A Bacterial Cell Wall Stain1

A new method is given to stain bacterial cell walls, especially of Escherichia coli and Bacillus cereus. The cells are smeared in water on a slide and, as soon as air-dry, are stained 3-4 minutes with a 1 % aqueous solution of new fuchsin. The smear is washed with water until the stain ceases to run and is then allowed to air dry. The slide is placed on a 50°C. warm plate for 10-20 seconds, and the smear is then covered with a thin film of a 1-2% solution of Congo red at a pH of about 9.5. The smear is ready for observation as soon as dry or it may be washed with water if desired before observation.     

A Modified Wright's Method for Staining Blood Smears

After the blood smear is treated for the proper length of time with Wright's stain, neutral distilled water is used for diluting the stain. After the slide has been treated with neutral distilled water until the smear becomes pinkish it is then treated with pure absolute methyl alcohol which destains the plasma. Neutral distilled water is again kept on the mount until the corpuscles are well stained. Then the slide is dehydrated with absolute ethyl alcohol, cleared with clove oil and completed in the usual way.

Blood smears of different groups of vertebrates were uniformly brilliantly stained with the above technic.

Several lots of Wright's dry stain have been tested with the modified technic and no difficulties have been encountered in its application.     

A Versatile Stain for Pollen Fungi, Yeast and Bacteria

A versatile stain has been developed for demonstrating pollen, fungal hyphae and spores, bacteria and yeasts. The mixture is made by compounding in the following order: ethanol, 20 ml; 1% malachite green in 95% ethanol, 2 ml; distilled water, 50 ml; glycerol, 40 ml; acid fuchsin 1% in distilled water, 10 ml; phenol, 5 g and lactic acid, 1-6 ml. A solution has also been formulated to destain overstained pollen mounts. Ideally, aborted pollen grains are stained green and nonaborted ones crimson red. Fungal hyphae and spores take a bluish purple color and host tissues green. Fungi, bacteria and yeasts are stained purple to red. The concentration of lactic acid in the stain mixture plays an important role in the differential staining of pollen. For staining fungi, bacteria and yeasts, the stain has to be acidic, but its concentration is not critical except for bacteria. In the case of pollen, staining can be done in a drop of stain on a slide or in a few drops of stain in a vial. Pollen stained in the vial can be used immediately or stored for later use. Staining is hastened by lightly flaming the slides or by storing at 55±2 C for 24 hr. Bacteria and yeasts are fixed on the slide in the usual manner and then stained. The stock solution is durable, the staining mixture is very stable and the color of the mounted specimens does not fade on prolonged storage. Slides are semipermanent and it is not necessary to ring the coverslip provided 1-2 drops of stain are added if air bubbles appear below the coverslip. The use of differentially stained pollen mounts in image analyzers for automatic counting and recording of aborted and nonaborted pollen is also discussed.     

Staining Mitochondria in Fixed Blood Smears

Wet blood smears are placed immediately in Helly's fluid for 24 hr, transferred directly to a saturated solution of potassium dichromate for 48 hr and washed in running water for 2-4 hr. The slides are then treated with iodine and sodium thiosulf ate and washed several hours or overnight. Excess water is removed by blotting the slide around the smear, Altmann's aniline fuchsin is placed on the smear and the slide is heated over a spirit lamp until white fumes appear. After the slide cools the stain is poured off and the excess removed by washing with distilled water. Methyl green (1% aqueous) is dropped on the smear and left for approximately 30 sec. It is then passed rapidly through 2 changes of absolute ethanol and into xylene, from which it is mounted in Permount. This stains mitochondria, red blood corpuscles and specific granules of eosinophilic granulocytes red on a green background.     

Staining Procedures for Ultrathin Sections of Tissues Embedded in Polyester Resin

Tissues were fixed for 30 min In cold (0-2° C) 1% OsO₄ (Palade) buffered at pH 7.7, to which 0.1% MgCl₂ was added. Dehydration was in a graded ethanol series (containing 0.5% MgCl₂) at 0-2° C, and terminated with 2 changes of absolute ethanol. Tissues were then transferred by a graded series to anhydrous acetone. Infiltration of the tissue with Vestopal-W (a polyester resin), is gradual with the aid of graded solutions of Vestopal-W in acetone. The infiltrated tissue is encapsulated and initial polymerization is done under ultraviolet light at room temperature for 8-16 hr. This is followed by final hardening at 60° C for 36-48 hr. Sections (0.2-1 μ) were cut, dried on slides, placed in acetone for 1 min and then treated by either of the following staining procedures: (1) Thionin-azure-fuchsin staining: Flood the preparation with 0.2% aqueous thionin and heat to 60-80° C for 3 min; if the preparation begins to dry, add stain. Rinse in distilled water. Flood the slide with 0.2% azure B in phosphate buffer at pH 9. Heat to 60-80° C for 3 min; do not permit the preparation to dry. Rinse in distilled water. Dip the slide in MacCallum's variant of Goodpasture's carbol-fuchsin stain for 1-2 sec. Rinse in distilled water. Check the preparation microscopically for intensity of the fuchsin stain. Repeat dips as may be needed to obtain the desired intensity. Rinse in distilled water. Dehydrate quickly in 95% and absolute alcohol; clear in 2 changes of xylene and cover in Permount or similar synthetic resin. (2) Thionin-azure counterstain for the periodic acid-Schiff reaction: Oxidize the tissue in 0.5% periodic acid for 15 min and transfer to Schiff's leucofuchsin solution for 30 min. Counterstain with 0.5% aqueous thionin for 3 min; wash in distilled water; stain in 0.2% azure B in phosphate buffer at pH 5.5; wash in distilled water; dehydrate; clear and cover as in the first method. For temporary preparations let dry after absolute alcohol and apply a drop of immersion oil directly on the section.     

SpermBlue®: A new universal stain for human and animal sperm which is also amenable to automated sperm morphology analysis

Abstract

Our study was aimed at exploring a simple procedure to stain differentially the acrosome, head, midpiece, and flagellum of human and animal sperm. A further prerequisite was that sperm morphology of the stained samples could be analyzed using automated sperm morphology analysis (ASMA). We developed a new staining process using SpermBlue® fixative and SpermBlue® stain, which are iso-osmotic in relation to semen. The entire fixation and staining processes requires only 25 min. Three main steps are required. First, a routine sperm smear is made by either using semen or sperm in a diluting medium. The smear is allowed to air dry at room temperature. Second, the smear is fixed for 10 min by either placing the slide with the dried smear in a staining tray containing SpermBlue® fixative or by adding 1 ml SpermBlue® fixative to the slide. Third, the fixed smear is stained for 15 min by either immersing the slide in a staining tray containing SpermBlue® stain or adding four drops of SpermBlue® stain to the fixed smear. The stained slide is dipped gently in distilled water followed by air drying and mounting in DPX® or an equivalent medium. The method is simple and suitable for field conditions. Sperm of human, three monkey species, horse, boar, bull, ram, mouse, rat, domestic chicken, fish, and invertebrate species were stained successfully using the SpermBlue® staining process. SpermBlue® stains human and animal sperm different hues or intensities of blue. It is possible to distinguish clearly the acrosome, sperm head, midpiece, principal piece of the tail, and even the short end piece. The Sperm Class Analyzer® ASMA system was used successfully to quantify sperm head and midpiece measurements automatically at either 600 × or 1000 × magnification for most of the species studied.     

Staining Germinating Pollen and Pollen Tubes

Germinating pollen on stigmas and pollen tubes in styles of Antirrhinum, Brassica, Oenothera, Raphanus, Rosa, solatium and Tagetes spp. were prepared for examination as follows: The styles were fixed in ethyl alcohol-acetic acid 3:1 for 1 hr, and hydrolyzed at 60°C for 5 to 60 min (depending on the species) in 45% acetic acid. The stigma with its attached strand(s) of stigmatoid tissue was then dissected out under a stereoscopic microscope, placed in a few drops of a staining solution made by dissolving 150 mg of safranin O and 20 mg of aniline blue in 25 ml of hot 45% acetic acid. After 5-15 min in this stain, the tissue was placed in a fresh drop of stain on a microscope slide and gently squashed under a cover glass. Because of a gradual precipitation of the aniline blue component, the stain had to be filtered regularly before use. However, a staining solution could be kept at room temperature for several weeks.     

Brilliant Cresyl Blue as a Stain for Plant Chromosomes

Methods are proposed for staining plant chromosomes with the dye brilliant cresyl blue, and for making these stained preparations permanent by using polyvinyl alcohol mounting medium.

The stain, which is composed of 2% brilliant cresyl blue in 45% aqueous acetic or propionic acid, is used with fixed material in making smear preparations. The technics for staining are similar to those employed in the aceto-carmine method.

The mounting medium is made by mixing 56% polyvinyl alcohol, which is diluted in water to the consistency of thick molasses, with 22% lactic acid and 22% phenol by volume. The permanent slides are made by floating off the cover slip of the temporary slide in 70% alcohol, then applying the mounting medium and replacing the cover slip.

The chief advantages of the methods described are:

1)The preparation of the stain is rapid and simple. The batch of stain will be good with the first try.

2)The staining procedure in some instances is shorter than when using aceto-carmine.

3)The stain shows a high degree of specificity for nuclear structures and gives better results than aceto-carmine when used on certain plant tissues.

4)A minimum number of cells is lost in making the slides permanent when using polyvinyl alcohol mounting medium as the slide and cover slip are run through only one solution prior to mounting.

5)The mounting medium dries rapidly and this shortens the time required before critical examination of the permanent mounts can be made.     

Making Autoradiographs of Individual Blood Cells

Single blood cell autoradiographs were made by smearing blood, diluted with serum, directly upon the emulsion of an Eastman NTB plate. After exposure, the plates were developed in D19. A 10% sodium sulfate solution was used for the photographic fixing to prevent lysis of the cells. After staining with Wright's stain the cells and autographs could be examined simultaneously under the microscope.     

Iodination-Coupled Tetrazonium and Ferric Ferricya-Nide Reduction Stains for Differentiating Red Blood Cells Containing Fetal or Adult Hemoglobin

Either the iodination-coupled tetrazonium reaction or the ferric ferricyanide reduction procedure can be used to differentiate red blood cells containing fetal hemoglobin (hemoglobin F) from those containing adult hemoglobin (hemoglobin A) in blood smears. Oxalated blood is diluted with 3 parts of physiological saline, and smears are made on slides. The air-dried slides are treated with absolute ethanol for 2 min, dried, and placed in phosphate-citrate buffer of pH 3.2-3.6 for 1 min at 37°C. They are then rinsed in distilled water, and dried for storage or stained at once by either the iodination-coupled tetrazonium or the ferric ferricyanide reduction procedure. Adult hemoglobin is extracted by the buffer, so that red blood cells containing fetal hemoglobin give a much darker stain than those containing adult hemoglobin. The hemoglobin S of patients with sickle-cell anemia behaves like adult hemoglobin.     

Sequential Use of Wright's and Ziehl-Neelsen's Stains for Demonstrating Phagocytosis of Bacterial Spores

Nongerminating spores, germinating spores, and vegetative cells of Clostridium botulinum type A were observed during phagocytosis in the peritoneal fluid of white mice. Since phagocytes are easily ruptured by heat, the method described avoids heating, as this has been employed in conventional spore staining methods. A thin smear of the fluid is air dried on the slide for 2 hr, and stained by Wright's method: stain, 2 min; dilution water, 2 min; and rinsed; then in 0.005% methylene blue for 30 sec, and rinsed. This is followed by Ziehl-Neelsen's stain for 3-4 min and destained with 1: acetone-95% ethanol for 10 sec. The slide is rinsed, and Wright's staining repeated: stain 1 min, dilution 2-3 min; and thereafter washed about 5 ml of Wright's buffer. Blotting and air drying completes the staining. Non-germinating spores stain light red with a red spore wall, germinating spores are deep red throughout, vegetative cells are blue, and leucocytes show a dark purple nucleus and light blue cytoplasm.     

A Urea Silver Nitrate Method for Nerve Fibers and Nerve Endings

A silver nitrate stain for nerve fibers and endings applicable to paraffin sections on the slide utilizes the properties of urea to accelerate the procedure and improve the specificity of the stain. After removal of the paraffin the sections are run through absolute, 95% and 80% alcohol and placed for 60-90 minutes at 50-60°C. in: 1% aqueous silver nitrate, 100 ml.; urea, 20-30 g.; 1g. mercuric cyanide and 1 g. picric acid in 100 ml. of distilled water, 1-3 drops. After the silver bath they are rinsed quickly in 2 changes of distilled water and reduced for 3-5 minutes at 25-30°C. in: water, 100 ml.; sodium sulfite, anhydrous, 10g.; hydroquinone, 1-2g.; urea, 20-30g. They are then washed thoroughly in 4-5 changes of distilled water, passed through graded alcohols into 80% alcohol and examined under the microscope. If nerve fibers are not distinct, the sections are returned to the same urea-silver-nitrate bath for 10-15 minutes, rinsed, reduced, washed and dehydrated as before. This process may be repeated until staining is adequate; then they are dehydrated, cleared, and mounted.

Nerve fibers show a color range from brown to black; nerve cells from yellow to brown; and the background, depending on the type of tissue and its fixation, from yellow to light brown.     

Hydrogen Chloride Gas for Effecting Staining of Internal Structures in Escherichia Coli.

The cells were smeared in water or water which had stood over about 10 mg. of magnesium powder per ml. for 30 minutes or longer. After the smear was dry and whitish in appearance it was held over a beaker of hot water (60-65° C.) until it was translucent or becoming translucent and exposed immediately to hydrogen chloride (gas) for a few seconds. After drying, it was covered with a 0.1% aqueous solution of neutral red for 5-8 minutes. The excess stain was washed from the slide with water and, while wet, placed in a saturated aqueous solution of mercuric nitrate for 5-15 seconds. The smear was rinsed in water and allowed to dry. When dry the slide was placed on a 50° C. warm plate and covered with a thin film of a 5% aqueous solution of nigrosin adjusted to a pH of about 3. The film dried quickly and upon cooling was ready for study. The stained material in the cells varied in shape and location with the moisture content of the smear and the time of exposure to hydrogen chloride. In the area of the smear directly exposed to the gas, the cells in general possessed a round or oval stained structure. Where there was little, if any, exposure to the gas the cells were uniformly stained. There were various gradations in the location and shape of the stained material in the cells from the one extreme to the other.     

A Nuclear Stain for Escherichia Coli And Related Organisms

The dye base of new fuchsin was precipitated by adding potassium hydroxide to the dye solution. The precipitate was filtered out and washed with water. It was then suspended in water, brought into solution and adjusted to a pH of about 5.0 with nitric acid. The staining solution was prepared by adding 0.3 ml. of a 14% aqueous solution of pyrogallol and 0.1 ml. of a 1% aqueous solution of boric acid to 3.0 ml. of the dye solution. Smears of cells were made in water on a slide and allowed to dry before covering with the staining solution which was also permitted to air dry. The smear was then washed in water and mordanted for 5-20 seconds in a 0.1% aqueous solution of mercuric nitrate. After rinsing in water, the smear was air dried. When dry, the slide was placed on a 50° C. warm plate for a few seconds before covering with a very thin film of a 5% aqueous solution of nigrosin which had a pH of about 5.0.     

Permanent-Type Mounting of Aceto-Iron-Haematoxylin Squashes in Corn Syrup

Preparations obtained by the aceto-iron-haematoxylin technique reported previously (Stain Tech., 37, 27-30, 1962) can be made relatively permanent either by ringing the cover slip with Karo corn syrup, or by mounting the squash in this syrup after separating slide and cover slip by the solid CO₂ freezing technique. In the latter procedure, both slide and cover slip may be placed briefly in 45% acetic acid for further differentiation of the stain, then recombined with a drop of syrup.     

An Improved Pollen Grain Smear Method for Wheat

Anthers containing actively dividing pollen grains were treated 1 hour at 18-20° C. with 0.2% solution of colchicine, washed 1 hour in water, soaked in 0.002 M aqueous solution of 8-oxyquinoline at 10-14° C. for 1 hour, washed in water for 1 hour and then fixed in Carnoy's solution (alcohol, chloroform, acetic acid, 6:3:1) for 6 hours to overnight. They were washed successively in acetic-alcohol (1:1) 10-15 minutes, 70% alcohol 10-15 minutes and in water 30 minutes before hydrolysing them in bulk in 1 N HCl at 60° C. for 10-15 minutes. “Finally, they were stained in leuco-basic fuchsin for 15-30 minutes. Pollen grains were squeezed out of a stained anther in a small drop of egg albumen on a slide and the albumen smeared uniformly on the slide. The slide was dipped successively for a few seconds in glacial acetic acid and 45% acetic acid respectively. The smear was covered by a cover glass in a drop of aceto-carmine and pressed gently between folded filter papers. The cover glass was sealed with paraffin and stored overnight. To make the preparation permanent the paraffin was removed and the cover glass separated in a 1:1 mixture of acetic acid and n-butyl alcohol. The slide and the cover glass were then passed through n-butyl alcohol, 2 changes, and finally remounted in balsam.     

Open-Face, Epoxy Embedding of Single Cells for Ultrathin Sections

Intact stamens of Tradescantia were fixed, dehydrated, and infiltrated with an epoxy resin. Each stamen was then put into a drop of resin on a microscope slide, which was transferred to the stage of a dissecting microscope so that individual hairs could be detached from the filament with fine tungsten needles. The detached hairs were transferred to drops of resin ca. 2 mm in diameter (6 or 7 in each of two rows) lying on a slide heavily coated with evaporated carbon. Polymerization was carried out in an oven until the resin attained a degree of viscosity that permitted orientation of the isolated hairs (by using a compound microscope) without their subsequent dislocation. When the small drops of resin had hardened after further polymerization, the positions of the hairs were marked by circumscribing the cells with India ink. The block was pried from the slide after rapid cooling with solid CO₂, and was then trimmed and sectioned. Cells suspended in culture medium were embedded in much the same way; they were centrifuged to obtain a pellet, which was fixed, dehydrated, and infiltrated. A small fragment of the pellet with a little resin was placed on a microscope slide, where the cells were dissociated under a dissecting microscope at ca. 100 × magnification. Individual cells were then picked up with tungsten needles and transferred to droplets of resin on a carbon-coated slide. The subsequent steps were similar to those described for the staminate hairs. Pieces of tissue in the 50-500 μ range were also handled by the foregoing technique. However, after infiltration they were put into large drops of resin on a slide coated with silicone mold-release rather than on a surface coated with carbon.     

Permanent Stained Preparations of Thick Blood Films

Fixing thick films in alcoholic solution of dye after the usual staining-and-laking procedure preserves the appearance of parasites and blood elements very similar to that of the usual thick films (not fixed) for the diagnosis of malaria and relapsing fever.

Procedure recommended: Films are stained and laked for 15 minutes in diluted Giemsa—1 to 3 drops of stock solution (0.4 g. in 60 ml. equal parts absolute methyl alcohol and glycerin) per ml. distilled water; rinsed in water and allowed to dry. They are then immersed in, or flooded with, May-Griinwald's stain (0.5% in absolute methyl alcohol) for 30 seconds, rinsed in water and allowed to dry. Solutions of MacNeal's tetrachrome stain in methyl alcohol and glycerin may be substituted for Giemsa and a solution in methyl alcohol may be substituted for May-Griinwald. With slight modification of the procedure, both thick and thin films on the same slide may be stained together.

Films stained and fixed as described, and mounted in Diaphane, have shown no evidence of fading in 3 years.     

A Smear Technic for the Cucurbitaceae

It is very difficult to make satisfactory smear preparations of species in the Cucurbitaceae by ordinary methods because, (a) the differentiation between chromosomes and cytoplasm is poor, (b) the pollen mother cells are held together in tissue-like masses, (c) the chromosomes are comparatively small and numerous. Special procedures have been devised to overcome these difficulties. Staminate buds selected at the proper stage of maturity are fixed for a period of 12 to 24 hours in a mixture composed of 3 parts of 100% ethyl alcohol and 1 part acetocarmine to which a small quantity of iron acetate has been added. After prefixation the material is rinsed in several changes of 100% alcohol. It can then be transferred directly to the slide for smearing, or hydrated to 70% alcohol for storage. For smearing the anthers are dissected out, the extraneous flower parts discarded, and the storage fluid removed. A drop of acetocarmine diluted to one-half strength with 45% glacial acetic acid is added, and the anthers are macerated into small pieces and smeared. The anther debris is removed, and the cover slip added. It is necessary to carry out the above operations with a low power binocular microscope. After heating the cover slip can be sealed with Pyseal for temporary storage.     

Method for Staining Bacterial Flagella

The following method of staining bacterial flagella is ecommended for use on smears made from suspensions of 10 to 16-tour agar slant cultures, incubated 30 minutes at 37°C before spreadng on thoroly cleaned and named slides:

1. Cover with fixative (100 cc. of 1/4 sat. aqu. solution picric acid, with 5 g. tannic acid and 7.5 g. ferrous sulfate).
2. Wash with tap water, dry and cover with Fontana spirochaete stain; heat to steaming and allow to act for 1 to 2 minutes. Wash in ap water. The stain is prepared as follows: To 25 cc. 2% AgNO₃ add dilute ammonia till the precipitate which forms redissolves; then add more AgNO₃ till a faint turbidity results. A clear solution is useess.