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.     

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.     

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.     

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.     

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.     

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.     

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.     

Staining the Nucleus in Yeasts

Yeast cells kept young by repeated subculturing were centrifuged, washed twice in distilled water and smeared on slides coated with a little egg albumen. The cells were treated with 0.002 M 8-hydroxyquinoline for 1 hr, fixed first in OsO, vapour for 30 sec and then in chloroform for 30 sec. The slides were passed through descending grades of alcohol, washed in distilled water, then immersed in 0.17 M NaCl solution for 2 hr. at 57°C. They were again washed in distilled water and later hydrolysed in 1 N HCl at 60°C for 5-7 min. This was followed by washing in distilled water and in buffer. The slides were then kept for 3 hr in Giemsa stain comprising 96 ml of phosphate buffer of pH 7.0 and 4 ml of the stain. After dehydration, mounting was done in balsam. Nuclei were brightly stained and well differentiated; centrosomes were clear, and the process of nuclear division and movement to daughter cells could be studied. Pretreatment with 8-hydroxyquinoline increased the viscosity of the cytoplasm, while NaCl treatment and acid hydrolysis led to the complete removal of ribonucleic acid and basophilic material. A selective staining of chromatin was thus achieved. Structures resembling chromosomes could be seen when fixed and stained cells were squashed, soon after the removal of the slides from the stain, under a cover glass by applying uniform pressure with a rubber stopper. Fixation in osmic acid vapor and chloroform followed by acid hydrolysis and staining in leucobasic fuchsin also helps to obtain bright staining of the nucleus; however, the preparations are inferior to those obtained after 8-hydroxyquinoline, NaCl treatment and Giemsa staining.     

A Pollen Grain Squash Technique for Saccharum and Related Genera

Anthers collected between 9 and 10 AM were treated for 1 hr at 26-28 C with a 0.5% solution of colchicine, washed for 2-4 min in water, placed in 0.002 M 8-hydroxyquinoline for 1 hr, washed in water for 10 min and fixed in: methanol, 60 ml; chloroform, 30 ml; distilled water, 20 ml; picric acid, 1 gm and mercuric chloride 1 gm, for 24 hr. After washing they were hydrolysed in 1 N HCl for 15 min at 60 C, stained in leuco basic fuchsin for 30 min, then smeared on a slide in a drop of acetocarmine. The slides were sealed, stored overnight, the paraffin was removed, and the slide passed through a 1:1 mixture of n-butyl alcohol and acetic acid, then through pure n-butyl alcohol and mounted in Canada balsam. The significant features of this procedure are: (1) use of chromosomes in the haploid condition for karyotype analysis, (2) better exaggeration of constrictions for easier interpretation of chromosome types and (3) good spreading in plants with a large chromosome number.     

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.     

Permanent Mounts Of Chromosomes After 8-Oxyquinoline and Squashing

Fresh young root tips or free-hand cross sections thereof were placed in 0.002 M 8-oxyquinoline (aq.) at 10-14^oC. for 3 hours. After rinsing in water 1-2 minutes, they were soaked in N HC1 at 55^oC. for 25 minutes, rinsed again and squashed under a cover glass on a dry slide. Slide and cover glass were separated by placing in 70% alcohol and allowed to remain therein at least 0.5 hour after separation. Both slide and cover glass were passed through 50% and 30% alcohol to water and stained by the Feulgen procedure (without further hydrolysis) or with crystal violet after mordanting in 1% chromic acid overnight and washing in running water 3-4 hours. Dehydration and mounting in balsam completed the process. The smear on the slide was covered with a clean cover glass and the cover glass, bearing stained material, mounted separately.     

An Acetic Acid Dissociation, Air-Drying Technique for Insect Chromosomes, With Aceto-Lactic Orcein Staining

The method differs from mammalian techniques for somatic chromosomes in that it uses very small amounts of material. Drosophila melanogaster and an ant, Dorymyrmex sp., are used as examples. Pretreatment with 0.05% Colcemid in insect Ringer solution is applied to mature Drosophila larvae for 5 hr, by feeding, but Dorymyrmex prepupae require puncture and a 15 hr exposure of the puncture to the solution. Organs are removed under 1% sodium citrate, tansferred to fresh citrate for 10-20 min, than fixed in acetic-methanol, 1:3, for 30 min. Transfer to a drop of 60% acetic acid on a clean warmed slide dissociates the cells, which are spread by adding a small drop of fixative and tilting the slide in all directions. After immersion in acetic ethanol, 1:3, for 4 hr, rinsing in the stain solvent and draining the slides then have 2-3 drops of aceto-lactic orcein placed on each, coverslips added, and warmed (at about 50 C) for about 12 hr or until staining is sufficient. They can then either be treated as semipermanent or made permanent by allowing the coverslips to slide off in acetic-ethanol, dehydrating, and mounting in Euparal, or a synthetic resin.     

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.     

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 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.     

Toluidine Blue Staining of Coccidia in Cultured Animal Cells

Cultured animal cells infected with various species of Eimeria (coccidia) from chickens are washed in Hanks balanced salt solution (HBSS) and fixed in 5% formalin in HBSS. The fixed cultures are washed briefly in distilled water to remove HBSS salts and then dehydrated in a series of mixtures of 40 to 50% ethanol with increasing concentrations of tertiary butanol (TB) and decreasing concentrations of distilled water. Cultures are placed for 1 min in a mixture of 2 parts ethanol :TB (25:75) and 1 part 0.05% toluidine blue O in McIlvaine buffer (pH 6.0), followed by 1 min in 0.05% toluidine blue O in McIlvaine buffer (pH 6.0). The stained cultures are dipped for 1-2 sec in TB, allowed to dry and mounted permanently on slides. Cover-slip cultures fixed and stained by this procedure 8 years ago have not faded or discolored. The alcohol mixtures, formalin in HBSS, stain and buffer can be prepared in large volumes and stored indefinitely. The staining procedure has proven to be rapid and dependable with a variety of cell types in monolayer cultures in research and teaching applications.     

Cleared Whole Mounts of Shoot Apices of Angiosperms for Topographic Study

Cleared and stained whole mounts of stem apices of two Labiates and of Phaseolus plumule giving a three-dimensional picture of the apical structure have been prepared as follows. Fix the buds in formalin-acetic acid-50% alcohol (5:5:90) for 24 hr or longer and then dissect under a binocular microscope to leave only the youngest leaves surrounding the apex. Wash for several minutes in distilled water and then clear the material in a 5% solution of sodium hydroxide at approximately 40° C for 24-48 hr. Wash thoroughly in several changes of distilled water, transfer to a solution of 1% tannic acid and 0.5% sodium salicylate for up to a minute. Wash briefly in distilled water and stain in a 1.5% solution of ferric chloride until blue-black. Wash in distilled water and dehydrate through 50%, 70%, 85%, 95% and 2 changes of absolute ethyl alcohol. If the xylem is not stained well, counter-stain for a few seconds in a 0.5% solution of safranin O in a 1:1 mixture of xylene and absolute alcohol and wash out the excess stain in the same mixture. Clear in 2 changes of xylene and place on a glass slide in thick Canada balsam. Orient with needles under low magnification and cover.     

Production of Carbon Films for Electron Microscopy; Hydrofluoric Acid Detachment from Glass

Chemically clean microscope slides are coated as usual by vaporized carbon. The carbon film is floated off the slide by slowly lowering it at an angle of 45° into 1% HF in distilled water containing 0.025% Tween 80. This solution fills completely (forming a positive meniscus at the edges) one chamber of a double-compartment Perspex trough; the other compartment being similarly filled with the Tween solution only. A Teflon bar, laid on top of the partition keeps the solutions from mixing. After the carbon film loosens, it is floated across the central partition into the second compartment with the aid of a second Teflon bar, using both bars to guide the film on the surface of the fluid. The HF is thus washed from the film. Grids are thinly coated with 0.5% poly isobutylene in toluene (as an adhesive) and previously placed on a rectangle of filter paper supported by wire screening about 1/2 inch from the bottom of the trough. While the Tween solution is drained away through a bottom opening, the carbon film is guided to cover the grids. The filter paper bearing the grids is then removed and caused to dry slowly (about 12-16 hr) to avoid cracking or distortion of the film.     

Chromosomes of Sequoia sempervirens; 8-Hydroxy-Quinoline-Castor Oil Pretreatment for Improving Preparation

Living root tips of coast redwood were pretreated for 6 hr at 10-14 C in a homogenized mixture of 0.003 M aqueous solution of about 5 ml of 8-hydroxyquinoline and a drop of castor oil. They were rinsed 2-3 times in Carnoy's alcohol-chloroform-acetic acid (3:2:1), and fixation allowed to continue in this fluid for 1 hr at 60 C. They were then hydrolysed in 1 N HC1 at 60 C for 45 min; washed with distilled water, and squashed in a drop of aceto-carmine or aceto-orcein to make a temporary slide, subsequently made permanent by a quick-freezing method. Our work to date seems to confirm 2n = 66 for S. sempervirens.     

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.