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

Thu Sep 23 18:28:03 CDT 2021

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:
> 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 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
> > **********************************************************
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-- 
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