Orthochromatic, Species-Limited Staining of Mast Cells With Night Blue

Night blue will stain the mast cells of rat, mouse and hamster selectively if alcohol differentiation is controlled. The technical steps are: Dewax paraffin sections with xylene, 2 changes; air dry; 2% Na₂SO₄, 3-5 sec; 0.5% night blue in 10% ethanol, 1 hr at 60°C; rinse in water; 9% HNO₃, 15 sec; water 1-5 min; 70% ethanol, 2 changes, 30 sec each; wash; 0.01% safranin, 3-5 sec; rinse, blot, air dry, mount in synthetic resin. A clear orthochromatic stain of the mast-cell granules occurs. Acid fixation prevents the staining reaction.     

Thick Sections for Embryology

Night blue will stain the mast cells of rat, mouse and hamster selectively if alcohol differentiation is controlled. The technical steps are: Dewax paraffin sections with xylene, 2 changes; air dry; 2% Na₂SO₄, 3-5 sec; 0.5% night blue in 10% ethanol, 1 hr at 60°C; rinse in water; 9% HNO₃, 15 sec; water 1-5 min; 70% ethanol, 2 changes, 30 sec each; wash; 0.01% safranin, 3-5 sec; rinse, blot, air dry, mount in synthetic resin. A clear orthochromatic stain of the mast-cell granules occurs. Acid fixation prevents the staining reaction.     

A Six-Dye Staining Schedule for Sections of Mesquite and Other Desert Plants

Materials are fixed in FPA (formalin, 2; propionic acid, 1; 70% ethanol, 17). Paraffin sections on slides are brought to 50% ethanol and stained as follows: (1) in Bismarck brown Y, a 0.02% solution in 0.1% aqueous phenol, 10-30 min; wash 30 sec in 0.7% acetic acid, and wash in distilled water 20-30 sec; (2) in crystal violet, 1% in 70% ethanol alkalinized with 1 drop of 1 N NaOH per 100 ml, 12-35 min; wash 30-60 sec in tap water to remove excess stain, and rinse 0.5 sec in 70% ethanol; then mordant in I₂-KI, 1% each in 70% ethanol, 40 sec, and rinse in 70% ethanol 2-5 sec; (3) in a mixture containing 0.4% acid fuchsin and 0.6% crythrosin B in 70% ethanol about 0.5 sec; rinse in 70% ethanol 5-15 sec to remove excess red; dehydrate in 70%, 95%, and absolute ethanol, 2-3 sec each; (4) in fast green FCF, 0.5% in a mixture of equal parts of methyl cellosolve, absolute ethanol, and clove oil, 5-15 sec; rinse in a mixture of clove oil, 10 ml; absolute ethanol, 100 ml; and methyl cellosolve, 10 ml, 5-7 sec; (5) in orange G, 0.75 gm in a mixture of clove oil, 40 ml; absolute ethanol, 40 ml; and methyl cellosolve, 60 ml, 5-30 sec; rinse clean in a 1:1 mixture of xylene and absolute ethanol, 5-20 sec Complete the clearing in pure xylene, 3 changes, 1.5 min in each, and apply a cover glass with synthetic resin. Slides are agitated in all steps except Bismark brown Y, crystal violet, and the xylenes. Contrast and staining intensity are adjusted by varying staining times in the dye solutions.     

Basic Fuchsin-Crystal Violet; A Rapid Staining Sequence for Juxtaglomerular Granular Cells Embedded in Epoxy Resin

A basic fuchsin-crystal violet staining sequence for demonstration of juxtaglomerular granular cells in epoxy-embedded tissues is rapid and results in slides with excellent contrast and intensity. Procedure: Cut sections 0.3-0.6 μ thick. Hydrate through xylene and alcohol to water. Stain in modified Goodpasture's stain (basic fuchsin, 1; aniline, 1; phenol, 1; 30% alcohol, 100) for 20-30 sec; rinse in tap water; stain in modified Stirling's (crystal violet, 5; alcohol, 10; aniline, 2; water, 88) for 20-30 sec; rinse in tap water and dry on a hotplate; mount in a synthetic resin. Granular cells of the juxtaglomerular apparatus are stained an intense dark blue by the crystal violet. Arterial elastic membranes and collagen are pale blue. Other structures are shades of red.     

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.     

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.     

Sequential Use of Wright's and Ziehl-Neelsen's Stains for Demonstrating Phagocytosis of Bacterial Spores

Nongerminating spores, germinating spores, and vegetative cells of Clostridium botulinum type A were observed during phagocytosis in the peritoneal fluid of white mice. Since phagocytes are easily ruptured by heat, the method described avoids heating, as this has been employed in conventional spore staining methods. A thin smear of the fluid is air dried on the slide for 2 hr, and stained by Wright's method: stain, 2 min; dilution water, 2 min; and rinsed; then in 0.005% methylene blue for 30 sec, and rinsed. This is followed by Ziehl-Neelsen's stain for 3-4 min and destained with 1: acetone-95% ethanol for 10 sec. The slide is rinsed, and Wright's staining repeated: stain 1 min, dilution 2-3 min; and thereafter washed about 5 ml of Wright's buffer. Blotting and air drying completes the staining. Non-germinating spores stain light red with a red spore wall, germinating spores are deep red throughout, vegetative cells are blue, and leucocytes show a dark purple nucleus and light blue cytoplasm.     

Hot, Acidified Rhodamine B and Methylene Blue; A Differential Stain Dichromatic for Hair Follicular Keratins, Achromatic for Epidermal Keratin

Procedure:Cut paraffin sections and float on a 45-50 C water bath; spread silicone-rubber adhesive (Clear Seal-General Electric) thinly and evenly over 2/3 of the slide; pick up the sections from the floatation water with the coated slide; dry for 1.5 hr at 25 C and at 60 C for 0.5 hr; deparaffinize, and hydrate to water. Place 150 mg of rhodamine B and 150 mg of methylene blue each in separate 100 ml beakers and add 80 ml of 10% HCl to each beaker. Bring both solutions to a boil on a hot plate in a fume hood; immerse tissue sections in the boiling rhodamine B exactly 2 min; rinse in a beaker of 10% HCl 5 sec; immerse in the boiling methylene blue exactly 0.5 min; rinse in distilled water; blot dry; and mount in a silicone-rubber medium (Glass and Ceramic Adhesive—Dow Corning Corp.). Hair shaft keratin stains red; inner root sheath keratin and keratogenous zone of the hair shaft, blue green; epidermal keratin remains unstained. Pilomatrixornas show foci of both red and blue green keratin; epidermal and hair sheath (“sebaceous”) cysts remain unstained.     

Staining Nerve Fibers After Sublimate-Acetic and After Bouin'S Fluid

A silver staining method for paraffin sections of material fixed in HgCl₂, sat. aq., with 5% acetic acid is as follows. Process the sections through the usual sequence of reagents, and including I-KI in 70% alcohol, thiosulfate (5% aq.), washing and back to 70% alcohol containing 5% of NH₄OH (conc. aq.). After 3 minutes in the ammoniated alcohol, wash through tap water and 2 changes of distilled water and silver 5-10 minutes at 25°C. in 15% AgNO₃ aq. to which 0.02 ml. of pyridine per 100 ml. has been added. Blot the slide, but not the section and do not rinse. Reduce at 45°C. in 0.1% pyrogallol in 55% alcohol, then rinse in 55% alcohol and wash in water. The remainder of the process consists of gold toning, intensifying in oxalic acid, fixing in 5% Na₂S₂O₃, washing, dehydrating, clearing and covering. When the specimen contains much smooth muscle, the I-KI solution is acidified before use by adding 2 ml. of 1N nitric acid per 100 ml., and the sections treated for 3 minutes instead of the usual 2 minutes. Formalin should not be added to sublimate-acetic, but specimens that do not contain strongly argyrophilic nonneural tissue may be fixed in formalin or, preferably, Bouin's fluid. Sections of tissue after the latter type of fixation will not require the I-KI and thiosulfate but can go from 95% alcohol to the ammoniated alcohol. The advantages of fixing in HgCl₂-acetic acid are suppression of the staining of connective tissue and intensifying the staining of nerve fibers.     

Controlled Differentiation of Cells and Connective Tissue Fibers in Silver Staining

Controlled silver staining of connective tissue fibers and sometimes of these fibers and cells simultaneously can be obtained. 1. Fix in 10% formalin. Embed in paraffin and cut sections as usual, but do not mount them on slides. Deparaffinize and hydrate through xylene, alcohols and distilled water and henceforth treat them the same as frozen sections. Real frozen sections can also be used. 2. Treat with a freshly prepared 1% solution of KMnO₄, usually 15-60 sec, sometimes up to 10 min. 3. Wash in distilled water, 5-10 sec. 4. Decolorize in 2% potassium metabisulfite, 10-20 sec. 5. Place in distilled water, 1 min. 6. Sensitize with 2% iron alum, 1 min. 7. Place in distilled water, 1 min. 8. Impregnate in Gomori's silver oxide solution, 2 min. 9. Wash in a 1.5% aqueous solution of pyridine, about 15 sec. 10. Reduce in a mixture containing 0.25% gelatin and 2% formalin 1 min. 11. Repeat steps 7 to 10 once or several times until the connective tissue fibers are completely stained. For cell staining (which may fail) proceed as follows: After the first insufficient staining of the connective tissue fibers, rinse in distilled water, dip for 1 sec in Gomori's solution and reduce immediately in gelatin-formalin without previous washing in pyridined water. This step can be repeated. 12. If the staining is too strong, decolorize as needed in 2% iron alum. 13. Toning in 0.2% gold chloride, 5 min or more, followed by fixation in 5% sodium thiosulfate, 1 min, is optional. Counterstain as desired. 14. Wash in tap water, dehydrate, clear in xylene and mount in balsam. The same technique applied to sections attached to slides gives good results but inferior to that obtained in paraffin sections processed in the loose, unmounted condition.     

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.     

Silver Staining, Featuring Rapid Reduction, For Whole Mounts of Retina and Optic Pathways in Chick Embryos

Developing and established nerve fibers in the retina and in superficial tracts of the brain can be stained and viewed en bloc. The method was developed on chick embryos of 2 days of incubation to several months post-hatching but could be used on other material provided that the objects of interest were within 35 μ of the surface. Procedure: (1) Place the entire eye or head in 50% pyridine for at least 16 hr. (2) Wash well for 5-7 hr with hourly changes of distilled water or with running tap water for 4-6 hr followed by several changes of distilled water. (3) Transfer to 95% ethanol for 16-48 hr. (4) Impregnate with 1.5% AgNO₃ for 2 days at 37 C. (5) Submerge in water and, when staining the retina, remove the vitreous body and apply an aqueous solution of 0.25% pyrogallic acid in 1.25% formalin by directing a narrow stream of this reducer against the retina for 2-5 sec. Wash the eye with distilled water 30-60 sec after applying the reducer. When staining the brain, remove the meninges under water, direct the stream of reducer against the brain for 20-30 sec, and rinse the brain immediately after the nerves have stained. (6) Dissect the specimen and make temporary mounts in glycerol; or, dehydrate and clear for resin mounting. The technique stains both mature and growing axons with their growth cones and sometimes their cell bodies. The fiber patterns show best on the surfaces of the retina and brain. The stain works consistently and is suited to the study of both normal and abnormal development.     

Studies on Textile Dyes for Biological Staining II. A Trichrome Stain for General Use

This trichrome staining procedure differentially stains elastic fibers, collagen fibers and mucin. Gomori's aldehyde-fuchsin is used for elastic fibers; fast yellow TN is the component used for collagen and cytoplasm; pontacyl blue black SX is the nuclear stain. Procedure: Paraffin sections to water; aldehyde-fuchsin, 30 min; 70% ethanol; distilled water; 0.75% pontacyl blue black SX in 1.5% K.₂Cr₂O₇, 15 min; tap water; 70% ethanol to wash off all free dye; 2% fast yellow TN in 95% ethanol, 5 min; dehydrate, clear and cover.     

The Demonstration of Beta Cells in Pancreatic Islets and Their Tumors

The stain is applied routinely to tissues fixed in 10% buffered formalin (pH near 7.0) or in Bouin's fluid. Bring paraffin section to water as usual and mordant 72 hr in 5% CrCl₃ dissolved in 5% acetic acid. Wash in water and in 70% alcohol and stain 6 hr. Formula of staining solution: new fuchsin, 1% in 70% alcohol, 100 ml; HCl, conc., 2 ml and paraldehyde, 2 ml, mixed together and added to the dye solution; let stand 24 hr before use. After staining, wash in running tap water 5-10 min, rinse in distilled water and counterstain if desired. Dehydration in alcohol, clearing and covering completes the process. When the paraldehyde is obtained from a freshly opened bottle, standardized staining times can be used and thus eliminate the necessity of differentiating individual slides. The granules of beta cells stained deep blue to purple and were demonstrated in the pancreatic islet of man, dog, mouse, frog, guinea pig and rabbit.     

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.     

Practical staining method: T-OPA triple stain

The following procedure has proven to be successful as routine trichrome stain on paraffin embedded material: 1) Mayer's hemalum for 10 min, followed by running tap water wash; 2) staining in 1% Orange G in 1% acqueous PTA for 5 min and rinsing a few seconds in distilled water; 3) Aniline blue 1% acqueous for 5 min, followed by few seconds distilled water wash. Dehidratation in ethanol, or by blotting followed by t-buthanol or 1:3 terpineol-xylene, clearing and mounting, completed the procedure.     

A Differential Staining Method for Elastic Fibers, Collagenic Fibers, and Keratin

A method allowing for the differential presentation of elastic fibers, other connective tissue fibers, epithelial and other types of cytoplasm, and keratin is described. The procedure is based on the affinity of orcein for elastic fibers, of anilin blue for collagenic material, and of orange G for keratin. Bouin-fixed, tissue-mat embedded sections are stained in Pinkus' acid orcein for 1 1/2 hours and rinsed in distilled water. The sections are differentiated in 50% alcohol containing 1% hydrochloric acid, washed in tap and then in distilled water. The sections are next transferred for I to 2 minutes to the anilin blue, orange G, phosphomolybdic acid combination known as solution No. 2 of Mallory's connective tissue stain, diluted 1:1 with distilled water. They are then rinsed in distilled water, quickly passed into 95% alcohol, and dehydrated in absolute alcohol containing some orange G, after which they are cleared and mounted. Within less than two hours sections may be stained and mounted with the following results: elastic fibers — red; collagenic fibers — blue; muscle fibers — yellow; keratin — orange.     

Serial Sectioning of Insects with Hard Exoskeleton by Dissolution of the Exocuticle

The method reported here was designed to produce paraffin serial sections as thin as 5 Mm of insects or other arthropods with a hard cuticle. Heads and abdomens of Apis mellifera, Eristalomyia tenax and Tenebrio molitor were fixed with Schaffer's liquid, dehydrated with 80% ethanol, 90% ethanol, two changes of 100% isopropanol (2 hr each) and 12 hr in a 1:1 mixture of paraffin (58 C melting point) at 60 C. They were molded in paraffin after 12 hr of infiltration under a partial vacuum at 60 C. Large body openings of objects were sealed with paraffin to prevent infiltration of solvents.

Thereafter, the outer paraffin was removed manually and with xylene (15 min); the cuticle was rehydrated with 100% isopropanol and 100% ethanol (15 min each). The objects were then treated with Sputofluol (Merck; a mixture of NaOH and NaCIO) until they became white or their colorless endocuticle was stainable with aniline blue WS (C.I. 42755) after rinsing in a 50% acetic acid solution (v/v). They were then dehydrated with 100% ethanol and 100% isopropanol (15 min each) and subsequently re-embedded in paraffin. They were molded, sectioned, stained and mounted as usual.     

A Differential Stain for Rubber in Guayule

The following schedule, which combines an intense blue stain for rubber with sharply contrasting red counterstains, has been found satisfactory for use in an anatomical study of rubber deposition in guayule: Cut fresh or fixed sections about 50 to 100 % thick, transfer to 50% ethanol. Extract with acetone 5 minutes, treat with 1% NaOCl 5 minutes, saponify with 10% KOH in 95% ethanol 15 minutes, rinse 3 times with 50% ethanol, stain in oil blue NA (Calco) with safranin and Congo red 30 minutes at 55° C. Rinse in 50% ethanol 2 (or more) times to remove excess stain and mount in Karo syrup.     

The Surface Staining of Embedded Tissues

The staining procedure is based on the theory that the freshly cut surface of embedded material will absorb stain only in the exposed tissue elements, provided that the embedding compound itself will not absorb the staining fluid. Concentrated stains are used for short intervals to insure minimum penetration. For paraffin embedded materials: (1) Cut block, preferably on microtome, to the desired tissue surface. (2) Rinse in absolute alcohol. (3) Float face down in stain. (Ripe, concentrated alum hematoxylin—Galigher's formula recommended—will stain in 10 to IS minutes. Heidenhain's iron hematoxylin works exceptionally well in some cases.) Mordant 20% alum 5 to 10 minutes, briefly rinse, and stain comparable 5 to 10 minutes in 1 to 1.5% hematoxylin. (4) Allow to become blue in tap water (for hematoxylin stains). (5) Counter-stain if desired. (6) Dehydrate in absolute alcohol for not more than 10 minutes. (7) Dry for 15 to 20 minutes. (8) Trim block to 2-3 mm. and mount between two cover glasses by use of microflame. Attach mount to slide with balsam. For celloidin embedded materials: (1) Dehydrate block with 90% alcohol, phenol-toluene, finally pure toluene. (2) Rinse cut surface with 90% alcohol, then apply stain. (3) Wash, after hematoxylin stains, counterstain if desired. (4) Dehydrate surface, 90% alcohol, phenol toluene, pure toluene, and mount in medium dissolved in toluene.

Possible applications of surface staining technic are suggested and illustrated.