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

Tony Henwood (SCHN) tony.henwood at health.nsw.gov.au
Thu Sep 23 18:13:40 CDT 2021

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.

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

Try the method given in StainsFile at:

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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