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 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.     

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 Serum-Film Technic for Bone-Marrow Smears

A chemically clean slide was covered with a thin film of serum by using a glass spreader. A drop of marrow suspension (bone-marrow in autogenous serum, about 1 to 20 dilution) was placed upon this film and spread by blowing gently on it until a central area of about 2 cm in diameter became dry. The peripheral rim of the smear was then wiped off and the slide stained. A statistical analysis showed that the variation in size of cells did not influence their distribution within the area and that improved cell morphology with fewer damaged cells favored quantitative studies.     

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.     

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.     

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.     

The Smear Technic in Plant Cytology

The following method of making permanent smears of pollen mother cells is in general use and gives excellent results. Determine the stage of meiosis from aceto-carmin mounts. Smear the pollen mother cells on a dry slide. Fix in Navaschin's or a modified Flemming's solution from 1 to 2 hours. Wash in 10 to 20% alcohol from 15 to 30 minutes. Stain in 1% aqueous crystal violet from 1 to 5 minutes. Rinse in water and pass thru 30 to 50% alcohol, about 15 to 20 seconds in each. Transfer to 80% alcohol containing 1% iodine and 1% potassium iodide for 30 seconds. Destain with absolute alcohol, followed by clove oil. xylol, balsam and cover.

Permanent smears for chromosome counts can be quickly made by smearing pollen mother cells on a dry slide, fix and stain with aceto-carmin, dehydrate with mixtures of absolute alcohol and acetic acid, follow with xylol, balsam, and cover.     

The Use of Fluorescein Diacetate and Ethidium Bromide as a Stain for Evaluating Viability of Mycobacteria

A method for evaluating the viability of mycobacterial cultures using fluorescein diacetate and ethidium bromide is described. Smears from 2-4 week old cultures of mycobacteria were stained with a 0.005% Dubos albumin broth dilution of fluorescein diacetate for thirty minutes at 37 C. The smears were counterstained with a 0.005% aqueous solution of ethidium bromide for ten minutes at room temperature and blotted dry. One drop of glycerol was placed on the smear and covered with a 22 × 40 mm coverslip. Stained smears were observed with fluorescence microscopy at 450 X. Viable organisms hydrolyzed the fluorescein diacetate and appeared yellow-green while dead organisms absorbed the ethidium bromide and appeared red-orange. Aliquots of material from which the slides were made were concurrently placed on Lowenstein-Jensen media and cultured for growth as a confirmation of viability. Investigation of 104 mycobacterial isolates using the described test showed that the viability of mycobacterial cultures can be determined by this method.     

Crystal Violet Contamination of Methyl Green and Purification of Methyl Green—a Historical Note

A method for evaluating the viability of mycobacterial cultures using fluorescein diacetate and ethidium bromide is described. Smears from 2-4 week old cultures of mycobacteria were stained with a 0.005% Dubos albumin broth dilution of fluorescein diacetate for thirty minutes at 37 C. The smears were counterstained with a 0.005% aqueous solution of ethidium bromide for ten minutes at room temperature and blotted dry. One drop of glycerol was placed on the smear and covered with a 22 × 40 mm coverslip. Stained smears were observed with fluorescence microscopy at 450 X. Viable organisms hydrolyzed the fluorescein diacetate and appeared yellow-green while dead organisms absorbed the ethidium bromide and appeared red-orange. Aliquots of material from which the slides were made were concurrently placed on Lowenstein-Jensen media and cultured for growth as a confirmation of viability. Investigation of 104 mycobacterial isolates using the described test showed that the viability of mycobacterial cultures can be determined by this method.     

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.     

Silver Staining of Nerve Fibers in Cardiac Tissue

Fresh hearts of dog were perfused through the coronary vessels with 1000 ml. of fixative (chloral hydrate, 5 g. per 100 ml. of 70% ethyl alcohol) and blocks of tissue 2 × 5 mm. from epicardium to endocardium fixed 48 hours in the same fixative. The blocks were placed in 95% alcohol containing 0.3% addition of strong ammonia for 4 hours, followed by 2 changes of plain 95% alcohol of 1 hour each, then cleared and infiltrated with paraffin. Mounted sections 12-15 µ thick were incubated in 1% silver proteinate (obtained from Serumvertrieb, Marburg, Germany)² at 38° C. for 48 hours in the presence of 10 g. of 15 gauge copper wire per 200 ml. of solution. The slides were rinsed gently in 3 changes of distilled water for 2 minutes, 1 minute and 1 minute, respectively, and reduced in 1% hydroquinone and 5% sodium sulfite for 5 minutes. They were washed 5 minutes in tap water and 5 minutes in 2 changes of distilled water and toned 3-5 minutes in 0.25% gold chloride, rinsed in distilled water 10 seconds, reduced 10 seconds in 1 % oxalic acid, rinsed 1 minute, fixed in 5% sodium thiosulfate 5 minutes, washed in tap water through 3 changes, dehydrated, cleared and covered. All solutions were made with distilled water except where otherwise specified. The results gave good impregnation of fine nerve fibers without the usual confusing staining of reticular tissue.     

Histochemical Use of Sodium Bismuthate

Histochemical 1,2-glycoI cleavage, similar to that obtained with periodic acid and lead tetraacetate, may be obtained with sodium bismuthate. Routinely prepared slide sections, from tissues fixed in 10% formalin, are run down through xylene and graded alcohols to water and then oxidized for three minutes in a 1% sodium bismuthate 20% aqueous phosphoric acid solution. The oxidizing solution must be freshly prepared and used immediately. Following oxidation, sections are rinsed 15 sec. in IN HC1 to remove bismuth pentoxide precipitate, a by-product of the reaction. The sections are then washed in distilled water and placed in leuco-fushsin for 10 min., or in a saturated 30%) alcoholic solution of p-nitrophenylhydrazine for 5 min. or 2,4-dinitrophenylhydrazine for 30 minutes. After staining, the sections are rinsed in 30% alcohol if the nitrophenylhydrazines were used, or in the standard dilute sulfite bath followed by running tap water for 5 min. if leucofuchsin were used. Sections are routinely dehydrated, cleared, and covered. On examination, the sites of 1,2-glycol linkages will be stained violet by leucofushsin or yellow by the nitrophenylhydrazines.     

Histochemical Use of Sodium Bismuthate

Histochemical 1,2-glycoI cleavage, similar to that obtained with periodic acid and lead tetraacetate, may be obtained with sodium bismuthate. Routinely prepared slide sections, from tissues fixed in 10% formalin, are run down through xylene and graded alcohols to water and then oxidized for three minutes in a 1% sodium bismuthate 20% aqueous phosphoric acid solution. The oxidizing solution must be freshly prepared and used immediately. Following oxidation, sections are rinsed 15 sec. in IN HC1 to remove bismuth pentoxide precipitate, a by-product of the reaction. The sections are then washed in distilled water and placed in leuco-fushsin for 10 min., or in a saturated 30%) alcoholic solution of p-nitrophenylhydrazine for 5 min. or 2,4-dinitrophenylhydrazine for 30 minutes. After staining, the sections are rinsed in 30% alcohol if the nitrophenylhydrazines were used, or in the standard dilute sulfite bath followed by running tap water for 5 min. if leucofuchsin were used. Sections are routinely dehydrated, cleared, and covered. On examination, the sites of 1,2-glycol linkages will be stained violet by leucofushsin or yellow by the nitrophenylhydrazines.     

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.     

A new technique for staining sections and recovering celloidin film in high resolution autoradiography

Ultrathin sections are stained immediately after cutting by placing them in contact with staining solution and then placed on a slide covered by a celloidin film. This method largely avoids precipitates of heavy metals. The recovering of celloidin film is improved using a stainless steel basket. This technique is far more reliable than that involving use of a filter paper.     

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.     

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 Aceto-Osmium Method for Demonstrating Nematodes in Citrus Roots

Difficulties are encountered in observing nematodes in citrus feeder roots because of the presence of suberin and other unsaturated compounds. To obviate these difficulties, infected citrus roots, either fresh or preserved, are immersed in a covered jar for 2 hr at 52° C in a solution composed of distilled water, 16 parts; 10% acetic acid, 10 parts; and 2% aqueous osmium tetroxide, 2 parts. The stained, blackened roots are washed in running water for at least 1 hr and then bleached in 10-30% hydrogen peroxide at 32°C for a few seconds until the color of the roots lightens perceptibly. After several washings in water to stop the oxidation reaction, roots are dehydrated in 70, 95, and absolute ethanol held at 52 °C for 30 min at each concentration. After dehydration, roots are cleared in methyl salicylate at 52°C. Examination for nematodes in most cases, can be made after 30 min.     

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.