Pseudoisocyanin Staining of Insulin and Specificity of Empirical Islet Cell Stains

Two closely related pseudoisocyanins, N,N'-diethyl-6,6'-dichlorpseudoisocyanin chloride and N, N'-diethylpseudoisocyanin chloride, were tested for their metachromatic staining behavior with oxidized insulin. N,N'-diethyl-6,6-dichlorpseudoisocyanin chloride gave nonspecific metachromasia with collagen, mucus, and mast cells of adult tissues; almost all tissues of rat embryos exhibited nonspecific staining. Nonspecific reactions were rarely observed in adult or fetal tissues with the extremely labile metachromasia of N, N'-diethylpseudoiso-cyanin chloride. When oxidation time and temperatures are carefully controlled, this reagent apears to be highly specific for insulin-containing cells and can be used as a selective stain for beta cells. Paraffin sections of formalin fixed material were oxidized 45 sec at 28-29 C in freshly prepared acidified permanganic (2.5% KMnO₄, 1; 5% H₂SO₄, 1; distilled water, 7—parts by volume), decolorized 30 sec in 5% oxalic acid, and washed 5 min in running tap water. After rinsing in 2 changes of distilled water, sections were stained 20 min in a 36 mg/100 ml aqueous solution of N, N'-diethylpseudoisocyanin chloride. Sections were then washed in running tap water until the albumen adhesive was decolorized, and mounted in Karo syrup diluted with an equal amount of distilled water. The insulin-containing cells are stained light to dark purple; all other tissue components, various shades of red. N, N'-diethylpseudoisocyanin chloride was used as a reference for evaluating the specificity of 5 commonly used empirical methods for demonstrating alpha and beta cells in pancreatic islets. Cells exhibiting pseudo isocyanin metachromasia were stained selectively by aldehyde-fuchsin, Heidenhain's azan, and chrome-hematoxylin. Aldehyde-Iuchsin was the only empirical stain tested which gave results comparable to pseudoisocyanin for clarity and definition of beta cells. After oxidation in acidified permanganate, azocarmine and phosphotungstic acid-hematoxylin differentially stained alpha cells; cells demonstrated by these two methods did not exhibit pseudoisocyanin metachromasia. This histochemical procedure can precede empirical methods which require preliminary oxidation in acidified permanganate or it can follow empirical methods which do not extract the insulin nor alter its intramolecular disulfide bonds.     

Adaptation of Mallory's Trichrome Stain to Embryonic and Fetal Material

Based upon results of an investigation of the role of phosphotungstic acid in connective tissue staining, the Mallory trichrome stain was adapted to sequential application of all three dyes, thus making it usable on embryonic and fetal material. Ten to twelve day postconception mouse fetuses were formalin fixed and paraffin embedded. Staining was as follows: (1) 1% aqueous acid fuchsin for 5 min followed by not more than 30 sec in running tap water; (2) 2% aqueous phosphomolybdic acid (PMA) for 10 min followed by a 2 min running tap water wash; (3) staining in 0.5% aniline blue in 8% acetic acid for 10 min, followed consecutively by 30 sec in running tap water, 2% aqueous PMA for 2 min, and 30 sec in running tap water; (4) 2% orange G in 8% acetic acid for 5 min, and rinsing for 30 sec in running tap water. Dehydration in ethanol, t-butanol, acetone, or by blotting followed by 1:3 terpineol-xylene, clearing in xylene and mounting, completed the procedure. The 30 sec tap water rinses can optionally be replaced by 1-2 min in 8% acetic acid. Sections can be made redder by increasing acid fuchsin staining time, or increasing time in the first PMA; red can be decreased by decreasing staining time, increasing time of the 2 min tap water wash, or decreasing time in the first PMA. Blue or orange staining can be increased or decreased by varying staining times in these solutions. Sharper differentiation may be obtained by increasing the time in PMA.     

Preparation of Brain Slices for Macroscopic Study by the Copper Sulfate-Phenol-Ferrocyanide Technique

The technique of Mulligan (J. Anat., 65: 468-72, 1931) for preparing gross brain slices has been modified to include improvements in fixation, slicing and staining. The iron alum-tannic acid steps have been eliminated and potassium ferrocyanide substituted. Washing times have been reduced and a detailed routine for fixation and slicing developed. After postmortem removal, human brains were perfused with 40% formalin and kept suspended in 10% formalin for 2-4 wk. Serial slices, 4 mm thick, were cut with an electrically driven rotary-blade meat slicer, washed in running water for 12 hr, immersed in CuSO₄-phenol solution (0.5%: 5%) at 60 C for 6 min, washed again in running water for 5 min, and then treated with a 2% solution of potassium ferrocyanide until a red-brown color developed in the gray matter, usually within 30-60 sec. The slices were then washed again for 5 min and stored in 10% formalin. Brains of patients over 60 yr of age frequently stained poorly and were not used.     

Controlled Differentiation of Cells and Connective Tissue Fibers in Silver Staining

Controlled silver staining of connective tissue fibers and sometimes of these fibers and cells simultaneously can be obtained. 1. Fix in 10% formalin. Embed in paraffin and cut sections as usual, but do not mount them on slides. Deparaffinize and hydrate through xylene, alcohols and distilled water and henceforth treat them the same as frozen sections. Real frozen sections can also be used. 2. Treat with a freshly prepared 1% solution of KMnO₄, usually 15-60 sec, sometimes up to 10 min. 3. Wash in distilled water, 5-10 sec. 4. Decolorize in 2% potassium metabisulfite, 10-20 sec. 5. Place in distilled water, 1 min. 6. Sensitize with 2% iron alum, 1 min. 7. Place in distilled water, 1 min. 8. Impregnate in Gomori's silver oxide solution, 2 min. 9. Wash in a 1.5% aqueous solution of pyridine, about 15 sec. 10. Reduce in a mixture containing 0.25% gelatin and 2% formalin 1 min. 11. Repeat steps 7 to 10 once or several times until the connective tissue fibers are completely stained. For cell staining (which may fail) proceed as follows: After the first insufficient staining of the connective tissue fibers, rinse in distilled water, dip for 1 sec in Gomori's solution and reduce immediately in gelatin-formalin without previous washing in pyridined water. This step can be repeated. 12. If the staining is too strong, decolorize as needed in 2% iron alum. 13. Toning in 0.2% gold chloride, 5 min or more, followed by fixation in 5% sodium thiosulfate, 1 min, is optional. Counterstain as desired. 14. Wash in tap water, dehydrate, clear in xylene and mount in balsam. The same technique applied to sections attached to slides gives good results but inferior to that obtained in paraffin sections processed in the loose, unmounted condition.     

Enhancement of the Affinity of Nucleoli and Chromosomes for Zinc by Treatment with Chromic Acid

Cultured mammalian cells and wet touch preparations from human organs were fixed for 10 min in 5:85:10 acetic-alcohol-formalin; placed in 5% aqueous CrO₃ for 30 min at 22-25 C; washed in running water 1 min; placed in 2 mM zinc acetate in 0.14 M veronal-acetate buffer, pH 6.5, at 37 C, 30 mm; rinsed 5 sec in 50% acetone; and stained 10 min in a solution dithizone. This results in selective staining of the nucleoli of interphase cells, and of the chromosomes of mitotic cells.     

Stain Technology: A Phenylhydrazine-Leucofuchsin Sequence for Staining Nerves and Nerve Endings in the Integument of Poultry

This sequence for staining cutaneous nerves and nerve endings uses 1% formic acid as a fixative for 1 hr, followed by two treatments of 5 min each in 6% H₂SO₄. The tissue is then submerged in fresh 5% phenylhydrazine hydrochloride for 30 min, washed in running tap water for 10 min, and given a 5 min soak in distilled water. The specimen is placed in Lillie's “cold Schiff” reagent for 4 hr; transferred to 6% H₂SO₄, 4 changes of 5 min each; washed in distilled water, 3 changes of 5 min each; dehydrated in acetone, 4 changes of 10 min each; and cleared in 2 changes of methyl benzoate, the 1st for 1 hr and the 2nd until the tissue clears. Nerve fibers stain pinkish-purple; muscles also take up the stain, yet the nerves are discernible from the muscles. All other tissue remain unstained.     

Histochemical Use of Sodium Bismuthate

Histochemical 1,2-glycoI cleavage, similar to that obtained with periodic acid and lead tetraacetate, may be obtained with sodium bismuthate. Routinely prepared slide sections, from tissues fixed in 10% formalin, are run down through xylene and graded alcohols to water and then oxidized for three minutes in a 1% sodium bismuthate 20% aqueous phosphoric acid solution. The oxidizing solution must be freshly prepared and used immediately. Following oxidation, sections are rinsed 15 sec. in IN HC1 to remove bismuth pentoxide precipitate, a by-product of the reaction. The sections are then washed in distilled water and placed in leuco-fushsin for 10 min., or in a saturated 30%) alcoholic solution of p-nitrophenylhydrazine for 5 min. or 2,4-dinitrophenylhydrazine for 30 minutes. After staining, the sections are rinsed in 30% alcohol if the nitrophenylhydrazines were used, or in the standard dilute sulfite bath followed by running tap water for 5 min. if leucofuchsin were used. Sections are routinely dehydrated, cleared, and covered. On examination, the sites of 1,2-glycol linkages will be stained violet by leucofushsin or yellow by the nitrophenylhydrazines.     

Histochemical Use of Sodium Bismuthate

Histochemical 1,2-glycoI cleavage, similar to that obtained with periodic acid and lead tetraacetate, may be obtained with sodium bismuthate. Routinely prepared slide sections, from tissues fixed in 10% formalin, are run down through xylene and graded alcohols to water and then oxidized for three minutes in a 1% sodium bismuthate 20% aqueous phosphoric acid solution. The oxidizing solution must be freshly prepared and used immediately. Following oxidation, sections are rinsed 15 sec. in IN HC1 to remove bismuth pentoxide precipitate, a by-product of the reaction. The sections are then washed in distilled water and placed in leuco-fushsin for 10 min., or in a saturated 30%) alcoholic solution of p-nitrophenylhydrazine for 5 min. or 2,4-dinitrophenylhydrazine for 30 minutes. After staining, the sections are rinsed in 30% alcohol if the nitrophenylhydrazines were used, or in the standard dilute sulfite bath followed by running tap water for 5 min. if leucofuchsin were used. Sections are routinely dehydrated, cleared, and covered. On examination, the sites of 1,2-glycol linkages will be stained violet by leucofushsin or yellow by the nitrophenylhydrazines.     

Protective Paraffin Infiltration of Soft Tissues in Insects to Facilitate Softening of Hard Exoskeletons

This technique can produce serial sections as thin as 5 μ from hard chitin-covered materials of insects or other arthropods. Procedures: Fix with alcoholic Bouin's fluid for 3 hr. Henceforth subject material to partial vacuum in each step to ensure a final proper embedding. Wash with 80% ethanol 2 or 3 times for 2 hr or until the picric acid is largely removed. Dehydrate to 90% ethanol and give 2 changes of n-butanol 2 hr each, and one of a 1:1 n-butanol-paraffin mixture in 56-57° oven for 12 hr. Finally, use 2 baths of pure paraffin, 3 hr each, to complete the infiltration. After the last bath, withdraw the specimen from the paraffin, and remove the superficial paraffin, first mechanically and then with a xylene bath for 4 min. Rinse first with n-butanol, and afterwards with absolute ethanol, 2 min each. The compound eyes are protected with a paraffin covering, the specimen is hydrated with a 1% aqueous solution of detergent for 1 hr and then washed with running tap water. The material is treated with a concentrated sulfuric-nitric mixture (H₂SO₄:HNO₃) for 4 hr to eliminate the exoskeleton. After this treatment, the specimen is washed with running tap water for 12 hr, dehydrated with acetone and then bathed in a 2% solution of celloidin in ethyl acetate to form a protective artificial cuticle. This coating is hardened with 2 quick baths of chloroform, the specimen reembedded in paraffin, and the block cast for sectioning.     

Histological Localization of Microelectrode Placement in Brain by Ferrocyanide and Silver Staining

After recordings had been taken from a microelectrode used for mapping nerve impulses, a current of 100 μa from the positive pole of a direct current generator was run through the electrode for 5 sec while it was still in place. On terminating the experiment, in which the use of several electrodes was possible, 50-75 ml of a 1:1 mixture of 4% potassium ferrocyanide and 4% acetic acid was injected into each common carotid artery, and the brain left in situ for 0.5 hr. It was then removed and the electrode-bearing part fixed 5-6 hr in a 1:1 mixture of 40% formalin and 95% ethyl alcohol at 55 °C. This specimen was washed in running water 5-10 min, the electrodes removed and frozen sections of 40-80 μ cut and placed in 95% alcohol. Sections were stained 5-10 min at 25-30°C in 10% silver nitrate solution in 75-80% alcohol acidified by 3-4 drops of glacial acetic acid per 50 ml, washed 4-5 sec in each of 2 baths of 95% alcohol, and reduced while being agitated constantly in a 2% solution of pyrogallol and 6-7% formalin in 75-80% alcohol. Washing in 95% alcohol, clearing in clove oil or methyl salicylate followed by xylene and mounting in synthetic resin or balsam completed the process. Sites of electrolysis at the tips of electrodes (under magnification) were blue before silver staining and black after staining. Axons stained brown to black on a yellow background.     

Silver Protein Staining of Nerve Fibers in Celloidin Sections

Celloidin sections from formalin-fixed brain and spinal cord of primates are stored in 70% alcohol after cutting, soaked in 2% pyridine in 50% alcohol for 6-8 hr at 37 C, and transferred to 1% concentrated NH₄OH in 50% alcohol 15-18 hr at 20-25 C. After washing and flattening, the sections are transferred to 1% silver protein solution containing 30 ml of 0.2 M H₃BO₃/100 ml. Impregnation is accomplished in 50 ml screw-top jars, 50 mm in diameter, which are filled to a depth of 35 mm, and have 1 gm of copper foil, 0.002 inch thick added. The foil is folded in loose accordion-fashion, pierced and threaded, cleaned in 5% HNO₃, rinsed in distilled water, and suspended in the solution just above the sections by fastening the thread to the jar lid. The sections are impregnated for 24 hr at 37 C, rinsed in distilled water, reduced in a solution of 5% Na₂SO₃ and 1% hydroquinone for 10 min, washed in distilled water and toned in 0.2% gold chloride for 5 min. After rinsing in distilled water, the sections are transferred to 1% oxalic acid for 45-60 sec, washed in distilled water and placed in 5% Na₂S₂O₃ for 5 min. Sections are then washed, dehydrated to 95% alcohol, cleared in terpineol, followed by 3 changes in xylene, and mounted.     

Differentiation of Articular and Epiphysial Cartilage, Based on Resorcin-CrystaeL Violet and Picro-Fuchsin Staining of Chondroitin Sulphate and Keratosulphate

Formalin-fixed, decalcified knee joints of young vertebrates were embedded in paraffin wax and cut at 4 μ. Sections were stained in Harris' Haematoxylin, washed in tap water, then immersed in the following staining solution for 60 min: crystal violet, 1 gm; resorcin, 2 gm; distilled water, 100 ml; boiled for 3 min, with constant stirring. After adding 30 ml of 30% FeCl₃, it was boiled for 3 min more. The solution was filtered. The precipitate was washed oil with 50 ml of distilled water and 100 ml of absolute alcohol added. This was combined with the original filtrate and boiled for 5 min. The solution was filtered once more, the precipitate discarded and 2 ml of cone. HC1 added. After cooling, the solution was ready for use. Sections were then washed briefly in tap water, stained in van Gieson's picro-fuchsin for 2 min, and differentiated as they were dehydrated and brought to Xylene. The sections were mounted in a synthetic resin (D.P.X.). Articular type cartilage stains red and growth cartilage blue.     

A Six-Dye Staining Schedule for Sections of Mesquite and Other Desert Plants

Materials are fixed in FPA (formalin, 2; propionic acid, 1; 70% ethanol, 17). Paraffin sections on slides are brought to 50% ethanol and stained as follows: (1) in Bismarck brown Y, a 0.02% solution in 0.1% aqueous phenol, 10-30 min; wash 30 sec in 0.7% acetic acid, and wash in distilled water 20-30 sec; (2) in crystal violet, 1% in 70% ethanol alkalinized with 1 drop of 1 N NaOH per 100 ml, 12-35 min; wash 30-60 sec in tap water to remove excess stain, and rinse 0.5 sec in 70% ethanol; then mordant in I₂-KI, 1% each in 70% ethanol, 40 sec, and rinse in 70% ethanol 2-5 sec; (3) in a mixture containing 0.4% acid fuchsin and 0.6% crythrosin B in 70% ethanol about 0.5 sec; rinse in 70% ethanol 5-15 sec to remove excess red; dehydrate in 70%, 95%, and absolute ethanol, 2-3 sec each; (4) in fast green FCF, 0.5% in a mixture of equal parts of methyl cellosolve, absolute ethanol, and clove oil, 5-15 sec; rinse in a mixture of clove oil, 10 ml; absolute ethanol, 100 ml; and methyl cellosolve, 10 ml, 5-7 sec; (5) in orange G, 0.75 gm in a mixture of clove oil, 40 ml; absolute ethanol, 40 ml; and methyl cellosolve, 60 ml, 5-30 sec; rinse clean in a 1:1 mixture of xylene and absolute ethanol, 5-20 sec Complete the clearing in pure xylene, 3 changes, 1.5 min in each, and apply a cover glass with synthetic resin. Slides are agitated in all steps except Bismark brown Y, crystal violet, and the xylenes. Contrast and staining intensity are adjusted by varying staining times in the dye solutions.     

A Sudan Black B Myelin Stain for Peripheral Nerves

Blocks of fresh issue were fixed 2 or more days in: cobalt sulfate (or nitrate), 1 gm; distilled water, 80 ml; 10% calcium chloride, 10 ml; and formalin, 10 ml. The fixed tissue was washed thoroughly in tap water, embedded in gelatin, frozen sections cut, and mounted on slides with gelatin adhesive. The sections were stained 15-30 min in a saturated, filtered solution of Sudan black B in 70% alcohol, differentiated in 50% alcohol under microscopic observation, and a cover glass applied with glycerol-gelatin. In thick (50-100 μ) sections, myelin stained green to gray-green and this allowed easy differentiation between nerves and other tissue elements.     

Differentiated Staining of Osmium Tetroxide-Fixed, Vestopal-w-Embedded Tissue in thin Sections

Tissues were fixed at 20° C for 1 hr in 1% OsO₄, buffered at pH 7.4 with veronal-acetate (Palade's fixative), soaked 5 min in the same buffer without OsO₄, then dehydrated in buffer-acetone mixtures of 30, 50, 75 and 90% acetone content, and finally in anhydrous acetone. Infiltration was accomplished through Vestopal-W-acetone mixtures of 1:3, 1:1, 3:1 to undiluted Vestopal. After polymerisation at 60° C for 24 hr, 1-2 μ sections were cut, dried on slides without adhesive, and stained by any of the following methods. (1) Mayer's acid hemalum: Flood the slides with the staining solution and allow to stand at 20°C for 2-3 hr while the water of the solution evaporates; wash in distilled water, 2 min; differentiate in 1% HCl; rinse 1-2 sec in 10% NH,OH. (2) Iron-trioxyhematein (of Hansen): Apply the staining solution as in method 1; wash 3-5 min in 5% acetic acid; restain for 1-12 hr by flooding with a mixture consisting of staining solution, 2 parts, and 1 part of a 1:1 mixture of 2% acetic acid and 2% H₂SO₄ (observe under microscope for staining intensity); wash 2 min in distilled water and 1 hr in tap water. (3) Iron-hematoxylin (Heidenhain): Mordant 6 hr in 2.5% iron-alum solution; wash 1 min in distilled water; stain in 1% or 0.5% ripened hematoxylin for 3-12 br; differentiate 8 min in 2.5%, and 15 min in 1% iron-alum solution; wash 1 hr in tap water. (4) Aceto-carmine (Schneider): Stain 12-24 hr; wash 0.5-1.0 min in distilled water. (5) Picrofuchsin: Stain 24-48 hr in 1% acid fuchsin dissolved in saturated aqueous picric acid; differentiate for only 1-2 sec in 96% ethanol. (6) Modified Giemsa: Mix 640 ml of a solution of 9.08 gm KH₂PO₄ in 1000 ml of distilled water and 360 ml of a solution of 11.88 gm Na₂HPO₄-2H₂O in 1000 ml of distilled water. Soak sections in this buffer, 12 hr. Dissolve 1.0 gm of azur I in 125 ml of boiling distilled water; add 0.5 gm of methylene blue; filter and add hot distilled water until a volume of 250 ml is reached (solution “AM”). Dissolve 1.5 gm of eosin, yellowish, in 250 ml of hot distilled water; filter (solution “E”). Mix 1.5 ml of “AM” in 100 ml of buffer with 3 ml of “E” in 100 ml of buffer. Stain 12-24 hr. Differentiate 3 sec in 25 ml methyl benzoate in 75 ml dioxane; 3 sec in 35 ml methyl benzoate in 65 ml acetone; 3 sec in 30 ml acetone in 70 ml methyl benzoate; and 3 sec in 5 ml acetone in 95 ml methyl benzoate. Dehydrated sections may be covered in a neutral synthetic resin (Caedax was used).     

Azure a-Schiff, Alcian Blue, HIO4-Schiff, Naphthol Yellow S—A Sequential Staining Method for Paraffin Sections

Paraffin sections from tissue fixed 4-12 hr in 10% formalin containing 0.5% cetyl pyridinium chloride, and washed 2 hr, were stained as follows: (1) Hydrolyze in 5 N HCl at room temperature for 8.5-9 min, or use standard Feulgen hydrolysis at 60 C. (2) Stain in azure A-Schiff, 0.5% in bisulfite bleach (1 N HCl, 5; 10% Na₂S₂O₅, 5; and distilled water 90—parts by volume) for 10 min. (3) Place in bisulfite bleach 2 changes, 2 min each; wash in water, 1-2 min. (4) Stain in Alcian blue (0.1% in 0.01 2V HCl, pH 2.0) for 10 min. (5) Place in 0.01 N HCl for 2-3 min; wash in water for 1-2 min. (6) Oxidize in 0.5% HIO₄ for 5 min; wash in water, 1-2 min. (7) Stain in Schiff's leucofuchsiu, 10 min. (8) Treat with bisulfite bleach as in step 3; wash in running water, 10 min. (9) Stain in naphthol yellow S (0.01% in 1% acetic acid) for 1-2 min. (10) Place in 1% acetic acid for 2 min, dehydrate in tertiary butanol, clear and cover. Result: DNA is deep blue; acidic mucins are light blue; neutral polysaccharides, red to magenta; and proteins, yellow. Proper timing of the hydrolysis for the Feulgen reaction is the most critical step. Overhydrolysis results in green nuclei (staining by naphthol yellow S) whereas purplish nuclei are the results of insufficient hydrolysis.     

Silver Staining of Nerve Fibers in Cardiac Tissue

Fresh hearts of dog were perfused through the coronary vessels with 1000 ml. of fixative (chloral hydrate, 5 g. per 100 ml. of 70% ethyl alcohol) and blocks of tissue 2 × 5 mm. from epicardium to endocardium fixed 48 hours in the same fixative. The blocks were placed in 95% alcohol containing 0.3% addition of strong ammonia for 4 hours, followed by 2 changes of plain 95% alcohol of 1 hour each, then cleared and infiltrated with paraffin. Mounted sections 12-15 µ thick were incubated in 1% silver proteinate (obtained from Serumvertrieb, Marburg, Germany)² at 38° C. for 48 hours in the presence of 10 g. of 15 gauge copper wire per 200 ml. of solution. The slides were rinsed gently in 3 changes of distilled water for 2 minutes, 1 minute and 1 minute, respectively, and reduced in 1% hydroquinone and 5% sodium sulfite for 5 minutes. They were washed 5 minutes in tap water and 5 minutes in 2 changes of distilled water and toned 3-5 minutes in 0.25% gold chloride, rinsed in distilled water 10 seconds, reduced 10 seconds in 1 % oxalic acid, rinsed 1 minute, fixed in 5% sodium thiosulfate 5 minutes, washed in tap water through 3 changes, dehydrated, cleared and covered. All solutions were made with distilled water except where otherwise specified. The results gave good impregnation of fine nerve fibers without the usual confusing staining of reticular tissue.     

Adaptation of the Millon, Sudan Black B, and Periodic Acid-Schiff Technics for Block Staining of Insect Tissues

Spermatophores and reproductive systems of the beetle, Lytta nuttalli Say, fixed in Bouin's aqueous picroformol or buffered 10% neutral formol were stained in toto by the Millon, Sudan black B and periodic acid-Schiff reactions as follows. Millon: after excess fixative is removed in 70% ethanol, specimens are brought to water, stained in Millon's reagent at 60 C for 1 hr, rinsed in 2% aqueous nitric acid at 40-50 C, dehydrated rapidly, cleared, embedded and sectioned as usual. Sudan black B: specimens are taken to absolute ethanol, stained in a saturated solution of Sudan black B in absolute ethanol at room temperature for 24-48 hr, rinsed and cleared in xylene, embedded and sectioned. PAS: specimens are brought to water, oxidized in 0.5 aqueous HIO₄ at 37 C for 30 min, washed in 2 changes of water, stained in Schiif reagent at room temperature for 1 hr, rinsed in 3 changes of 0.5% aqueous potassium metabisulfite, washed in running water for 10-15 min, dehydrated, cleared, embedded and sectioned. All 3 methods produced their characteristic staining in specimens up to 3 mm thick     

Selective Silver Impregnation of Degenerating Axons In The Central Nervous System

Rat and rabbit brains containing surgical lesions of 5-10 days' duration were fixed in 10% formalin (neutralized with calcium carbonate) for 1 week to 6 months. Frozen sections (15-20 n) were rinsed and then soaked 7 minutes in a 1.7% solution of strong ammonia in distilled water. Subsequent treatment was as follows: rinse; 0.05% aqueous potassium permanganate 5-15 minutes; 0.5% aqueous potassium metabisulfite, 2 changes of 2.5 minutes each; wash thoroughly in 3 changes distilled water; 1.5% aqueous silver nitrate, 0.5-1.0 hr.; 1% citric acid, 5-10 sec.; 2 changes distilled water; 1% sodium thiosulfate, 30 see.; 3 changes distilled water. Each section is then processed separately. Ammoniacal silver solution (450 mg. silver nitrate in 10 ml. distilled water; add 5 ml. ethanol; let cool to room temperature; add 1 ml. strong ammonia water and 0.9 ml. of 2.5% aqueous sodium hydroxide), 0.5-1.0 min. with gentle agitation. Reduction of about 1 minute is accomplished in: distilled water, 45 ml.; ethanol, 5 ml.; 10% formalin, 1.5 ml.; 1% citric acid, 1.5 ml. Rinsing; 1% sodium thiosulfate, 10 sec.; thorough washing followed by dehydration through graded alcohol and 3 changes of xylene or toluene complete the staining process. Normal nerve fibers are slightly stained to unstained, degenerating fibers, black. The treatment in potassium permanganate is critical since too little favors overstaining of normal fibers and too much abolishes staining of degenerating fibers.     

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.