Staining Sections of Peripheral Nerves for Axis Cylinders and for Myelin Sheaths

Pieces of mammalian nerves 1 to 2 cm. long were placed under moderate tension and fixed 24-48 hours in: picric acid, saturated aqueous, 90 ml.; formalin, 10 ml.; and trichloracetic acid, 25% aqueous, 2 ml. They were washed in water, cut in two and one end stained with 0.04-0.06% osmic acid solution, while the other was dehydrated, embedded in paraffin, and mounted sections from it stained with protargol. The fixing solution used was selected from a number of combinations of acidified picro-formalin as the one most likely to give satisfactory results when followed by both silver and osmic acid. The use of osmic acid solutions of less than 0.1% concentration avoided the overstaining of myelin sheaths seen frequently when stronger solutions were used with material that had been fixed previously. Protargol, 0.5% solution with fast green FCF added to make 0.05% dye in the final concentration, was used to impregnate sections for axis cylinders. Reduction and toning were done as in Bodian's method.     

Modification of the Marchi Technic

The formula proposed by Swank and Davenport (1935) was modified and applied to human and macaque nervous material. Three groups of experiments were performed and the following observations were made. (1) Diluting the osmic acid component, without altering the relative concentration of the other constituents of the solution resulted in practically no staining of the degenerated fibers. (2) When all constituents of the staining solution were used in much lower concentration than previously suggested, enhancement of staining of the degenerating fibers occurred and the different structures of the normal tissue were more easily identified. (3) At low concentrations of osmic acid and potassium chlorate, the contrast was diminished and artifacts produced by increasing the concentration of acetic acid or formalin or both. The new formula, based on the present results, consists of osmic acid, 0.5%, 11 ml.; potassium chlorate, 1%, 16 ml.; formalin (cone), 3 ml.; acetic acid, 10%, 3 ml.; and distilled water to make 100 ml. (All solutions are aqueous). Good staining after a long period of fixation in formalin, following degeneration of 8-80 days, was obtained and the cost of staining solution greatly reduced.     

Staining Methods For Studying Ingredient Distribution In Baked Cakes

A method is described for preparing cake crumb for sectioning and staining. Previous to embedding, the fat was stained and fixed by exposing small blocks of cake to the fumes from a 5%, freshly-prepared, aqueous solution of osmic acid (OsO₄). This was followed by dehydration in ethyl alcohol and tertiary butyl alcohol, removal of air under vacuum and infiltration with paraffin.

Sections were cut 20 and 9Op thick and mounted with water.

Wax was removed by immersion in xylene. The sections were rehydrated in a series of ethyl alcohol dilutions, from concentrated to dilute, then transferred to distilled water.

Protein was then stained pink by immersion of the slides in an acidified 0.04% water solution of eosin Y, or starch was stained blue with a dilute aqueous solution of iodine. Ten grams iodine and 10 g. KI were dissolved in 25 ml. distilled water. This stock solution was diluted for use one to two hundred times.

The relationship between protein and starch was demonstrated by staining the sections with eosin, differentiating in 50% alcohol and staining with iodine.

When slides of cake crumb were prepared in this way, the fat was stained black, the protein bright pink and the starch granules a dark blue.     

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.     

1. Effects of Embedding 2. Experiments with Modifications

Paraffin embedding was found to be satisfactory for brain stained by a modification of the Golgi dichromate-silver method. Nitrocellulose embedding caused fading in a few specimens. Several modifications in which the tissue was impregnated with silver nitrate before treating it with potassium dichromate were investigated. The following one is recommended. Fix pieces of brain 5-6 mm. thick for 2 days in: silver nitrate;0.5%, 90 ml.; formalin, comml. unneutralized (37-40% gas), 10 ml.; pyridine, pure, 0.05-0.1 ml. Mix in the order given and test for pH with brom cresol purple. A pH of 5.5-6.0 is about optimum and the amount of pyridine added can be varied to adjust it. A slight turbidity of the fixing fluid may be disregarded, but precipitation indicates too much alkalinity. Rinse the tissues with distilled water and place them in a mixture of potassium dichromate, 2.5%, 100 ml. and osmic acid, 1%, 1 ml., for 3-5 days. Wash in water, dehydrate with alcohol and embed in soft paraffin for thick sectioning. Greater intensity of staining (but with an increase in precipitate) can be secured by rinsing the blocks after the dichromate treatment and resilvering in a 0.5% solution of silver nitrate for a day or two, then washing, dehydrating and embedding. This modification of the Golgi method was worked out on brain of adult rat, guinea pig, cat and monkey. Results with fetal material were not good. All solutions used were aqueous, and staining was done at room temperature.     

1. Effects of Embedding 2. Experiments with Modifications

Paraffin embedding was found to be satisfactory for brain stained by a modification of the Golgi dichromate-silver method. Nitrocellulose embedding caused fading in a few specimens. Several modifications in which the tissue was impregnated with silver nitrate before treating it with potassium dichromate were investigated. The following one is recommended. Fix pieces of brain 5-6 mm. thick for 2 days in: silver nitrate;0.5%, 90 ml.; formalin, comml. unneutralized (37-40% gas), 10 ml.; pyridine, pure, 0.05-0.1 ml. Mix in the order given and test for pH with brom cresol purple. A pH of 5.5-6.0 is about optimum and the amount of pyridine added can be varied to adjust it. A slight turbidity of the fixing fluid may be disregarded, but precipitation indicates too much alkalinity. Rinse the tissues with distilled water and place them in a mixture of potassium dichromate, 2.5%, 100 ml. and osmic acid, 1%, 1 ml., for 3-5 days. Wash in water, dehydrate with alcohol and embed in soft paraffin for thick sectioning. Greater intensity of staining (but with an increase in precipitate) can be secured by rinsing the blocks after the dichromate treatment and resilvering in a 0.5% solution of silver nitrate for a day or two, then washing, dehydrating and embedding. This modification of the Golgi method was worked out on brain of adult rat, guinea pig, cat and monkey. Results with fetal material were not good. All solutions used were aqueous, and staining was done at room temperature.     

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.     

Staining Raw and Cooked Apple Tissues for Study of Cell Wall Structure

Permanent preparations were made of paraffin sections from raw and cooked apple tissues stained with microchemical color reagents for pectins and pentosans. Sections stained with ruthenium red to show pectins were dehydrated and covered in balsam, and sections stained with diphenylene diamine acetate (DDA) to show pentosans were washed with water and covered in Clearcol.

Cooking was accomplished by steaming cubed histological samples. Both raw and steamed specimens were fixed in FAA in a vacuum chamber, dehydrated and cleared in tertiary butyl alcohol, and embedded in paraffin. Paraffin sections first fixed to slides with Haupt's adhesive were further stabilized by immersing in a 1% celloidin solution after dissolving the paraffin.

Ruthenium oxychloride flakes were dissolved in a Coplin jar of water containing 2 drops of ammonium hydroxide. Rehydrated sections were stained in ruthenium red 30 minutes and rinsed in water. Three methods of further preparation follow: (1) Flood sections with 10% gum arabic; drain and air-dry thoroughly; immerse in xylene 5 minutes; cover in balsam. (2) Drain and air-dry sections; if desired, counterstain dry sections with Johansen's fast green solution; immerse in xylene; cover in balsam. (3) Dehydrate by dipping in 70%, 95%, and absolute ethyl alcohol; immerse in xylene; cover in balsam.

DDA was made by heating 15 g. of benzidine in 150 ml. of glacial acetic acid and 450 ml. of water until dissolved, then adding water to make 750 ml. of solution. Rehydrated sections were stained 4 hours in DDA, washed, stained 5 minutes in Congo red (Congo red, 5 g.; NaOH, 5 g.; water, 100 ml.), washed, and covered in Clearcol.

An Autotechnicon was used for dehydration, clearing, infiltration, deparaffinizing sections, and staining. Procedures that necessarily remained manual were fixation in a vacuum chamber, and all operations that followed staining.

Ruthenium red, though the best available indicator for pectins, may not be specific for these substances. DDA and ruthenium red stained identical structures in hypodermis and cortex. DDA also stained cuticle, hence was more useful than ruthenium red for delineating that portion. DDA sections were better for photomicrography, and for measuring thickness of cell walls. Neither stain prevented the study of cell walls in polarized light.     

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.     

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.     

Azophloxine in Two-Dye and Four-Dye Staining Combination

Tissues are fixed in either 10% formalin or Lavdow-sky's mixture. After the tissues are sectioned and mounted, they are stained in hematoxylin, then counterstained for 2 minutes in 0.1% aqueous azophloxine to which 4 drops of acetic acid have been added to each 100 ml. of stain. Sections are then rinsed in 0.2% acetic acid and dehydrated. Azophloxine GA can be used also in a tetrachrome method. Sections are stained in Harris' hematoxylin, washed, and placed in 0.2% acidified aqueous azophloxine for 2 minutes. They are then rinsed in 0.2% acetic acid, stained 1 minute in an aqueous mixture of 4% phosphotungstic acid and 2% orange G solution and rinsed again in 0.2% acetic acid. Finally, they are stained in 0.2% light green for 2 minutes, and differentiated in 0.2% acetic acid for 5 minutes. The advantage in using azophloxine is that it is clear and delicate and when used in a constant concentration, does not overstain if the recommended procedure is followed.     

Azophloxine in Two-Dye and Four-Dye Staining Combination

Tissues are fixed in either 10% formalin or Lavdow-sky's mixture. After the tissues are sectioned and mounted, they are stained in hematoxylin, then counterstained for 2 minutes in 0.1% aqueous azophloxine to which 4 drops of acetic acid have been added to each 100 ml. of stain. Sections are then rinsed in 0.2% acetic acid and dehydrated. Azophloxine GA can be used also in a tetrachrome method. Sections are stained in Harris' hematoxylin, washed, and placed in 0.2% acidified aqueous azophloxine for 2 minutes. They are then rinsed in 0.2% acetic acid, stained 1 minute in an aqueous mixture of 4% phosphotungstic acid and 2% orange G solution and rinsed again in 0.2% acetic acid. Finally, they are stained in 0.2% light green for 2 minutes, and differentiated in 0.2% acetic acid for 5 minutes. The advantage in using azophloxine is that it is clear and delicate and when used in a constant concentration, does not overstain if the recommended procedure is followed.     

Staining of Nerve Fibers with Methylene Blue Factors Improving the Staining

Several factors influencing the staining of nerve fibers with methylene blue, especially the influence of chloralhydrate and carbamylcholine chloride (as parasympathicotonics), and of some anesthetics were studied. The intestines of mouse, rat, and guinea pig were used. The following immersion technic is suggested: Tissue from animals anesthetised by chloralhydrate is immersed in: zinc free methylene blue, 0.03%; sodium tartrate, 0.5%; sodium pyruvate, 0.05% carbamylcholine, 0.00005%; 0.2 M Na₂HPO₄, 0.77%; 0.1 M citric acid, 0.18%; NACl, 0.79%; also an anesthetic which varies with the animal selected. Air is kept bubbling through the staining solution and microscopic examination is made at 6 min. intervals. After 0.5-1 hr. the tissue is fixed in: ammonium molyb-date, 10 g.; sucrose, 35 g.; distilled water, 100 ml.; to which is added just before use, 1% platinum chloride, 3 ml.; 2% osmic acid, 3 drops. Washing is in ice cold water and dehydration at 0°C. in Lang's fluids (varying mixtures of ethanol and n-butanol). The tissues thus prepared are stored in liquid paraffin.     

Staining Paraffin Sections with Protargol: 1. Experiments with Bodian's Method. 2. Use of Jvpropyl and N-Butyl Alcohol in Hofker's Fixative:

Paraffin sections of nervous tissue, which had been fixed in Hofker's fluid, stained readily with protargol solution without the addition of metallic copper or other activator. Amidolsulfite mixtures reduced the protargol more rapidly and completely than hydroquinone-sulfite. Intensification of the stain could be secured by reducing with 0.5% amidol (or pyrogallol) solution after gold toning. The completeness of staining of unmyelinated fibers of the dorsal roots of cat spinal nerves was checked by estimating the number of fibers in a root and the cells of its associated ganglion. A fiber cell ratio of 1:1 was found hi 4 specimens, indicating within limits of error that all fibers were stained. An improvement of die original Hofker's mixture as a fixative was obtained by using a mixture of formic acid, 5 cc.; trichloracetic acid, 10 g.; n-propyl alcohol, 20 cc.; and n-butyl alcohol, 60 cc. (instead of the acetic, trichloracetic, ethyl alcohol mixture used hi the original formula). The following arbitrary method is suggested. Fix 12 to 24 hours, pass to water thru graded ethyl alcohol, wash several hours, dehydrate and embed in paraffin. Cut, mount, and remove the paraffin, pass to water and impregnate 2 or 3 days at 27 to 30$$C. in a 0.5% aqueous solution of protargol (Winthrop Chemical Co.). Rinse 2 or 3 seconds and reduce with 0.5% amidol (Agfa brand used) in 5% sodium sulfite solution. Wash, tone with 0.1% gold chloride, wash and reduce with 0.5% amidol (no sulfite), wash, dehydrate and cover. The method works well on spinal nerve roots, cerebrum, cerebellum, and spinal cord, and moderately well on nerve trunks including sympathetic nerves. Tissues from cat and guinea pig were used.     

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.     

Effect of phosphomolybdic acid on the binding of Sudan black B

Paraffin sections of tissues fixed in absolute alcohol or Carnoy's fluid were mordanted in a 1% aqueous solution of phosphomolybdic acid, stained in saturated solutions of Sudan black B, acetylated Sudan black, various solvent and basic dyes in 70% ethyl alcohol for 5 min at room temperature, dehydrated in alcohol and covered in Permount. Sudan black B and other dyes with basic groups stained basement membranes, reticulum and collagen fibers intensely. Acetylated Sudan black, Sudan IV and oil red 0 did not color any tissue structures. Control sections, without pretreatment, did not bind Sudan black B. These findings indicate interaction between basic groups of the dye and free acid groups of phosphomolybdic acid.     

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.     

Improved Procedures in Fungal Cytology Utilizing Giemsa

To study nuclear events in fructifications of the Basidiomycetes, material was fixed 24 hr in a saturated aqueous solution of HgCl₂ containing 1% glacial acetic acid, and embedded in Aquax (G. T. Gurr Ltd.). Following a 4 hr hydrolysis at 20 C in 60% H₃PO₄, sections were stained for 30 min in a mixture of 4 ml Giemsa R66 (G T. Gurr Ltd.) and 100 ml phosphate buffer at pH 6.5. Differentiation was carried out in sodium cacodylate-HCl buffer at pH 5.8 when required. Preparations were dehydrated in an acetone-xylene series prior to mounting in Euparal. The use of paraffin wax as the embedding medium and HCl as the hydrolysing agent yielded preparations of an inferior quality.     

A Modified One-Step Trichrome Stain for Demonstration of Fine Connective Tissue Fibers

Gomori's one-step trichrome procedure was modified to improve coloration of fine connective tissue fibers. Paraffin sections from tissues fixed in alcohol, acetone, Zenkerformol, 10% formalin, Kaiserling's or Carnoy's fluid were mordanted 1 hr at 56 C in Bouin's solution, stained 1 min in a trichrome solution (chromotrope 2R-phosphomolybdic acidaniline blue WS) adjusted to pH 1.3 with HCl, rinsed in 1% aqueous acetic acid, dehydrated and covered. Collagen, reticulum fibers, basement membranes, ring fibers around splenic sinuses, intercalated discs in cardiac muscle and cartilage were colored blue. Nuclei, cytoplasm, fibrin, muscle fibers and elastic fibers were stained red. Pretreatment of sections with Bouin's solution enhanced the affinity of tissues for chromotrope 2R and was found essential for satisfactory coloration of material fixed in alcohol, acetone, formalin or Carnoy's fluid. Because this method does not require differentiation, it gave uniform results even in the hands of inexperienced laboratory trainees. No fading was observed in sections stored for more than 8 yr.     

The Demonstration of Beryllium Oxide in Paraffin Sections with Chromoxane Stains

Aqueous solutions of the arylmethane dyes Chromoxane pure blue BLD (C.I. No. 43825) and Chromoxane pure blue B (C.I. No. 43830) will stain beryllium oxide. In the presence of EDTA the staining of other metals is masked. As a specific stain for BeO, formol saline fixed paraffin sections are hydrated and stained for 1 hr with either 0.1 gm of pure blue BLD in 100 ml of pH 4.0 Na-acetate buffer or with 0.1 gm of pure blue B in 1 N NaOH adjusted to pH 9.0 with HCl. To mask interference from other metal ions, 9 gm of Na₂-EDTA is added to 100 ml of the stain solution. BeO is stained blue, organic tissue components are either unstained or pink. Results of tests against other materials show that a high degree of specificity may be expected from these dyes. A 1% aqueous solution of neutral red may be used as a counterstain.