Staining Juxtaglomerular Granules with Basic Fluorescent Stains

Many basic fluorescent dyes stain juxtaglomerular granules to produce characteristic colors in ultraviolet light. The stain is applied to paraffin sections of tissues fixed in 2% calcium acetate-10% formalin or in phosphate-buffered 10% formalin. Procedure: Bring section to water, stain 0.5 min in Delafield hematoxylin, wash in tap water, stain 3 min in a 0.1% aqueous solution of basic fluorescent dye (auramine O, acriflavine, acridine orange, coriphosphine O, acridine yellow, phosphine E, thioflavine T, berberine sulfate, atebrine or rivanol) and differentiate 1 min in 0.1% acetate acid (or omit this step). After washing in tap water, air dry with or without subsequent mounting in a resin. Juxtaglomerular granules stain bright fluorescent yellow or orange against a dark background.     

Quick Staining Method For Frozen Sections

Since the advent and general acceptance of frozen sections in histological and pathological laboratories it has been necessary to devise methods for staining these sections. The usual method is fixing the tissue to a slide by the use of celloidin. This paper is an attempt to describe a permanent, quick method of staining frozen sections without distortion or mechanical tearing of the tissues.     

A Protargol Method for Staining Nerve Fibers in Frozen or Celloidin Sections

Extensive experimentation with protargol staining of neurons in celloidin and frozen sections of organs has resulted in the following technic: Fix tissue in 10% aqueous formalin. Cut celloidin sections IS to 25 μ, frozen sections 25 to 40 μ. Place sections for 24 hours in 50% alcohol to which 1% by volume of NH₄OH has been added. Transfer the sections directly into a 1% aqueous solution of protargol, containing 0.2 to 0.3 g. of electrolytic copper foil which has been coated with a 0.5% solution of celloidin, and allow to stand for 6 to 8 hours at 37° C. Caution: In this and the succeeding step the sections must not be allowed to come in contact with the copper. From aqueous protargol, place the sections for 24 to 48 hours at 37° C. directly into a pyridinated solution of alcoholic protargol (1.0% aqueous solution protargol, 50 ml.; 95% alcohol, 50 ml.; pyridine, 0.5 to 2.0 ml.), containing 0.2 to 0.3 g. of coated copper. Rinse briefly in 50% alcohol and reduce 10 min. in an alkaline hydroquinone reducer (H₃BO₃, 1.4 g.; Na₂SO₃, anhydrous, 2.0 g.; hydroquinone, 0.3 g.; distilled water, 85 cc; acetone, 15 ml.). Wash thoroly in water and tone for 10 min. in 0.2% aqueous gold chloride, acidified with acetic acid. Wash in distilled water and reduce for 1 to 3 min. in 2% aqueous oxalic acid. Quickly rinse in distilled water and treat the sections 3 to 5 min. with 5% aqueous Na₂S₂O₃+5H₂O. Wash in water and stain overnight in Einarson's gallocyanin. Wash thoroly in water and place in 5% aqueous phosphotungstic acid for 30 min. From phosphotungstic acid transfer directly to a dilution (stock solution, 20 ml.; distilled water, 30 ml.) of the following stock staining solution: anilin blue, 0.01 g.; fast green FCF, 0.5 g.; orange G, 2.0 g.; distilled water, 92.0 ml.; glacial acetic acid, 8 ml.) and stain for 1 hour. Differentiate with 70% and 95% alcohol; pass the sections thru butyl alcohol and cedar oil; mount.     

Differential Staining of Tannin in Sections of Epoxy-Embedded Plant Cells

A staining procedure is described for the light microscopic localization of ergastic tannins in epoxy sections of plant cells embedded for study by transmission electron microscopy. Callus and cell suspensions of Pseudotsuga menziesii and Pinus taeda fixed in glutaraldehyde:acrolein and then OsO₄, followed by epoxy embedding, were sectioned 0.5 μn thick, stained on a glass slide with ethanolic Sudan black B at 60 C as described by Bronner, and then mounted in Karo syrup. Tannin deposits stained brownish-orange and were easily distinguished from lipid bodies of similar size, which stained dark blue to black, and from starch grains, which were unstained. The significance of this differential polychromasia was confirmed by transmission electron microscopy. This staining proadure should prove valuable in the cytoplasmic evaluation of the plant cell ergastics (especially tannins) via light microscopy whether or not electron microscopic examination is intended.     

Enumerative Fluorescent Vital Staining of Live and Dead Pathogenic Yeast Cells

A quantitative technique is presented for differentiating live and dead yeast cells grown in culture through the use of the fluorescent dye acridine orange. the method gives results that correlate well with those of other commonly used vital staining techniques and is free of certain interpretative errors inherent in them. Vital staining of yeasts with acridine orange also allows for more precise assessment of the physiological state of individual cells and the culture as a whole. the progressive senescence of yeast cells in culture can be monitored by the changing staining characteristics of several subcellular organelles. the method is simple and reliable.     

Selective and Contrast Staining of Granules in the Juxtaglomerular Complex

A review of four methods for staining juxtaglomerular cells revealed that one method may be highly selective for juxtaglomerular granules (JGG) whereas another may stain general cytological features in addition to the granules. The kind of research undertaken would determine the particular method to be used. Harada's (1952) method, which uses a 1:400,000 solution of gentian violet is recommended as the highly selective stain, and the Masson-Goldner stain after a Ciaccio type fixation is best for cytological detail combined with clear tinctorial contrast of the JGG.     

Ninhydrin Staining of Tissue Sections

This method represents a considerable improvement over earlier ninhydrin procedures. Celloidin sections were stained after mounting in a medium which clears with incubation at 55 °C. There appears to be no reason why paraffin section cannot be used. The sections were not placed in a large volume of ninhydrin (0.25% triketohydrindene hydrate in n-butanol) but only a small volume was sprayed onto the slide. Distortion resulting from heating in boiling water to develop the color was avoided by a slower treatment of 3 days' incubation at 55 °C. The use of water as a solvent in staining is also avoided, thus minimizing the possibility of color migration and insuring against the development of the intensely colored products of the ninhydrin reaction that occur in aqueous solution. Slides need not be observed upon the day of preparation, since the color was stable for about a week after its formation.     

Detection of Active Yeast Cells (Saccharomyces cerevisiae) in Frozen Dough Sections

A new method based on fluorescence microscopy was developed to detect active yeast cells in cryosections of wheat dough. The sections were stained with 4′,6-diamidino-2-phenylindole (DAPI) and counterstained with Evans blue. The active yeast cells in the sections appeared brilliant yellow and were readily distinguished from the red dough matrix. The dead cells allowed penetration of the Evans blue through the cell membrane, which interfered with the DAPI staining and caused the dead cells to blend into the red environment. The number of active yeast cells in fermenting dough sections containing different proportions of living and dead yeast cells correlated well with the gas-forming capability of the yeast in the dough but not with the results of the conventional plate count method. The new method allows the study of yeast activity not only during the different stages of frozen dough processing but also during the fermentation of doughs.     

Mounting Frozen Sections with Gelatin

Frozen sections, 15-50 µ thick, are soaked for 5 minutes or longer in a mixture of equal parts of 1.5% aqueous gelatin and 80% alcohol, and teased onto a slide. After allowing excess fluid to evaporate, sections will be moist and can be blotted with filter paper that may require dampening with 95% alcohol. Immersed in 95% alcohol, the remaining gelatin will congeal, anchoring the section to the slide. If necessary, the sections can subsequently be coated with celloidin.     

Producing Frozen Sections of Calcified Bone

We describe a procedure for the rapid production and maintenance of fresh frozen bone biopsies which can be used for a variety of immunohistochemical techniques. Within 5 min of excision. tissue is placed in cold 5% polyvinyl alcohol, surrounded with 3% carboxymethylcel-lulose in a hand made aluminum foil embedding mold and frozen by immersion in an absolute ethanol/dry ice slurry at -70 C. The tissue block is attached to the specimen stub with cryocom-pound and installed in a -32 C cryostat whose tungsten carbide D profile knife is maintained at -70 C. Automatic controls are set at a slow cutting speed and the “sectioning window” is adjusted to fit the biopsy size. Knife angle, thickness gauge and antiroll bar are changed to produce a complete section. The block face is smoothly “papered” with a polyvinylpyrrolidone (PVP) impregnated Ross lens paper strip. A single section is cut and positioned on a sequentially numbered, acid cleaned, double dipped chrome-alum gelatin coated slide: adhesion is aided by “press-blotting” with bibulous paper. Sections are stored at -20 C or in a desiccator at room temperature. A brief fixation followed by removal of the water soluble PVP and lens paper generates fresh frozen bone sections suitable for further analysis.     

A Technic for Preparing Frozen Sections

A combination of the Gram-Pappenheim stains for the examination of gonorrheal pus, cellular exudate and paraffin sections of formalin-fixed tissues has been described elsewhere (Scudder and Lisa, 1931). The crystal violet solution was made stable for the first time by employing phosphate buffers on the acid side of neutrality, and a stable counterstain was prepared for the first time from National Aniline dyes, ethylated methyl green and pyronin yellowish. Original findings were demonstrated by means of color plate I and II (Scudder, 1931) to show gonococci, pneumococci and cells in smears, and formalin-fixed tissue brought down to water in the usual way. A new color plate is published herewith to show the microscopic appearance of cells, Gram-positive and Gram-negative bacteria, higher bacteria, fungi and spermatozoa in the study of genitourinary and gynecological cases. The method has a value in the field of medical jurisprudence. Crystals were well demonstrated, especially those resulting from sulfa drug therapy. The National Aniline methyl green batches numbered NG 10, 11, 13 to 19, and their batches of pyronin numbered NP 5 to 10 were found consistently stable. Earlier dyes were found either too purple or too blue for the technic and the most satisfactory dyes were found to require a ripening time of several days and could be prepared in amounts of from 1 to 4 liters and stored indefinitely without preservatives.     

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.     

An Efficient Staining Method for Ultrathin Sections

A staining method to handle simultaneously as many as 20 electron microscope grids is described. The devices used are easily constructed of readily obtained inexpensive materials. The volumes of stain and wash water required are very small and drying grids is simplified.