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).     

Counterstaining Golgi-Cox Impregnations with Luxol Fast Blue as a Myelin Stain

Immerse pieces of brain tissue 4 wk in solutions A and B, mixed just before use: A. K₂Cr₂O₇, 1 gm; HgCl₂, 1 gm; boiling distilled water, 85 ml. Boil A for 15 min, cool to 2 C and add: B. K₂CrO₄, 0.8 gm; Na₂WO₄, 0.5 gm; distilled water, 20 ml. Rinse in water and immerse 24 hr in LiOH, 0.5 gm; KNO₃, 15 gm; distilled water, 100 ml. Wash 24 hr in several changes of 0.2% acetic acid and then for 2 hr in tap water. Dehydrate and embed in celloidin. Process a 60 μ section through 70 and 95% ethanol, a 3:1 mixture of absolute ethanol and chloroform, and toluene. Immerse it for 5 min in a solution containing methyl benzoate, 25 ml; benzyl alcohol, 100 ml; chloroform, 75 ml. Orient the section on a chemically clean slide and let air-dry 5-10 min. Process through toluene, 3:1 ethanol-chloroform and 95% ethanol. Place the section for 5-60 min at 60 C in a solution made up of: Luxol fast blue G (Matheson, Coleman and Bell), 1 gm; 95% ethanol, 1000 ml; 10% acetic acid, 5 ml. Hydrate to water and immerse in 0.05% Li₂CO₃ for 3-4 min. Differentiate in 70% ethanol and place in water. Immerse for 5-15 min in a mixture of two solutions: A. cresylechtviolet (Otto C. Watzka, Montreal), 2 gm; 1 M acetic acid, 185 ml; B. 1 M sodium acetate, 15 ml; distilled water, 400 ml; absolute ethanol, 200 ml. Dehydrate to 3:1 ethanol-chloroform. Clear in toluene and apply a coverslip. The technique produces fast Golgi-Cox impregnated neurons against a background of counterstained myelinated fibers. Patterns of the myelinated fibers can be used to localize impregnated neurons.     

Staining Tissue of the Central Nervous System with Luxol Fast Blue and Neutral Red

The tissue is fixed in 10% neutral saline formalin for 1 day to 3 wk depending on the size of the block, dehydrated and embedded in paraffin. The sections are stained at 57° C for 2 hr, then at 22° C for 30 min, in a 0.0125% solution of Luxol fast blue in 95% alcohol acidified by 0.1% acetic acid. They are differentiated in a solution consisting of: Li₂CO₃, 5.0 gm; LiOH-H₂O, 0.01 gm; and distilled water, 1 liter at 0-1° C, followed by 70% alcohol, and then treated with 0.2% NaHSO₃. They are soaked 1 min in an acetic acid-sodium acetate buffer 0.1 N, pH 5.6, then stained with 0.03% buffered aqueous neutral red. Sections are washed in distilled water, 1 sec, then treated with the following solution: CuSO₄·5H₂O, 0.5 gm; CrK(SO₄)₂·12H₂O, 0.5 gm; 10% acetic acid, 3 ml; and distilled water, 250 ml. Dehydration, clearing and covering complete the process. Myelin sheaths are stained bright blue; meninges and the adventitia of blood vessels are blue; red blood cells are green. Nissl material is stained brilliant red; axon hillocks, axis cylinders, ependyma, nuclei and some cytoplasm of neuroglia, media and endothelium of blood vessels are pink.     

Mercuric Bromphenol Blue Staining of Precipitin Lines in Agar

Lines formed by antibody-organ antigen reactions are stained particularly well by a modification utilizing the mercuric bromphenol blue (MBB) mixture of Mazia et al. (Biol. Bull., 104: 57-67, 1953). The agar covered slides are placed overnight in 0.85% NaCI at 4 C, followed by washing for 2 hr in 0.85% NaCI at 25 C. They are then rinsed for 10 min in distilled water, and dried overnight at 37 C. The precipitin lines are fixed by immersing the slides for 25 min in 95% alcohol, followed by 5 min hydration in distilled water. They are stained for 25 min in MBB mixture (HgCI₂, 10 gm; bromphenol blue, 0.1 gm; 95% ethanol, 100 ml). Excess stain is removed by immersing in acidified alcohol (95% ethanol, 98 ml; glacial acetic acid, 2 ml). Finally, the slides are passed through alcohol and xylene, and resin-mounted under coverslips.     

Handling Germinated Pollen on Millipore Membrane During the Feulgen and Autoradiographic Procedures

To prevent loss of pollen during the Feulgen's procedure, the pollen was grown on an autoclaved membrane filter (Millipore AA WP 025 00) in contact with a sterilized medium containing agar 0.5-1%, sucrose according to the genus (Malus 0.3-0.5 M; Persica and Tulipa 0.4 M), and H₃BO₃, 0.01%. To fix the germinated pollen of most species, the membrane was placed for 2 hr to overnight at 2-4 C on filter paper wet with the following mixture: OsO₄, 1 gm; CrO₃, 1.66 gm; and distilled water, 233 ml. To fix Persica pollen, 10% of glacial acetic acid had to be added to the fixative. Washing with distilled water and bleaching with a mixture of 3% H₂O₂ and sat. aq. ammonium oxalate, 1:1, were performed also on filter paper. Similarly, the preparation was processed for Feulgen staining by use of pieces of filter paper wet with the required fluids. Hydrolysis preceding the Schiff's reagent was performed at room temperature with 5 N HCl for 18 min. The differentiation after the Schiff's action was with 2% K₂S₂O₅ buffered to pH 2.3 with 9 ml of phosphate buffer (KH₂PO₄, 1.4 gm; conc. HCl, 0.35 ml and distilled water to make 100 ml). The stained pollen was floated off the membrane with a drop of glacial acetic acid to a gelatinized or an albumenized slide, and squashed. When the coverslip is removed the preparation may be either dehydrated and mounted or coated with autoradiographic film.     

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.     

Use of Gallocyanin as a Myelin Stain for Brain and Spinal Cord

Tissue fixed in 10% formalin, formol saline, CaCO₃ or phosphate buffer neutralized formalin, Baker's formol calcium, Cajal's formol ammonium bromide, formalin-95% ethanol 1:9, formalin-methanol 1:9, Lillie's methanol-chloroform or Salthouse's formol cetyltrimethylammonium bromide was dehydrated and embedded in paraffin. Sections were attached to slides with either albumen or gelatine adhesive and processed throughout at room temperature of 22-25 C. Mordanting 30-60 min in 1% iron alum was followed by a 10 min wash in 4 changes of distilled water. Myelin was stained in a gallocyanin self-differentiating solution for 1-2.5 hr; thick sections requiring the longer time. The staining solution (pH approximately 7.4) consisted of Na₂CO₃, 90 mg; distilled water, 100 ml; gallocyanin, 250 mg; and ethanol, 5 ml. The ethanol was added to this mixture last, and after the other ingredients had been boiled and then cooled to room temperature. After a staining and thorough washing, Nissl granules were stained for 5-10 min in a solution consisting of: 0.1 M acetic acid, 60 ml; 0.1 M sodium acetate, 40 ml; methyl green, 500 mg. Washing, dehydration, clearing and mounting completed the process. Myelin sheaths were stained dark violet; neuronal nuclei, light green with dark granules of chromatin; nucleoli of motor cells and erythrocytes, dark violet; cytoplasm, green with dark green Nissl granules. The simple and reliable method can be adapted easily for use with automatic tissue processors.     

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.     

A Phloxine-Azure-Hematoxylin Sequence for Differential Staining of Cells in Pancreatic Islets

Sections of 6 μ from tissues fixed in Susa or in Bouin's fluid (without acetic acid) and embedded in paraffin were attached to slides with Mayer's albumen, dried at 37 C for 12 hr, deparaffinized and hydrated. The sections fixed in Susa were transferred to a I₂-K₁ solution (1:2:300 ml of water); rinsed in water, decolorized in 5% Na₂S₂O₃; washed in running water, and rinsed in distilled water. Those fixed in Bouin's were transferred to 80% alcohol until decolorized, then rinsed in distilled water. All sections were stained in 1% aqueous phloxine, 10 min; rinsed in distilled water and transferred to 3% aqueous phosphotungstic acid, 1 min; rinsed in distilled water; stained 0.5 min in 0.05 azure II (Merck), washed in water; and finally, nuclear staining in Weigert's hematoxylin for 1 min was followed by a rinse in distilled water, rapid dehydration through alcohols, clearing in xylene and covering in balsam or a synthetic resin. In the completed stain, islet cells appear as follows: A cells, purple; B cells, weakly violet-blue; D cells, light blue with evident granules; exocrine cells, grayish blue with red granules.     

Pyronin Y in the Methyl-Green-Pyronin Histological Stain

Two samples of pyronin Y were found which, with the exception of eosinophilic granules and osteoid, stained only nucleic acids in animal tissues. Good differentiation was obtained. with n-butyl alcohol. It was therefore possible to prepare a differentially staining mixture of either of these pyronins combined with methyl green. This mixture stains polymerized desoxyribose nucleic acid (DNA) clear green, depolymerized DNA and ribonucleic acid red. The red staining of eosinophilic granules and osteoid is readily distinguished by its persistence after ribonuclease or warm-buffer extraction. The staining mixture consists of: (1) pyronin Y (Edward Gurr or G. T. Gurr), CHCl₃ extracted, 2% aq, 12.5 ml; (2) methyl green, CHCl₃ extracted, 2% aq, 7.5 ml; (3) distilled water, 30 ml. The staining procedure is as follows. (1) Immerse slides 6 min in the dye mixture. (2) Blot with filter paper. (3) Immerse in 2 changes of n-butyl alcohol, 5 min each. (4) Xylene, 5 min. (5) Cedar oil, 5 min. (6) Apply Permount and cover.     

Application of Methods for Mammalian Chromosome Spreads to the Cells of Cestodes

Tapeworm cells obtained by physical maceration between ground-glass surfaces are incubated for 3 hr in Hanks' balanced salt solution (BSS) supplemented with colchicine to a concentration of 10-⁴ M. After washing in BSS, the cells are incubated for 10 min in 1/4 strength BSS then centrifuged 10 min. Fixation of the intact button of cells (or alternatively, by squirting the cells directly into the fixative) in Carnoy's alcohol-chloroform-acetic acid (6;3:1) for 30 min follows, and cells, dispersed and washed in the fixative, are flattened by dropping the suspension on clean, water-wet slides which are then air-dried and stained with Giemsa diluted 1 ml;47 ml with distilled water to which 2 ml of buffer—M/15 KH₂PO₄, 32 ml, mixed with M/15 Na₂HPO₄, 68 ml—is added. After staining 15 min and washing in distilled water, slides are air-dried and mounted with resin. Well separated and well stained chromosomes have resulted.     

The Quantitative Determination of Osmium Tetroxide in Fixatives

This rapid spectrophotometric method for determining the OsO₄ concentration in fixative and stock solutions is based on the reduction of OsO₄ by acidified KI to the blue species of OsI₆ =, which is then determined at 649 mµ. The salt K₂OsI₆ has been isolated from the reaction mixture and characterized. Method: A I ml aliquot of the solution, containing up to 3% OsO₄, is diluted to 100 ml with distilled water. To 1 ml of the diluted solution is added, in order: distilled water, 2 ml; 1 M HCI, 1 ml; and 1 M KI, 1 ml. Optical density at 649 mµ is read from 10-120 min thereafter. OsO₄ concentration is calculated from the measured molecular extinction coefficient of OsI₆ =, 4400 liter/mole cm.     

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.     

Adaptation of the Nauta Method for Use on Degenerating Fibers in Rat Cortex and Thalamus

Lesions produced in the cerebral cortex of rats were studied by Nauta's method for degeneration. The brains were perfused with physiological NaCl solution, followed by 10% neutral (CaCO₃) formalin. The brains were removed and stored in the formalin for 2 wk to 1 yr. Experimental modifications of the staining method showed that its sensitivity for fine degenerating fibers could he enhanced by the following changes: (a) omitting 0.05% potassium permanganate; (b) replacing the hydroquinone-oxalic acid mixture with 0.1% pyrogallol. Procedure: (1) frozen sections to water; (2) 0.5% phosphomolybdic acid, 45 min; (3) distilled water, 1 min; (4) 0.1% pyrogallol (aq.), 2 min; (5) distilled water, 3 washes of 1 min each; (6) 1.5% silver nitrate (aq.), 30 min; (7) distilled water, 1 min, (8) Laidlaw's ammoniated silver carbonate, 10110 sec; (9) Nauta's reducer, 1-2 min; (10) distilled water, 1 min; (11) 1.0% Na₂S₂O₃, 2 min; (12) distilled water 3 changes, 1 min each; (13) dehydrate, clear, and cover. This method gave equally good results on degenerating axons in both cortex and thalamus.     

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.     

A Protargol Method for Staining Nerve Fibers in Frozen or Celloidin Sections

Extensive experimentation with protargol staining of neurons in celloidin and frozen sections of organs has resulted in the following technic: Fix tissue in 10% aqueous formalin. Cut celloidin sections IS to 25 μ, frozen sections 25 to 40 μ. Place sections for 24 hours in 50% alcohol to which 1% by volume of NH₄OH has been added. Transfer the sections directly into a 1% aqueous solution of protargol, containing 0.2 to 0.3 g. of electrolytic copper foil which has been coated with a 0.5% solution of celloidin, and allow to stand for 6 to 8 hours at 37° C. Caution: In this and the succeeding step the sections must not be allowed to come in contact with the copper. From aqueous protargol, place the sections for 24 to 48 hours at 37° C. directly into a pyridinated solution of alcoholic protargol (1.0% aqueous solution protargol, 50 ml.; 95% alcohol, 50 ml.; pyridine, 0.5 to 2.0 ml.), containing 0.2 to 0.3 g. of coated copper. Rinse briefly in 50% alcohol and reduce 10 min. in an alkaline hydroquinone reducer (H₃BO₃, 1.4 g.; Na₂SO₃, anhydrous, 2.0 g.; hydroquinone, 0.3 g.; distilled water, 85 cc; acetone, 15 ml.). Wash thoroly in water and tone for 10 min. in 0.2% aqueous gold chloride, acidified with acetic acid. Wash in distilled water and reduce for 1 to 3 min. in 2% aqueous oxalic acid. Quickly rinse in distilled water and treat the sections 3 to 5 min. with 5% aqueous Na₂S₂O₃+5H₂O. Wash in water and stain overnight in Einarson's gallocyanin. Wash thoroly in water and place in 5% aqueous phosphotungstic acid for 30 min. From phosphotungstic acid transfer directly to a dilution (stock solution, 20 ml.; distilled water, 30 ml.) of the following stock staining solution: anilin blue, 0.01 g.; fast green FCF, 0.5 g.; orange G, 2.0 g.; distilled water, 92.0 ml.; glacial acetic acid, 8 ml.) and stain for 1 hour. Differentiate with 70% and 95% alcohol; pass the sections thru butyl alcohol and cedar oil; mount.     

Brazilin-Toluidine Blue O and Hematoxylin-Darrow Red Methods for Brain and Spinal Cord

Tissue fixed in 10% formalin, formalin-95% ethanol 1:s CaCO₂ or phosphate buffer neutralized formalin, or methanol-chloroform 2:1, was dehydrated and embedded in paraffin or double-embedded by infiltration in 1% celloidin followed by a chloroform-paraffin sequence. Sections were attached to slides with either albumen or gelatine adhesive and processed throughout at room temperature of 24-26 C. For either method, mordanting 30-60 min in 1% iron alum was followed by a 10 min wash in 4 changes of distilled water. For brazilin-toluidne blue O, myelin was stained for 20-60 min, depending upon section thickness, in a self-differentiating solution consisting of: 0.15% Li₂CO₃ 75 ml; 6% brazilin in 95% ethanol, 25 ml; and NaIO₃ 75 mg. After a thorough washing, Nissl material was stained for 3-8 min in a solution consisting of: 0.1 M acetic acid, 90 ml; 0.1 M sodium acetate, 10 ml; and 1% toluidine blue 0, 2.5 ml. For hematoxylin-Darrow red, myelin was stained for 2-6 hr in a self-differentiating solution consisting of: 0.15% Li₂,CO₃ 95 ml; 10% hematoxylin in 95% ethanol, 5 ml; and NaIO₃ 25 mg. After a thorough washing, Nissl material was stained for 20 min or less in a solution consisting of: 0.1 M acetic acid, 90 ml; 0.1 M sodium acetate, 10 ml; Darrow red, 25 mg. This mixture was first boiled, cooled to room temperature and filtered. In both methods, washing, dehydration, clearing, and mounting completed the process. In the brazilin-toluidine blue technic, myelin sheaths were stained reddish purple; neuronal nuclei light blue with dark granules of chromatin; nucleoli dark blue; and cytoplasm blue with dark blue Nissl granules. In the hematoxylin-Darrow red procedure, myelin sheaths were blue-black; nuclei light red with dark granules of chromatin; nucleoli almost black; and cytoplasm red with bright red Nissl granules.     

Bromphenol Blue as A Stain for Muscle Striations

Sections from formalin-fixed muscle are stained 4-24 hr with a 0.05% solution of bromphenol blue in 2% acetic acid, rinsed with water and placed in 0.5% acetic acid for 5-10 min. They are then treated 30 sec with tap water substitute (KHCO₃, 0.2 gm; MgSO₄, 2 gm; distilled water, 100 ml), rinsed, dehydrated in alcohol, cleared in xylene and covered in a polystyrene mountant Striatums of both cardiac and skeletal muscle fibers are fully resolved under oil immersion, against the blue background of the other parts of the fibers.     

Staining Reticulin with Gold

This bromine-iodine-gold chloride-reduction sequence stains reticulin in formalin-fixed paraffin sections without risk of sections becoming detached. After hydration, sections are exposed to 0.2% bromine water containing 0.01% KBr for 1 hr, then rinsed and placed for 5 min in a solution consisting of KI, 2 gm; iodine crystals, 1 gm; and distilled water, 100 ml. After this the sections are well washed in distilled water, immersed for 5 min in 1% w/v aqueous solution of chloro-auric acid, again rinsed in distilled water, and the gold is reduced by placing in freshly made 3% H₂O₂ for 2-4 hr at 37 C, or in 2% oxalic acid for 1-3 hr at the same temperature.     

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.