A Tribasic Stain for Thin Sections of Plastic-Embedded, OsO4-Fixed Tissues

Thin (0.5-1 μ) sections of plastic-embedded, OsO₄-fixed tissues were attached to glass slides by heating to 70 C for 1 min. A saturated solution combining toluidine blue and malachite green was prepared in ethanol (8% of each dye) or water (4% of each dye). Methacrylate or epoxy sections were stained in the ethanol solution for 2-5 min. The water solution was more effective for some epoxy sections (10-80 min). Epoxy sections could be mordanted by 2% KMnO₄, in acetone (1 min) before use of the aqueous dye, reducing staining time to 5-10 min and improving contrast. Aqueous basic fuchsin (4%) was used as the counter-stain in all cases; staining time varied from 1-30 min depending upon the embedding medium and desired effects, methacrylate sections requiring the least time. In the completed stain, nuclei were blue to violet; erythrocytes and mitochondria, green; collagen and elastic tissue, magenta; and much and cartilage, bright cherry red. Sections were coated with an acrylic resin spray and examined or photographed with an oil-immersion lens.     

The staining of elastic fibres with Direct Blue 152. A general hypothesis for the staining of elastic fibres

Summary Elastic fibres may be stained by a number of dyes, e.g. Direct Blue 1 (C.I. 24410), Direct Blue 10 (C.I. 24340), Direct Blue 15 (C.I. 24400), Direct Blue 152 (C.I. 24366) and Direct Violet 37 (C.I. 24370). A convenient method using Direct Blue 152 has been developed which is specific for elastic fibres. The method is simple and allows the demonstration of other connective tissue fibres. Staining of elastic fibres is unimpaired by numerous blocking procedures or by changes in dyebath pH. These properties are shared by several standard elastic fibre stains.As the Direct dyes and several of the standard elastic fibre stains possess numerous aromatic rings a wide range of dyes containing varying numbers of aromatic rings were examined for ability to stain elastic fibres. No association was observed between the ability to stain elastic fibres and dye class, formal charge or the presence of hydrogen bonding groups. Staining was, however, definitely associated with the presence in the dye molecule of 5 or more aromatic rings. This suggested that van der Waals forces of attraction may be responsible for elastic fibre staining both by Direct dyes and the standard elastic fibre stains. Staining of elastic fibres as a side-effect in many procedures is similarly explicable.     

Improved Coomassie Blue Dye-Based Fast Staining Protocol for Proteins Separated by SDS-PAGE

The time required to visualize proteins using Coomassie Blue dye has been significantly reduced with the introduction of fast staining protocols based on staining with a Coomassie Blue dye solution at boiling temperatures. However, fast stainings suffer from high gel backgrounds, reducing the signal-to-noise ratio and limiting the number of detectable spots in the case of 2D SDS-PAGE. The aim of this work was to eliminate the high gel background, and thus improve fast staining protocols based on Coomassie Blue dye. We show that merely replacing water with a 4 mM EDTA washing solution at boiling temperatures, results in a transparent gel background within 50 to 60 minutes of destaining. Moreover, when a combination of imidazole-zinc reverse staining and Coomassie Blue-based fast staining is used the sensitivity is improved significantly; nanogram amounts of proteins can be detected using 1D SDS-PAGE, and about 30% to 60% more spots can be detected with 2D SDS-PAGE in plasma, platelet, and rat brain tissue samples. This work represents an optimized fast staining protocol with improved sensitivity, requiring between 60 to 75 minutes to complete protein visualization.     

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.     

Current applications of orcein in histochemistry. A brief review with some new observations concerning influence of dye batch variation and aging of dye solutions on staining

Current uses of orcein to demonstrate elastic fibers and, following permanganate oxidation (Shikata's modification), hepatitis B surface antigen, copper associated protein, and sulfated mucins, are reviewed. Variations in staining performance with batch of dye and age of dye solution is also discussed. Additional experimental findings support the view that the orcein stain for elastic tissue and Shikata's modification produces consistent, high quality results as long as appropriate controls and suitable dye batches, e.g., Biological Stain Commission certified dyes, are used.     

Effects of Ph on Post-Mortem Retention of Trypan Blue Vital Staining in Tissues of Mice

Vital staining of aortas from mice injected subcutaneously (daily for 5 days) with trypan blue was studied. In routine paraffin sections elastic membranes were observed to be well stained and other medial elements unstained following fixation in 10% formaldehyde (25% formalin) at pH 7-9. An identical pattern of vital staining was observed in specimens that had been immersed for 48 hr in saline solutions at pH 7-11. Elastic membranes were not stained, but intermembranous connective tissue was stained after the following: (1) fixation in 10% formaldehyde at pH 1-4 and in Lavdowsky's solution (ethanol, formaldehyde, water and glacial acetic acid), pH 2.3-2.8; and (2) immersion in saline for 48 hr at pH 14. Aortic elastic membranes were vitally stained after fixation by intracardiac perfusion with 10% formaldehyde (pH 7-8) but not after perhion with Lavdowsky's fixative (pH 2.3-2.8). Vital staining was limited to medial elastic membranes in sections of fresh aorta made in a cryostat or by a regular freezing microtome. The vital staining (coarse cytoplasmic granules of dye) within macrophages (Kupffer cells and others) and in cytoplasm of renal tubular epithelium was well demonstrated following use of all methods discussed above     

Thermodynamics of Orcein staining of elastic fibres

Synopsis Elastic fibres in histological sections have only a slightly higher affinity (than chromatin or cartilage matrix) for unpurified Orcein in acidified 70% ethanol, but the staining of elastic fibres is more exothermic (the heat of staining being in good agreement with publishedin vitro measurements), has a considerably higher activation energy, and is probably accompanied by a greater decrease in entropy. Experiments with purified dye fractions, and unpurified dye in 10% ethanol, were inconclusive, as it was not possible to prove unequivocally that equilibrium between dyebath and substrate had been achieved under these conditions.The results are consistent with the selectivity of orcein for elastic fibres under practical conditions being due to (a) the presence in elastic fibres of a relatively large number of dye-binding sites per unit volume, which probably bind by some non-ionic mechanism, (b) the relatively non-polar nature of elastic fibres, which repel cationic dye particles less than do tissue components that at low pH carry a positive charge, and (c) the low permeability of elastic fibres, so that dyeing, once achieved, is relatively resistant to alcoholic extraction. An alcoholic solvent for the dye enables strong solutions, and hence short staining times, to be used.     

Current applications of orcein in histochemistry. A brief review with some new observations concerning influence of dye batch variation and aging of dye solutions on staining

Current uses of orcein to demonstrate elastic fibers and, following permanganate oxidation (Shikata's modification), hepatitis B surface antigen, copper associated protein, and sulfated mucins, are reviewed. Variations in staining performance with batch of dye and age of dye solution is also discussed. Additional experimental findings support the view that the orcein stain for elastic tissue and Shikata's modification produces consistent, high quality results as long as appropriate controls and suitable dye batches, e.g., Biological Stain Commission certified dyes, are used.     

A Note on the Purification of Alcian Blue

Alcian blue 8GX is a copper phthalocyanin dye that shows a high degree of specificity for polyanionic substances such as hyaluronic acid, sialic acid and the chondroitin sulfates. This dye has proved useful for both histochemical and electrophoretic staining of these substances. The Biological Stain Commission has recently begun to certify Alcian blue (Schenk 1981). Commercially available lots contain approximately 50% dye. The remaining constituents have been identified as primarily boric acid, as well as sulfates and dextrins (Scott 1972, Horobin and Goldstein 1972). Horobin and Goldstein (1972) have pointed out that these contaminants may adversely affect staining in the critical electrolyte concentration procedure. Scott (1972), while not ascribing any adverse effects to the presence of boric acid, recommends its removal by differential precipitation with acetone. In this procedure one part of a 2-5% aqueous solution of the dye is added to 5-10 parts of acetone. The precipitated dye is approximately 80% pure. While this method is relatively simple, it does have several drawbacks. Low concentrations of Alcian blue (i.e., 2%) must be used to obtain purities near 80%. Thus a minimum of 250 ml of acetone is needed to purify 1 gram of dye. Furthermore, Horobin and Goldstein (1972) have reported that contamination by dextrin or unknown organic substances (detergent?) interferes with precipitation of the dye enough to make purification by Scott's method impossible. When difficulty in the precipitation of Alcian blue by Scott's method was encountered, the following simple method for the purification of the dye was developed.     

A Note on the Purification of Alcian Blue

Alcian blue 8GX is a copper phthalocyanin dye that shows a high degree of specificity for polyanionic substances such as hyaluronic acid, sialic acid and the chondroitin sulfates. This dye has proved useful for both histochemical and electrophoretic staining of these substances. The Biological Stain Commission has recently begun to certify Alcian blue (Schenk 1981). Commercially available lots contain approximately 50% dye. The remaining constituents have been identified as primarily boric acid, as well as sulfates and dextrins (Scott 1972, Horobin and Goldstein 1972). Horobin and Goldstein (1972) have pointed out that these contaminants may adversely affect staining in the critical electrolyte concentration procedure. Scott (1972), while not ascribing any adverse effects to the presence of boric acid, recommends its removal by differential precipitation with acetone. In this procedure one part of a 2-5% aqueous solution of the dye is added to 5-10 parts of acetone. The precipitated dye is approximately 80% pure. While this method is relatively simple, it does have several drawbacks. Low concentrations of Alcian blue (i.e., 2%) must be used to obtain purities near 80%. Thus a minimum of 250 ml of acetone is needed to purify 1 gram of dye. Furthermore, Horobin and Goldstein (1972) have reported that contamination by dextrin or unknown organic substances (detergent?) interferes with precipitation of the dye enough to make purification by Scott's method impossible. When difficulty in the precipitation of Alcian blue by Scott's method was encountered, the following simple method for the purification of the dye was developed.     

The use of a basic dye (azure A or toluidine blue) plus a cationic surfactant for selective staining of RNA: a technical and mechanistic study.

Selective purple staining of RNA-rich structures such as basophilic cytoplasms of exocrine pancreas and plasma cells, Nissl substance, and nucleoli was achieved by treating tissue sections as follows. Stain dewaxed sections for 1/2 hour in a dyebath containing 0.1% w/v axure A or toluidine blue and 1% cationic surfactant (Hyamine 2389, a 50% w/v aqueous solution of diisobutylphenoxyethoxyethyldimethylbenzylammonium chloride; or benzyldimethylammonium chloride, or cetylpyridinium bromide, or cetyltrimethylammonium bromide) buffered to pH 7 with phosphate. Rinse in water, blot, air dry and mount in synthetic resin. Intense purple staining of RNA-rich regions occurred after fixation in neutral formalin or in Carnoy's or Gendre's fluids, though satisfactory results were also found after fixation in acetone or alcohol. Chromatin generally stained a very pale azure after all fixations, though occasionally nuclei were unstained (Gendre's or Zenker's fluids). Subjecting tissue sections to acid hydrolysis or to digestion by RNAase eliminated or reduced the purple staining, but left the azure staining of nuclei unaffected. Satisfactory staining of RNA-rich structures was not critically dependent on the precise concentrations of dye, surfactant or inorganic salts in the dyebath, nor on pH, staining time or chemical nature of the surfactant. The staining patterns can be rationalized with a tissue model that considers both surface charge and permeability factors, since present in the dyebath are small dye cations and large cationic surfactant micelles. As micelles and dye will both quickly penetrate basophilic structures considered to be porous, such as chromatin, competition will then greatly reduce staining of such substrates. But the large micelles will only slowly penetrate regions considered to be more impermeable, such as basophilic cytoplasms, so consequently small fast moving dye ions may enter and stain without competition.     

Iron gallein elastic method---a substitute for Verhoeff's elastic tissue stain.

A method for staining elastic fibers in formalin fixed, paraffin embedded sections is described. After deparaffinizing and dehydration, sections are stained for 30 minutes in a solution prepared by mixing equal parts of 1% gallein dissolved in ethylene glycol and absolute alcohol (1:4), and 1.16% aqueous ferric chloride in 1% hydrochloric acid. The sections are washed in water and then differentiated in 2% ferric chloride for 2 minutes. After washing in water, the sections are counterstained with a variant of Van Gieson's picric acid-acid fuchsin for 1 minute. The results are similar to Verhoeff's elastic stain with elastic fibers staining black. An advantage to this staining procedure is that visually controlled differentiation is not necessary.     

Methods of Staining Plant Tissues for Differentiating the Natural Plant Oils from Petroleum Spray Oils

Petroleum, spray oils in sections of plant tissue have been distinguished from the plant oils by staining the fresh sections in the following dye solution: To a saturated aqueous solution of Nile blue sulfate, 0.5% sulfuric acid is added and the mixture is boiled under a reflux condenser for 4 or 5 hours. It should be as nearly alkaline as possible without a change of color. A solution of 50% alcohol and 50% acetone is then saturated with oil red O. One part of the Nile blue sulfate solution is then added to two parts of the oil red O solution. Allow to settle over night and filter. Stain several hours. Rinse in water and mount in glycerin jelly. A short discussion of the merits of this method and the differentiation of the spray oils by means of indophenol blue are also given.     

Iron Gallein Elastic Method—A Substitute for Verhoeff'S Elastic Tissue Stain

A method for staining elastic fibers in formalin fixed, paraffin embedded sections is described. After deparaffinizing and dehydration. sections are stained for 30 minutes in a solution prepared by mixing equal parts of 1% gallein dissolved in ethylene glycol and absolute alcohol (1:4), and 1.16% aqueous ferric chloride in 1% hydrochloric acid. The sections are washed in water and then differentiated in 2% ferric chloride for 2 minutes. After washing in water, the sections am counterstained with a variant of Van Girson's picric acid-acid fuchsin for 1 minute. The results are similar to Verhoeff s elastic stain with elastic fibers staining black. An advantage to this staining procedure is that visually controlled differentiation is not necessary.     

A Rhodocyan Technic for Staining the Anterior Pituitary

A reproducible, one-step, differential staining technic which uses routine formalin-fixed tissue and gives brilliantly contrasting results is produced by incubating sections for 1 hr in a 60° C oven in the following dye mixture: 1% eosin B (CI#771), 8 ml; 1% anilin blue (CI#707), 2 ml; and buffer solution (0.1M citric acid, 1.1 ml; 0.2M Na₂HPO₄, 0.9 ml; distilled water, 28.0 ml) at pH 4.5. No differentiation is necessary. The method can be modified for duodenal enterochromaffin cells and alpha cells of pancreatic islets by adjusting the buffer to pH 3.6 and staining for only 3 min at 60° C.     

A Simplified Method for Staining Mast Cells with Astra Blue

The copper phthalocyanin dye astra blue has been used to stain differentially mast cells of the intestine; however, the procedure has not been used widely because of the difficulty in preparing and using the dye solution. Described here is a simple, reliable, and consistent method for selectively staining mast cells using a dye solution that may be prepared in any laboratory without the aid of sophisticated pH metering equipment. Astra blue is mixed with an alcoholic solution containing MgCl₂ · 6H₂O and the pH indicator pararosaniline hydrochloride. Concentrated hydrochloric acid is added dropwise, changing the dye mixture from purple to violet and then to blue. In this low range the weakly ionizing ethanol provides a more stable hydrogen ion concentration than the corresponding aqueous solutions used previously. Alcoholic acid fuchsin is a convenient counterstain, and this simple procedure then provides good contrast between the blue staining mast cell granules and the red tissue background.     

A simplified method for staining mast cells with astra blue

The copper phthalocyanin dye astra blue has been used to stain differentially mast cells of the intestine; however; the procedure has not been used widely because of the difficulty in preparing and using the dye solution. Described here is a simple, reliable, and consistent method for selectively staining mast cells using a dye solution that may be prepared in any laboratory without the aid of sophisticated pH metering equipment. Astra blue is mixed with an alcoholic solution containing MgCl2-6H2O and the pH indicator pararosaniline hydrochloride. Concentrated hydrochloric acid is added dropwise, changing the dye mixture from purple to violet and then to blue. In this low range the weakly ionizing ethanol provides a more stable hydrogen ion concentration than the corresponding aqueous solutions used previously. Alcoholic acid fuchsin is a convenient counterstain, and this simple procedure then provides good contrast between the blue staining mast cell granules and the red tissue background.     

Demonstration of Elastic Tissue and Iron or Elastic Tissue and Calcium Simultaneously

For the concomitant demonstration of iron and elastic tissue Perls' test solution was used, followed by Verhoeff's stain or Gomori's aldehyde fuchsin. When Perls' and Verhoeff's stain were used in sequence, the iron deposits were greenish blue and the elastic lamellae were black. When Perls' test solution was combined with aldehyde fuchsin the iron deposits were blue and elastic tissue purple. Calcium salts and elastic tissue were demonstrated concomitantly by using von Kossa's method followed by Gomori's aldehyde fuchsin. With such combined staining, the calcium salts appeared brownish black and elastic tissue purple. With these procedures, it was possible to see the exact relationship of calcium and iron deposits to the elastic tissue.     

On the mechanism of Verhoeff's elastica stain: a convenient stain for myelin sheaths.

Verhoeff (1908) recommended an iron-hematein formula containing Lugol's solution for demonstration of elastic tissue; sections are differentiated until desired staining patterns are obtained. Verhoeff's stain colored a variety of tissue structures and showed higher substantivity for myelin sheaths than for elastin. Addition of HCL or omission of Lugol's solution decreased or abolished coloration of pseudo-elastica and thus enhanced selectivity for elastin. Substitution of Fe++ for Fe+++ abolished dye binding by elastin. A review of chemical data indicated interaction of components of Lugol's solution with the dye. Hematein and Fe+++ form a variety of cationic, anionic and non-ionic chelates; the ratio of these compounds changes with time. Dye binding apparently occurs mainly via van der Waals forces and hydrogen bonds. Verhoeff's elastica stain is definitely not specific for elastin and is inferior to orcein and resorcin-fuchsin because of the required differentiation with its inherent bias to produce patterns which conform to expectations. However, Verhoeff's elastica stain is far superior to other metal-hematein technics for myelin sheaths. The combined Verhoeff-picro-Sirius Red F3BA stain can be performed in 30 min and does not require differentiation. It is therefore suggested to reclassify Verhoeff's elastica stain as a method for myelin sheaths.     

Propylene and Ethylene Glycol as Solvents for Sudan IV and Sudan Black B

Propylene or ethylene glycol is recommended as a solvent for Sudan IV and Sudan black B to replace the commonly used alcohol-acetone mixtures for general lipid staining in tissue sections. Either glycol is used as a dehydrating agent, dye solvent, and differentiating solution. They offer the advantages of a stable solution, inert with respect to solubilities of lipid material in it, and excellent control of differentiation without loss of dye from lipid particles. Sections remain pliable and are not shrunken by the glycols. Counterstains may be used after staining with Sudan IV but are generally not necessary after staining with Sudan black B. With the use of propylene glycol as a solvent, Sudan IV appears to equal the staining ability of Sudan black B as regards the type of lipid material detected, and the choice of dye to be used would depend on the color contrast desired.