Myelin impregnation: an improved Golgi-Cox modification.

Optic tecta of goldfish were coated with egg yolk and immersed for only one week in one of the following impregnation fluids: a) Solution A + B; A = 1 g K2Cr2O7 and 1 g HgCl2 boiled for 15 min in 85 ml distilled water and allowed to cool; B = 0.8 g K2Cr2O4 and 0.5 g KWO4 dissolved in 20 ml distilled water. b) Solution A + B two volumes diluted with boiled distilled water. c) Solution A + B four volumes diluted with boiled distilled water. Each tectum was immersed 6 hr in 100 ml distilled water containing 0.5 g LiOH and 15 g KNO3, washed 18 hr in 500 ml 0.2% acetic acid, dehydrated with ethanol, and embedded in low viscosity nitro cellulose. Sections were cut at 100 micron with a rotary microtome after clearing with cedarwood oil. Methods b) and c) have two advantages compared with method a), the original Golgi-Cox method. First, more cells are impregnated, especially in the layers extending 200-400 micron below the surface, and dendrites as well as unmyelinated axons are well impregnated. Second, myelin sheaths are impregnated and can be recognized by their peculiar chain-like appearance. The described Golgi-Cox modification offers an appropriate method to study the morphology of superficially located nervous tissue.     

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.     

A Tungstate Modification of the Golgi-Cox Method

Pieces of fresh nervous tissue 4-5 mm thick are put into the following solution: HgCl₂, 1 gm; K₂Cr₂O₇, 1 gm; K₂CrO₄, 0.8 gm; K₂WO₄ (or Na₂WO₄), 0.5 gm; distilled water 100 ml. They are kept undisturbed in the dark at room temperature for 20-30 days, then transferred to the following alkaline solution: LiOH (or NaOH), 1 gm; KNO₃, 15 gm; distilled water, 100 ml. After 12-24 hr in this solution they are washed for 12-24 hr in several changes of distilled water. (If sodium hydroxide was used, 0.5 ml of acetic acid should be added per 100 ml of wash water.) Embedding in celloidin follows dehydration. Sections are dehydrated in 3 parts of absolute alcohol and 1 part of chloroform, cleared in iodobenzene and mounted with a cover slip using a mounting medium with a refractive index around 1.61. The use of tungstate improves the general results and allows especially successful impregnations in very young animals, when the usual technic fails.     

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.     

Zinc Chloride Methylene Blue, Ii. Preparation of Giesma and Wright Type Low Azure B Blood Stains from Dichromate Oxidized Commercial Dye

TO determine the amount of K₂Cr₂O₇ required to produce optimal Giemsa type staining, six 1 g amounts (corrected for dye content) of zinc methylene blue were oxidized with graded quantities of K₂Cr₂O₇ to produce 4, 8, 12, 16, 20 and 24% conversion of methylene blue to azure B. These were heated with a blank control 15 minutes at 100 C in 60-65 ml 0.4 N HCI. cooled, and adjusted to 50 ml to give 20 mg original dye/ml. Aliquots were then diluted to 1% and stains were made by the “Wet Giemsa” technic (Lillie and Donaldson 1979) using 6 ml 1% polychrome methylene blue, 4 ml 1% cosin (corrected for dye content), 2 ml 0.1 M pH 6.3 phosphate buffer, 5 ml acetone, and 23 ml distilled water. The main is added last and methanol fixed blood films are stained immediately for 20-40 min.

For methylene blue supplied by MCB 12-H-29, optimal stains were obtained with preparations containing 20 and 24% conversion of methylene blue to azure B. With methylene blue supplied by Aldrich (080787), 16% conversion of methylene blue to azure B was optimal. Eosinates prepared from a low azure B/methylene blue preparation selected in this way give good stains when used as a Wright stain in 0.3% methanol solution. However, when the 600 mg eosinate solution in glycerol methanol is supplemented with 160 mg of the same azure B/methylene blue chloride the mixture fails to perform well. The HCI precipitation of the chloride apparently produces the zinc methylene blue chloride salt which is poorly soluble in alcohol. It appears necessary to have a zinc-free azure B/methylene blue chloride to supplement the probably zinc-free eosinate used in the Giemsa mixture.     

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 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.     

Studies on Polychrome Methylene Blue II. Acid Oxidation Methods of Polychroming

The action of K₂Cr₂O₇, Ag₂O, KMnO₄, HgO and NaIO₃ in polychroming methylene blue is explored. The last two have no action in neutral or acid methylene blue solutions. With the other three reagents the amount of polychroming, as measured by the shift in the absorption spectrum, is roughly proportional to the amount of oxidant used. Various lots of methylene blue produce similar products with similar proportions of K₂Cr₂O₇. With similar quantities of this reagent similar products are produced by polychroming at 100°, 80°, 70° or 60° C. At 100° C. the action of K₂Cr₂O₇ or of Ag₂O appears to be completed in 15 minutes. In K₂Cr₂O₇ polychroming, H₂SO₄ can be substituted for HCl, and subsequent BaCO₃ neutralization removes the salts formed and prevents accidental alkali polychroming. K₂Cr₂O₇ polychroming produces products with narrower absorption bands than alkali polychroming.     

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.     

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.     

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.     

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.     

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.     

Dichromate-Chlorate Perfusion Prior to Staining Degeneration in Brain and Spinal Cord

A method of fixation compatible with both the Nauta-Gygax and Swank-Davenport procedures for degenerating nerve fibers, which shortens the time required by the former procedure, is as follows: The central nervous system is perfused with a 0.9% aqueous solution of NaCl followed by an aqueous solution containing 5% K₂Cr₂O₇ and 2.5% KClO₃. The central nervous system is then hardened in 10% formalin for 1-3 days. Tissue for Marchi-type staining can be taken at this stage. For silver staining, the processing is continued by immersion overnight in 10% formalin in 20% alcohol, and frozen sections cut the next day. Sections, up to 50μ in thickness, are collected in 10% formalin and impregnated by the Nauta-Gygax technique. Best results are obtained by impregnating within 24-48 hr after sectioning.     

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.     

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.     

Basic Dye Counterstaining of Sections Impregnated by the Golgi-Cox Method

Celloidin blocks of Golgi-Cox impregnated material are cut at 50 μ, the sections collected in 70% alcohol, transferred to a 3:1 mixture of absolute alcohol and chloroform for 2 min, and then stored in xylene or toluene for at least 3 min, or up to 2 wk until processed further. Mounting is done on glass slides which have been coated with fresh egg albumen diluted in 0.2% ammonia water (or a 0.5% solution of dry powdered egg albumen) and then dried at 60°C overnight. For attachment to these coated slides, sections are first soaked for 2-3 min in a freshly prepared mixture of methyl benzoate, 50 ml; benzyl alcohol, 200 ml; chloroform, 150 ml; and then transferred quickly to the slides by means of a brush. After 2-3 min the chloroform evaporates and the celloidin softens. The slides are then immersed in toluene which hardens the celloidin and anchors the sections to the slides. Alcohols of descending concentrations to 40% are followed by alkalinizations, first in: absolute alcohol, 40 ml; strong ammonia water 60 ml, for 2 min, then in: absolute alcohol, 70 ml; strong ammonia water, 30 ml, for 1 hr. Excess alkali is then removed by 70% and 40% alcohol, 2 min each, and a 10 min wash in running tap water. Bleaching in 1% Na₂S₂O₃, for 10 min and washing again in tap water for 10 min completes the process preliminary to staining. The preparations are then stained for 90 min in an aqueous solution of either 0.5% cresylecht violet, neutral red, or Darrow red, buffered at pH 3.6. Dehydration and differentiation in ascending grades of alcohol, clearing with toluene or xylene, and applying a cover glass with a mounting medium having a refractive index of about 1.61 completes the process.     

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.     

A Rapid Silver Impregnation Method for Nervous Tissue: A Modified Protargol-Peroxide Technic

A rapid, reliable silver impregnation method is described for nervous tissue fixed in formol-saline, Bouin or Sum. Sections are impregnated for 10-15 minutes at room temperature or 37 C in a solution containing 0.5 g Protargol-S, 0.005-0.01 g allantoin, 1 ml of 1% Cu[NO₂]₂, 1 ml of 1% AgNO₃. and 1-2 drops of 30% H₂O₂ in 100 ml distilled water. Thereafter the dons arc reduced in a hydroquinone-formalin solution. This is followed by gold toning and subsequent reduction and mounting. Alternatively. following the first reduction, the silver image can be intensified by placing sections in a silver-allantoin bath which is followed by reduction and mounting. This method is very reliable and selective, making it suitable for general routine and research use.     

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.