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.     

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 naphthol yellow S and erythrosin B staining procedure for use in studies of the acrosome reaction of rabbit spermatozoa.

Rabbit spermatozoa suspended in Krebs-Ringer-phosphate containing 0.25% glucose were smeared on polylysine-coated slides and dried in air at room temperature for 30 min at room temperature, blotted, rinsed in 1.0% aqueous acetic acid for 10-15 sec, drained and stained for 7 min in a mixture of equal parts of aqueous naphthol yellow S and erythrosin B (final concentration of each dye 0.1% w/v) at pH 4.6-5.0 (pH adjusted with acetic acid). Stained slides were well rinsed in distilled water adjusted to pH 4.6-5.0 with acetic acid, blotted, allowed to dry completely, rinsed in xylene and mounted in synthetic resin. Acrosomal caps were stained cherry-red (apical ridge) to pink (dorsal and ventral aspects); postnuclear caps stained pale pink; nuclei were either unstained or stained a very faint yellowish-pink. The mid-piece and flagellum were stained different shades of pink. The procedure is simple, rapid, and gives highly reproducible results. When present, acrosomes are easily detected regardless of the density of the smear.     

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.     

A Naphthol Yellow S and Erythrosin B Staining Procedure for Use in Studies of the Acrosome Reaction of Rabbit Spermatozoa

Rabbit spermatozoa suspended in Krebs-Ringer-phosphate containing 0.25% glucose were smeared on polylysine-coated slides and dried in air at room temperature for 2 hr to overnight. Smears were stained in 0.1% naphthol yellow S in 1.0% acetic acid for 30 min at room temperature, blotted, rinsed in 1.0% aqueous acetic acid for 10-15 sec, drained and stained for 7 min in a mixture of equal parts of aqueous naphthol yellow S and erythrosin B (final concentration of each dye 0.1% w/v) at pH 4.6-5.0 (pH adjusted with acetic acid). Stained slides were well rinsed in distilled water adjusted to pH 4.65.0 with acetic acid, blotted, allowed to dry completely, rinsed in xylene and mounted in synthetic resin. Acrosomal caps were stained cherry-red (apical ridge) to pink (dorsal and ventral aspects); postnuclear caps stained pale pink; nuclei were either unstained or stained a very faint yellowish-pink. The mid-piece and flagellum were stained different shades of pink. The procedure is simple, rapid, and gives highly reproducible results. When present, acrosomes are easily detected regardless of the density of the smear.     

Staining of Animal Tissues with the Dye Base of Methylene Green in Benzene to Facilitate Identification and Selection of Material

Methylene green as the free dye base dissolved in benzene can be used to stain animal tissues sufficiently during clearing for paraffin embedding to distinguish morphological features. It can therefore be used to allow trimming off unwanted tissues, to orient specimens when casting in wax and to select sections from a wax ribbon. The stain is removed very quickly on contact with water, and has no effect on subsequent staining techniques. The dye base is prepared by adding 2.5 ml of 10% NaOH to 100 ml of 1% aqueous methylene green, mixing well and allowing to stand at 20-30 C for 24 hr. The precipitate is separated by filtration, washed on the paper and dried at 50-55 C. The dry precipitate is dissolved in 200 ml of benzene to make the stock solution, which is diluted with 1 or 2 parts of benzene for tissue blocks, but used undiluted on sections.     

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.     

The Gallocyanin-Chrome Alum Stain; Influence of Methods of Preparation on Its Activity and Separation of Active Staining Compound

A dye, which is probably a cationic chelate, has been separated from a gallocyanin-chrome alum staining solution and prepared in the dry form. This dye is apparently the major staining compound. To prepare the chelate or dye, dissolve 150 mg of gallocyanin and 15 gm of chrome alum in 100 ml of distilled water and boil for 10-20 min, cool, filter, wash the precipitate with sufficient distilled water to restore the volume of the filtrate to 100 ml, then add concentrated NH₄OH until the pH is raised to 8-8.5. Filter, with suction, through a medium porosity fritted glass funnel. Wash with 100-200 ml of anhydrous ethyl ether and air dry the precipitate. This ratio of chrome alum to gallocyanin and the 10-20 min boiling time are optimal for preparation of the staining solution, which may be used either for staining or for separation of the chelate in its dry form. From the dried chelate, the staining solution is prepared as a 3% solution in1 N H₂SO₄ and a staining time of 16-24 hr is required. No differentiation is needed; the stain is self-limiting.     

Technic for Studying Chromosome Structure

The methods described are modifications of various technics for the study of spiral structure in chromosomes. They enable permanent preparations to be made with better fixation and allow the use of stains which give clear and more critical definition. The first method described involves the use of ammonium, hydroxide (880 vols.) fumes for the treatment of pollen mother cells before fixation. Anthers of Tradescantia are smeared on a slide and wet in a 3% cane sugar solution. The preparation is then immediately placed in a dish of fixative where it remains for two hours. The slide can then be washed, bleached and stained with gentian violet or hematoxylin. It was found that fumes of nitric acid, hydrochloric acid and glacial acetic acid gave similar results. For the second method, boiling water is used for pre-treatment. A smear is made on a slide and immersed in boiling water for five to ten seconds. The smear is then fixed and treated in the usual manner.     

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.     

Differential Stain for acid fast Bacteria and Spores

A modified method of staining acid-fast organisms is described. After staining with carbol-fuchsin as usual in the Ziehl-Neelson method, wash with water and while the slide is still wet cover with a saturated acetone solution of malachite green for three to five minutes. Wash and examine. The acid-fast organisms and spores are red in a green background. If the smear is thick and appears too dense, dry for three minutes and hold over the mouth of a bottle of ammonia until decolorized to suit. Upon exposure to the air the green returns. This can be prevented by keeping the smear alkaline, by the addition of sodium bicarbonate.

A second method is described for use with sputum in which acid-fast organisms are scarce. It permits the examination of thick smears and therefore increases the chances of finding tubercle organisms when few in number. Stain with carbol fuchsin as in the Ziehl-Neelson method. Decolorize with 30% phenol-disulfonic acid in water for a few seconds or until decolorized. Wash and examine at once. If color returns upon washing decolorize again. The tubercle organisms appear red in a colorless background.     

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.     

Preparation and Use of Aldehyde Fuchsin Stain in the Dry Form

A three-day old aldehyde fuchsin staining solution (Gomori, 1950) was precipitated in a separatory funnel by adding 50 ml. of chloroform and 200 ml. of distilled water to each 100 ml. of the staining solution. The mixture was shaken, allowed to settle and the precipitate-bearing layer filtered through paper. The precipitate was dried at 50°C, scraped from the paper and stored in a stoppered vial. To use, 0.5 g. of the dry stain was dissolved in 100 ml. of 70% ethanol containing 1 ml. of concentrated hydrochloric acid. In the dry form, the dye has retained its property of staining thyrotroph cells and neurosecretory substance in the hypophysis of the rat for several months.     

Improving the Staining Qualities of Toluidin Blue

To a 1.0% filtered aqueous solution of toluidin blue add drop by drop 4-5 ml of either a saturated aqueous solution of HgCl₂ or of KI. Collect the resulting dark precipitate on a filter paper and wash it with numerous small quantities of distilled water applied to both inside and outside of the filter paper. Wash until the drippings are distinctly blue (equivalent to about a 0.05% dye solution). Remove the paper and its contents from the funnel and dry either at room temperature or at 37°C. When dry, the treated dye can be brushed off the paper and stored. To prepare a staining solution add a weighed amount (0.12 gm if derived from the HgCl₂ treatment, or 0.3 gm if from KI) to 100 ml of distilled water. This insures a saturated solution in either case and gives a satisfactory stain with most sections in 10-30 min. Thionin and other members of the thiazine dyes also showed improvement in staining qualities after this treatment.     

Concentrating and Staining Bone Marrow; An On-the-Slide Technique

A 0.5-1 ml sample of bone marrow is aspirated into a syringe containing 3 drops of 15% K₂-EDTA and an additional 1-2 drops of the EDTA solution previously placed on a slide, is then drawn into the syringe. All of the contents are ejected onto this slide, which is carefully tilted 2 or 3 times to an angle of 5-10°, and the edge brought to the center of another slide. The slide with the aspirate is then slowly tilted to 80-90°. Most of the blood and part of the marrow will drain off, leaving spicules of marrow and some blood on the original slide. A small drop of this concentrated marrow is dragged off with the edge of a third slide and deposited about 2 cm from the edge of a fourth slide on which the smear is to be made. The smear is made by bringing a clean (smearing) slide to the slide with the deposited marrow with flat surfaces parallel and the edges at a 90° angle. With gentle pressure, the smearing slide is pushed toward the empty end of the slide upon which the smear is made. This separates the marrow from the circulating blood. Before staining the smear is air dried and heated in an oven at 120-125 C for 2 min; or alternately for satisfactory but less uniform results the smear is heated over a microburner for 10 sec; then the smear is covered with 1 part of undiluted Wright's stain for 30—45 sec which is then diluted with 2 parts of a solution of 0.1-0.2 gm of Na₂S₂O₃ in 1 liter of distilled water and stained for 10-13 min with this diluted stain. Smears made in this manner have 3 concentric zones; the central zone contains the myeloid tissue; the middle, erythropoetic tissue; the outer, a mixture of blood and marrow.     

Specific Staining of Glycogen with Haematoxylin and Certain Anthraquinone Dyes

Specific staining of glycogen in rat liver fixed in chilled 80% alcohol, chilled formol alcohol or 10% neutral formalin has been accomplished with acid alizarin blue SWR, alizarin brilliant blue BS, alizarin red S, gallein, haematein, and haematoxylin solutions. TO prepare a staining solution, 1 gm dye, 1 gm K₂CO₃ and 5 gm KCl were dissolved by heating in 60 ml of water. Concentrated NH₄OH (0.880 sp.gr.), 15 ml, followed by 15 ml of dry methanol were added to 20 ml of the cooled solution. Paraffi sections were stained for 5 min, rinsed in dry methanol, cleared in xylene, and mounted in D.P.X. The high specificity obviated the need for counterstaining: nuclei and cytoplasm were unstained. Precipitation of stain onto the slide was rare. As all the dyes carried, like carminic acid, numerous groups capable of forming hydrogen bonds, it is suggested that the staining mechanism involved hydrogen bonding.     

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.     

Further Studies on a Modification of the Gram Stain

In perfecting the modification of the Gram-stain previously proposed, the following points are of interest:

1. Acetone is too strong a decolorizer for Gram-positive organisms and alcohol too weak for Gram-negative organisms. Consequently, it is now recommended that equal parts of acetone (100% c.p.) and ethyl alcohol (95%) be used as a decolorizing agent. The time of application should not ordinarily exceed 10 seconds.

2. Aqueous basic fuchsin (0.1%) serves as a strongly contrasting counterstain. Prolonged application renders Gram-positive organisms doubtful or Gram-negative, while short application renders Gram-negative organisms doubtful or Gram-positive. Twenty (20) seconds is therefore recommended as the time of application of the counterstain.

3. The method here described, with due regard for its limitations, is of value in Gram-staining pure or mixed cultures as well as for organic materials, such as Acidophilus milk, feces, etc., either for research purposes or classroom use. The method is as follows:

Air-dry film and fix with least amount of heat necessary.

Flood with dye for 5 minutes. Previously mix 30 drops of a 1% aqueous solution of crystal violet or methyl violet 6B with 8 drops of a 5% solution of sodium bicarbonate. Allow the mixture to remain for 5 minutes or more.

Flush with iodine solution for 2 minutes. Two grams iodine dissolved in 10 cc. normal sodium hydroxide solution and 90 cc. water added.

Drain without blotting but do not allow film to dry.

Add a mixture of equal parts of acetone and alcohol drop by drop until the drippings are colorless. (10 seconds or less.)

Air-dry slide.

Counterstain for 20 seconds with 0.1% aqueous solution of basic fuchsin.

Wash off excess stain by short exposure to tap water and air-dry. If slide is not clear immersion in xylol is recommended.     

The Use of Pyrogallol to Demonstrate Peroxidase in Mammalian Blood Eosinophils

Canine blood films were fixed in a mixture of formaline (40% HCHO) and 95% ethyl alcohol, 1:9, for 30 sec, washed in distilled water and air dried. A mixture of 10% aqueous pyrogallol 6 ml and H₂O₂ (6% or 20 vols.) 0.1 ml were applied to the film, allowed to react for 6 min and then washed off with distilled water. The film was counterstained with May-Grünwald Giemsa, Leishman, or Giemsa stain. This method stains canine eosinophils specifically for the presence of peroxidase, but has variable effects on eosinophils of other mammalian species, depending on the type of fixative used. Modified techniques are described for 4 other mammalian species and the possible causes of the staining variations are discussed.     

A Cresyl Fast Violet Stain for Bacteria and Fungi in Tissue

The cresyl fast violet staining method was modified to eliminate differentiation. Paraffin sections from tissues fixed in Zenker-formol were stained in a 1% aqueous solution of cresyl fast violet (Chroma), adjusted to pH 3.7 with acetic acid, washed in running tap water, dehydrated and covered. Because basophilia increases with time of fixation or storage in formalin or Kaiserling's fluid, dilution of the dye solution to 0.5-0.1% is recommended for such material. Bacteria, nuclei, Nissl substance, and lipofuscin were colored dark blue; fungi, blue to purple; and cytoplasm and muscle fibers, light blue. Collagen and reticulum fibers were only faintly stained. Thus, microorganisms were easily visible against the lightly colored background. In formalin-fixed material, bile pigment was colored olive green. Because this method does not require differentiation, it gave uniform results even in the hands of different users. Little or no fading was observed in sections stored for more than 2 yr.