utsouthwestern.edu
Subject: Re: [Histonet] Bone marrow clot IHC tissue sections washing
Message-ID:
	<CAMrtFXQSWEQOHEsC=tU4VN9aYxbfAW6npvuWEh5KL3PiHgernQ at mail.gmail.com>
Content-Type: text/plain; charset="UTF-8"

Martha,

We reported our study of 15 brands of adhesive slides for "wash off" and found little difference among the different slides when well-fixed cell culture material was examined.
On the other hand, poorly-fixed breast cancer tissues did appear to adhere more strongly to some slides than others (TOMO being among the best).
Additional factors need to be considered, though, and I note that your baking procedure is different from what is recommended by many slide vendors.
In general, baking at 60-65 deg C for 1 hour is said to be optimal, although we did not examine that parameter specifically.

Please see our poster at https://urldefense.com/v3/__https://www.arrayscience.com/publications*Posters__;Iw!!OToaGQ!6kZFsSGmGAqtHSUCXGvBt6QPRHlQc7P4geZI9L6HlrVmY_FPmCDYnGHGszj4cR01HGjCd-A$ 

Best regards,

Regan
Regan Fulton, M.D., Ph.D.
CEO and Co-Founder
Array Science, LLC
475 Gate 5 Road, #100
Sausalito, CA 94965
(415) 577-7360
email: fulton at arrayscience.com

https://urldefense.com/v3/__http://www.arrayscience.com__;!!OToaGQ!6kZFsSGmGAqtHSUCXGvBt6QPRHlQc7P4geZI9L6HlrVmY_FPmCDYnGHGszj4cR01q5l4Oys$ 

On Thu, Sep 23, 2021 at 1:02 PM Martha Ward-Pathology via Histonet < histonet at lists.utsouthwestern.edu> wrote:

> It was brought to my attention that we had significant washing on  3 
> of 8 bone marrow clot sections the other day; this is not the first time so we
> would like to get to the bottom of this.   We use positively charged slides
> and all 8 cases were cut and run the same morning but allowed to air 
> dry and then bake at 60C for 20 minutes before being run on our Bond 3
> stainer.   Has anyone out there experienced this type of problem and if so,
> what were your solutions?    The repeat of the 3 cases today showed similar
> washing of tissue.
>
> This hasn't just started but has occurred periodically but the 
> pathologists have tried to live with it and usually we can finally get
> enough tissue to stay on after 1-2 attempts.   Suggestions include cutting
> and drying the slides overnight and/or going to a gelatinated  slide versus
> a sialylated slide.   We have been using this particular brand of positive
> charged slide with good results for several years and rarely have 
> issues with other tissue types unless they are particularly bloody.
>
> Thoughts or suggestions are greatly appreciated.
>
>
> Martha Ward MT(ASCP) QIHC
> Atrium Health Wake Forest Baptist
>
>
>
> _______________________________________________
> Histonet mailing list
> Histonet at lists.utsouthwestern.edu
> https://urldefense.com/v3/__http://lists.utsouthwestern.edu/mailman/li
> stinfo/histonet__;!!OToaGQ!6kZFsSGmGAqtHSUCXGvBt6QPRHlQc7P4geZI9L6HlrV
> mY_FPmCDYnGHGszj4cR01z2-BarQ$
>

------------------------------

Message: 5
Date: Thu, 23 Sep 2021 14:47:24 -0700
From: Bryan Llewellyn <llewllew at shaw.ca>
To: Jordan <jordhood at med.umich.edu>,	Histonet
	<histonet at lists.utsouthwestern.edu>
Subject: Re: [Histonet] Jones' Methenamine Silver Stain for Basement
	Membranes of Kidney - Issues and Questions
Message-ID: <9e1fe5b8-be5e-ae7f-ca29-e57dac959071 at shaw.ca>
Content-Type: text/plain; charset=UTF-8; format=flowed

Hi,
Try the method given in StainsFile at:
https://urldefense.com/v3/__http://stainsfile.info/stain/metallic/jones.htm__;!!OToaGQ!6kZFsSGmGAqtHSUCXGvBt6QPRHlQc7P4geZI9L6HlrVmY_FPmCDYnGHGszj4cR01CJSb57Q$ 

Bryan Llewellyn

Hood, Jordan via Histonet wrote:
> Hello,
> 
> I'm new to histology (and new to histonet), and I work in a small histology lab specializing in animal tissues that receives requests/submissions from researchers. I tried (and failed) to perform a Jones' Methenamine Silver stain on a client's submission of pig kidneys (formalin-fixed, paraffin-embedded, cut at 2.5 microns), and I need some help troubleshooting this stain since my co-workers are stumped, too.  I used the following procedure from Rowley Biochemical:
> 
> 
> ~~~~~
> "Fixation: 10% Buffered Neutral Formalin (F-113) or Bouin's Solution 
> (F-40) or Zenker's (F-155)
> 
> Sections: Paraffin, 2 microns
> 
> Procedure: Acid washed glassware must be used!!!!
> 1. Deparaffinize and hydrate to distilled water.
> 2. Oxidize in Periodic Acid 0.5% (F-396-1) 11 minutes. Wash in chloride-free water.
> 3. Prepare Methenamine Silver solution by mixing: 42.5 ml Methenamine 3% (F-396-2), 2.5 ml Silver Nitrate, 5% (F-396-3) and 12.0 ml Borate Buffer, pH 8.2 (F-396-4).
> 4. Place slides in the solution and the entire jar in a water bath at 70?C for approx. 60-75 minutes. Check under microscope when slides appear medium brown microscopically. Every 10 minutes, once the medium brown color has been established, rinse a slide in 70?C, chloride free water and check under a microscope. Rinse again in hot water and return to the hot staining solution. As the staining time approaches the end point, check the slides, as above, every 1-2 minutes. The entire procedure must be performed quickly to prevent an uneven staining of the tissues. The slides should exhibit a brownish- yellow background, intense black reticulum fibers, and black basement membranes. If the slides become oversaturated, i.e. too black, destain in a dilute Potassium Ferricyanide Solution (F-396-11) for one or two dips.
> 5. Rinse well in distilled water. Tone in Gold Chloride 0.2% (F-396-5), 1 minute. If sections are overtoned place in Sodium Metabisulfite, 3% (F-396-12) for 1-3 minutes. Rinse well in distilled water.
> 6. Sodium Thiosulfate 3% (F-396-9), 1-2 miutes. Wash in running tap water, 10 minutes. Rinse well in distilled water.
> 7. Stain in Harris' Hematoxylin (F-396-6) containing 2-4ml of Glacial Acetic Acid per 100 ml for 5-15 minutes. Wash in water.
> 8. Differentiate in Acid Alcohol 1% (F-396-13) until the sections turn red.
> 9. Blue section in Ammonia Water, 0.3% (F-396-14). Wash thoroughly.
> 10. Counterstain in Eosin Y, 1%, Alcoholic Solution (F-396-7).
> 11. Dehydrate in 95% alcohol, absolute alcohol and clear in xylene 3 changes each. Mount.
> 
> Stain Results:
> Basement membranes, reticulum fibers: Black
> Nuclei: Blue
> Cytoplasm, collagen, connective tissue: Pink-orange
> 
> References: Jones, D.B., Amer.J.Path. 27:99 (1951). AFIP Manual of Histolocical Staining Methods, 3rd ed., Ed. L. Luna: NY: McGraw-Hill Publications, c. 1968, p. 97."
> ~~~~~
> 
> 
> It became apparent that something went wrong during Step 4 when the slides were in the glass container (not a coplin jar - we have ten slides that we need to stain so we're using a rectangular glass container that holds ten slides on their sides - it does require a metal handle to move, but the handle is flexible and easy to remove after the glass slide rack has been transferred between containers) of silver solution in the water bath because there was lots of precipitate on the slides and floating on the surface of the silver solution.
> 
> In my first test, I used five test slides (extra slides that we cut from the same blocks that were submitted to us). I deparaffinized them in coplin jars (moving them with plastic forceps) and hydrated them to deionized water. I transferred the slides to a glass slide rack that holds ten slides on their sides, added five blank slides that were rinsed in deionized water (so that the displacement of reagents would be equivalent to when we stain our ten "real" slides after testing is complete), and completed Step 2. I don't recall exactly how long the glass container of silver solution and the glass container of deionized water had been heating up in the water bath, but I would estimate ~15-30 minutes. The thermometer said that the water in the bath (not inside the containers) reached ~60-65 degrees Celsius. The silver solution was clear and colorless when I made it up, but by the time I put the slides into the warm silver solution, the solution was beginning to turn a light brown color (though it was still clear and I did not see any precipitate floating around). I removed the metal handle of the glass slide rack after the rack was transferred into the silver solution, but the metal handle did dip into the silver solution briefly. At some point, I noticed precipitate floating around of the surface of the silver solution. After ~80 minutes, I used plastic forceps to remove one test slide from the warm silver solution, dipped it several times into the warm deionized water to rinse it, and wiped off the back of the slide with gauze. The amount of precipitate was so extreme that the gauze did nearly nothing. I showed the slide to one of our pathologists and they could hardly see beyond the precipitate, but said that they couldn't see any staining of the structure that they were looking for (I forget exactly what it was, but I know it's supposed to turn black).
> 
> In my second test (to see if the metal holder was the problem) that I performed immediately after the first test, I used one test slide. I deparaffinized it in the same coplin jars as before (moving it with plastic forceps) and hydrated it to deionized water. I used new glass containers for the periodic acid and deionized water rinse in Step 2, for making the silver solution in Step 3, and for the warm silver solution and warm deionized water in Step 4. I used plastic forceps to move the slide into the periodic acid, and propped it up in the container so that no glass rack or metal handle was used at all. I used plastic forceps to transfer the slide to the deionized water rinse, and dunked it several times and swished the slide around a bit. I used plastic forceps to transfer the slide into the warm(-ish) silver solution and propped it up against the side again. After approximately 20 minutes, I saw precipitate floating around, and I used plastic forceps to remove the slide from the silver solution. I dipped the slide into the warm(-ish) deionized water several times, and saw that the precipitate was again covering the slide and the tissue so I stopped there for the day.
> 
> We purchased all of the reagents listed in the above procedure from Rowley Biochemical (except for the Glacial Acetic Acid mentioned in Step 7, but I didn't even get that far).
> 
> Questions:
> 
> 1. Could this indicate that the acid-washing was not done correctly? I made up a ~1% Hydrochloric Acid solution (with deionized water) and filled a plastic bin with the solution (I rinsed the bin with deionized water first). I then submerged all glassware (in several batches) for at least five minutes, then rinsed well with deionized water (not by filling a bin - I just used the hose of deionized water in our lab sink and poured it over the glassware) and left them to air-dry overnight.
> 
> 2. Are using acid-washed glassware and avoiding metal even necessary precautions after the sodium thiosulphate in Step 6? I read that sodium thiosulphate "stops the reaction," and the procedure stops specifically saying to use deionized water after Step 6 and starts saying to use just "water" or "tap water." My lab refers to our waters as either "tap" or "deionized," so I'm assuming that using my deionized water is fine when the procedure calls for "distilled" or "dechlorinated."
> 
> I don't even know enough to ask more questions, but I'm sure many more will arise after I test the stain again next week, so I welcome any and all advice about silver stains, acid-cleaning glassware, and literally anything else...
> 
> Thank you!!!
> 
> Jordan H.
> University of Michigan
> Ann Arbor, MI
> **********************************************************
> Electronic Mail is not secure, may not be read every day, and should 
> not be used for urgent or sensitive issues 
> _______________________________________________
> Histonet mailing list
> Histonet at lists.utsouthwestern.edu
> https://urldefense.com/v3/__http://lists.utsouthwestern.edu/mailman/li
> stinfo/histonet__;!!OToaGQ!6kZFsSGmGAqtHSUCXGvBt6QPRHlQc7P4geZI9L6HlrV
> mY_FPmCDYnGHGszj4cR01z2-BarQ$
> 

------------------------------

Message: 6
Date: Thu, 23 Sep 2021 23:13:40 +0000
From: "Tony Henwood (SCHN)" <tony.henwood at health.nsw.gov.au>
To: Bryan Llewellyn <llewllew at shaw.ca>, Jordan
	<jordhood at med.umich.edu>
Cc: "'histonet at lists.utsouthwestern.edu'"
	<histonet at lists.utsouthwestern.edu>
Subject: Re: [Histonet] Jones' Methenamine Silver Stain for Basement
	Membranes of Kidney - Issues and Questions
Message-ID:
	<5a34030c402f4f4b97dec87e4856d6f6 at SVDCMBX-MEX024.nswhealth.net>
Content-Type: text/plain; charset="utf-8"

I agree with Bryan,

The introduction of thiosemicarbazide before the silver step improves the staining immensely.

I would also look at the periodic acid. Is it too dilute, though 0.5% should work? I usually cover this by using a 1% solution for 20 minutes.

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

-----Original Message-----
From: Bryan Llewellyn via Histonet [mailto:histonet at lists.utsouthwestern.edu]
Sent: Friday, 24 September 2021 7:47 AM
To: Jordan <jordhood at med.umich.edu>; Histonet <histonet at lists.utsouthwestern.edu>
Subject: Re: [Histonet] Jones' Methenamine Silver Stain for Basement Membranes of Kidney - Issues and Questions

Hi,
Try the method given in StainsFile at:
https://urldefense.com/v3/__http://stainsfile.info/stain/metallic/jones.htm__;!!OToaGQ!6kZFsSGmGAqtHSUCXGvBt6QPRHlQc7P4geZI9L6HlrVmY_FPmCDYnGHGszj4cR01CJSb57Q$ 

Bryan Llewellyn

Hood, Jordan via Histonet wrote:
> Hello,
> 
> I'm new to histology (and new to histonet), and I work in a small histology lab specializing in animal tissues that receives requests/submissions from researchers. I tried (and failed) to perform a Jones' Methenamine Silver stain on a client's submission of pig kidneys (formalin-fixed, paraffin-embedded, cut at 2.5 microns), and I need some help troubleshooting this stain since my co-workers are stumped, too.  I used the following procedure from Rowley Biochemical:
> 
> 
> ~~~~~
> "Fixation: 10% Buffered Neutral Formalin (F-113) or Bouin's Solution
> (F-40) or Zenker's (F-155)
> 
> Sections: Paraffin, 2 microns
> 
> Procedure: Acid washed glassware must be used!!!!
> 1. Deparaffinize and hydrate to distilled water.
> 2. Oxidize in Periodic Acid 0.5% (F-396-1) 11 minutes. Wash in chloride-free water.
> 3. Prepare Methenamine Silver solution by mixing: 42.5 ml Methenamine 3% (F-396-2), 2.5 ml Silver Nitrate, 5% (F-396-3) and 12.0 ml Borate Buffer, pH 8.2 (F-396-4).
> 4. Place slides in the solution and the entire jar in a water bath at 70?C for approx. 60-75 minutes. Check under microscope when slides appear medium brown microscopically. Every 10 minutes, once the medium brown color has been established, rinse a slide in 70?C, chloride free water and check under a microscope. Rinse again in hot water and return to the hot staining solution. As the staining time approaches the end point, check the slides, as above, every 1-2 minutes. The entire procedure must be performed quickly to prevent an uneven staining of the tissues. The slides should exhibit a brownish- yellow background, intense black reticulum fibers, and black basement membranes. If the slides become oversaturated, i.e. too black, destain in a dilute Potassium Ferricyanide Solution (F-396-11) for one or two dips.
> 5. Rinse well in distilled water. Tone in Gold Chloride 0.2% (F-396-5), 1 minute. If sections are overtoned place in Sodium Metabisulfite, 3% (F-396-12) for 1-3 minutes. Rinse well in distilled water.
> 6. Sodium Thiosulfate 3% (F-396-9), 1-2 miutes. Wash in running tap water, 10 minutes. Rinse well in distilled water.
> 7. Stain in Harris' Hematoxylin (F-396-6) containing 2-4ml of Glacial Acetic Acid per 100 ml for 5-15 minutes. Wash in water.
> 8. Differentiate in Acid Alcohol 1% (F-396-13) until the sections turn red.
> 9. Blue section in Ammonia Water, 0.3% (F-396-14). Wash thoroughly.
> 10. Counterstain in Eosin Y, 1%, Alcoholic Solution (F-396-7).
> 11. Dehydrate in 95% alcohol, absolute alcohol and clear in xylene 3 changes each. Mount.
> 
> Stain Results:
> Basement membranes, reticulum fibers: Black
> Nuclei: Blue
> Cytoplasm, collagen, connective tissue: Pink-orange
> 
> References: Jones, D.B., Amer.J.Path. 27:99 (1951). AFIP Manual of Histolocical Staining Methods, 3rd ed., Ed. L. Luna: NY: McGraw-Hill Publications, c. 1968, p. 97."
> ~~~~~
> 
> 
> It became apparent that something went wrong during Step 4 when the slides were in the glass container (not a coplin jar - we have ten slides that we need to stain so we're using a rectangular glass container that holds ten slides on their sides - it does require a metal handle to move, but the handle is flexible and easy to remove after the glass slide rack has been transferred between containers) of silver solution in the water bath because there was lots of precipitate on the slides and floating on the surface of the silver solution.
> 
> In my first test, I used five test slides (extra slides that we cut from the same blocks that were submitted to us). I deparaffinized them in coplin jars (moving them with plastic forceps) and hydrated them to deionized water. I transferred the slides to a glass slide rack that holds ten slides on their sides, added five blank slides that were rinsed in deionized water (so that the displacement of reagents would be equivalent to when we stain our ten "real" slides after testing is complete), and completed Step 2. I don't recall exactly how long the glass container of silver solution and the glass container of deionized water had been heating up in the water bath, but I would estimate ~15-30 minutes. The thermometer said that the water in the bath (not inside the containers) reached ~60-65 degrees Celsius. The silver solution was clear and colorless when I made it up, but by the time I put the slides into the warm silver solution, the solution was beginning to turn a light brown color (though it was still clear and I did not see any precipitate floating around). I removed the metal handle of the glass slide rack after the rack was transferred into the silver solution, but the metal handle did dip into the silver solution briefly. At some point, I noticed precipitate floating around of the surface of the silver solution. After ~80 minutes, I used plastic forceps to remove one test slide from the warm silver solution, dipped it several times into the warm deionized water to rinse it, and wiped off the back of the slide with gauze. The amount of precipitate was so extreme that the gauze did nearly nothing. I showed the slide to one of our pathologists and they could hardly see beyond the precipitate, but said that they couldn't see any staining of the structure that they were looking for (I forget exactly what it was, but I know it's supposed to turn black).
> 
> In my second test (to see if the metal holder was the problem) that I performed immediately after the first test, I used one test slide. I deparaffinized it in the same coplin jars as before (moving it with plastic forceps) and hydrated it to deionized water. I used new glass containers for the periodic acid and deionized water rinse in Step 2, for making the silver solution in Step 3, and for the warm silver solution and warm deionized water in Step 4. I used plastic forceps to move the slide into the periodic acid, and propped it up in the container so that no glass rack or metal handle was used at all. I used plastic forceps to transfer the slide to the deionized water rinse, and dunked it several times and swished the slide around a bit. I used plastic forceps to transfer the slide into the warm(-ish) silver solution and propped it up against the side again. After approximately 20 minutes, I saw precipitate floating around, and I used plastic forceps to remove the slide from the silver solution. I dipped the slide into the warm(-ish) deionized water several times, and saw that the precipitate was again covering the slide and the tissue so I stopped there for the day.
> 
> We purchased all of the reagents listed in the above procedure from Rowley Biochemical (except for the Glacial Acetic Acid mentioned in Step 7, but I didn't even get that far).
> 
> Questions:
> 
> 1. Could this indicate that the acid-washing was not done correctly? I made up a ~1% Hydrochloric Acid solution (with deionized water) and filled a plastic bin with the solution (I rinsed the bin with deionized water first). I then submerged all glassware (in several batches) for at least five minutes, then rinsed well with deionized water (not by filling a bin - I just used the hose of deionized water in our lab sink and poured it over the glassware) and left them to air-dry overnight.
> 
> 2. Are using acid-washed glassware and avoiding metal even necessary precautions after the sodium thiosulphate in Step 6? I read that sodium thiosulphate "stops the reaction," and the procedure stops specifically saying to use deionized water after Step 6 and starts saying to use just "water" or "tap water." My lab refers to our waters as either "tap" or "deionized," so I'm assuming that using my deionized water is fine when the procedure calls for "distilled" or "dechlorinated."
> 
> I don't even know enough to ask more questions, but I'm sure many more will arise after I test the stain again next week, so I welcome any and all advice about silver stains, acid-cleaning glassware, and literally anything else...
> 
> Thank you!!!
> 
> Jordan H.
> University of Michigan
> Ann Arbor, MI
> **********************************************************
> Electronic Mail is not secure, may not be read every day, and should 
> not be used for urgent or sensitive issues 
> _______________________________________________
> Histonet mailing list
> Histonet at lists.utsouthwestern.edu
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> mY_FPmCDYnGHGszj4cR01z2-BarQ$
> 
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This message is intended for the addressee named and may contain confidential information. If you are not the intended recipient, please delete it and notify the sender.

Views expressed in this message are those of the individual sender, and are not necessarily the views of NSW Health or any of its entities.

------------------------------

Message: 7
Date: Thu, 23 Sep 2021 18:28:03 -0500
From: Colleen Forster <cforster at umn.edu>
To: "Tony Henwood (SCHN)" <tony.henwood at health.nsw.gov.au>
Cc: Bryan Llewellyn <llewllew at shaw.ca>, Jordan
	<jordhood at med.umich.edu>,	"histonet at lists.utsouthwestern.edu"
	<histonet at lists.utsouthwestern.edu>
Subject: Re: [Histonet] Jones' Methenamine Silver Stain for Basement
	Membranes of Kidney - Issues and Questions
Message-ID:
	<CAGW4+=pJZFYjmyg-+6fNC4y6MN9KogKnKCSWRLBCiW2cFOPhcw at mail.gmail.com>
Content-Type: text/plain; charset="UTF-8"

Make sure the periodic acid is made fresh EACH time you run  the stain.
That can also make a big difference in the stain quality.

Colleen Forster HT(ASCP)QIHC

On Thu, Sep 23, 2021 at 6:14 PM Tony Henwood (SCHN) via Histonet < histonet at lists.utsouthwestern.edu> wrote:

> I agree with Bryan,
>
> The introduction of thiosemicarbazide before the silver step improves 
> the staining immensely.
>
> I would also look at the periodic acid. Is it too dilute, though 0.5% 
> should work? I usually cover this by using a 1% solution for 20 minutes.
>
> Regards
> Tony Henwood JP, MSc, BAppSc, GradDipSysAnalys, CT(ASC), FFSc(RCPA) 
> Principal Scientist, the Children?s Hospital at Westmead Adjunct 
> Fellow, School of Medicine, University of Western Sydney
> Tel: 612 9845 3306
> Fax: 612 9845 3318
> Pathology Department
> the children's hospital at westmead
> Cnr Hawkesbury Road and Hainsworth Street, Westmead Locked Bag 4001, 
> Westmead NSW 2145, AUSTRALIA
>
>
> -----Original Message-----
> From: Bryan Llewellyn via Histonet [mailto:
> histonet at lists.utsouthwestern.edu]
> Sent: Friday, 24 September 2021 7:47 AM
> To: Jordan <jordhood at med.umich.edu>; Histonet < 
> histonet at lists.utsouthwestern.edu>
> Subject: Re: [Histonet] Jones' Methenamine Silver Stain for Basement 
> Membranes of Kidney - Issues and Questions
>
> Hi,
> Try the method given in StainsFile at:
> https://urldefense.com/v3/__http://stainsfile.info/stain/metallic/jone
> s.htm__;!!OToaGQ!6kZFsSGmGAqtHSUCXGvBt6QPRHlQc7P4geZI9L6HlrVmY_FPmCDYn
> GHGszj4cR01CJSb57Q$
>
> Bryan Llewellyn
>
>
> Hood, Jordan via Histonet wrote:
> > Hello,
> >
> > I'm new to histology (and new to histonet), and I work in a small
> histology lab specializing in animal tissues that receives 
> requests/submissions from researchers. I tried (and failed) to perform 
> a Jones' Methenamine Silver stain on a client's submission of pig 
> kidneys (formalin-fixed, paraffin-embedded, cut at 2.5 microns), and I 
> need some help troubleshooting this stain since my co-workers are 
> stumped, too.  I used the following procedure from Rowley Biochemical:
> >
> >
> > ~~~~~
> > "Fixation: 10% Buffered Neutral Formalin (F-113) or Bouin's Solution
> > (F-40) or Zenker's (F-155)
> >
> > Sections: Paraffin, 2 microns
> >
> > Procedure: Acid washed glassware must be used!!!!
> > 1. Deparaffinize and hydrate to distilled water.
> > 2. Oxidize in Periodic Acid 0.5% (F-396-1) 11 minutes. Wash in
> chloride-free water.
> > 3. Prepare Methenamine Silver solution by mixing: 42.5 ml 
> > Methenamine 3%
> (F-396-2), 2.5 ml Silver Nitrate, 5% (F-396-3) and 12.0 ml Borate 
> Buffer, pH 8.2 (F-396-4).
> > 4. Place slides in the solution and the entire jar in a water bath 
> > at
> 70?C for approx. 60-75 minutes. Check under microscope when slides 
> appear medium brown microscopically. Every 10 minutes, once the medium 
> brown color has been established, rinse a slide in 70?C, chloride free 
> water and check under a microscope. Rinse again in hot water and 
> return to the hot staining solution. As the staining time approaches 
> the end point, check the slides, as above, every 1-2 minutes. The 
> entire procedure must be performed quickly to prevent an uneven 
> staining of the tissues. The slides should exhibit a
> brownish- yellow background, intense black reticulum fibers, and black 
> basement membranes. If the slides become oversaturated, i.e. too 
> black, destain in a dilute Potassium Ferricyanide Solution (F-396-11) 
> for one or two dips.
> > 5. Rinse well in distilled water. Tone in Gold Chloride 0.2% 
> > (F-396-5),
> 1 minute. If sections are overtoned place in Sodium Metabisulfite, 3%
> (F-396-12) for 1-3 minutes. Rinse well in distilled water.
> > 6. Sodium Thiosulfate 3% (F-396-9), 1-2 miutes. Wash in running tap
> water, 10 minutes. Rinse well in distilled water.
> > 7. Stain in Harris' Hematoxylin (F-396-6) containing 2-4ml of 
> > Glacial
> Acetic Acid per 100 ml for 5-15 minutes. Wash in water.
> > 8. Differentiate in Acid Alcohol 1% (F-396-13) until the sections 
> > turn
> red.
> > 9. Blue section in Ammonia Water, 0.3% (F-396-14). Wash thoroughly.
> > 10. Counterstain in Eosin Y, 1%, Alcoholic Solution (F-396-7).
> > 11. Dehydrate in 95% alcohol, absolute alcohol and clear in xylene 3
> changes each. Mount.
> >
> > Stain Results:
> > Basement membranes, reticulum fibers: Black
> > Nuclei: Blue
> > Cytoplasm, collagen, connective tissue: Pink-orange
> >
> > References: Jones, D.B., Amer.J.Path. 27:99 (1951). AFIP Manual of
> Histolocical Staining Methods, 3rd ed., Ed. L. Luna: NY: McGraw-Hill 
> Publications, c. 1968, p. 97."
> > ~~~~~
> >
> >
> > It became apparent that something went wrong during Step 4 when the
> slides were in the glass container (not a coplin jar - we have ten 
> slides that we need to stain so we're using a rectangular glass 
> container that holds ten slides on their sides - it does require a 
> metal handle to move, but the handle is flexible and easy to remove 
> after the glass slide rack has been transferred between containers) of 
> silver solution in the water bath because there was lots of 
> precipitate on the slides and floating on the surface of the silver solution.
> >
> > In my first test, I used five test slides (extra slides that we cut 
> > from
> the same blocks that were submitted to us). I deparaffinized them in 
> coplin jars (moving them with plastic forceps) and hydrated them to 
> deionized water. I transferred the slides to a glass slide rack that 
> holds ten slides on their sides, added five blank slides that were 
> rinsed in deionized water (so that the displacement of reagents would 
> be equivalent to when we stain our ten "real" slides after testing is 
> complete), and completed Step 2. I don't recall exactly how long the 
> glass container of silver solution and the glass container of 
> deionized water had been heating up in the water bath, but I would 
> estimate ~15-30 minutes. The thermometer said that the water in the 
> bath (not inside the containers) reached ~60-65 degrees Celsius. The 
> silver solution was clear and colorless when I made it up, but by the 
> time I put the slides into the warm silver solution, the solution was 
> beginning to turn a light brown color (though it was still clear and I 
> did not see any precipitate floating around). I removed the metal 
> handle of the glass slide rack after the rack was transferred into the 
> silver solution, but the metal handle did dip into the silver solution 
> briefly. At some point, I noticed precipitate floating around of the 
> surface of the silver solution. After ~80 minutes, I used plastic 
> forceps to remove one test slide from the warm silver solution, dipped 
> it several times into the warm deionized water to rinse it, and wiped 
> off the back of the slide with gauze. The amount of precipitate was so 
> extreme that the gauze did nearly nothing. I showed the slide to one 
> of our pathologists and they could hardly see beyond the precipitate, 
> but said that they couldn't see any staining of the structure that they were looking for (I forget exactly what it was, but I know it's supposed to turn black).
> >
> > In my second test (to see if the metal holder was the problem) that 
> > I
> performed immediately after the first test, I used one test slide. I 
> deparaffinized it in the same coplin jars as before (moving it with 
> plastic
> forceps) and hydrated it to deionized water. I used new glass 
> containers for the periodic acid and deionized water rinse in Step 2, 
> for making the silver solution in Step 3, and for the warm silver 
> solution and warm deionized water in Step 4. I used plastic forceps to 
> move the slide into the periodic acid, and propped it up in the 
> container so that no glass rack or metal handle was used at all. I 
> used plastic forceps to transfer the slide to the deionized water 
> rinse, and dunked it several times and swished the slide around a bit. 
> I used plastic forceps to transfer the slide into the warm(-ish) silver solution and propped it up against the side again.
> After approximately 20 minutes, I saw precipitate floating around, and 
> I used plastic forceps to remove the slide from the silver solution. I 
> dipped the slide into the warm(-ish) deionized water several times, 
> and saw that the precipitate was again covering the slide and the 
> tissue so I stopped there for the day.
> >
> > We purchased all of the reagents listed in the above procedure from
> Rowley Biochemical (except for the Glacial Acetic Acid mentioned in 
> Step 7, but I didn't even get that far).
> >
> > Questions:
> >
> > 1. Could this indicate that the acid-washing was not done correctly? 
> > I
> made up a ~1% Hydrochloric Acid solution (with deionized water) and 
> filled a plastic bin with the solution (I rinsed the bin with 
> deionized water first). I then submerged all glassware (in several 
> batches) for at least five minutes, then rinsed well with deionized 
> water (not by filling a bin - I just used the hose of deionized water 
> in our lab sink and poured it over the glassware) and left them to air-dry overnight.
> >
> > 2. Are using acid-washed glassware and avoiding metal even necessary
> precautions after the sodium thiosulphate in Step 6? I read that 
> sodium thiosulphate "stops the reaction," and the procedure stops 
> specifically saying to use deionized water after Step 6 and starts 
> saying to use just "water" or "tap water." My lab refers to our waters 
> as either "tap" or "deionized," so I'm assuming that using my 
> deionized water is fine when the procedure calls for "distilled" or "dechlorinated."
> >
> > I don't even know enough to ask more questions, but I'm sure many 
> > more
> will arise after I test the stain again next week, so I welcome any 
> and all advice about silver stains, acid-cleaning glassware, and 
> literally anything else...
> >
> > Thank you!!!
> >
> > Jordan H.
> > University of Michigan
> > Ann Arbor, MI
> > **********************************************************
> > Electronic Mail is not secure, may not be read every day, and should 
> > not be used for urgent or sensitive issues 
> > _______________________________________________
> > Histonet mailing list
> > Histonet at lists.utsouthwestern.edu
> > https://urldefense.com/v3/__http://lists.utsouthwestern.edu/mailman/
> > listinfo/histonet__;!!OToaGQ!6kZFsSGmGAqtHSUCXGvBt6QPRHlQc7P4geZI9L6
> > HlrVmY_FPmCDYnGHGszj4cR01z2-BarQ$
> >
> _______________________________________________
> Histonet mailing list
> Histonet at lists.utsouthwestern.edu
> https://urldefense.com/v3/__http://lists.utsouthwestern.edu/mailman/li
> stinfo/histonet__;!!OToaGQ!6kZFsSGmGAqtHSUCXGvBt6QPRHlQc7P4geZI9L6HlrV
> mY_FPmCDYnGHGszj4cR01z2-BarQ$
>
> This message is intended for the addressee named and may contain 
> confidential information. If you are not the intended recipient, 
> please delete it and notify the sender.
>
> Views expressed in this message are those of the individual sender, 
> and are not necessarily the views of NSW Health or any of its entities.
> _______________________________________________
> Histonet mailing list
> Histonet at lists.utsouthwestern.edu
> https://urldefense.com/v3/__http://lists.utsouthwestern.edu/mailman/li
> stinfo/histonet__;!!OToaGQ!6kZFsSGmGAqtHSUCXGvBt6QPRHlQc7P4geZI9L6HlrV
> mY_FPmCDYnGHGszj4cR01z2-BarQ$
>

--
Colleen Forster HT(ASCP)QIHC
BLS Histology and IHC Laboratory
Jackson Hall, Room 2-155
321 Church St. SE
Minneapolis, MN 55455
612-626-1930

------------------------------

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------------------------------

End of Histonet Digest, Vol 214, Issue 14
*****************************************