Direct Staining of Reticular Fibers with Gold Chloride

Reticular fibers are selectively stained in paraffin sections of formalin-fixed or Bouin's-fixed tissue as follows: 1% aqueous solution of gold chloride for 20 min, followed by a 10 min immersion in an aqueous solution containing 5% Na₂CO₃ and 0.5% KOH. The sections then are placed in a 5% aqueous solution of KI for 2 min. Counterstaining with a 0.25% aqueous solution of methylene blue chloride is optional. The reticular fibers stain dark pink; the collagen bundles are a light pink to straw color without the counterstain, or a light blue color when the methylene blue is used.     

A polychromatic staining method for epoxy embedded tissue: a new combination of methylene blue and basic fuchsine for light microscopy

A simple and rapid method is described for staining semithin sections of material embedded in epoxy resin for observing tissues prior to transmission electron microscopy. The method is suitable for tissue fixed with a glutaraldehyde-formaldehyde mixture and postfixed in osmium tetroxide. No etching or oxidizing procedures are necessary. Sections 0.5-0.8 µm thick are dried onto a slide and stained with either 0.75% methylene blue and 0.25% azure B or 0.5% methylene blue and 0.5% azure II in 0.5% aqueous borax and heated over a flame for 8-10 sec. The slides are rinsed with water, then stained the same way with 0.1% basic fuchsine in 5% aqueous ethanol. Cytoplasm stains blue; nuclei darker blue; collagen, mucus and elastin pink to red; fat and intracellular lipid droplets gray-green.     

A polychromatic staining method for epoxy embedded tissue: a new combination of methylene blue and basic fuchsine for light microscopy

A simple and rapid method is described for staining semithin sections of material embedded in epoxy resin for observing tissues prior to transmission electron microscopy. The method is suitable for tissue fixed with a glutaraldehyde-formaldehyde mixture and postfixed in osmium tetroxide. No etching or oxidizing procedures are necessary. Sections 0.5–0.8 µm thick are dried onto a slide and stained with either 0.75% methylene blue and 0.25% azure B or 0.5% methylene blue and 0.5% azure II in 0.5% aqueous borax and heated over a flame for 8–10 sec. The slides are rinsed with water, then stained the same way with 0.1% basic fuchsine in 5% aqueous ethanol. Cytoplasm stains blue; nuclei darker blue; collagen, mucus and elastin pink to red; fat and intracellular lipid droplets gray-green.     

A polychromatic staining method for epoxy embedded tissue: a new combination of methylene blue and basic fuchsine for light microscopy.

A simple and rapid method is described for staining semithin sections of material embedded in epoxy resin for observing tissues prior to transmission electron microscopy. The method is suitable for tissue fixed with a glutaraldehyde-formaldehyde mixture and postfixed in osmium tetroxide. No etching or oxidizing procedures are necessary. Sections 0.5-0.8 microm thick are dried onto a slide and stained with either 0.75% methylene blue and 0.25% azure B or 0.5% methylene blue and 0.5% azure II in 0.5% aqueous borax and heated over a flame for 8-10 sec. The slides are rinsed with water, then stained the same way with 0.1% basic fuchsine in 5% aqueous ethanol. Cytoplasm stains blue; nuclei darker blue; collagen, mucus and elastin pink to red; fat and intracellular lipid droplets gray-green.     

A Phloxine-Azure-Hematoxylin Sequence for Differential Staining of Cells in Pancreatic Islets

Sections of 6 μ from tissues fixed in Susa or in Bouin's fluid (without acetic acid) and embedded in paraffin were attached to slides with Mayer's albumen, dried at 37 C for 12 hr, deparaffinized and hydrated. The sections fixed in Susa were transferred to a I₂-K₁ solution (1:2:300 ml of water); rinsed in water, decolorized in 5% Na₂S₂O₃; washed in running water, and rinsed in distilled water. Those fixed in Bouin's were transferred to 80% alcohol until decolorized, then rinsed in distilled water. All sections were stained in 1% aqueous phloxine, 10 min; rinsed in distilled water and transferred to 3% aqueous phosphotungstic acid, 1 min; rinsed in distilled water; stained 0.5 min in 0.05 azure II (Merck), washed in water; and finally, nuclear staining in Weigert's hematoxylin for 1 min was followed by a rinse in distilled water, rapid dehydration through alcohols, clearing in xylene and covering in balsam or a synthetic resin. In the completed stain, islet cells appear as follows: A cells, purple; B cells, weakly violet-blue; D cells, light blue with evident granules; exocrine cells, grayish blue with red granules.     

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 simple, rapid staining procedure for methacrylate embedded tissue sections using chromotrope 2R and methylene blue

Sections of tissue embedded in glycol methacrylate can be stained in rapid sequence with solutions of 1% aqueous chromotrope 2R adjusted to pH 3 and 0.1% methylene blue to produce sufficient contrast and cellular detail to permit quick visual inspection and/or photomicrography. Solutions of these stains are simple to prepare and are stable over long periods. Staining of sections may be accomplished within six minutes.     

A Simple, Rapid Staining Procedure for Methacrylate Embedded Tissue Sections Using Chromotrope 2R and Methylene Blue

Sections of tissue embedded in glycol methacrylate can be stained in rapid sequence with solutions of 1% aqueous chromotrope 2R adjusted to pH 3 and 0.1% methylene blue to produce sufficient contrast and cellular detail to permit quick visual inspection and/or photomicrography. Solutions of these stains are simple to prepare and are stable over long periods. Staining of sections may be accomplished within six minutes.     

A Simple Polychrome Stain for Conventionally Fixed Epon-Embedded Tissues

The described technique, based upon a one-step Mallory-Heidenhain stain, can be applied as a routine stain for glutaraldehyde or OsO₄ fixed, Epon embedded tissues of various organs. The technique consists of a short treatment of the sections with H₂O₂, a nuclear staining with celestine blue B and a final staining in a modified Cason's solution. The different tissue and cell components are displayed as follows: dark brown nuclei, yellow cytoplasm, red collagen fibers and blue elastic' fibers. Intra cytoplasmic components as glycogen and mucus are stained respectively blue and violet, whereas other inclusions such as leucocyte granules are colored orange to red.     

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.     

Keratin in the Egg Shells of Amphistomes; Histochemical Differentiation from Quinonetanned Protein in Other Trematoda

Various combinations of the oxidation method for demonstrating keratin in shell material of amphistomes were tried. Acidified permanganate worked more efficiently than performic and peracetic acids, and Alcian blue and aldehyde fuchsin excelled other basic dyes for subsequent staining. For the permanganate-Alcian blue reaction, sections of material fixed in Susa or Bouin were oxidized in 0.3% permanganate in 0.3% H₂SO₄ for 5 min., decolourized in 1% oxalic acid, stained in 3% Alcian blue in 2 N H₂SO₄ and counterstained with eosin. The shell globules stained a deep blue. For permanganate aldehyde fuchsin staining, the sections were stained in aldehyde fuchsin for 1 hr, after oxidation with permanganate. The shell globules then stained a deep magenta. The catechol and fast red reactions were negative in amphistomes and the specimens lack the characteristic amber colour due to quinone tanning.     

Staining Paraffin Embedded Sections of Scald of Barley before Paraffin Removal

Staining of paraffin embedded sections with periodic acid-Schiff reagent and fast green before paraffin removal resulted in differentiation of barley seed and leaf tissue from fungal structures of Rhynchosporium secalis. Crystal violet, toluidine blue O and aniline blue also successfully stained fungal structures of R. secalis in barley leaf tissues. Staining of embedded sections before paraffin removal allows simple processing of a series of sections, saves time and reduces solvent consumption.     

Chlorantine fast green BLL as a stain for callose in oak phloem

Sections of oak bark were stained with chlorantine fast green BLL, used as a 0.25% aqueous solution. All tissues were unstained, except for local deposits of material associated with phloem cell walls, which stained deep green. This green-staining material also stained specifically with resorcinol blue and with the aniline blue fluorescence technique, the usual histochemical tests for callose. The chlorantine fast green-staining material was removed from sections by treatment with a beta-1,3-glucan hydrolase. It is concluded that chlorantine fast green BLL stains callose in plant sections and is a useful additional stain for the histochemical detection of this polymer.     

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.     

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.     

Staining Methods For Studying Ingredient Distribution In Baked Cakes

A method is described for preparing cake crumb for sectioning and staining. Previous to embedding, the fat was stained and fixed by exposing small blocks of cake to the fumes from a 5%, freshly-prepared, aqueous solution of osmic acid (OsO₄). This was followed by dehydration in ethyl alcohol and tertiary butyl alcohol, removal of air under vacuum and infiltration with paraffin.

Sections were cut 20 and 9Op thick and mounted with water.

Wax was removed by immersion in xylene. The sections were rehydrated in a series of ethyl alcohol dilutions, from concentrated to dilute, then transferred to distilled water.

Protein was then stained pink by immersion of the slides in an acidified 0.04% water solution of eosin Y, or starch was stained blue with a dilute aqueous solution of iodine. Ten grams iodine and 10 g. KI were dissolved in 25 ml. distilled water. This stock solution was diluted for use one to two hundred times.

The relationship between protein and starch was demonstrated by staining the sections with eosin, differentiating in 50% alcohol and staining with iodine.

When slides of cake crumb were prepared in this way, the fat was stained black, the protein bright pink and the starch granules a dark blue.     

Selective Staining of Neurosecretory Material in Semithin Epoxy Sections by Gomori's Aldehyde Fuchsin

The epoxy resin was removed from semithin (1 μm) sections by immersing them for 30 sec in sodium methoxide (Mayor et al., J. Biophys. Biochem. Cytol., 9: 909-10, 1961) and then processed as follows: (1) left for 1-3 hr at 60 C in a mixture of formalin, 25 ml; glacial acetic acid, 5 ml; CrO₃, 3 gm; and distilled water, 75 ml: (2) oxidized 10 min in a 1:1:6 v/v mixture of 2.5% KMnO₄, 5% H₂SO₄ and distilled water: (3) bleached in 1% oxalic acid, and (4) stained for 15 min in aldehyde fuchsin, 0.125% in 70% alcohol, or in a 1% aqueous solution of toluidine blue. The neurosecretory material is selectively stained.     

Epoxy Embedding, Sectioning and Staining of Plant Material for Light Microscopy

By using a formula which gives a relatively soft epoxy embedding medium, it is possible to cut sections of plant material with a sliding microtome equipped with a regular steel knife. Blocks having a cutting face of 10 × 10 mm, giving sections of 4-10 μm, can be used. Tissues are fixed in Karnovsky's fluid, postfixed in 1 or 2% OsO₄, embedded in Spurr's soft epoxy resin, Araldite, or Epon mixtures. 5% KMnO₄, followed by 5% oxalic acid, then neutralized in 1% LiCO₃, are used to mordant the sections. Some of the stains used are Mallory's phosphotungstic acid-hemotoxylin, acid fuchsin and toluidine blue, or toluidine blue. Mounting is done with whichever soft epoxy resin was used in casting the blocks.     

Staining Rickettsiae in Yolk-Sac Cultures

Rickettsiae in yolk sacs are not stained well by the Macchiavello technique, and experiments were undertaken to understand the mechanisms involved. It was found that the citric acid destaining step was not effective and that most of the basic fuchsin was lost from the rickettsiae during the application of methylene blue, another basic dye. A staining technique was then evolved with carbol basic fuchsin in pH 7.45 phosphate buffer (0.4% dye, 0.4% phenol, 0.07 M buffer), followed directly by 0.8% aqueous malachite green oxalate. This technique worked well for R. mooseri, R. prowazeki, R. rickettsii, R. akari, and R. buretii, but for R. tsutsugumushi a modification was needed, whereby 4% aqueous Fe(NO₃)₃·9H₂O was used as destaining solution, and 0.5% aqueous fast green as the counter-stain.     

Improved Procedures in Fungal Cytology Utilizing Giemsa

To study nuclear events in fructifications of the Basidiomycetes, material was fixed 24 hr in a saturated aqueous solution of HgCl₂ containing 1% glacial acetic acid, and embedded in Aquax (G. T. Gurr Ltd.). Following a 4 hr hydrolysis at 20 C in 60% H₃PO₄, sections were stained for 30 min in a mixture of 4 ml Giemsa R66 (G T. Gurr Ltd.) and 100 ml phosphate buffer at pH 6.5. Differentiation was carried out in sodium cacodylate-HCl buffer at pH 5.8 when required. Preparations were dehydrated in an acetone-xylene series prior to mounting in Euparal. The use of paraffin wax as the embedding medium and HCl as the hydrolysing agent yielded preparations of an inferior quality.