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 <jordh...@med.umich.edu>; 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 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@lists.utsouthwestern.edu > > http://lists.utsouthwestern.edu/mailman/listinfo/histonet > > > _______________________________________________ > Histonet mailing list > Histonet@lists.utsouthwestern.edu > http://lists.utsouthwestern.edu/mailman/listinfo/histonet > > 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@lists.utsouthwestern.edu > http://lists.utsouthwestern.edu/mailman/listinfo/histonet > -- 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 _______________________________________________ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
