Enhanced Reliability in Silver Impregnation of Terminal Axonal Degeneration-Original Nauta Method

Brains of rat with surgical lesions 3-5 days old are fixed in 10% neutralized formalin (excess of CaCO₃), 20 μ serial frozen sections cut therefrom and kept in neutralized formalin for an additional 24-48 hr. The sections are soaked in distilled water 12-24 hr, transferred to 50% alcohol containing 0.75 ml of concentrated NH₄OH (sp. gr. 0.91) per 100 ml 12-24 hr, placed in distilled water 2-3 hr and then in silver-pyridine solution (AgNO₃ 3% aq., 20 ml; pyridine, 1 ml) for 48 hr. Test sections are transferred directly to each one of 3 ammoniated silver-solutions, pH 12.8, 13.0 and 13.2, made as follows: To 200 ml of solution 1 (silver nitrate, 6.4 gm; alcohol 96%, 220 ml; NH₄OH (sp. gr. 0.91), 28 ml and distilled water, 440 ml) is added respectively 8-12 ml, 12-16 ml and 16-20 ml of solution 2 (2% NaOH) to give the pH desired. The test sections are studied and the optimal ammoniated silver solution chosen. Two baths of ammoniated silver are used, the section placed with continuous agitation into the first bath for 30 sec and the second bath for 60 sec. The sections are then transferred directly into a reducing bath (formalin 10%, 2ml; alcohol 96%, 5 ml; citric acid 1%, 1.5 ml and distilled water, 4.5 ml) for 2 min and from there to 5% Na₂S₂O₃ for 1 min, rinsed in 3 changes of distilled water, dehydrated and mounted.     

Staining of Oligodendroglia with Silver Ammino Tungstate in Unembedded and Embedded Material

Specimens of brain or spinal cord fixed in formalin, Cajal's formol-bromide, or Koenig, Groat and Windle's formalin-acacia can be used to stain oligodendrocytes in frozen, in paraffin, or in celloidin sections. The sections are soaked 3-5 min in 0.02% acetic acid, pH 3.4, then rinsed 2-3 sec in 3% H₂O₂ and transferred to a silver bath prepared as follows: Mix equal parts of 10% AgNO₃ and 10% Na₂WO₄, and dissolve the precipitate with concentrated NH₄OH; avoid an excess of ammonia. Silver at room temperature for 15-20 sec, develop in 1% formalin, dehydrate, and mount. For embedded material, prepare a mixture consisting of 1 part of 10% aqueous Aerosol MA and 4 parts of 10% Aerosol OT in 95% alcohol. Add 5 drops of this mixture to each 50 ml of dilute acetic acid and 3% H₂O₂; 5 drops to each 20 ml of the silver bath.     

A Copper Sulfate-Silver Nitrate Method for Nerve Fibers of Planarians

For staining in toto, planarians are fixed in a mixture of 10 ml of commercial formalin, 45 ml of 95% ethanol and 2 ml of glacial acetic acid. After treatment with 70% ethanol 3-10 days, they are washed in distilled water and immersed in 10% CuSO₄. 5H₂O for 3 hr at 50° C, transferred without washing to 1% AgNO₃ for 1.0-1.5 hr at 50° C; and then developed in: 10 ml of 1% pyrogallol, 100 ml of 56% ethanol and 1 ml of 0.2% nitric acid. Gold toning, 5% Na₂S₂O₃ and dehydration follow as usual. For staining sections, material is fixed in the same fixative, embedded in paraffin and sectioned at 10 μ. After bringing sections to water, they are immersed in 20% CuSO₄. 5H₂O for 48 hr at 37° C; then rinsed briefly in distilled water and placed in 7% AgNO₃ for 24 hr at 37° C. They are washed briefly in distilled water and reduced in: hydroquincne, 1 gm; Na₂SO₃, 5 gm and distilled water 100 ml. Gold toning, followed by 5% Na₂S₂O₃ and dehydration completes the process. Any counterstaining may follow.     

Application of the Nauta-Gygax Technic for Degenerating Axons to Mounted Sections

Frozen sections of formalin-fixed brains containing lesions were mounted on slides that had been coated first with albumen-glycerol (1:1) then 4% gelatin and blotted. The slides were placed in formaldehyde vapor at 56° C for 40-60 min, washed, and stored (optional) in 10% formalin-saline. The staining technic was as follows: after washing, soak 30-40 min in 0.5% phosphomolybdic acid, rinse; put in 0.05% potassium permanganate 9-16 min (usually 12 min); decolorize in a 1:1 mixture of 1% hydroquinone and 1% oxalic acid; wash thoroughly; soak in 1.5% AgNO₃ at about 20° C for 25-35 min; rinse; put into an ammino-silver solution (4.5% AgNO₃, 20 ml; pure ethanol, 10 ml; ammonia, sp. gr. 0.880, 2.4 ml; 2.5% NaOH, 1 ml) for 1-2 min; reduce in acidified formalin (distilled water, 400 ml; pure ethanol, 45 ml; 1 % citric acid, 13.5 ml; 10% formalin, 13.5 ml) for 1-3 min; wash; dehydrate through ascending grades of alcohol, including absolute; coat with 0.5% collodion, allow to dry slightly and harden in absolute alcohol-chloroform (2:1); rehydrate and put into 1% Na₂S₂O₃ for 1 min; dehydrate and cover.     

Silver Impregnation of Nerve Fibers in Teeth After Decalcification With Ethylenediaminetetraacetic ACID

The difficulties in impregnating bony tissues, which occur after decalcification with acids or electrolysis are avoided by decalcification with ethylenediaminetetraacetic acid at pH 8.2-8.5. The decalcification of adult human teeth which have been cut to a thickness of 2-5 mm takes 1-2 mo. If frozen sections of the decalcified teeth are impregnated 24 hr in 20% AgNo₃, rinsed through 6 changes of 20% neutralized (CaCO₃) formalin, blotted thoroughly with a cloth and placed in an ammoniated silver solution for 15-20 min, reliable impregnation of nerve fibers is obtained. The stock ammoniated silver solution is prepared by adding concentrated NH₄OH to 10-20 ml of 20% AgNO₃ until the precipitate formed by it is dissolved and then adding a few drops of the silver solution until the first permanent opalescence of the mixture is obtained. From this 2 ml are diluted directly before use with 6 ml of distilled water and 4 drops of concentrated NH₄OH added. The diluted stock solution should be used for few (5-10) sections only. The rest of the technic is done in the routine manner.     

A HYDROXYL-DEPENDENT SILVER METHOD FOR NERVE AXOPLASM

Axoplasm is selectively impregnated by the following steps: (1) fixation in 10% formalin or in 10% formalin with added sucrose, 15%, and concentrated NH₄OH, 1%, for 1-7 days; (2) frozen sections; (3) extraction of the sections in 95% ethyl alcohol, absolute alcohol, xylene, and 95% ethyl alcohol and absolute alcohol, 1 hr each; (4) distilled water, 3 changes of 10 min each; (5) 20% AgNO₃ (aq.) at 25°C, 30 min; (6) distilled water, 3 changes of 1-2 sec each; (7) 6.9% K₂CO₃, 1 hr; (8) water, 3 changes of about 1 min each; (9) 0.2%AuCl₃, 2 min; (10) distilled water; (11) 5% Na₂S₂O₃, 2 min; (12) washing, clearing and mounting. This procedure is proposed as a simplified stain for axoplasm, with other tissue components remaining unstained. The few reagents necessary suit this method for histochemical investigation of the mechanism of silver staining.     

A Chlorate-Formaldehyde Modification of the Golgi Method

Pieces of fresh nervous tissue 3-5 mm thick are put into a mixture of: 6% K₂Cr₂O₇, 40 ml; 5% KClO₃, 20 ml; 20% chloral hydrate, 30 ml; and concentrated formalin (38% HCHO), 10 ml; allowed to fix 3 days, with a daily change of fluid; transferred to 3% K₂Cr₂O₇ for 3 days, with twice daily changes; then to 1% AgNO₃ for 3 days at 20-25° C. Frozen sections are cut, dehydrated, cleared and mounted in Permount with a cover glass. The method gives good results for microglia and oligodendroglia in addition to the usual staining of nerve cells and their processes.     

Correlation of CrystaL Growth with the Staining of Axons by the Golgi Procedure

The surface of the specimens subjected to a modified Golgi technique (formalin fixed material; specimens in the following solution for 8-10 days at 27 C: 3% K₂Cr₂O₇, 100 ml, with the addition of 2.5-10 ml of 10% formalin and 6-25 gm of sucrose; then in 0.75% AgNO₃ for at least 2 days at 27 C) is sometimes covered with a fur of filamentous crystals and sometimes with a powdery precipitate of laminar crystals. In a series of experiments in which about 500 blocks of tissue were treated with variations of the staining procedure, good axonal stain was positively correlated with the appearance of filamentous crystals. These filaments have a thickness of 1-4 μ and grow at a rate of 160-330 μ/hr, reaching a length of 2-7 mm.     

A New Method of Silver Impregnation for Nerve Cells and Fibers

Fragments of tissue, immediately after death, are fixed in Debaisieux's modification of the Duboscq-Brazil picro-aceticformol fluid, and treated as follows: Hydrate by soaking 2-6 hr. in distilled water with 30 drops of cone. NH₄OH per 100 cc. Freeze and cut sections about 25μ in thickness. Bleach sections about 15 min. in ammoniacal water (52 drops cone. NH₄OH per 100 cc. water). Transfer to 20% AgNO₃ solution and heat at 45° C. till light brown. Add cone. NH₄OH drop by drop till the Ag precipitates and then redisolves into an opalescent solution. Pour solution and sections into a little distilled water and transfer sections quickly to formaldehyde solution (3 cc. formalin to 100 cc. water). Dip sections in distilled water and transfer to 1% aqueous gold chloride till deep blue. Place for about 10 minutes in 5% aqueous sodium thiosulfate solution for fixing and clearing. Wash thoroly in tap water, dehydrate and mount. Special directions are given for applying this technic to delicate material such as insects, and for use with serial sections.     

A Silver Impregnation Method for Nervous Tissue Suitable for Routine use with Mounted Sections

A simple, reliable silver impregnation method for nervous tissue is described for tissues fixed in various fixatives including formalin, Bouin, and Sum. Sections are impregnated in a solution containing 1 g Protargol, 2 ml of a 1% Cu(NO₃)₂ solution, 2 ml of a 1% AgNO₃ solution, and 2-4 drops 30% H₂O₂ in 100 ml distilled water. Sections are impregnated 4-5 days at 37 C and thereafter reduced in a hydroquinone-formalin solution. This is followed by gold toning and subsequent reduction, dehydration and mounting. This method has been found to be very reliable and selective.     

Ammoniated Silver Carbonate for the Demonstration of Lysosomes

Hortega's ammoniated silver carbonate method was used to demonstrate lysosomes in the central nervous system and kidney of adult rats. Formol-CaCl₂, (10%:1%) fixed, frozen sections were impregnated for 10 min in Hortega's solution: 30 ml of 10% AgNO₂ and 90 ml of 5% Na₂CO₃, with concentrated NH₄OH added until the precipitate dissolved, then distilled water to make 400 ml. This procedure revealed silver-positive cytoplasmic structures whose form, shape and distribution were similar to that seen by staining adjacent sections for acid phosphatase. A short fixation of 18-24 hr appears to be essential. A useful, nonenzymatic method for the demonstration of lysosomes is thereby available.     

Biological Stain Commission Membership,June 1959

Frozen sections of avian tissue fixed 7 days or longer in 10% formalin or formol-saline are cut at 20-50 μ, left in distilled water for 2 hr, and placed in 0.002% aqueous AgNO₃ for 3-4 days. Subsequent procedure is essentially that of Weddell and Glees. Sections are placed in 20% AgNO₃ for 30 min, then carried through 3 baths of 3% formalin in less than 10 min. Immediately thereafter they are washed 1-2 sec in a 0.1% solution of NH₄OH (cone) and placed in the ammoniacal silver solution (made with 20% AgNO₃) until the nerves become distinct, as seen under a microscope; usually, in about 15 min. After washing briefly, the sections are fixed in 5% Na₂S₂O₃ for 3-10 min, dehydrated, cleared, and mounted in the usual way.     

Staining Pericellular Boutons of Glutaraldehyde-Fixed Nervous Tissue; A Chromation-Silvering Sequence for Frozen Sections

After fixing in phosphate-buffered 5% glutaraldehyde, pH 6.8, by perfusion, brains were sliced to 3-5 mm pieces which were placed in the fixative for 5-7 days. The pieces were washed through several changes of 2.26% NaH₂PO₄ for 12 hr, 30 μ frozen sections cut, and mordanted 2 days in an equal-parts mixture of 3.5% CrO₃ and 5% Na-tartrate, which had been aged at 20-25 C for 20 days prior to use. After washing in distilled water, the sections were put into a solution containing AgNO₃, 20 gm; and KNO₃, 15 gm, in distilled water, 80 ml; at 30 C for 1.5-2 hr, then reduced at 40-45 C in three pyrogallol solutions as follows: 1-2 sec in 1% pyrogallol in 55% alcohol; 3-4 sec in a 0.67% solution in 33% alcohol, and 5-7 sec in a 0.5% solution in 25% alcohol. Gold toning is optional; dehydration, clearing and covering, routine. The technic shows particularly the perisomatic fibers, boutons en passant and boutons termineaux. Fibers in nerve tracts may be visible but lightly stained; cell nuclei may be dark, but the cytoplasm remains pale.     

Application and Evaluation of the Lead-Adenosine Triphosphatase Method in Skeletal Tissue

Application and evaluation of the lead-ATPase histochemical method in skeletal tissue has demonstrated an intracellular localization of enzyme activity. The skeletal tissue was demineralized for 72 hr in cold 10% aqueous EDTA adjusted to pH 7.2. Frozen sections were cut and placed on cold albumenized slides, oriented, thawed, dried in a cool air stream, and fixed for 10 min in cold (-2 to -3 C) 10% formalin buffered with Na-acetate and adjusted to pH 7.2. The sections were washed, treated with 10% EDTA for 20 min at room temperature, rewashed, and incubated for an optimal period of 30 min at 37 C. in the lead-ATP medium of Wachstein and Meisel. Following incubation the sections were washed, treated for 1 min with 1% (NH₄)₂S, rewashed, immersed for 30 min in 10% buffered formalin, dehydrated, cleared, and mounted. Evaluation of the substrate specificity suggests that other phosphatases associated with skeletal tissue do not complicate the ATPase reaction.     

Impregnation of Oligodendroglia in Nervous Tissue Kept in Formalin for Many Years

A method for impregnating oligodendroglia in nervous tissue (monkey) fixed and preserved in formalin for many years is described. This tissue is reconditioned by placing 12 to 30μ frozen sections of it in concentrated ammonia (sp. gr. 0.90) and by washing them slowly for 24 hours with a 1 mm. stream of water. The fluid is then poured off the sections; the jar is refilled with concentrated ammonia; and washing is repeated for another 24 hours. The sections are then plunged into concentrated ammonia for 7 minutes.

After treatment in ammonia, the sections are incubated for one hour at 38^oC. in Globus' 5% hydrobromic acid solution. They are washed again, in distilled water, and then impregnated in a “medium” strength ammoniacal silver carbonate solution (5 ml. of 10% AgNO₃ added to 15 ml. of 5% Na₂CO₃. The precipitate is dissolved in concentrated ammonia and diluted to SO ml. with distilled water). Impregnation is followed by reduction in 1% formalin without agitation; fixation in 5% Na₂S₂O₃; dehydration, and mounting in clarite.

Typical oligodendroglia (Fig. 1) were made visible by use of the method outlined in this paper.     

A Glutaraldehyde-Dichromate-Silver Sequence for Differentiating Norepinephrine Cells from Epinephrine Cells in Neonatal and Adult Rats—Paraffin Sections

A glutaraldehyde-K₂Cr₂O₇ procedure intensified by silver staining enabled norepinephrine and epinephrine cells to be distinguished readily in paraffin sections of the adrenal glands of rats 8 days after birth. The technique involved fixation in 0.1 M cacodylate-buffered 5% glutaraldehyde (6-24 hr), treatment with 3.5% K₂Cr₂O₇ (6-12 hr) and routine preparation of paraffin sections. The sections were deparaffinised, brought to water and immersed in Fontana's solution (24 hr), prepared by adding concentrated NH₄OH drop by drop to 5% AgNO₃ until the precipitate formed just redissolved; more 5% AgNO₃ was then added until a permanent cloudiness just developed. After a rinse in distilled water, the sections were treated with 0.5% gold chloride (5 min) and Na₂S₂O₃ (5 min), then mounted in Depex. This sequence resulted in an intense black cytoplasmic colouration in norpinephrine-containing cells of both the adult and 8-day-old animals whereas epinephrine-containing cells remained colourless. The glutaraldehyde-K₂CrO₇ procedure, without intensification, gave very clear results in the adult: a yellow cytoplasmic colour in the norepinephrine cells with epinephrine cells colourless. A glutaraldehyde-OsO₄ sequence gave a less well defined separation of these cell types in the adult and failed to distinguish the cell types in the neonate.     

The Histochemical Detection of Thallium in Paraffin-Embedded Tissues by the Sulfide-Selenium-Silver Method

Thallium can be histochemically localized in formalin-fixed, paraffin-processed tissues by treating sections, after passing them through xylene and graded alcohols to water, with gaseous H₂S for 15 min and with 20% ammonium sulfide saturated with powdered selenium for 10 min. Sections are then washed, treated 10 min with 20% H₂O₂, and incubated in darkness for 20-30 min in the following mixture: 25% gum acacia, 10 ml; 2% hydroquinone in 5% citric acid, 1 ml; and 10% AgNO₃, 0.1 ml. Tissues and cells, which contain thallium, are demonstrated by small black granules of silver.     

Glutaraldehyde-Ammoniacal Silver Carbonate-Formaldehyde Staining of Histones of Mitotic Chromosomes

Cells from monolayer culture of Chinese hamster line Don were treated by Colcemid (0.1 μg/ml) for 2 hr, trypsinized and spun; resuspended in 0.5% sodium citrate solution for 10 min, respun, and then resuspended in a small volume of the supernatant. Slide preparations were made by smearing, followed by air drying for 1 min at room temperature. They were fixed and stained by the following sequence: 2.5% glutaraldehyde in Millonig's buffer, 30 min; distilled water, 6 min, 5 changes; ammoniacal silver at 18-26 C, 10 sec; distilled water, 30 min, 5 changes; 2.5% formalin, 2 min; and distilled water, 3 changes during 15 min. Staining solution: add 225 ml of 5% Na₂CO₃ to 75 ml of 10% AgNO₃, then add concentrated NH₄OH slowly, drop by drop, until the solution is transparent. Finally add 300 ml of dstilled water. Cells treated with cold 0.25 N HCl before fixation were not stained. Sequence modifications show that chromatin does not reduce silver by itself. This method stains the sites of high histone concentrations in mitotic chromosomes of cytogenetic preparations.     

Histochemical Demonstration of Pyrophosphatase

Fresh tissue slices were fixed in 5% formalin containing 0.9% NaCl for 10-20 min and frozen sections therefrom floated for 3 hr at 37°C on an incubating mixture made as follows. Sodium pyrophosphate (Na₄P₂O₇-12H₂O), 1.088 gm was dissolved in 20-30 ml of distilled water and to this was added ferric chloride (FeCl₃-6H₂O), 0.61 gm dissolved in 10-15 ml of water. The precipitate was just dissolved by cautiously adding 5-10% aqueous Na₂CO₃ solution and the pH adjusted to 7.2 with 1N HCl. The volume was made up to 100 ml and 0.9 gm of NaCl added. Before use, 1 ml of 10% Mg(NO₃) was added. After incubation, sections were washed 10-15 min in 0.9% NaCl, then mounted on glass slides and air-dried. When dry, the slides were immersed in 0.9% NaCl containing 0.2-0.5% ammonium sulfide for 2-3 min, then dehydrated rapidly through graded alcohols, cleared, and covered in balsam. Sites of pyrophosphatase activity stained in various shades of green. Acid pyrophosphatase also was histochemically demonstrated by the same principle, excepting that the substrate solution was adjusted to pH 3.7-4.0 with acetate buffer. The pattern of distribution of pyrophosphatase and glycerophosphatase was almost identical.     

Acceleration of Mallory's Phosphotungstic Acid-Hematoxylin Staining for Skeletal Muscle Fixed in Formalin

The staining time for mammalian skeletal muscle fixed in neutral phosphate-buffered formalin was shortened from 12-24 hr to 10-30 min. The permanganate-oxalate sequence was omitted although oxidation by periodic acid or with iodine was found to be necessary. The material was embedded in paraffin and cut 6 μ or less. Deparaffinized sections were treated with 1% alcoholic iodine for 10 rain followed by 5% Na₂S₂O₃ for 2 min and placed in an oven at 60 C for 10-30 min to stain in a preheated mixture of 50 ml of ripened Mallory's phosphotungstic acid-hematoxylin and 1 ml of 2% phosphomolybdic acid. Experiments with fixation showed that the staining procedure followed Zenker's fluid successfully but not Bouin's fluid. Oxidation by KMnO₄ was effective only after Zenker fixation; oxidation by CrO₃ was unsuccessful.