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.     

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.     

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.     

Techniques for Preparing and Staining Phytophthora Oospores in Soybean Rootlets

Nylon mesh tissue carriers were constructed to hold soybean rootlets through fixing, dehydrating and embedding. Mesh pieces three centimeters square were doubled and sealed at each end by heat. Tissue samples were placed inside with an identifying piece of aluminum foil and the carrier sealed. Rootlets were fixed in Karpechenko's solution, dehydrated in an alcohol series and infiltrated with paraffin. They were embedded in paraffin after removal from the carrier, and sectioned on a microtome. Sections were mounted on glass slides and deparaffinized. A new stain was developed to differentiate oospores of Phytophthora megasperma var. sojae formed in these rootlets. The stain was prepared by dissolving 100 mg bromphenol blue in 50 ml of 95% ethanol and adding 3 g silver nitrate. Procedure: 5 sec in 95% ethanol, 30 min in silver stain, tap water rinse, 5 sec in 95% ethanol, 1 sec in saturated methylene blue in ethanol, immediate rinse in tap water, dehydration in absolute ethanol, rinse in tertiary butanol and xylene and mount. Previous clearing of the tissue was not required, and no air bubbles accumulated within the mesh carrier. This low cost, permeable carrier preserved the minute tissue specimens throughout processing, and the simple, progressive stain clearly differentiated oospores from surrounding tissue.     

Staining Procedures for Ultrathin Sections of Tissues Embedded in Polyester Resin

Tissues were fixed for 30 min In cold (0-2° C) 1% OsO₄ (Palade) buffered at pH 7.7, to which 0.1% MgCl₂ was added. Dehydration was in a graded ethanol series (containing 0.5% MgCl₂) at 0-2° C, and terminated with 2 changes of absolute ethanol. Tissues were then transferred by a graded series to anhydrous acetone. Infiltration of the tissue with Vestopal-W (a polyester resin), is gradual with the aid of graded solutions of Vestopal-W in acetone. The infiltrated tissue is encapsulated and initial polymerization is done under ultraviolet light at room temperature for 8-16 hr. This is followed by final hardening at 60° C for 36-48 hr. Sections (0.2-1 μ) were cut, dried on slides, placed in acetone for 1 min and then treated by either of the following staining procedures: (1) Thionin-azure-fuchsin staining: Flood the preparation with 0.2% aqueous thionin and heat to 60-80° C for 3 min; if the preparation begins to dry, add stain. Rinse in distilled water. Flood the slide with 0.2% azure B in phosphate buffer at pH 9. Heat to 60-80° C for 3 min; do not permit the preparation to dry. Rinse in distilled water. Dip the slide in MacCallum's variant of Goodpasture's carbol-fuchsin stain for 1-2 sec. Rinse in distilled water. Check the preparation microscopically for intensity of the fuchsin stain. Repeat dips as may be needed to obtain the desired intensity. Rinse in distilled water. Dehydrate quickly in 95% and absolute alcohol; clear in 2 changes of xylene and cover in Permount or similar synthetic resin. (2) Thionin-azure counterstain for the periodic acid-Schiff reaction: Oxidize the tissue in 0.5% periodic acid for 15 min and transfer to Schiff's leucofuchsin solution for 30 min. Counterstain with 0.5% aqueous thionin for 3 min; wash in distilled water; stain in 0.2% azure B in phosphate buffer at pH 5.5; wash in distilled water; dehydrate; clear and cover as in the first method. For temporary preparations let dry after absolute alcohol and apply a drop of immersion oil directly on the section.     

Mercuric Bromphenol Blue Staining of Precipitin Lines in Agar

Lines formed by antibody-organ antigen reactions are stained particularly well by a modification utilizing the mercuric bromphenol blue (MBB) mixture of Mazia et al. (Biol. Bull., 104: 57-67, 1953). The agar covered slides are placed overnight in 0.85% NaCI at 4 C, followed by washing for 2 hr in 0.85% NaCI at 25 C. They are then rinsed for 10 min in distilled water, and dried overnight at 37 C. The precipitin lines are fixed by immersing the slides for 25 min in 95% alcohol, followed by 5 min hydration in distilled water. They are stained for 25 min in MBB mixture (HgCI₂, 10 gm; bromphenol blue, 0.1 gm; 95% ethanol, 100 ml). Excess stain is removed by immersing in acidified alcohol (95% ethanol, 98 ml; glacial acetic acid, 2 ml). Finally, the slides are passed through alcohol and xylene, and resin-mounted under coverslips.     

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.     

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.     

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.     

üBisch Granules on Tapetal Membranes in Anthers; Rapid Selective Staining by Spirit Blue

Decapitate the anther and squeeze out its contents into a drop of water on a clean slide coated with Haupt's adhesive. Let slides air dry and stain the preparations for 4-6 hr in 0.005% spirit-soluble aniline blue, prepared in 50% ethanol. Pass the slides through acetone, 10 min; 1:1 mixture of acetone and xylene, 5 min; and xylene. Mount in a resinous medium. The technique is effective for both fresh anthers and anthers fixed in FAA, Carnoy's fluid, 1:3 acetic alcohol, and 10% formalin (commercial). For fixed anthers, follow customary methods of paraffin embedding and microtomy.     

The Peracetic-Orcein-Halmi Stain: A Stain for Connective Tissues

A selective stain useful for the study of connective tissues is described. The stain demonstrates elastic and oxytalan fibers as well as fibrils in mucous connective tissues previously undescribed. Reticular fibers are not stained. The stain may be used on sections that have been fresh frozen or fixed in formalin or ethanol. Sections are deparaffinized, washed in absolute ethanol, oxidized in peracetic acid 30 min, washed in running water, stained in Taenzer-Unna orcein 15 min, 37°C, differentiated in 70% ethanol, washed in running water, stained in Lillie-Mayer alum hematoxylin 4 min, blued in running water, and counterstained 20 sec in a modified Halmi mixture of 100 ml distilled water, 0.2 gm light green SF, 1.0 gm orange G, 0.5 gm phosphotungstic acid and 1.0 ml glacial acetic acid. Sections are rinsed briefly in 0.2% acetic acid in 95% ethanol, dehydrated and mounted.     

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 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. H2SO4 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 Department Devoted to Abstracts of Books and Papers from Other Journals Dealing With Stains and Microscopic Technic in General

TO enable staining of insoluble calcium salts with glyoxal bis(2-hydroxyanil) (GBHA), the original solution containing 2 ml of 0.4% GBHA in absolute ethanol, and 0.3 ml of aqueous 5% NaOH, and limited to staining only soluble calcium salts, was modified as follows: 1, 2 ml of 0.4% GBHA in absolute ethanol in 0.6 ml of 10% aqueous NaOH; 11, 0.1 gm GBHA in 2 ml of 3.4% NaOH in 75% ethanol. To prevent diffusion and loss of calcium, the tissues were processed by the freeze-substitution or freeze-dry method and sections stained without removing the paraffin. Modification I is effective only when 1 or 2 drops placed on the section are evaporated gradually to dryness, concentrating the GBHA and NaOH on the insoluble calcium salts. Modification II is effective when dried or poured on the the section and allowed to stain for 5 min. The stained slides are immersed for 15 min in 90% ethanol saturated with KCN and Na₂CO₃ for specificity to calcium; rinsed and counterstained in 95% ethanol containing 0.1% each of fast green FCF and methylene blue; rinsed and dehydrated in ethanol; deparaffinized and cleared in xylene; and mounted in neutral synthetic resin. Although the modified methods tested on models failed to stain reagent grade CaCO₃ and Ca₃(PO₄)₂ crystals completely, apatite in developing vertebrae and calcified plaques in soft tissues were stained intensely red. The distribution of gross deposits of insoluble calcium salt in tissue sections corresponded with that shown in adjacent sections by the alizarin red S, ferrocyanide, and von Kossa methods. The modified GBHA method revealed smaller quantities of insoluble as well as soluble calcium salts discretely within cells where the other methods failed; also, calcium in cytoplasm of hypertrophied cartilage cells of developing vertebrae, and in cytoplasm of renal tubular cells of magnesium-deficient rats, not described previously, was demonstrated.     

The Glyoxal BIS(2-Hydroxyanil) Method Modified for Localizing Insoluble Calcium Salts

TO enable staining of insoluble calcium salts with glyoxal bis(2-hydroxyanil) (GBHA), the original solution containing 2 ml of 0.4% GBHA in absolute ethanol, and 0.3 ml of aqueous 5% NaOH, and limited to staining only soluble calcium salts, was modified as follows: 1, 2 ml of 0.4% GBHA in absolute ethanol in 0.6 ml of 10% aqueous NaOH; 11, 0.1 gm GBHA in 2 ml of 3.4% NaOH in 75% ethanol. To prevent diffusion and loss of calcium, the tissues were processed by the freeze-substitution or freeze-dry method and sections stained without removing the paraffin. Modification I is effective only when 1 or 2 drops placed on the section are evaporated gradually to dryness, concentrating the GBHA and NaOH on the insoluble calcium salts. Modification II is effective when dried or poured on the the section and allowed to stain for 5 min. The stained slides are immersed for 15 min in 90% ethanol saturated with KCN and Na₂CO₃ for specificity to calcium; rinsed and counterstained in 95% ethanol containing 0.1% each of fast green FCF and methylene blue; rinsed and dehydrated in ethanol; deparaffinized and cleared in xylene; and mounted in neutral synthetic resin. Although the modified methods tested on models failed to stain reagent grade CaCO₃ and Ca₃(PO₄)₂ crystals completely, apatite in developing vertebrae and calcified plaques in soft tissues were stained intensely red. The distribution of gross deposits of insoluble calcium salt in tissue sections corresponded with that shown in adjacent sections by the alizarin red S, ferrocyanide, and von Kossa methods. The modified GBHA method revealed smaller quantities of insoluble as well as soluble calcium salts discretely within cells where the other methods failed; also, calcium in cytoplasm of hypertrophied cartilage cells of developing vertebrae, and in cytoplasm of renal tubular cells of magnesium-deficient rats, not described previously, was demonstrated.     

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.     

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.     

Safranin and Anilin Blue with Delafield's Hematoxylin for Staining Cell Walls in Shoot Apexes

The following technic is suggested for staining cell walls in shoot apexes: After the usual preliminary steps through 50% ethyl alcohol, stain in 1 % safranin 0 for 24 hours. Rinse in tap water and place in 2% aqueous tannic acid for 2 minutes. After rinsing in tap water, stain for 2 minutes in 1 part Delafield's hematoxylin to 2 parts distilled water and rinse in tap water. Remove excess hematoxylin with acidified water (1 drop cone. HC1 in 200 ml. water), then place slides in 0.5% lithium carbonate for 5 minutes. Dehydrate through an ethyl alcohol series, then transfer from absolute alcohol to a saturated solution of anilin blue in “methyl cellosolve” for 5-10 minutes. Wash in absolute alcohol, rinse in a solution of 25% methyl salicylate, 33% xylene, 42% absolute ethyl alcohol and clear for 10 minutes in a solution of 2 parts methyl salicylate, 1 part xylene, 1 part absolute ethyl alcohol. Transfer through two changes of xylene and mount in “clarite” or suitable alternate. The resulting preparations will have clearly defined, dark-staining cell walls and will photograph well when “Super Panchro-Press, Type B” film (Eastman Kodak Co.) is used in conjunction with suitable Wratten filters.     

Adaptation of the Millon, Sudan Black B, and Periodic Acid-Schiff Technics for Block Staining of Insect Tissues

Spermatophores and reproductive systems of the beetle, Lytta nuttalli Say, fixed in Bouin's aqueous picroformol or buffered 10% neutral formol were stained in toto by the Millon, Sudan black B and periodic acid-Schiff reactions as follows. Millon: after excess fixative is removed in 70% ethanol, specimens are brought to water, stained in Millon's reagent at 60 C for 1 hr, rinsed in 2% aqueous nitric acid at 40-50 C, dehydrated rapidly, cleared, embedded and sectioned as usual. Sudan black B: specimens are taken to absolute ethanol, stained in a saturated solution of Sudan black B in absolute ethanol at room temperature for 24-48 hr, rinsed and cleared in xylene, embedded and sectioned. PAS: specimens are brought to water, oxidized in 0.5 aqueous HIO₄ at 37 C for 30 min, washed in 2 changes of water, stained in Schiif reagent at room temperature for 1 hr, rinsed in 3 changes of 0.5% aqueous potassium metabisulfite, washed in running water for 10-15 min, dehydrated, cleared, embedded and sectioned. All 3 methods produced their characteristic staining in specimens up to 3 mm thick