Re: [Histonet] Jones' Methenamine Silver Stain for Basement Membranes of Kidney - Issues and Questions

[Hood, Jordan via Histonet]

I would like to thank all of you for your advice --

Using Thiosemicarbazide after the periodic acid did the trick, along with 
making up the silver solution immediately before use. It only took ~30 minutes 
in the warm silver solution before the basement membranes reached the desired 
staining intensity. Excellent results on my first try using the TSC, especially 
when compared to my first two disastrous tests.

I am sincerely grateful to you all for helping me so much!!

Best wishes,
Jordan

From: Colleen Forster 
Sent: Thursday, September 23, 2021 7:28 PM
To: Tony Henwood (SCHN) 
Cc: Bryan Llewellyn ; Hood, Jordan ; 
histonet@lists.utsouthwestern.edu
Subject: Re: [Histonet] Jones' Methenamine Silver Stain for Basement Membranes 
of Kidney - Issues and Questions

External Email - Use Caution
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 
mailto:histonet@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@lists.utsouthwestern.edu]
Sent: Friday, 24 September 2021 7:47 AM
To: Jordan mailto:jordh...@med.umich.edu>>; Histonet 
mailto:histonet@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:
http://stainsfile.info/stain/metallic/jones.htm

Bryan Llewellyn


Hood, Jordan via Histonet wrote:
> Hello,
>
> I'm new to histology (and new to histonet), and I work in a small histology 
> lab specializing in animal tissues that receives requests/submissions from 
> researchers. I tried (and failed) to perform a Jones' Methenamine Silver 
> stain on a client's submission of pig kidneys (formalin-fixed, 
> paraffin-embedded, cut at 2.5 microns), and I need some help troubleshooting 
> this stain since my co-workers are stumped, too.  I used the following 
> procedure from Rowley Biochemical:
>
>
> ~
> "Fixation: 10% Buffered Neutral Formalin (F-113) or Bouin's Solution
> (F-40) or Zenker's (F-155)
>
> Sections: Paraffin, 2 microns
>
> Procedure: Acid washed glassware must be used
> 1. Deparaffinize and hydrate to distilled water.
> 2. Oxidize in Periodic Acid 0.5% (F-396-1) 11 minutes. Wash in chloride-free 
> water.
> 3. Prepare Methenamine Silver solution by mixing: 42.5 ml Methenamine 3% 
> (F-396-2), 2.5 ml Silver Nitrate, 5% (F-396-3) and 12.0 ml Borate Buffer, pH 
> 8.2 (F-396-4).
> 4. Place slides in the solution and the entire jar in a water bath at 70°C 
> for approx. 60-75 minutes. Check under microscope when slides appear medium 
> brown microscopically. Every 10 minutes, once the medium brown color has been 
> established, rinse a slide in 70°C, chloride free water and check under a 
> microscope. Rinse again in hot water and return to the hot staining solution. 
> As the staining time approaches the end point, check the slides, as above, 
> every 1-2 minutes. The entire procedure must be performed quickly to prevent 
> an uneven staining of the tissues. The slides should exhibit a brownish- 
> yellow background, intense black reticulum fibers, and black basement 
> membranes. If the slides become oversaturated, i.e. too black, destain in a 
> dilute Potassium Ferricyanide Solution (F-396-11) for one or two dips.
> 5. Rinse well in distilled water. Tone in Gold Chloride 0.2% (F-396-5), 1 
> minute. If sections are overtoned place in Sodium Metabisulfite, 3% 
> (F-396-12) for 1-3 minutes. Rinse well in distilled water.
> 6. Sodium Thiosulfate 3% (F-396-9), 1-2 miutes. Wash in running tap water, 10 
> minutes. Rinse well in distilled water.
> 7. Stain in Harris' Hematoxylin (F-396-6) containing 2-4ml of Glacial Acetic 
> Acid per 100 ml for 5-15 minutes. Wash in water.
> 8. Differentiate in Acid Alcohol 1% (F-396-13) until the sections turn red.
> 9. Blue section in Ammonia Water, 0.3% (F-396-14). Wash thoroughly.
> 10. Counterstain in Eosin Y, 1%, Alcoholic Solution (F-396-7).
> 11. Dehydrate in 95% alcohol, absolute alcohol and clear in xylene 3 changes 
> each. Mount.
>
> Stain Results:
> Basement membranes, reticulum fibers: Black
> Nuclei: 

Re: [Histonet] Jones' Methenamine Silver Stain for Basement Membranes of Kidney - Issues and Questions

[Mayer,Toysha N via Histonet]

Hi Jordan,

So wow!  You are doing a Jones with an H
What you are seeing is common when using a water bath and preheating.  While 
you can perform the stain with this many sections, I would start off with 1-3 
slides until you have perfected the technique.
I know that makes you have multiple runs, but you don't want to waste the 
tissue or have a lot of precipitate.
1. I would only preheat silver  for 5-10 min to avoid excess precipitation and 
agitate the solution before immersing your slides.
2. If you use the glass staining rack, make sure no excess water from the water 
bath gets in.
3. Rinse silver stains 6x with de-I or distilled water (my original trainers 
taught me that).
4. If you can, use an oven, not water bath to heat. Same temp settings as on 
the water bath.
5. No metal at all.  Use plastic hemostats and rinse them well between uses.
6. Mix your silver and methenemine first, then filter your sliver.  Add the 
borax sol'n just  before use, cut it down to 5ml.
7. The glomeruli take a long while to get to the point where they start to turn 
and they have to be dark.  Paper bag brown is the what we say.
8. Always use de-I after periodic acid.

When I teach this stain to students, we sometimes use the oven and sometimes 
the water bath.  The oven cuts down on the precip and we can agitate the slides 
better.  The gold chloride and hypo will help with the precip, but wipe the 
back and sides of the slides well between each change. Be careful not to push 
the solution back onto the section.
The new Carson's version uses 50 mL of methenamine silver (5% silver in 3% 
methenamine) and 5mL of Borax. It also uses 1% periodic. I was always taught 
that 0.5% would give me non-specific staining because it was not sensitive 
enough.  I trained using the formulation you did, but we also microwaved.  We 
were specifically animal where I trained, and I have the microwave procedure 
somewhere if you want.
Also, others mentioned the procedure on stainsfile.  It works as well. I taught 
that method for a few years as well.

Good luck and keep me posted.


Sincerely,


Toysha N. Mayer, DHSc, MBA, HT (ASCP)
Assistant Professor/Associate Program Director
HTL Program School of Health Professions
MD Anderson Cancer Center
tnma...@mdanderson.org
713-563-3481 wk
832-710-1837 cell








--

Message: 2
Date: Thu, 23 Sep 2021 19:49:52 +
From: "Hood, Jordan" 
To: "'histonet@lists.utsouthwestern.edu'"

Subject: [Histonet] Jones' Methenamine Silver Stain for Basement
Membranes of Kidney - Issues and Questions
Message-ID: <48bd31f00b074079baca10d963e1b...@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) 

Re: [Histonet] Jones' Methenamine Silver Stain for Basement Membranes of Kidney - Issues and Questions

[Colleen Forster via Histonet]

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@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@lists.utsouthwestern.edu]
> Sent: Friday, 24 September 2021 7:47 AM
> To: Jordan ; Histonet <
> histonet@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:
> http://stainsfile.info/stain/metallic/jones.htm
>
> Bryan Llewellyn
>
>
> Hood, Jordan via Histonet wrote:
> > Hello,
> >
> > I'm new to histology (and new to histonet), and I work in a small
> histology lab specializing in animal tissues that receives
> requests/submissions from researchers. I tried (and failed) to perform a
> Jones' Methenamine Silver stain on a client's submission of pig kidneys
> (formalin-fixed, paraffin-embedded, cut at 2.5 microns), and I need some
> help troubleshooting this stain since my co-workers are stumped, too.  I
> used the following procedure from Rowley Biochemical:
> >
> >
> > ~
> > "Fixation: 10% Buffered Neutral Formalin (F-113) or Bouin's Solution
> > (F-40) or Zenker's (F-155)
> >
> > Sections: Paraffin, 2 microns
> >
> > Procedure: Acid washed glassware must be used
> > 1. Deparaffinize and hydrate to distilled water.
> > 2. Oxidize in Periodic Acid 0.5% (F-396-1) 11 minutes. Wash in
> chloride-free water.
> > 3. Prepare Methenamine Silver solution by mixing: 42.5 ml Methenamine 3%
> (F-396-2), 2.5 ml Silver Nitrate, 5% (F-396-3) and 12.0 ml Borate Buffer,
> pH 8.2 (F-396-4).
> > 4. Place slides in the solution and the entire jar in a water bath at
> 70°C for approx. 60-75 minutes. Check under microscope when slides appear
> medium brown microscopically. Every 10 minutes, once the medium brown color
> has been established, rinse a slide in 70°C, chloride free water and check
> under a microscope. Rinse again in hot water and return to the hot staining
> solution. As the staining time approaches the end point, check the slides,
> as above, every 1-2 minutes. The entire procedure must be performed quickly
> to prevent an uneven staining of the tissues. The slides should exhibit a
> brownish- yellow background, intense black reticulum fibers, and black
> basement membranes. If the slides become oversaturated, i.e. too black,
> destain in a dilute Potassium Ferricyanide Solution (F-396-11) for one or
> two dips.
> > 5. Rinse well in distilled water. Tone in Gold Chloride 0.2% (F-396-5),
> 1 minute. If sections are overtoned place in Sodium Metabisulfite, 3%
> (F-396-12) for 1-3 minutes. Rinse well in distilled water.
> > 6. Sodium Thiosulfate 3% (F-396-9), 1-2 miutes. Wash in running tap
> water, 10 minutes. Rinse well in distilled water.
> > 7. Stain in Harris' Hematoxylin (F-396-6) containing 2-4ml of Glacial
> Acetic Acid per 100 ml for 5-15 minutes. Wash in water.
> > 8. Differentiate in Acid Alcohol 1% (F-396-13) until the sections turn
> red.
> > 9. Blue section in Ammonia Water, 0.3% (F-396-14). Wash thoroughly.
> > 10. Counterstain in Eosin Y, 1%, Alcoholic Solution (F-396-7).
> > 11. Dehydrate in 95% alcohol, absolute alcohol and clear in xylene 3
> changes each. Mount.
> >
> > Stain Results:
> > Basement membranes, reticulum fibers: Black
> > Nuclei: Blue
> > Cytoplasm, collagen, connective tissue: Pink-orange
> >
> > References: Jones, D.B., Amer.J.Path. 27:99 (1951). AFIP Manual of
> Histolocical Staining Methods, 3rd ed., Ed. L. Luna: NY: McGraw-Hill
> Publications, c. 1968, p. 97."
> > ~
> >
> >
> > It became apparent that something went wrong during Step 4 when the
> slides were in the glass container (not a coplin jar - we have ten slides
> that we need to stain so we're using a rectangular glass container that
> holds ten slides on their sides - it does require a metal handle to move,
> but the handle is flexible and easy to remove after the glass slide rack
> has been transferred between containers) of silver solution in the water
> bath because there was lots of precipitate on the slides and floating on
> the surface of 

Re: [Histonet] Jones' Methenamine Silver Stain for Basement Membranes of Kidney - Issues and Questions

[Tony Henwood (SCHN) via Histonet]

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@lists.utsouthwestern.edu] 
Sent: Friday, 24 September 2021 7:47 AM
To: Jordan ; Histonet 

Subject: Re: [Histonet] Jones' Methenamine Silver Stain for Basement Membranes 
of Kidney - Issues and Questions

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

Bryan Llewellyn


Hood, Jordan via Histonet wrote:
> Hello,
> 
> I'm new to histology (and new to histonet), and I work in a small histology 
> lab specializing in animal tissues that receives requests/submissions from 
> researchers. I tried (and failed) to perform a Jones' Methenamine Silver 
> stain on a client's submission of pig kidneys (formalin-fixed, 
> paraffin-embedded, cut at 2.5 microns), and I need some help troubleshooting 
> this stain since my co-workers are stumped, too.  I used the following 
> procedure from Rowley Biochemical:
> 
> 
> ~
> "Fixation: 10% Buffered Neutral Formalin (F-113) or Bouin's Solution 
> (F-40) or Zenker's (F-155)
> 
> Sections: Paraffin, 2 microns
> 
> Procedure: Acid washed glassware must be used
> 1. Deparaffinize and hydrate to distilled water.
> 2. Oxidize in Periodic Acid 0.5% (F-396-1) 11 minutes. Wash in chloride-free 
> water.
> 3. Prepare Methenamine Silver solution by mixing: 42.5 ml Methenamine 3% 
> (F-396-2), 2.5 ml Silver Nitrate, 5% (F-396-3) and 12.0 ml Borate Buffer, pH 
> 8.2 (F-396-4).
> 4. Place slides in the solution and the entire jar in a water bath at 70°C 
> for approx. 60-75 minutes. Check under microscope when slides appear medium 
> brown microscopically. Every 10 minutes, once the medium brown color has been 
> established, rinse a slide in 70°C, chloride free water and check under a 
> microscope. Rinse again in hot water and return to the hot staining solution. 
> As the staining time approaches the end point, check the slides, as above, 
> every 1-2 minutes. The entire procedure must be performed quickly to prevent 
> an uneven staining of the tissues. The slides should exhibit a brownish- 
> yellow background, intense black reticulum fibers, and black basement 
> membranes. If the slides become oversaturated, i.e. too black, destain in a 
> dilute Potassium Ferricyanide Solution (F-396-11) for one or two dips.
> 5. Rinse well in distilled water. Tone in Gold Chloride 0.2% (F-396-5), 1 
> minute. If sections are overtoned place in Sodium Metabisulfite, 3% 
> (F-396-12) for 1-3 minutes. Rinse well in distilled water.
> 6. Sodium Thiosulfate 3% (F-396-9), 1-2 miutes. Wash in running tap water, 10 
> minutes. Rinse well in distilled water.
> 7. Stain in Harris' Hematoxylin (F-396-6) containing 2-4ml of Glacial Acetic 
> Acid per 100 ml for 5-15 minutes. Wash in water.
> 8. Differentiate in Acid Alcohol 1% (F-396-13) until the sections turn red.
> 9. Blue section in Ammonia Water, 0.3% (F-396-14). Wash thoroughly.
> 10. Counterstain in Eosin Y, 1%, Alcoholic Solution (F-396-7).
> 11. Dehydrate in 95% alcohol, absolute alcohol and clear in xylene 3 changes 
> each. Mount.
> 
> Stain Results:
> Basement membranes, reticulum fibers: Black
> Nuclei: Blue
> Cytoplasm, collagen, connective tissue: Pink-orange
> 
> References: Jones, D.B., Amer.J.Path. 27:99 (1951). AFIP Manual of 
> Histolocical Staining Methods, 3rd ed., Ed. L. Luna: NY: McGraw-Hill 
> Publications, c. 1968, p. 97."
> ~
> 
> 
> It became apparent that something went wrong during Step 4 when the slides 
> were in the glass container (not a coplin jar - we have ten slides that we 
> need to stain so we're using a rectangular glass container that holds ten 
> slides on their sides - it does require a metal handle to move, but the 
> handle is flexible and easy to remove after the glass slide rack has been 
> transferred between containers) of silver solution in the water bath because 
> there was lots of precipitate on the slides and floating on the surface of 
> the silver solution.
> 
> In my first test, I used five test slides (extra slides that we cut from the 
> same blocks that were submitted to us). I deparaffinized them in coplin jars 
> (moving them with plastic forceps) and hydrated them to deionized water. I 
> transferred the slides to a glass slide rack that holds ten slides on their 
> sides, added five blank slides that were rinsed 

Re: [Histonet] Jones' Methenamine Silver Stain for Basement Membranes of Kidney - Issues and Questions

[Bryan Llewellyn via Histonet]

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

Bryan Llewellyn


Hood, Jordan via Histonet wrote:

Hello,

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


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

Sections: Paraffin, 2 microns

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

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

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


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

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

[Histonet] Jones' Methenamine Silver Stain for Basement Membranes of Kidney - Issues and Questions

[Hood, Jordan via Histonet]

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