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 Versatile New Mineralized Bone Stain for Simultaneous Assessment of Tetracycline and Osteoid Seams

A versatile mineralized bone stain (MIBS) for demonstrating osteoid seams and tetracycline fluorescence simultaneously in thin or thick undecalcified sections has been developed. Bone specimens are fixed in 70% ethanol, but 10% buffered formalin is permissible. Depending upon one's preference, these specimens can be left unstained or be prestained before plastic embedding. Osteoid seams are stained green to jade green, or light to dark purple. Mineralized bone matrix is unstained or green. Osteoblast and osteoclast nuclei are light to dark purple, cytoplasm varies from slightly gray to pink. The identification of osteoid seams by this method agrees closely with identification by in vivo tetracycline uptake using the same section from the same biopsy. The method demonstrates halo volumes, an abnormal, lacunar, low density bone around viable osteocytes in purple. This phenomenon is commonly seen in vitamin D-resistant rickets, fluorosis, renal osteodystrophy, hyperparathyroidism, and is sometimes seen in fluoride treated osteoporotic patients. In osteomalacic bone, most osteoid seams are irregularly stained as indicated by the presence of unmineralized osteoid between mineralized lamellae. The method has been used effectively in staining new bone formation in hydroxyapatite implants and bone grafts. Old, unstained, plastic embedded undecalcified sections are stained as well as fresh sections after removal of the coverslip. This stain also promises to be valuable in the study of different metabolic bone diseases from the point of view of remodeling, histomorphometry, and pathology.     

Solochrome Cyanine R As An Indicator Dye of Bone Morphology

Sections of undemineralized bone embedded in a polyester resin and cut at 6 μ are stained for 10 min, without removal of the embedding matrix, in an aqueous solution composed of Solochrome cyanine R, 1 gin; glacial acetic acid, 2 ml; and distilled water, 98 ml. A pH about 2 is obtained by the acetic acid. The sections are washed and differentiated in tap water at 30 C, dehydrated in ascending alcohols, cleared and mounted in synthetic resin. “Young osteoid” stains light orange and, in the rest of an osteoid seam, two types of lamellae can be distinguished: one blue layer of ground substance or collagen and one orange layer of fibrillar collagen. The “calcification front” is sharply demarcated by its dark blue color.     

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.     

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.     

Basic Two-Dye Stains for Epoxy-Embedded 0.3-1 μ Sections

Dyes used in the 3 methods recommended are: I, thionin and acridine orange (T-AO); II, Janus green and Darrow red (JG-DR); III, methyl green and methyl violet (MG-MV). The first 2 methods were two-solution stains, applied in sequence; the third, required only one solution since methyl violet is present in commercial methyl green. Staining solution and timing was as follows: Method I. 0.1% thionin in a 45% ethanolic solution of 0.01 N NaOH, 5 min at 70 C; rinsing in water and followed by 1 min in a 1% aqueous solution of acridine orange made up in 0.02 N NaOH, also at 70 C, then washed, and dried on slides. Method II. 0.5% Janus green in aqueous 0.05 N NaOH, 5 min at 70 C; rinsing in water then into 0.5% Darrow red in 0.05 N NaOH (aq.), 2 min at 70 C., washing, and drying on slides. Method III. 1% methyl green (commercial, unpurified) in 1% aqueous borax for 15-20 min at 20-25 C, washing and attaching to slides. All staining was performed by floating the sections on the staining solutions, all drying at 70 C, and mounting in a resinous medium. T-AO gave blue to violet cytoplasmic structures, darker nuclei which contrasted strongly with yellow connective tissue and the secretion of goblet cells. JG-DR resembled a hematoxylineosin stain, but by shortening the staining time in DR to 0.5-1 min, collagenous and elastic tissue retained more of the green dye. MG-MV gave dark green nuclei in light green cytoplasm, with collagenous and elastic tissues in blue to violet. As with most methods for staining ultrathin sections, thicknesses of less than 1 μ required longer staining times.     

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.     

A Technic for Differential Staining of Nucleoli and Chromosomes

A procedure is described which enables a stain to be definitely located in the substance of the nucleolus. Material is fixed in either Navashin or Levitsky; the chromatin is stained by means of the improved Feulgen technic introduced by de Tomasi, and preparations brought thru the washing solutions down to distilled water. From distilled water the material is transferred to a mordant solution, 5% sodium carbonate in water, in which it is left for at least one hour. After mordanting wash well with water then stain for ten minutes in light green solution (90% alcohol, 100 cc, light green SFY, 0.5 g, aniline oil, 2 drops, well shaken); differentiate in alcoholic sodium carbonate solution, (70% alcohol saturated with carbonate); treat with 95% alcohol, absolute alcohol, equal parts xylene and absolute alcohol, clear in pure dry xylene and mount in neutral balsam. Cytoplasm and karyolymph should be quite clear, with magenta chromatin and well defined green nucleoli. The light green does not behave like a simple counterstain as in previous technics but as a definite stain for nucleolar material.     

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.     

Suggested Counterstains for Davenport Reduced Silver Preparations of Peripheral Nerves

—Peripheral nerves which have been fixed in a mixture of formaldehyde and acetic acid and stained according to the method of Davenport can be successfully counterstained for demonstration of myelin sheaths and stroma. After mounted sections have been silvered, reduced and toned, the coating of nitrocellulose is removed by passing thru two changes of acetone. Following brief washes in 100,95,85 and 75% alcohols they are stained in an acidified aqueous solution of azo carmine for 30 to 60 minutes. Excess azo carmine is extracted with anilin alcohol followed by acetic alcohol after which the sections are mordanted for 15 to 60 minutes in a 5% aqueous solution of phosphotungstic acid. Without washing they are transferred to a stain mixture of either anilin blue and orange G (acidified) or light green and orange G (acidified) where they remain from 1 to 5 hours. After destaining in 95% alcohol and dehydration in absolute alcohol the sections are mounted in dammar. Result: axons stain black; sheath and fibroblast nuclei, red; myelin sheaths, orange; and connective tissue, blue or green. When the counterstains are applied to ganglia, cytological details of individual cells are demonstrated.     

Orcein-Hematoxylin in Iodized Ferric Chloride as a Stain for Elastic Fibers, With Metanil Yellow Counterstaining

Autopsy and biopsy specimens of human skin were fixed overnight in alcoholic Bouin's solution, embedded in paraffin, cut at 7 μ, deparaffinized, hydrated to 70% alcohol, and treated as follows—stained 2 hours in a mixture consisting of: 0.2% orcein in 70% alcohol and 1% HC1 (conc.), 125 ml; 5% hematoxylin in absolute alcohol, 40 ml; 6% FeCl₃ in water, 25 ml; and aqueous I₂-KI (1:2:100), 25 ml—rinsed in distilled water until the excess stain was removed—differentiated in 1.2% FeCl₃, 5-15 sec—washed in running water, 5 min—differentiation completed in 0.01% HC1 acid-alcohol, 1 min—a dip in 95% alcohol—distilled water, 2 min—0.25% aqueous metanil yellow, 5-10 sec—a 95% alcohol dip—dehydrated in absolute alcohol, xylene, and mounted in a resinous medium. The technic combines the orcein of Pinkus' stain and the hematoxylin mixture of Verhoeff into a single staining solution and gives sharp and reliable results for both coarse and extremely delicate elastic fibers. These stain purple; nuclei, violet; and background, yellow. The stain allows the use of formalin, Bouin's fluid and Zenker-formol fixation. The results have been consistent in other primates as well as in man.     

Staining Sex Chromatin; Biebrich Scarlet and Fast Green Fcf as a Mixture Versus Their Use in Sequence

Human skin was fixed in Davidson's solution (95% alcohol, 35; formalin, 20; glacial acetic acid, 10; and distilled water, 35—parts by volume) and sections prepared through paraffin embedding in the usual manner. Stock stains were: I(BS)—Biebrich scarlet, 1 gm in 100 ml of 50% alcohol to which 0.3 gm of phosphotungstic acid and 5 ml of glacial acetic acid were added—and II(FG)—fast green, 0.5 gm in 85 ml of 50% alcohol to which 0.3 gm of phosphotungstic acid, 0.3 gm of phosphomolybdic acid, and 15 ml of glacial acetic acid were added. Experimental staining solutions were prepared in the following proportions of stock BS to stock FG—1:1, 2:1, 3:1, 1:2 and 1:3. Sections were brought to 50% alcohol and stained for 15, 20, 25 and 30 min in each of the five BS-FG mixtures, rinsed in 50% alcohol, then dehydrated in 70%, 95%, and absolute alcohol, 2 min each; cleared in xylene, and covered in balsam. The 2:1 (optimum proportion) combination of BS with FG, acting for 20 min, yielded 97% sex chromatin-positive nuclei in female material. If sections were stained in stock solution BS for 2 min, they could be differentiated by a 20 min treatment in the mordanting component of stock FG (without dye) to give a one-color stain. Such stains gave about the same percentage of sex chromatin-positive nuclei as those obtained by the regular two-color procedure. These modifications are simpler, more rapid, and yield results comparable to previously employed techniques.     

A Thionin Stain for Visualizing Bone Cells, Mineralizing Fronts and Cement Lines in Undecalcified Bone Sections

A staining method is described using thionin, for undecalcified deacrylated bone sections. RNA is stained purplish violet, allowing still active osteoblasts to be distinguished from lining cells. Staining intensity of mineralized bone is related to the degree of mineralization. Mineralizing fronts and cement lines are visualized clearly. Lamellae show an alternate pattern. Histomorphometric parameters such as osteon thickness and interstitial bone thickness can be measured without using polarized light. The mineralizing front can be assessed and expressed as a percentage of the osteoblast-covered interface between osteoid and mineralized bone. The stain is also useful for qualitative assessment of metabolic bone disease. Thionin stained sections can be kept for at least one year when stored hi the dark at 7 C.     

A Nile Blue Staining Technic for the Differentiation of Melanin and Lipofuscins

When paraffin sections are stained in 0.05-.01% Nile blue in 1 % sulfuric acid, washed thoroughly in water and mounted in aqueous media, lipofuscins color deep blue, melanins dark green, myelin and red cells lighter greens and background pale green. If, immediately after staining, the preparations are at once extracted with acetone with or without a 1% sulfuric acid rinse, melanins remain dark green, mast cells color purple, lipofuscins and background decolorize and nuclei may stain light green.     

A Nile Blue Staining Technic for the Differentiation of Melanin and Lipofuscins

When paraffin sections are stained in 0.05-.01% Nile blue in 1 % sulfuric acid, washed thoroughly in water and mounted in aqueous media, lipofuscins color deep blue, melanins dark green, myelin and red cells lighter greens and background pale green. If, immediately after staining, the preparations are at once extracted with acetone with or without a 1% sulfuric acid rinse, melanins remain dark green, mast cells color purple, lipofuscins and background decolorize and nuclei may stain light green.     

Staining Pericellular Boutons of Glutaraldehyde-Fixed Nervous Tissue; A Chromation-Silvering Sequence for Frozen Sections

After fixing in phosphate-buffered 5% glutaraldehyde, pH 6.8, by perfusion, brains were sliced to 3-5 mm pieces which were placed in the fixative for 5-7 days. The pieces were washed through several changes of 2.26% NaH₂PO₄ for 12 hr, 30 μ frozen sections cut, and mordanted 2 days in an equal-parts mixture of 3.5% CrO₃ and 5% Na-tartrate, which had been aged at 20-25 C for 20 days prior to use. After washing in distilled water, the sections were put into a solution containing AgNO₃, 20 gm; and KNO₃, 15 gm, in distilled water, 80 ml; at 30 C for 1.5-2 hr, then reduced at 40-45 C in three pyrogallol solutions as follows: 1-2 sec in 1% pyrogallol in 55% alcohol; 3-4 sec in a 0.67% solution in 33% alcohol, and 5-7 sec in a 0.5% solution in 25% alcohol. Gold toning is optional; dehydration, clearing and covering, routine. The technic shows particularly the perisomatic fibers, boutons en passant and boutons termineaux. Fibers in nerve tracts may be visible but lightly stained; cell nuclei may be dark, but the cytoplasm remains pale.     

Differentiated Staining of Osmium Tetroxide-Fixed, Vestopal-w-Embedded Tissue in thin Sections

Tissues were fixed at 20° C for 1 hr in 1% OsO₄, buffered at pH 7.4 with veronal-acetate (Palade's fixative), soaked 5 min in the same buffer without OsO₄, then dehydrated in buffer-acetone mixtures of 30, 50, 75 and 90% acetone content, and finally in anhydrous acetone. Infiltration was accomplished through Vestopal-W-acetone mixtures of 1:3, 1:1, 3:1 to undiluted Vestopal. After polymerisation at 60° C for 24 hr, 1-2 μ sections were cut, dried on slides without adhesive, and stained by any of the following methods. (1) Mayer's acid hemalum: Flood the slides with the staining solution and allow to stand at 20°C for 2-3 hr while the water of the solution evaporates; wash in distilled water, 2 min; differentiate in 1% HCl; rinse 1-2 sec in 10% NH,OH. (2) Iron-trioxyhematein (of Hansen): Apply the staining solution as in method 1; wash 3-5 min in 5% acetic acid; restain for 1-12 hr by flooding with a mixture consisting of staining solution, 2 parts, and 1 part of a 1:1 mixture of 2% acetic acid and 2% H₂SO₄ (observe under microscope for staining intensity); wash 2 min in distilled water and 1 hr in tap water. (3) Iron-hematoxylin (Heidenhain): Mordant 6 hr in 2.5% iron-alum solution; wash 1 min in distilled water; stain in 1% or 0.5% ripened hematoxylin for 3-12 br; differentiate 8 min in 2.5%, and 15 min in 1% iron-alum solution; wash 1 hr in tap water. (4) Aceto-carmine (Schneider): Stain 12-24 hr; wash 0.5-1.0 min in distilled water. (5) Picrofuchsin: Stain 24-48 hr in 1% acid fuchsin dissolved in saturated aqueous picric acid; differentiate for only 1-2 sec in 96% ethanol. (6) Modified Giemsa: Mix 640 ml of a solution of 9.08 gm KH₂PO₄ in 1000 ml of distilled water and 360 ml of a solution of 11.88 gm Na₂HPO₄-2H₂O in 1000 ml of distilled water. Soak sections in this buffer, 12 hr. Dissolve 1.0 gm of azur I in 125 ml of boiling distilled water; add 0.5 gm of methylene blue; filter and add hot distilled water until a volume of 250 ml is reached (solution “AM”). Dissolve 1.5 gm of eosin, yellowish, in 250 ml of hot distilled water; filter (solution “E”). Mix 1.5 ml of “AM” in 100 ml of buffer with 3 ml of “E” in 100 ml of buffer. Stain 12-24 hr. Differentiate 3 sec in 25 ml methyl benzoate in 75 ml dioxane; 3 sec in 35 ml methyl benzoate in 65 ml acetone; 3 sec in 30 ml acetone in 70 ml methyl benzoate; and 3 sec in 5 ml acetone in 95 ml methyl benzoate. Dehydrated sections may be covered in a neutral synthetic resin (Caedax was used).     

Combined Stain for Fish Pituitary

A new combined stain is described for the study of cell types in the fish pituitary. Tissues are prepared by fixing in formol-sublimate and then embedded in win wax. Tissue is sectioned at 5 μm and then stained sequentially with performic acid-alcian blue, periodic acid-Schiff, orange G, and acid fuchsin As a result of this procedure acidophils stain as follows: lactotropes, red; corticotropes, light pink melanotropes, bright pink and somatotropes, orange. Cyanophils stain either magenta red (gonadotropes) or blue (thyrotropes). Neurosecretory material and the fibers of the pars nervosa which penetrate the pars intermedia stain light blue.     

Differential Staining of Aborted and Nonaborted Pollen

A single staining solution was made by compounding it in the following order (dyes were from British Drug Houses): ethanol, 10 ml; 1% malachite green in 95% ethanol, 1 ml; distilled water, 50 ml; glycerol 25 ml; phenol, 5 gm; chloral hydrate, 5 gm; acid fuchsin 1% in water, 5 ml; orange G, 1% in water 0.5 ml; and glacial acetic acid, 1-4 ml. For best results in differentiation to give green pollen walls and red protoplasm, the staining solution should be acidified with glacial acetic acid. The amount of acid to be added depends upon thickness of the pollen walls: for very thin-walled pollen, 1 ml; for moderately thin walls, 2 ml; and for thick-walled or spiny-walled pollen, 3 ml of acid. For pollen inside non-dehiscent anthers, 4 ml of acid should be used. Staining is hastened by flaming the slide (for loose thin-walled pollen) or by immersing thick-walled pollen or anthers for 24-48 hr at 50 C. In the typical stain, aborted pollen grains are green; nonaborted, red. The method is useful for pollen inside nondehiscent anthers if these are small and not too deeply coloured naturally. The stain is very durable, especially if the coverslips are sealed with param wax. The staining solution will keep well for about a month. It is useful both for angiosperms and gymnosperm microgametes.     

Calcium in Cells of Fresh Bone Stained with Glyoxal Bis(2-Hydroxyanil)

Fresh ileum of adult rats and vertebrae and calvariae of newborn rats were immersed in a staining solution containing 0.1 gm of glyoxal bis(2-hydroxyanil) (GBHA) per 2 ml of 3.4% NaOH in 75% ethanol, dehydrated in absolute ethanol, cleared in xylene, and embedded in paraffin. Paraffin sections of stained material, 7 μ thick, were affixed to albumenized slides, immersed in 90% ethanol saturated with Na₂CO₄ and KCN to ensure specificity for calcium, rinsed in 95% ethanol, counterstained in 50% ethanol containing 0.1% methylene blue, dehydrated in absolute ethanol, deparaffinized and cleared in xylene, and mounted in neutral synthetic resin. By this procedure, red Ca-GBHA granules were deposited in goblet and Paneth cells, and in the cytoplasm of osteoblasts, osteocytes, chondrocytes, and periosteal cells of developing bones. Calcium in apatite did not stain. In osseous tissues sectioned in a cryostat or processed by the freeze-dry or freeze-substitution method, epiphyseal chondrocyte calcium was removed, and apatite stained so intensely red that it obscured calcium in the bone cells. Failure of control osseous tissues to stain after immersion in a 1% solution of disodium salt of ethylenediaminetetraacetic acid or in the alcoholic, alkaline solvent of the GBHA solution, indicated that the red granules in the cells of developing bone were due to calcium present in the cells in vivo and not due to absorption of GBHA by tissue components other than calcium, or to diffusion of Ca⁺⁺ during staining. Calcium localized in the cytoplasm and processes of the osteogenic cells suggests the need to re-evaluate the role of osteoblasts as depositors of calcium during osteogenesis.