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.     

A method for staining pollen tubes in pistil

A quadruple staining procedure has been developed for staining pollen tubes in pistil. The staining mixture is made by adding the following in the order given: lactic acid, 80 ml; 1% aqueous malachite green, 4 ml; 1% aqueous acid fuchsin, 6 ml; 1% aqueous aniline blue, 4 ml; 1% orange G in 50% alcohol, 2 ml; and chloral hydrate, 5 g. Pistils are fixed for 6 hr in modified Carnoy's fluid (absolute alcohol:chloroform:glacial acetic acid 6:4:1), hydrated in descending alcohols, transferred to stain and held there for 24 hr at 45 +/- 2 C. They were then transferred to a clearing and softening fluid containing 78 ml lactic acid, 10 g phenol, 10 g chloral hydrate and 2 ml 1% orange G. The pistils were held there for 24 hr at 45 +/- 2 C, hydrolyzed in the clearing and softening fluid at 58 +/- 1 C for 30 min, then stored in lactic acid for later use or immediately mounted in a drop of medium containing equal parts of lactic acid and glycerol for examination. Pollen tubes are stained dark blue to bluish red and stylar tissue light green to light greenish blue. This stain permits pollen tubes to be traced even up to their entry into the micropyle.     

Differential Staining of Gastric Glandular Cells

Fundus of stomach is fixed in 10% formalin (aqueous), Bouin's fluid or 5% trichloracetic acid (aqueous). It is embedded in paraffin, and 7μ sections are cut, mounted, deparaffinized and passed to 70% alcohol and then stained as follows: Mordant 3 min. in saturated Bismarck brown in 70% alcohol. Rinse in 70% alcohol, pass to distilled water, then overstain (2 hr.) in aniline blue, 0.5% solution in 2.5% acetic acid (aqueous). Precipitate the anilin blue with 0.5 ml. of 0.1% methyl violet solution (aqueous) dropped on die slide. Leave on 2 min. or less. Wash and differentiate in 70% alcohol. (Parietal cells dark blue). Stain 30 min. in a mixture of hematein, 0.10g.; A1C13 cryst., 0.05g.; and 70% alcohol 50 ml., prepared just before use and not filtered. Rinse in 70% alcohol and differentiate with an alcoholic extract of saffron (2 g. saffron pistils in 100 ml. 90% alcohol at 60°C. for 6 hr.) while observing the progress of differentiation microscopically. Dehydrate by dropping a 0.1 % solution of acetic acid in absolute alcohol on the section for 30 sec., followed by pure absolute alcohol, xylene, and covering in balsam.     

Basic Two-Dye Stains for Epoxy-Embedded 0.3-1 μ Sections

Dyes used in the 3 methods recommended are: I, thionin and acridine orange (T-AO); II, Janus green and Darrow red (JG-DR); III, methyl green and methyl violet (MG-MV). The first 2 methods were two-solution stains, applied in sequence; the third, required only one solution since methyl violet is present in commercial methyl green. Staining solution and timing was as follows: Method I. 0.1% thionin in a 45% ethanolic solution of 0.01 N NaOH, 5 min at 70 C; rinsing in water and followed by 1 min in a 1% aqueous solution of acridine orange made up in 0.02 N NaOH, also at 70 C, then washed, and dried on slides. Method II. 0.5% Janus green in aqueous 0.05 N NaOH, 5 min at 70 C; rinsing in water then into 0.5% Darrow red in 0.05 N NaOH (aq.), 2 min at 70 C., washing, and drying on slides. Method III. 1% methyl green (commercial, unpurified) in 1% aqueous borax for 15-20 min at 20-25 C, washing and attaching to slides. All staining was performed by floating the sections on the staining solutions, all drying at 70 C, and mounting in a resinous medium. T-AO gave blue to violet cytoplasmic structures, darker nuclei which contrasted strongly with yellow connective tissue and the secretion of goblet cells. JG-DR resembled a hematoxylineosin stain, but by shortening the staining time in DR to 0.5-1 min, collagenous and elastic tissue retained more of the green dye. MG-MV gave dark green nuclei in light green cytoplasm, with collagenous and elastic tissues in blue to violet. As with most methods for staining ultrathin sections, thicknesses of less than 1 μ required longer staining times.     

Differential Staining of Gastric Glandular Cells

Fundus of stomach is fixed in 10% formalin (aqueous), Bouin's fluid or 5% trichloracetic acid (aqueous). It is embedded in paraffin, and 7μ sections are cut, mounted, deparaffinized and passed to 70% alcohol and then stained as follows: Mordant 3 min. in saturated Bismarck brown in 70% alcohol. Rinse in 70% alcohol, pass to distilled water, then overstain (2 hr.) in aniline blue, 0.5% solution in 2.5% acetic acid (aqueous). Precipitate the anilin blue with 0.5 ml. of 0.1% methyl violet solution (aqueous) dropped on die slide. Leave on 2 min. or less. Wash and differentiate in 70% alcohol. (Parietal cells dark blue). Stain 30 min. in a mixture of hematein, 0.10g.; A1C13 cryst., 0.05g.; and 70% alcohol 50 ml., prepared just before use and not filtered. Rinse in 70% alcohol and differentiate with an alcoholic extract of saffron (2 g. saffron pistils in 100 ml. 90% alcohol at 60°C. for 6 hr.) while observing the progress of differentiation microscopically. Dehydrate by dropping a 0.1 % solution of acetic acid in absolute alcohol on the section for 30 sec., followed by pure absolute alcohol, xylene, and covering in balsam.     

A Method for Staining Pollen Tubes in Pistil

A quadruple staining procedure has been developed for staining pollen tubes in pistil. The staining mixture is made by adding the following in the order given: lactic acid, 80 ml; 1% aqueous malachite green, 4 ml; 1% aqueous acid fuchsia, 6 ml; 1% aqueous aniline blue, 4 ml; 1 % orange G in 50% alcohol, 2 ml; and chloral hydrate, 5 g. Pistils are fixed for 6 hr in modified Carnoy's fluid (absolute alcohol:chloroform:glacial acetic acid 6:4:1), hydrated in descending alcohols, transferred to stain and held there for 24 hr at 45±2 C They were then transferred to a clearing and softening fluid containing 78 ml lactic acid, 10 g phenol, 10 g chloral hydrate and 2 ml 1% orange G. The pistils were held there for 24 hr at 45±2 C, hydrolyzed in the clearing and softening fluid at 58±1 C for SO min, then stored in lactic acid for later use or immediately mounted in a drop of medium containing equal parts of lactic acid and glycerol for examination. Pollen tubes are stained dark blue to bluish red and stylar tissue light green to light greenish blue. This stain permits pollen tubes to be traced even up to their entry into the micropyle.     

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.     

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).     

Staining and Mounting Helminths

The following procedure is recommended: Fix ces-todes and trematodes (while held flat between glass slides) 0.5-2.0 hr. in the following mixture: formalin, 15; acetic acid (gl.), 5; glycerol, 10; 95% ethyl alcohol, 24; distilled H₂O, 46; all proportions by volume. After freeing them from the slides, wash thoroughly in running water and stain immediately thereafter. Stock staining solution: ferric ammonium alum (violet cryst.), 2 g.; distilled H₂O (cold) 100 ml.; after solution, add 2 ml. concentrated H₂SO₄, bring to a boil; add 1 g. coelestin blue B (Nat. Aniline), boil 3-5 min.; cool and add 10 ml. absolute methyl alcohol and 10 ml. glycerol. Dilute 1 vol. with 3 vol. distilled H20 for use. Stain 5-30 min., depending on size of specimens. Wash with 2 changes 0.5 hr. each of distilled H₂O, then 50% isopropyl alcohol 12-16 hr., 50% isopropyl alcohol 2 hr., followed by graded isopropyl alcohol for dehydration. Ether: ethyl alcohol (equal parts), 1 hr., is followed by embedding in celloidin in a sheet just thick enough to cover the specimens. Trim embedded specimens and dehydrate with isopropyl alcohol, 80%, 90% and absolute. Clear in beechwood creosote. Mount in balsam with cover glasses that overlap the edges of the celloidin 1-2 mm. While drying at 37°C, refill edges of mount with fresh balsam as needed. When dry, remove excess balsam and ring the edges with ordinary gloss enamel paint.     

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.     

Toluidine Blue-Alizarin Red S Staining of Cartilage and Bone in Whole-Mount Skeletons in Vitro

Cartilage and bone of the developing skeleton can be reliably differentiated in whole-mount preparations with toluidine blue-alizarin red S staining after FAA fixation. The recommended staining procedure is based chiefly on the use of newborn white and Swiss-Webster mice, 4-9 days postnatal, but was tested also on mice and rats 3-8 wk of age. Procedure: Sacrifice, skin, eviscerate, remove body fat, and place specimens in FAA (formalin, 1; acetic acid, 1; 70% alcohol, 8) for approximately 40 min. Stain in 0.06% toluidine blue made in 70% ethyl alcohol for 48 hr at room temperature. Use 20 volumes of stain solution to the estimated volume of the specimen. Destain soft tissues in 35% ethyl alcohol, 20 hr; 50%, 28 hr; and 70%, 8 hr. Counterstain in a freshly prepared 1% aqueous solution of KOH to which is added 2-3 drops of 0.1% alizarin red S per 100 ml of solution. Each day for 3 days, transfer the specimen to a fresh 1% KOH-alizarin mixture, or until the bones have reached the desired intensity of red and soft tissues have cleared. Rinse in water, and place in a 1:1 mixture of glycerol and ethyl alcohol for 1-2 hr, then transfer the specimen to fresh glycerol-alcohol for final clearing and storage. Older mice and rats require procedural modifications: (1) fixation for 2 hr, (2) 0.12% toluidine blue, (3) maceration for 4 days in 3% KOH-alizarin, and (4) preliminary clearing for 24 hr in a mixture of glycerol, 2; 70% ethyl alcohol, 2; and benzyl alcohol, 1 (v/v) before placing in a 1:1 alcohol-glycerol mixture.     

Epon-Embedded Tissue Stained with Aldehyde-Fuchsin for Radioautography

Four-week-old Holtzman rats were injected intraperitoneally with 20 μCi ¹²⁵I. Six or eight weeks later, they were killed by intracardiac perfusion with glutaraldehyde; thyroid and adrenal glands were excised, postfixed in osmic acid, and embedded in Epon. Steps in the staining procedure of 0.5-1 μm thick sections are: oxidation in 0.3% potassium permanganate in 0.625% sulfuric acid, 2-5 min at 70 C; brief rinse; bleaching with 2.5% NaHSO₃, 4-5 min; brief r-utse; let dry completely; aldehydefuchsm, 15-20 min at 50 C; 95% alcohol; rinse in absolute alcohol; let dry completely. seaions were coated with Kodak NTB2 emulsion and exposed for 3 to 8 weeks. Results indicate that (1) tissues are well stained even after an 8-week exposure, (2) aldehyde-fuchsin pduces no chemographic effect, and (3) structures underneath the emulsion are easily identified.     

A One-Solution Triacid General Stain Which Differentiates Oxygenated from Non-Oxygenated Red Blood Corpuscles

Tissues from representative mammals, amphibia and invertebrates were fixed for 5-24 hr in either an aqueous solution of 8% p-toluene sulfonic acid (PTSA) or in 10% formalin to which 5 gm PTSA/100 ml had been added, and processed through embedding in polyethylene glycol 400 distearate in the usual manner. Sections cut at 4-6 μ were floated on 0.2% gelatin containing 1.25% formalin, and spread and dried on slides at a temperature not exceeding 25 C. Wax was removed with xylene, and the sections brought to water through ethanol as usual. The working staining solution was made from three stock solutions: A. Chlorantine fast blue 2RLL, 0.5%; B. Cibacron turquoise blue G-E, 0.5%; C. Procion red M-P, 0.5%—each of which was dissolved in 98.5 ml of distilled water to which 0.5 ml of glacial acetic acid and 0.5 ml of propylene glycol monophenyl ether (a fungicide) had been added. For use, the three solutions were mixed in the proportions: A, 3; B, 4; and C, 3 volumes. Staining time was uncritical, 10-30 min usually sufficing for 6 μ, sections. The chief feature of the staining is the differentiation of oxygenated and nonoxygenated red blood corpuscles, in reds and blues respectively. Connective tissue stained blue or blue-green and mucin, green. Nuclei and cytoplasm stain according to their condition at the time of fixation. The mixed stain keeps well, remaining active after 2 yr of storage.     

Haematoxylin-Phloxine-Alcian Blue-Orange G Differential Staining of Prekeratin, Keratin and Mucin

This is a modification of Kreyberg's stain with Alcian blue 8GS used to stain acid much while phloxine B and orange G stain keratin and prekeratin. Procedure: Dewax formalin-fixed paraffin sections in xylene and hydrate through alcohol. Stain in Mayer's haemalum, 10 min; blue in tap water; wash in distilled water; stain in 1% phloxine, 3 min; wash in running water, 1 min; wash in distilled water; stain in 0.5% aqueous Alcian blue in 0.5 acetic acid, 5 min; wash in distilled water; stain in 0.5% orange G dissolved in 2.0% phosphotungstic acid, 13 min; dehydrate quickly in 2 changes of 95% alcohol and 2 changes of absolute alcohol; clear in several changes of xylene; mount in a synthetic resin. Acid mucopolysaccharides are stained turquois blue; prekeratin and keratin are orange to red orange.     

Nile Blue Histochemical Method for Phospholipids

The technic recommended is: Fix 6-12 hr. in 10% formalin containing 1% CaCl₂. Cut frozen sections without embedding or after gelatin or carbowax. Stain 90 min. at 60°C. in saturated aqueous Nile blue sulfate, 500 ml. plus 50 ml. of 0.5% H₂SO₄, boiled 2 hr. before use. Rinse in distilled water, and place in acetone heated to 50°C. Remove the acetone from the source of heat and allow the sections to remain 30 min. Differentiate in 5% acetic acid 30 min., rinse in distilled water, and refine the differentiation in 0.5% HCl for 3 min. Wash in several changes of distilled water and mount in glycerol jelly. Results: phospholipids - blue; everything else - unstained. Counterstaining nuclei with safranin is optional, but if done, it preferably precedes the Nile blue and is then differentiated by the acetic acid. The histochemical principles on which the method is based are as follows: (1) The calcium compounds of phospholipids combine with the oxazine form of Nile blue sulfate and survive subsequent treatment; (2) neutral lipids are dissolved out by acetone; (3) proteins and other interfering substances are destained by the acetic acid and hydrochloric acid baths.     

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.     

Staining Paraffin Sections with Protargol 6. Impregnation and Differentiation of Nerve Fibers in Adrenal Glands of Mammals

A series of experiments with protargol staining of nerve fibers in mammalian adrenal glands has yielded the following procedure: Fix-1-2 days in a mixture of formamide (Eastman Kodak Company) 10 cc, chloral hydrate 5 g., and 50% ethyl alcohol 90 cc. Wash, dehydrate and embed in paraffin. Cut sections about 15 and mount on slides. Remove the paraffin and run down to distilled water. Mordant 1-2 days in a 1% aqueous solution of thallous (or lead) nitrate at 56-60°C. Wash thru several changes of distilled water and impregnate in 1% aqueous protargol (Winthrop Chemical Company) at 37-40°C. for 1 to 2 days. Rinse quickly in distilled water and differentiate 7-15 seconds in a 0.1% aqueous solution of oxalic acid. Rinse thru several changes of distilled water for a total time of 0.5 to 1.0 rain. Reduce 3-5 rain, in Bodian's reducer: hydroquinone 1 g., sodium sulfite 5 g., distilled water 100 cc. Wash in running water 3-5 min. and tone 5-10 min. in a 0.2% gold chloride solution. Wash 0.5 min. or more and reduce in a 2% oxalic acid solution to which has been added strong formalin, 1 cc. per 100. (Caution. This last reduction is critical and over-reduction can spoil an otherwise good stain; 15-30 seconds usually suffices, and the sections should show only the beginning of darkening to a purplish or gray color.) Wash, fix in hypo, wash, dehydrate and cover.     

Staining Sex Chromatin; Biebrich Scarlet and Fast Green Fcf as a Mixture Versus Their Use in Sequence

Human skin was fixed in Davidson's solution (95% alcohol, 35; formalin, 20; glacial acetic acid, 10; and distilled water, 35—parts by volume) and sections prepared through paraffin embedding in the usual manner. Stock stains were: I(BS)—Biebrich scarlet, 1 gm in 100 ml of 50% alcohol to which 0.3 gm of phosphotungstic acid and 5 ml of glacial acetic acid were added—and II(FG)—fast green, 0.5 gm in 85 ml of 50% alcohol to which 0.3 gm of phosphotungstic acid, 0.3 gm of phosphomolybdic acid, and 15 ml of glacial acetic acid were added. Experimental staining solutions were prepared in the following proportions of stock BS to stock FG—1:1, 2:1, 3:1, 1:2 and 1:3. Sections were brought to 50% alcohol and stained for 15, 20, 25 and 30 min in each of the five BS-FG mixtures, rinsed in 50% alcohol, then dehydrated in 70%, 95%, and absolute alcohol, 2 min each; cleared in xylene, and covered in balsam. The 2:1 (optimum proportion) combination of BS with FG, acting for 20 min, yielded 97% sex chromatin-positive nuclei in female material. If sections were stained in stock solution BS for 2 min, they could be differentiated by a 20 min treatment in the mordanting component of stock FG (without dye) to give a one-color stain. Such stains gave about the same percentage of sex chromatin-positive nuclei as those obtained by the regular two-color procedure. These modifications are simpler, more rapid, and yield results comparable to previously employed techniques.     

Staining Sex Chromatin; Biebrich Scarlet and Fast Green Fcf as a Mixture Versus Their Use in Sequence

Human skin was fixed in Davidson's solution (95% alcohol, 35; formalin, 20; glacial acetic acid, 10; and distilled water, 35—parts by volume) and sections prepared through paraffin embedding in the usual manner. Stock stains were: I(BS)—Biebrich scarlet, 1 gm in 100 ml of 50% alcohol to which 0.3 gm of phosphotungstic acid and 5 ml of glacial acetic acid were added—and II(FG)—fast green, 0.5 gm in 85 ml of 50% alcohol to which 0.3 gm of phosphotungstic acid, 0.3 gm of phosphomolybdic acid, and 15 ml of glacial acetic acid were added. Experimental staining solutions were prepared in the following proportions of stock BS to stock FG—1:1, 2:1, 3:1, 1:2 and 1:3. Sections were brought to 50% alcohol and stained for 15, 20, 25 and 30 min in each of the five BS-FG mixtures, rinsed in 50% alcohol, then dehydrated in 70%, 95%, and absolute alcohol, 2 min each; cleared in xylene, and covered in balsam. The 2:1 (optimum proportion) combination of BS with FG, acting for 20 min, yielded 97% sex chromatin-positive nuclei in female material. If sections were stained in stock solution BS for 2 min, they could be differentiated by a 20 min treatment in the mordanting component of stock FG (without dye) to give a one-color stain. Such stains gave about the same percentage of sex chromatin-positive nuclei as those obtained by the regular two-color procedure. These modifications are simpler, more rapid, and yield results comparable to previously employed techniques.     

Hematoxylin-Eosin Staining of OsO4-Fixed Epon- Embedded Tissue; Prestaining Oxidation by Acidified H2O2

Successful application of hematoxylin-eosin staining to 0.5-1 μ sections of OsO₄-fixed Epon-embedded mammalian tissue is made possible by first treating the sections for approximately 1 min at 25-30 C with 10% H₂O₂ acidified with 0.1 or 0.01 N H₂SO₄ to pH 3.2. Subsequent steps are: washing; drying; Hams hematoxylin at 50 C, 1-2 min; washing; drying; 0.2-0.3% NH₄OH in 70% ethanol, 3-5 sec, drying at 50 C; 5% aqueous eosin for 3 & 45 sec at 25-30 C, washing; drying; clearing in xylene and mounting in resin. The use of acidified H₂O₂ prevents the staining of Epon and permits the characteristic staining picture to be obtained. Sections were attached to glass slides without adhesive and processed horizontally on a rack. Slides should be well drained and blotted before each drying step, to prevent formation of precipitate on the section.