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.     

Basic Dye Counterstaining of Sections Impregnated by the Golgi-Cox Method

Celloidin blocks of Golgi-Cox impregnated material are cut at 50 μ, the sections collected in 70% alcohol, transferred to a 3:1 mixture of absolute alcohol and chloroform for 2 min, and then stored in xylene or toluene for at least 3 min, or up to 2 wk until processed further. Mounting is done on glass slides which have been coated with fresh egg albumen diluted in 0.2% ammonia water (or a 0.5% solution of dry powdered egg albumen) and then dried at 60°C overnight. For attachment to these coated slides, sections are first soaked for 2-3 min in a freshly prepared mixture of methyl benzoate, 50 ml; benzyl alcohol, 200 ml; chloroform, 150 ml; and then transferred quickly to the slides by means of a brush. After 2-3 min the chloroform evaporates and the celloidin softens. The slides are then immersed in toluene which hardens the celloidin and anchors the sections to the slides. Alcohols of descending concentrations to 40% are followed by alkalinizations, first in: absolute alcohol, 40 ml; strong ammonia water 60 ml, for 2 min, then in: absolute alcohol, 70 ml; strong ammonia water, 30 ml, for 1 hr. Excess alkali is then removed by 70% and 40% alcohol, 2 min each, and a 10 min wash in running tap water. Bleaching in 1% Na₂S₂O₃, for 10 min and washing again in tap water for 10 min completes the process preliminary to staining. The preparations are then stained for 90 min in an aqueous solution of either 0.5% cresylecht violet, neutral red, or Darrow red, buffered at pH 3.6. Dehydration and differentiation in ascending grades of alcohol, clearing with toluene or xylene, and applying a cover glass with a mounting medium having a refractive index of about 1.61 completes the process.     

A Simple Toluidine Blue—Basic Fuchsin Stain for Spermatozoa in Epoxy Sections

Differential staining of cell components of spermatozoa is readily accomplished in Epon or Araldite sections 0.5-1 μ thick from rat and hamster testis and epididymis, and stained as follows: 1% aqueous toluidine blue buffered at pH 6, 0.5-3 min at 90 C; washed in distilled water; 1% basic fuchsin in 50% alcohol, 3-5 min at 20-25 C; differentiated with 70% alcohol; allowed to dry; and mounted in a resin of high refraction (DPX was used). Results: acrosome, bright magenta; nucleus, deep blue; mitochondrial sheath of the middle-piece, pinkish purple; and tail, pale red. This procedure combined with staining of collagen by applying 2% aqueous phosphotungstic acid 1-2 min as a mordant, followed by 1% light green in 50% alcohol containing 1% acetic acid, 1-2 min at 20-25 C, gives polychromatic staining and is useful as a general stain for other epoxy-embedded tissues.     

Brazilin-Toluidine Blue O and Hematoxylin-Darrow Red Methods for Brain and Spinal Cord

Tissue fixed in 10% formalin, formalin-95% ethanol 1:s CaCO₂ or phosphate buffer neutralized formalin, or methanol-chloroform 2:1, was dehydrated and embedded in paraffin or double-embedded by infiltration in 1% celloidin followed by a chloroform-paraffin sequence. Sections were attached to slides with either albumen or gelatine adhesive and processed throughout at room temperature of 24-26 C. For either method, mordanting 30-60 min in 1% iron alum was followed by a 10 min wash in 4 changes of distilled water. For brazilin-toluidne blue O, myelin was stained for 20-60 min, depending upon section thickness, in a self-differentiating solution consisting of: 0.15% Li₂CO₃ 75 ml; 6% brazilin in 95% ethanol, 25 ml; and NaIO₃ 75 mg. After a thorough washing, Nissl material was stained for 3-8 min in a solution consisting of: 0.1 M acetic acid, 90 ml; 0.1 M sodium acetate, 10 ml; and 1% toluidine blue 0, 2.5 ml. For hematoxylin-Darrow red, myelin was stained for 2-6 hr in a self-differentiating solution consisting of: 0.15% Li₂,CO₃ 95 ml; 10% hematoxylin in 95% ethanol, 5 ml; and NaIO₃ 25 mg. After a thorough washing, Nissl material was stained for 20 min or less in a solution consisting of: 0.1 M acetic acid, 90 ml; 0.1 M sodium acetate, 10 ml; Darrow red, 25 mg. This mixture was first boiled, cooled to room temperature and filtered. In both methods, washing, dehydration, clearing, and mounting completed the process. In the brazilin-toluidine blue technic, myelin sheaths were stained reddish purple; neuronal nuclei light blue with dark granules of chromatin; nucleoli dark blue; and cytoplasm blue with dark blue Nissl granules. In the hematoxylin-Darrow red procedure, myelin sheaths were blue-black; nuclei light red with dark granules of chromatin; nucleoli almost black; and cytoplasm red with bright red Nissl granules.     

Silver Impregnation of Alzheimer's Neurofibrillary Changes Counterstained for Basophilic Material and Lipofuscin Pigment

A method is described in which selective silver staining of Alzheimer's neurofibrillary changes is combined with staining of cell nuclei, Nissl material, and lipofuscin granules. Formalin fixed, paraffin embedded sections of human autopsy tissue are silver stained according to a method proposed by Gallyas. Lipofuscin is stained by crotonaldehyde fuchsin following performic acid oxidation. Nissl substance is visualized by either Darrow red or gallocyanin-chrome alum staining. Architectonic units showing the specific pathology and the neuronal types prone to develop the neurofibrillary changes can be recognized using this technique.     

Silver impregnation of Alzheimer's neurofibrillary changes counterstained for basophilic material and lipofuscin pigment

A method is described in which selective silver staining of Alzheimer's neurofibrillary changes is combined with staining of cell nuclei, Nissl material, and lipofuscin granules. Formalin fixed, paraffin embedded sections of human autopsy tissue are silver stained according to a method proposed by Gallyas. Lipofuscin is stained by crotonaldehyde fuchsin following performic acid oxidation. Nissl substance is visualized by either Darrow red or gallocyanin-chrome alum staining. Architectonic units showing the specific pathology and the neuronal types prone to develop the neurofibrillary changes can be recognized using this technique.     

A Simple Stain for Myelin in Frozen Sections: A Modification of Mahon's Method

A simple and rapid method for demonstrating myelinated nerve fibers in frozen sections of the central and peripheral nervous system is described. Material fixed by perfusion with mixed aldehydes gives the best results but the method also works on specimens fixed by immersion in formaldehyde. Frozen sections varying in thickness from 15-50 μm are mounted on slides subbed with chrome alum-gelatin. After hydration (60-140 min), Sections are mordanted (20-40 min) in 2.5% iron alum and rinsed briefly in three changes of distilled H₂O (total 2 min). Staining is for 60-180 min in 40 cc freshly made 10% alcoholic hematoxylin diluted with 165 cc distilled H₂O to which 15 cc saturated Li₂CO₂is added. the sections are washed in distilled H₂O (5-15 min) and dehydrated in graded alcohols without differentiation in mordant, and covered. Myelin stains a dark blue-purple against a light grey background. Fiber tracts, as well as individual myelinated fibers, are clearly demonstrated.     

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.     

Simultaneous Formalin Fixation and Edta Decalcification, With Carbowax Embedding for Preservation of Acid Phosphatase

Carbowax serial sections from pubic symphyses of female mice, fixed and decalcified in a 10% formalin-5% Versenate solution for 18 hr at 4 C, pH 5.2, were incubated for 30 min with Burstone's simultaneous coupling reagent (pH 5.2); substrate: naphthol AS-TR and the diazonium salt, fast red violet L.B. All sections were counterstained with 1% methyl green at pH 4.0 in a phospho-citrate buffer. Inhibition by 0.01 M NaF, 0.0002 M CuCl₂, 10% tartaric acid and 0.01 M NaCN, as well as substrate-deficient and heat-inactivated controls, demonstrated conclusively that acid phosphatase was functionally preserved. Strong enzymatic activity was exhibited by osteoclasts, chondroclasts and free multinucleated giant cells. In addition, megakaryocytes, histiocytes, plasma cells, and monocytes exhibited moderate activity. The results demonstrated the technique to be consistently reproducible.     

Silver Protein Staining of Nerve Fibers in Celloidin Sections

Celloidin sections from formalin-fixed brain and spinal cord of primates are stored in 70% alcohol after cutting, soaked in 2% pyridine in 50% alcohol for 6-8 hr at 37 C, and transferred to 1% concentrated NH₄OH in 50% alcohol 15-18 hr at 20-25 C. After washing and flattening, the sections are transferred to 1% silver protein solution containing 30 ml of 0.2 M H₃BO₃/100 ml. Impregnation is accomplished in 50 ml screw-top jars, 50 mm in diameter, which are filled to a depth of 35 mm, and have 1 gm of copper foil, 0.002 inch thick added. The foil is folded in loose accordion-fashion, pierced and threaded, cleaned in 5% HNO₃, rinsed in distilled water, and suspended in the solution just above the sections by fastening the thread to the jar lid. The sections are impregnated for 24 hr at 37 C, rinsed in distilled water, reduced in a solution of 5% Na₂SO₃ and 1% hydroquinone for 10 min, washed in distilled water and toned in 0.2% gold chloride for 5 min. After rinsing in distilled water, the sections are transferred to 1% oxalic acid for 45-60 sec, washed in distilled water and placed in 5% Na₂S₂O₃ for 5 min. Sections are then washed, dehydrated to 95% alcohol, cleared in terpineol, followed by 3 changes in xylene, and mounted.     

New Uses for Calcium Chloride Solution as a Mounting Medium

Fresh cross sections of stems [Psilotum nudum, Coleus blumei, and Pelargonium peltatum] and roots (Setcreasea purpurea) 120 μm thick were fixed in FPA₅₀ (formalin: propionic acid: 50% ethanol, 5:5:90, v/v) for 24 hr and stored in 70% ethanol. The sections were transferred to water and then to 1% phloroglucin in 20% calcium chloride solution plus either hydrochloric, nitric, or lactic acid in the following ratios of phloroglucin-CaCl₂ solution:acid: 25:4, 20:2, or 15:5. The sections were mounted on slides either in one of the three mixtures or in fresh 20% calcium chloride solution. A rapid reaction of the acid-phloroglucin with lignin produced a deep red color in tracheary elements and an orange-red color in sclerenchyma. Fixed and stored leaf pieces from Nymphaea odorata were autoclaved in lactic acid, washed in two changes of 95% ethanol, transferred to water, and treated with the three acid-phloroglucin-calcium chloride mixtures. The abundant astrosclereids stained an orange-red color similar to that of sclerenchyma in the sections. In addition, a new method is reported for specifically staining lignified tissues. When sections or leaf pieces are stained in aqueous 0.05% toluidine blue O, then placed in 20% calcium chloride solution, all tissues destain except those with lignified or partially lignified cell walls. Thus, toluidine blue O applied as described becomes a reliable specific test for lignin comparable to the acid-phloroglucin test.     

A Simple Histochemical Reaction for Aldehydes

A study has been made on the possibility of replacing leucofuchsin by colored basic fuchsin for the histochemical demonstration of aldehydes. Several tissues from mammals and various pertinent fixatives were used. Aldehydes were freed from carbohydrates by oxidation and from thymonucleic acid by hydrolysis.

It was found that the colored form and not necessarily the leucoform of basic fuchsin can be used histochemically in demonstrating aldehydes. The technic used is as follows: (1) Treat with 1.0-0.5% H₅IO₆ (or in 1% KIO₄ in M/1 H₂SO₄) for 5 to 10 min. and wash thoroughly. For thymonucleic acid hydrolize with N HCl 5 min. at room temperature, 10 min. at 60°C. and 5 min. at room temperature. (2) Stain for 2-3 min. with 0.05% basic fuchsin in 5% ethanol, 3% phenol. (3). Transfer immediately to 1 or 2 changes of 1% sodium bisulphite or potassium metabisulphite in 0.1-0.2 N H₂SO₄ for a total of 5 min. (4) Rinse with water and treat with M H₂SO₄ in 95% ethanol for 3-5 min. 6. Wash thoroughly in water and dehydrate, clear, and mount. For glycogen and mucin the following counterstaining solution is recommended: orange G, 0.25 g.; light green SFY, 0.10 g.; phosphotungstic acid 0.50 g.; 50% ethanol, 100 ml.; glacial acetic acid, 0.25 ml.     

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.     

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.     

Acid Orcein-Iron and Acid Orcein-Copper Stains for Elastin

Paraffin sections of formol-fixed tissues stained 4-18 hr in 70% alcohol containing 1% orcein and 1% of concentrated (12 N) HCl by volume yield the familiar purple brown elastin and red nuclei on a pink background. When sections so stained are transferred directly from the stain to 70% alcohol containing 0.02% ferric chloride (FeCl₃·6 H₂O) or 0.02% copper sulfate (CuSO₄·5 H₂O) for a 15 sec to 3 min period, elastin coloration is changed to black or reddish black and chromatin staining to reddish black. The procedure can be counterstained with picro-methyl blue to yield blue collagen and reticulum or with our flavianic acid, ferric chloride, acid fuchsin mixture to give deep yellow background and deep red collagen.     

Cytochemical Staining of Sections from Plastic-Embedded Flagellates

Dinoflagellate chromosomes in sections of plastic-embedded cells were stained without removing the plastic. Azur B and Feulgen procedures were used to localise DNA. Azur B was used with Araldite or methacrylate sections by staining in 0.2% stain in 0.05 M citrate buffer at pH 4 for 1 hr at 50 C followed by rinsing in tertiary butyl alcohol to differentiate the chromosomes. Feulgen stain was used with Araldite sections by hydrolyzing in 1 N HCl at 60 C for 10 min, rinsing in water, staining for 24 hr, washing well, drying and covering. Fast green was used with methacrylate sections to stain proteins by flooding the slide with a 0.1% solution of stain in 0.06 M phosphate buffer at pH 8, allowing the stain to dry out at 40-50 C, washing well, drying and covering. Controls were carried out on material fixed in formalin and treated with nucleases or proteolytic enzymes prior to embedding, and staining.     

Maxilon Blue Rl: A New Metachromatic Monoazo Dye

Maxilon blue RL, a basic monoazo dye developed by Geigy S. A., stains metachromatically acid mucopolysaccharide-containing elements in histological sections. This property is due to a chromatographically pure blue fraction which is the main component of the dye. The following routine has been developed for staining sections of formalin-calcium fixed paraffin-embedded tissues. Dewax in xylene and hydrate the sections through alcohol; stain in 0.05% aqueous Maxilon blue RL, from 30-60 sec; wash in distilled water; dehydrate either in tertiary butanol, 2 to 3 min, after removing excess water from the slide by blotting; or, rinse in 70% ethyl alcohol and dehydrate in 2 changes, 2 min each, of absolute alcohol; clear in xylene; mount in DPX or in Canada balsam. Acid mucopolysaccharides are colored red to violet; other basophilic elements, blue.     

An Evaluation of the Metachromasy of Anionic Dyes. I. Visual Observations on Tissue Sections

Thirteen dyes of the azo (benzopurpurin, Congo red, trypan blue, chromotrope 2R, orange G), indigoid (indigocarmine), triphenylmethane (acid fuchsin, aniline blue, light green, methyl blue), and xanthene (eosin B, eosin Y, erythrosin B) groups were applied under standard conditions to a variety of human, rabbit, rat, mouse and frog tissues in paraffin sections. Sections were examined for color changes which might indicate metachromatic reactions analogous to the metachromasy of cationic dyes. Disazo and xanthene dyes showed shifts in hue, with some qualification on the shifts of xanthenes. Metachromatic shifts of anionic dyes were generally of low order compared to those of cationic dyes. Nuclei, erythrocytes, inner elastic laminae of arteries, keratinous structures, and certain areas in the ground substance of connective tissue most often elicited metachromasy. It is suggested that basic proteins are responsible for the metachromatic reactions. Equally interesting areas were those staining poorly (cartilage matrix, most types of mucus), since these are sites of highly acidic substances capable of binding proteins.     

A Gelatin Film Method for Improved Histochemical Localization of Dehydrogenases in Plant Cells

Glucose-6-phosphate and 6-phosphogluconate dehydrogenases diffused from frozen sections of Vicia faba embryos during histochemical incubation. In the liquid incubation medium, the dehydrogenases catalysed the oxidation of substrate and reduction of NADP. NADPH₂ thus formed could lead to artifactual deposition of formazan in frozen sections. The addition of 20% polyvinyl alcohol to the incubation medium was found unsatisfactory in preventing this loss which appeared to be overcome by incorporating the reaction mixture into a gelatin film. Equal volumes of 10% gelatin solution in 0.05 M phosphate buffer at pH 7.8, and the enzyme reaction medium containing twice the normal concentration of substrate (0.014 M), of 0.007 M pyridine nucleotide, of 0.02 M KCN and of 0.0024 M NBT in the buffer, were mixed and layered onto polyethylene, and allowed to set in the dark at room temperature for 30-60 min. The solidified medium and its support were cut into strips and layed onto unfixed, frozen sections of plant tissues which were incubated at 20 C. Evidence is presented to support the supposition that the enzymes are retained in the sections during the reaction.     

Romanowsky-type staining: Enzymatic analysis of nuclear metachromasia in formalin-fixed paraffin sections

The red color of nuclei produced in formol-fixed paraffin sections stained with toluidine blue has been investigated by using deoxyribonuclease (DNase), ribonuclease (RNase) and 0.1 M Tris buffer. The action of DNase on formol-fixed material is not fully reliable, but clear-cut when positive. Nuclear basophilia and metachromasia is removed, nucleolar and cytoplasmic RNA is preserved. The picture produced by RNase depends to some extent on the concentration and acidity of the toluidine blue used for subsequent staining. Cytoplasmic RNA is always removed, while the red stain in nuclei usually remains intact. With 0.1% toluidine blue in 1% acetic acid, a nuclear color change from red to pale green is observed. Using this same staining solution, it can be shown that 0.1 M Tris buffer (overnight extraction at 37° C) will remove cytoplasmic RNA but leave intact the nuclear material that stains red. A red to green shift can subsequently be produced by RNase. From this it is deduced that there is a chromatin-associated nuclear RNA fraction which can be removed by the enzyme, but is stable to the buffer solution.