Staining of Microorganisms

STAINING (of microorganisms). A smear on a slide or coverslip is dried in the air; drying over a flame is not recommended, and placing the preparation near a flame is alone permitted. The dried preparation is fixed by passing it through the burner flame three times. Fixation using chemically active substances is rarely employed; absolute alcohol and ether a.a. (10–15 min.), methyl alcohol (2–3 min.), acetone (5 min.), concentrated aqueous mercuric chloride solution, alcoholic mercuric chloride, and others are recommended as such substances. Basic aniline dyes (fuchsin, methylene blue, safranin, methyl violet, gentian violet, malachite green, vesuvin, neutral red), which possess an affinity for the nuclein-rich protoplasm of microbial bodies, are most frequently used for staining. Acidic aniline dyes (eosin, orange G, acid fuchsin, picric acid) find application for contrast staining or background staining, thanks to which microbes stained in a different color stand out clearly. The staining of microbes is based on the same principles and obeys the same regularities as the staining of tissues (see Histological technique, as well as Dyes). Methods of contrast staining, when a mixture of dyes is used such that the microbes are stained one color and the background another, are of particular value. Water-alcohol or aqueous solutions of dyes are used for staining; for the preparation of the former, pre-formulated basic saturated alcoholic solutions of dyes are used [10–15 g of fuchsin (or 10–15 g of methylene blue) + 100 cm³ of 96% alcohol, 6–8 g of gentian violet + 100 cm³ of 96% alcohol, etc.], filtering them and diluting with 4 parts of distilled water; weaker dilutions (1 part of stock solution to 9–10 parts of distilled water) are also employed. For the preparation of aqueous solutions, 1–2 g of dye per 100 cm³ of water is taken. Special solutions in which the action of the dye is enhanced by the addition of certain substances having the character of a mordant find great application. Of such solutions, the most commonly used are Ziehl-Neelsen carbol-fuchsin (Ziehl, Neelsen), frequently used diluted with water (1:5 or 1:10); Loeffler's stains (for the preparation of stains according to Loeffler, see Loeffler's methods of staining, media); Kühne's carbol methylene blue (Kühne) (1.5 g of methylene blue, 10 cm³ of absolute alcohol, 100 cm³ of 5% carbolic water); Manson's stain (Manson) (2 g of methylene blue, 5 g of borax, 100 cm³ of boiling water). The staining of smears is carried out in such a way that the coverslip (resp. glass slide) is grasped with a Cornet forceps, and the staining solution is poured onto the preparation with a pipette, covering the entire coverslip (or the preparation on the slide); staining lasts 3–5 minutes in the cold, 10–60 seconds when heated over a flame; the procedure ends with washing with water. If the preparation is made on a coverslip, it is placed stained-side down on a glass slide, carefully drying its upper surface with filter paper. Gram's method (see) belongs among the special staining methods. Modifications of this method are: 1) Nicolle's method—staining for 1–5 min. with heating by carbol-gentian violet (10 cm³ of alcoholic dye solution per 100 cm³ of 1% carbolic water), treatment for 4–6 min. with Lugol's solution (1 g of iodine, 2 g of KI, 200 cm³ of distilled water), repeated several times, decolorization in a mixture of 3 parts of absolute alcohol and 1 part of acetone; 2) Claudius's method—staining for 1 min. with carbol-methyl violet resp. gentian violet (as in Gram's), washing with water, drying with filter paper, treatment with a saturated solution of picric acid and distilled water in equal parts, washing in water, drying with filter paper, treatment with chloroform, drying. Gram's method is a method of double staining (bacteria not decolorized by Gram are stained dark violet, while other microbes take up the color of the counterstain). The Ziehl-Neelsen method (see) and other staining methods based on acid-fastness (see Sputum) also belong to the methods of double staining. The Pick-Jacobsohn method is a contrast method: staining for 8–10 seconds with a mixture of 15 drops of Ziehl's carbol-fuchsin, 8 drops of a saturated alcoholic solution of methylene blue, and 20 cm³ of distilled water; with this method, bacteria are dark blue, cell nuclei light blue, and everything else (tissues) red. For staining bacterial capsules, the following methods are recommended: 1) Johne's method (see Capsule bacteria); 2) Klett's method—staining to boiling with a mixture of 10 cm³ of a saturated alcoholic solution of methylene blue and 100 cm³ of distilled water, washing with water, counterstaining for 5 seconds with a fuchsin solution prepared according to the same recipe as the methylene blue; bacteria are blue, capsules pink; 3) Friedländer's method—after fixation, washing the preparation for 2 minutes in a 1% solution of acetic acid, drying, staining for a few seconds in a solution of gentian violet in saturated aniline water. For staining spores, Møller's method is used: fixation for 2 min. in absolute alcohol and for the next 1/2 min. in chloroform, treatment without prior washing with a 5% solution of chromic acid for from 5 sec. to 10 min. depending on the species of the stained microbe (to be tested beforehand), washing in water, staining with Ziehl's carbol-fuchsin with heating for 5 min. or in the cold for 5 hours, decolorization with 5% sulfuric acid for 5 sec., washing in water, and counterstaining with an aqueous solution of methylene blue. For staining flagella, the following have been proposed: 1) Loeffler-Nicolle-Morax method—treatment with heating until vapors form using a mordant of the following composition: 10 cm³ of a 25% aqueous solution of tannin, 5 cm³ of a cold-saturated solution of ferrous sulfate, and 1 cm³ of a saturated alcoholic solution of fuchsin; staining with Ziehl's fuchsin with heating until vapors form for 10–15 seconds, washing in distilled water, and drying. 2) Van Ermengem's method—treatment for 1/2 hour in the cold or 5–10 min. at a temperature of 50–60° with a mordant of the following composition: 1 cm³ of 2% osmic acid, 2 cm³ of 20% tannin solution, 4 drops of acetic acid; thorough washing in distilled water, treatment for 2–3 min. in a 0.5–2% silver nitrate solution, treatment without washing in a solution of the following composition: Acid. gallici 5 g, tannin 3 g, Natr. acet. fus. 10 g, distilled water 350 cm³; repeated treatment in silver nitrate solution until slight darkening of the solution, abundant washing with water, rapid drying; if staining is insufficient, the entire procedure is repeated starting from treatment with Acid. gallici solution, etc. 3) Pitfield's method as modified by Benignetti and Gino: to 3 cm³ of a saturated alcoholic solution of gentian violet are added 5 cm³ of an aqueous saturated solution of alum and 5 cm³ of the following solution: 1 g of zinc sulfate, 10 g of tannin, and 100 cm³ of water; a few drops of this stain are poured onto a drop of microbial suspension applied to a slide, heated until vapors form, thoroughly washed in water, and dried in air. 4) Lanceraux's method—treatment at 60° with a mordant of the following composition: antimony chloride 1 g, tannin 5 g, formalin 10 cm³, distilled water 100 cm³; after leaving the preparation in the cold for 10 minutes, washing first in tap water and then in distilled water, treatment with a silver solution (silver nitrate 1 g, distilled water 20 cm³) with slight heating until a metallic sheen appears, abundant washing first in tap water and then in distilled water, treatment for 1 min. with a solution of 1 g of metol (paramethylaminophenol) in 20 cm³ of distilled water, washing in water, drying. When staining flagella, it is necessary to use perfectly clean coverslips, washed first in alcohol and then in ether. A drop of sterile water is placed on the coverslip, into which a loop of the studied culture is then introduced. Capsules and flagella can also be studied using methods based on the use of Indian ink. Such is Gins's method (a modification of Burri's method): a drop of Chinese ink is placed on a slide and then mixed with an equal volume of a drop of water into which a loop of the studied culture has been preliminarily introduced; the mixture is smeared with the edge of a coverslip and examined using immersion; bacilli, their capsules, and flagella clearly stand out unstained against the dark background of the ink. A similar method for capsules was proposed by Borin. A drop of normal human, horse, etc. serum is placed on a glass slide; it is distributed in a thin layer with the edge of a coverslip and dried over a burner flame.

After this, a drop of Chinese ink with a culture of the microbes under study is applied to the slide, distributing the material with a platinum loop. The smear is moistened with xylene, the slide is turned downward to remove excess liquid, a drop of cedar oil is applied to the smear, and examination is carried out using immersion; the background is black, the capsules are light gray. For post-vital staining, the Sabrazes method is used: staining is performed with toluidine blue (0.5 g of toluidine blue, 10-15 cm³ of 95% alcohol, 3 g of carbolic acid, sterile distilled water up to 100 cm³); the smear on the glass slide must be thin and well dried, a drop of the stain is applied to a coverslip and inverted onto the smear; the method is used for staining sputum, urine sediments, etc.; cell protoplasm is pale blue, cell nuclei are violet with a red tint. Fungi are stained for 5-10 minutes with Loeffler's methylene blue or carbolic thionine; after staining, washing with water, rapid treatment with absolute alcohol, drying, and examination in a drop of cedar oil are performed. Staining of bacteria in sections. For the technique of preparing specimens, see Histological technique. 1) Loeffler's method: staining for 5-30 minutes in an alkaline solution of methylene blue, differentiation for several seconds up to ¹/₂ minute (depending on the thickness of the section) in ¹/₄-1% acetic acid, dehydration in absolute alcohol, clearing in cedar oil (see also Nikiforov's methods). 2) Pfeiffer's method: staining with diluted (1 : 3) Ziehl's carbol-fuchsin for 15-30 minutes, treatment with absolute alcohol + 1-2 drops of acetic acid, cedar oil or xylene (as soon as the section begins to take on a pink-violet tint). 3) Nicolle's method: staining with carbolic thionine for ¹/₂-1 minute, washing in water, treatment with absolute alcohol to extract water, then cedar oil. 4) Gram's method: staining for 5-30 minutes with carbolic methyl- or, respectively, gentian violet, treatment for 1-2 minutes in Lugol's solution, absolute alcohol until the section is decolorized, cedar oil; additional staining can also be added, as in the staining of smears (see Gram's method). Even better results are achieved by staining according to Weigert's method for fibrin (see Weigert's staining methods). 5) Ehrlich-Ziehl-Neelsen method for staining acid-fast bacteria: staining in fuchsin on aniline water for 1-24 hours, decolorization for 10 seconds in 5% sulfuric or 25% nitric acid, washing in 70% alcohol until the specimen is decolorized, additional staining with diluted Loeffler's serum (1 : 3) for 2-5 minutes, washing in ¹/₄-¹/₂% acetic acid, dehydration in absolute alcohol, clearing, etc. Stains on aniline water are prepared in the following manner: aniline oil is poured into a test tube so that its lower part is filled, filled up to ³/₄ of the test tube with water and shaken vigorously, filtered through a moist filter, and enough saturated alcoholic solution of the stain is added so that the staining solution is translucent.

V. Lyubarsky. Staining of protozoa. The study of fixed and stained preparations must be preceded by examining the parasite in a living state with a semi-closed diaphragm or in a dark field. If a fresh preparation is made on an ordinary glass slide, then, to avoid damaging large protozoa, hairs, bristles, thin glass rods, and also strips of filter paper are placed under the edges of the coverslip, which furthermore prevent the too rapid movement of the protozoa. The addition of various reagents to a fresh preparation makes it possible to carry out microchemical reactions (addition of Lugol's solution—a reaction for starch, a reaction for glycogen, etc.). The main requirement in the staining of protozoa is to preserve the natural shape and structure of the protozoon as perfectly as possible. The use of dry smears has a limited significance here and is suitable only for protozoa that endure drying without significant alteration of their morphology. This relates chiefly to blood parasites, in respect to which dry smears are used very widely for diagnostic purposes. The preparation of dry smears, their staining according to the Giemsa method, as well as the preparation and processing of a thick drop—see Giemsa and Malaria—clinic. For large, water-rich protozoa, as well as for protozoa that exist in their natural state in fluids poor in coagulable protein (aquatic and intestinal protozoa), dry fixation does not yield good results. Here, wet fixation and subsequent wet processing of the smear must be applied, paying attention that the smear does not dry out before mounting in Canada balsam. Wet fixation is performed in the following manner. Fresh thin smears on coverslips, before drying, are quickly immersed with the smeared side down into a Petri dish or watch glass with the fixing fluid. After a few minutes, the slides are turned with the smeared side upward and placed on the bottom of the dish, thanks to which the surface of the dish is freed for new smears. Very many liquids have been proposed for fixation. Sublimate fixation perfectly preserves the structure of the nucleus and protoplasm. Sublimate is ordinarily used in the form of sublimate alcohol according to Schaudinn: 2 parts of a concentrated aqueous solution of sublimate and 1 part of 96% alcohol. To this mixture, 0.5–5.0% of glacial acetic acid is added. Subsequent processing: 70% iodine alcohol (tincture of iodine is added until the color of strong tea), which frees the smears from an excess of sublimate. The excess iodine is removed with hyposulfite or by prolonged washing in 70% alcohol. Smears are preserved in 70% alcohol for an indefinite time until staining. In sublimate fixation, instead of metal instruments, which are spoiled by sublimate, glass, horn, wooden instruments, etc., are used. Osmic acid is also frequently used as a fixing agent, which fixes the forms of movement of protozoa particularly well and is used in the form of vapors or various mixtures. Fixation with vapors is performed in the following manner. On the bottom of a glass cylinder with a well-ground stopper, 0.25–0.5 of osmic acid is placed; the entire bottom is covered from above with glass fragments or beads. To avoid the evaporation of vapors that strongly irritate mucous membranes, the stopper is smeared with vaseline from above. For fixation, the stopper is opened with all precautions, and fresh, still moist smears are placed into the cylinder. After a few seconds, the material is already fixed. Further processing is alcohol of ascending concentration. Among the most usable mixtures are Hermann's fluid (4 cm³ of a 2% aqueous solution of osmic acid, 15 cm³ of a 1% aqueous solution of platinum chloride, and 1 cm³ of glacial acetic acid); Flemming's mixture: chromic acid 0.75 g, osmic acid 0.4 g, glacial acetic acid 5 cm³, water 95 cm³. In addition, a pure aqueous solution of osmic acid is used. Among mixtures with picric acid, one should note: Bouin's mixture—concentrated aqueous solution of picric acid 15 parts, formalin 5 parts, acetic acid 1 part; Boveri's mixture—concentrated aqueous solution of picric acid 100 cm³, water 200 cm³, glacial acetic acid 3 cm³. Mixtures with chromic acid, formalin, etc., are also used. One of the usable stains in the study of protozoa is Heidenhain's iron hematoxylin staining in various modifications. One of the frequently used variants is the following. Fixed smears from 70% alcohol are transferred to a mordant—a freshly prepared 2¹/2% aqueous solution of iron alum—for 12–24 hours. Water; Heidenhain's hematoxylin 12–24 hours. Differentiation under the control of a microscope in a 2¹/2% aqueous solution of iron alum. Prolonged (not less than 2 hours) washing under the tap with running water. Alcohols of ascending concentration, xylene, balsam. For additional staining, eosin, acid fuchsin, chromotrope, lichtgrün, etc., are used. The modification by Nöller is also recommended: fixation by Schaudinn or Bouin. After prolonged washing in 70% alcohol—mordanting in a fresh 4% aqueous solution of iron alum. Water. Heidenhain's hematoxylin for 60 minutes in a thermostat at 37°. Differentiation in a 2% alum solution with constant shaking. Duration of differentiation: for intestinal amoebae 3¹/₂–4¹/₂ min., for small flagellates 1–2 min., for medium-sized infusoria 5 min. According to Nöller, staining succeeds without microscopic control. In the history of the study of protozoa, especially the malaria parasite, a large role was played by the staining of Romanovsky (1893), who drew attention to the fact that the nucleus of the malaria parasite is stained red by a mixture of methylene blue and eosin. Nocht showed that from old solutions of methylene blue, a substance is extracted with chloroform which upon evaporation appears in the form of a red-violet powder and which also gives the "Romanovsky effect"; it was named by Nocht "Rot aus Methylenblau"; later it received the name azur (see). As a result of various improvements of the Romanovsky method, the Giemsa staining was obtained (see). Alcohol is excluded during the processing of smears, and the differentiation of smears is performed in mixtures of acetone and xylene (see Giemsa). Mounting in acid-free balsam. The rapid volatility of acetone makes it inconvenient when controlling differentiation under the microscope. This prompted Fleischer and Arndt to insert amyl alcohol into the differentiating series, which evaporates and differentiates more slower, making microscopic control possible.—Among other stains used in the study of protozoa are Delafield's hematoxylin, alizarin + toluidine blue, Twort's staining (neutral red and lichtgrün), and others. To reveal flagella and cilia, besides the usual methods with a mordant, examination in a dark field is used, as well as Burri's method and corresponding negative methods (Opalblau). At the edge of a coverslip, a drop of fluid containing protozoa is mixed with a drop of Chinese ink or 10% Opalblau, and a smear is made from the mixture. The dried smear is examined without a coverslip. The presence of various reserve substances in the protozoan cell is determined by usual microchemical reactions. Glycogen, besides the usual reaction with tincture of iodine, is determined by Best's stain: fixation with alcohol or according to Carnoy. Embedding in celloidin or celloidin-paraffin. 1) Intensive staining of the nucleus with hematoxylin. 2) Thorough washing. 3) Staining for 5 min. in the following mixture: Best's carmine solution 20.0 (filter), Liq. Ammonii caustici 30.0, methyl alcohol 30.0. 4) Differentiation without washing in the following mixture: methyl alcohol 40, absolute alcohol 80, distilled water 100 cm³ for 1–5 min., until the renewed fluid becomes transparent. 5) Washing with 80% alcohol: alcohols of ascending concentration. Balsam. Staining of protozoa in sections is performed according to the general rules of histological technique. Staining of spirochetes. For the majority of spirochetes, examination in a fixed and stained form is preceded by examination in a dark field, as well as the use of negative methods (Burri, Opalblau). Dry fixation and the usual thick drop with subsequent routine Giemsa staining are used mainly for blood parasite spirochetes (Leptospira obermeieri). For tissue spirochetes, prolonged staining according to Giemsa is recommended, and for Treponema pallida, in addition, formalin fixation, in which the number of treponemes significantly increases compared to control fixation with methyl alcohol. For spirochetes in smears, Fontana's method is recommended. A 5% tannin solution is heated for ¹/₂ min. until vapors appear. Water. Heating until vapors appear in a 5% silver nitrate solution for 20–30 seconds. SKA

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Water. Ammonia is added to a solution of acidic silver in a test tube until the precipitate that initially appears is dissolved. To avoid an excess of alkali, a solution of silver nitrate is added again until opalescence appears. For staining spirochetes in sections, their silver impregnation is used chiefly according to one of the modifications of the Levaditi method (see Levaditi method).