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.     

Stain Technology a Bone Stain for Osteoid Seams in Fresh, Unembedded, Mineralized Bone

Fresh, ground, mineralized bone sections 75-100 μ thick are stained 90 minutes or 48 hours in the Bone Stain, a preparation containing fast green FCF, orange G, basic fuchsin, and azure II. Surface stain is then removed by grinding under running water. Sections are washed in 0.1% zephiran chloride (benzalkonium chloride) or in 0.01% mild soap and again washed in tap water, followed with distilled water. Sections are next differentiated in 0.01% acetic acid in 95% methanol, dehydrated in 95% ethanol and 100% ethanol, cleared in alcohol:xylene 1:1, 1:4, 1:9 and 2 changes of xylol, and then mounted permanently in Eukitt's mounting media.

Osteoid seams stain either green to jade green or red to dark red, incompletely mineralized bone red or orange yellow, and the zone of demarcation light green. The walls of lacunae, canaliculae, feathered bone, procedural artifacts and periosteocyte lacunar low-density versions stain red.

The method helps in the differential diagnosis of certain metabolic bone diseases in human biopsy and autopsy material.     

A Durable Nissl Stain for Frozen and Paraffin Sections

Frozen sections, 25-50 /j. thick, of formalin-fixed nervous tissues are mounted following the Albrecht gelatin technic. Paraffin sections, 15 p., are deparaffinized and transferred to absolute ethanol. The slides are then coated with celloidin. Both frozen and paraffin sections subsequently follow the same steps: absolute ethanol-chloroform (equal parts) for at least 20 min, 95% ethanol, 70% ethanol (1-3 min), then rinsed in distilled water. Sections are stained in Cresylechtviolett (Chroma) 0.5% aqueous solution containing 4 drops of glacial acetic acid per 100 ml, rinsed in distilled water, agitated in 70% ethanol until excess stain leaves the slide, and rinsed in 95% ethanol. Sections are then dehydrated in absolute ethanol, followed by butanol, cleared in xylene, and enclosed in permount.     

Modified Whipf's Polychrome: A Connective Tissue Stain with Special Application for Demonstrating Leishmania

A polychrome stain procedure was developed to demonstrate amastigotes of the protozoan parasite Leishmania braziliensis as well as cytoplasmic and other tissue components in cutaneous lesions of infected animals. The procedure is as follows: stain nuclei for 10 minutes with an iron hematoxylin containing 0.5% hematoxylin and 0.75% ferric ammonium sulfate dissolved in 1:1 0.6 N H₂SO₄:95% ethanol; rinse 4 minutes in distilled water. Cytoplasmic staining is achieved by exposing tissues for 10 minutes to a solution containing 0.25% Biebrich scarlet, 0.45% orange G, 0.5% phosphomolybdic acid and 0.5% phosphotungstic acid in 1% aqueous acetic acid. These first two solutions are modified from Whipf's polychrome stain. Sections are differentiated for 10 seconds in 50% ethanol, rinsed in water, stained 3 minutes in 0.1% aniline blue WS in saturated aqueous picric acid, rinsed in water and differentiated for 1 minute in absolute ethanol containing 0.05% acetic acid. Mordanting overnight in 6% picric acid in 95% ethanol produced optimal results.

This procedure was applied to sectioned material from experimental animals with various protozoa. Trypanosoma cruzi, Besnoitia Jellisoni, Toxoplasma gondii and especially Leishmania braziliensis were well demonstrated. Combining cytoplasmic dyes and phosphomolybdic-phosphotungstic acids into one solution afforded differential staining of tissues by Biebrich scarlet and orange G; connective tissues were stained by this solution. Substantially improved definition of connective tissues resulted after counterstaining. This procedure differs from the Massou sequence in which connective tissues are first stained by cytoplasmic dyes along with other tissues and then destained prior to specific counter-staining. in comparing dyes structurally related to Biebrich scarlet, it was found that Crocein scarlet MOO, but not Poncenu S, was an acceptable substitute. Sirius supra blue GL and Sirius red FSBA were not useful as replacements for aniline blue WS in this procedure.     

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.     

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.     

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

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

A Selective Stain for Elastic Tissue (Orcinol-New Fuchsin)

A selective stain for elastic tissue (designated orcinol-new fuchsin) is described. Two grams of new fuchsin (C.I. No. 678) and 4 gm of orcinol (highest purity) are added to 200 ml of distilled water and the solution boiled for 5 min. Then 25 ml ferric chloride solution (U.S.P. IX) are added and the solution is boiled 5 min longer. The precipitate is collected and dissolved in 100 ml 95% ethanol. This is the staining solution. Sections are deparaffinized and brought to absolute ethanol, stained for 15 min at 37 °C with orcinol-new fuchsin, differentiated for 15 min in 70% ethanol, dehydrated, cleared and covered as usual.     

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.     

Digestion of Collagen and Reticulin in Paraffin Sections by Collagenase

Tissues are fixed in ethanol or in Carnoy's 6:3:1 mixture and embedded in paraffin after routine ethanol dehydration. Sections are taken to water and then covered with 0.2 ml of a 0.9% NaCl solution containing 1 mg/ml of collagenase, and incubated at 50° C for 45 min. After this, they were washed and then stained by the usual methods for connective tissue fibers. Control sections were made by substituting plain 0.9% NaCl solution for the collagenase solution. The collagenase used was from bacteria and obtained from Nutritional Biochemicals Corporation, Cleveland 28, Ohio.     

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.     

Azophloxine Ga, A Selective Stain for Light and Fluorescence Microscopy of Myoepithelial Cells

Paraffin sections from human lingual glands fixed in Carnoy's fluid No. 2 were dewaxed, hydrated and treated as follows: Mayer's acid hemalum, 5-10 min; running water, 15 min; 5% aqueous tannic acid, 10 min; distilled water, 3 changes; 1% aqueous phosphomolybdic acid, 10 min; distilled water, 3 changes; azophloxin GA, 2 gm in 9:1 methanol-acetic acid (mixed 16-20 hr before use), 5 min; 9:1 methanol-acetic acid, 2 changes; absolute alcohol, 1 dip; and apply a cover with nonfluorescent medium. Myoepithelial cells and muscle fibers were stained deep red; connective tissue fibers and serous cells, yellow; mucous cells, unstained. Only myoepithelial cells and muscle fibers were strongly fluorescent. This selective fluorescence greatly facilitated study of very fine fibers in myoepithelial cells and of the basket-like meshworks. This stain does not require differentiation and is useful in general histopathology. No fading was observed in sections stored for 1 yr.     

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.     

Carbol Fuchsin as A Stain for Human Chromosomes

Cells derived from cultures of bone marrow or leucocytes were treated with hypotonic citrate solution, squashed in 45% acetic acid frozen with CO₂ to allow removal of the cover glass without disturbing the smear, and stained by the following schedule: absolute alcohol, 5 min; coat with 0.2% parlodion and air dry; 70% alcohol, 5 min; distilled water, 5 min; stain 2-5 min in a mixture of 45 ml of a 0.3% solution of basic fuchsin in 5% phenol, 6 ml of glacial acetic acid, and 6 ml of 37% formaldehyde. Differentiate and dehydrate in absolute alcohol, clear in xylene and cover. The stain is durable for several weeks if slides are stored in darkness when not in use. Results resemble those obtained by Feulgen or aceto-orcein methods.     

A Consistent Phosphotungstic Acid Hematoxylin Stain for Glial Fibers

After deceration, celloidinization and hydration, oxidize 10 micron paraffin sections for 15 min in a solution containing 0.3 g KMnO₄, and 0.1 ml conc. H₂SO₂, per 100 ml distilled water. Wash in water and reduce in 5% oxalic acid until the sections are colorless. Wash thoroughly in water and place in 4% iron alum solution for two hours. Wash briefly in water and stain for two hours in phosphotungstic acid hematoxylin. Rinse briefly in 95% ethanol and dehydrate in n-butyl alcohol or absolute ethanol for 4 min with two changes, clear and mount. Glial fibers, myofibrils, red blood cells, etc. are stained blue while astrocyte cell bodies, collagen, etc. are stained red. This stain has proven highly consistent in a wide variety of astrocytic derangements. Despite the intensity of this PTAH modification, false positive staining was not observed.     

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.     

Acetic Acid as a Diluent and Dehydrant in the Preparation of Whole, Stained Helminths

Aqueous 45% acetic acid can be used successfully as a diluent for Ehrlich's haematoxylin and for Horen's trichrome stain (chromotrope 2 R, 0.6 gm; phosphotungstic acid, 0.7 gm; glacial acetic acid, 1.0 ml; water, 100 ml). Glacial acetic acid is used for dehydration of the stained helminths, and followed by a glacial acetic acid-methyl salicylate series for clearing. The whole process can be completed within 1 hr, from fixation to the cleared specimen, with helminths up to 5 mm in length. A satisfactory fixative for Monogenea, Digenea and Acanthocephala is: 85% ethanol, 85; formalin (40% HCHO), 10; and glacial acetic acid, 5—parts by volume. For Cestoda, 5% aqueous formalin is preferable because they are hardened excessively by the alcoholic fixative.     

Oil Blue Na as A Stain for Rubber in Sectioned or Ground Plant Tissues

Oil blue NA (Calco), a stain which colors rubber bright blue, has been used effectively in studying the distribution of rubber in several plant species. Fresh or fixed sections are cut, bleached with Javelle water or NaOCl solution, treated with 9% KOH in 95% ethanol, washed with several changes of water and finally with 95% ethanol, and stained with 0.05% oil blue NA in 70% ethanol. Sections are rinsed in 50%' ethanol, placed in 40% glycerin, and mounted in glycerin jelly.

For the detection of changes in the distribution and character of rubber in milled or ground tissues, much the same staining procedure is followed. The stained tissues usually are examined and dissected under a stereoscopic microscope, a procedure which permits rubber to be recognized by both its staining reaction and by a more specific property, elastic elongation.

A microscopic technic is presented whereby it is possible to determine approximately the relative proportion of dispersed and coagulated rubber latex in unstained tissues.     

Luxol Fast Blue Mbs: A Stain for Phase Contrast Microscopy

A staining method to increase the contrast of sectioned material for phase contrast microscopy is described. Two stock solutions of the stain are required. The first is made by dissolving 2 gm of luxol fast blue MBS in 100 ml of 95% ethanol. The second solution is made up of 4 ml of a 29% aqueous solution of FeCl₃, 95 ml of 95% ethanol, and 1 ml of concentrated HCl. The staining solution is made by mixing equal parts of the two solutions. Sections are deparaffinized and taken to 70% alcohol, stained for 1.5 hr, dehydrated, cleared and covered as usual.     

A Ribonuclease-Gallocyanin Stain for Sexing Skin Biopsies

Skin biopsies for sexing can be fixed best in 10-15% aqueous formalin or this solution saturated with HgCl₂. Bouin's fluid and all chromate mixtures should be avoided. Celloidin-paraffin double embedding is recommended but not essential. Sections are brought to water, mercurial residues removed if necessary, and then washed in distilled water. They are incubated at 37°C in a ribo-nuclease solution: approximately 1 mg of ribonuclease powder (Light's) in 100 ml of glass-distilled water; boiled 3-5 sec after dissolving, and kept in a refrigerator (usable about a week). The sections are rinsed and incubated at 37°C overnight in gallocyanin-chromalum (Einarson, 1951) made as follows: Dissolve 5 gm of chromalum in 100 ml of distilled water, add 0.15 gm of gallocyanin, shake thoroughly, heat slowly and boil 5 min; cool, filter, and wash through the filter with distilled water until the filtrate reaches 100 ml. This solution is usable at once and keeps at least a month. Sections should be dipped in acid alcohol to clean (optional), but no attempt made to differentiate them, and washed in tap water. Dehydration, clearing and covering complete the process. The method is nearly as precise as the Feulgen and more convenient and reliable for routine use on miscellaneous material.