MORPHOLOGICAL AND TINCTORIAL PROPERTIES OF BACTERIA - E. L. Zaitseva - 2015

CHAPTER V. METHODS OF STAINING MICROORGANISMS

Studying Bacteria in a stained preparation not only reveals their Morphology but also provides insight into certain details of their chemical Structure. This is achieved using specialized stains. Microorganism Cells are stained primarily with aniline Dyes, which are divided into acidic and basic dyes. In acidic dyes, the color-bearing ion (the chromophore) is an anion, whereas in basic dyes, it is a cation.

Acidic dyes include eosin, acid fuchsin, erythrosin, and others. They bind intensively to cytoplasmic Cell components.

Basic dyes — methylene blue, basic fuchsin, gentian violet, crystal violet, and safranin — bind more actively to the nuclear Components of the cell. The high concentration of DNA and ribosomal RNA in bacterial cells makes them particularly sensitive to basic dyes. Consequently, only basic dyes are used in microbiological practice.

The intensity of a dye's staining capacity depends on the pH of the medium: basic dyes stain objects more intensely as the alkalinity of the medium increases, while acidic dyes do so in a more acidic environment. Staining Methods for microorganisms are classified into simple and complex (differential) techniques.

SIMPLE STAINING METHODS

Simple staining allows for the detection of microbes in the material under Microscopy, the determination of their quantity, and a rapid assessment of their morphological features. This is accomplished using basic and neutral aniline dyes. Simple staining employs only a single dye—most commonly red, such as a fuchsin solution (stained for 10–30 seconds); violet, such as gentian violet (stained for 1–2 minutes); or blue, such as methylene blue (stained for 3–5 minutes). Preparations for staining are prepared on Glass slides, the thickness of which should not exceed 1.1–1.4 mm.

When staining smears, dye solutions or the staining paper proposed by A. I. Sinev are used. To use the paper, a 2x4 cm piece of staining paper is placed over the dried and fixed preparation, and a few drops of Water are applied. The staining duration depends on the specific method. After staining is complete, the paper is removed with forceps, the smear is rinsed with tap water, air-dried, and examined under a Microscope.

Staining with diluted fuchsin.

Materials:

1) glass slide;

2) Ziehl's carbol fuchsin;

3) water;

4) bacteriological loop or sterile Pasteur pipette;

5) alcohol burner.

Staining technique:

Diluted (1:10) Ziehl's carbol fuchsin is applied to the prepared, flame-fixed, and cooled test slide (either directly or onto filter paper) for 10–30 seconds. The preparation is then rinsed with water, air-dried, and examined under a microscope.

Staining with methylene blue. A method for the metachromatic detection of volutin granules in corynebacteria and accumulations of nucleic compounds in bacteria.

Materials:

1) glass slide;

2) aqueous-alcoholic solution of methylene blue;

3) alcohol burner;

4) pipette or pre-stained filter paper;

5) bacteriological loop or sterile Pasteur pipette.

Staining technique:

A water-alcohol solution of methylene blue is pipetted onto the prepared, dried, and fixed smear for 3–5 minutes, after which the slide is rinsed with water, dried, and examined under a microscope.

Evaluation of results. The bacterial protoplasm stains light blue, while volutin granules stain dark blue.

Loeffler’s methylene blue staining (Loeffler’s method). Methylene blue is a basic dye, meaning its chromophore is a cation. Due to this property, the dye binds intensely to cellular nuclear components by forming a complex with the anions of bacterial DNA and ribosomal RNA. The presence of alkali in the solution enhances the interaction between cellular components (DNA and RNA) and the chromophore.

Materials:

1) glass slide;

2) water for rinsing smears;

3) staining tray and staining racks for smear preparations;

4) 96% ethanol for smear fixation;

5) spirit lamp;

6) bacteriological loop;

7) Loeffler’s methylene blue solution.

Staining technique:

1. Apply the test material onto clean, degreased glass slides, spread it into a thin, uniform layer across the glass surface, and air-dry.

2. Fix the preparation in 96% ethanol for 3 minutes and dry.

3. Apply a drop of 1% alkaline Loeffler’s methylene blue solution to the dried preparation and stain for 3–10 minutes.

3. Rinse by immersing in a beaker of tap water.

4. Air-dry and examine under a microscope using an immersion oil system.

Evaluation of results. When stained correctly, the microscopic preparation reveals blue bacteria against a colorless or faint blue Background.

COMPLEX STAINING METHODS

Specific dyes differing in Chemical Composition and color, mordants, alcohols, acids, etc., are applied sequentially to the preparation.

This approach allows for the identification of specific cell structures and the differentiation of one microorganism species from another.

Gram staining method. This method was developed in the 19th century by the Danish bacteriologist Hans Christian Gram (Gram, 1884). When introducing his method in 1884 for staining bacteria in tissue sections and organ impression smears, he failed to recognize its immense differential-diagnostic significance, although he did note that certain microorganisms, such as the typhoid bacillus, become decolorized and take on a counterstain. The differential diagnostic value of Gram staining for bacteria was first highlighted in 1886 by E. Roux.

Principle of the method. Susceptibility to Gram staining is determined by the thickness of The Cell wall and its chemical structure. The Cell wall of Gram-positive bacteria consists of a multilayered peptidoglycan layer 20–60 nm thick. Peptidoglycan microfibrils intertwine to form a dense network permeated with pores. Teichoic and lipoteichoic acids are covalently linked to the cell wall peptidoglycan and protrude onto the cell surface through the pores of the peptidoglycan framework.

In Gram-positive bacteria, the cell wall lacks aromatic and Sulfur-Containing Amino Acids and has a low lipid content; by contrast, Gram-negative bacteria contain these substances in large amounts. Furthermore, Gram-positive bacteria contain a magnesium salt of ribonucleic acid, which is absent in Gram-negative bacteria. This salt forms a stable chemical complex with protein, gentian violet, and iodine that resists destruction during brief exposure to alcohol. Such a complex does not form in Gram-negative bacteria, making them easily decolorized by alcohol. Fuchsin then counterstains the Gram-negative microorganisms red. Additionally, the pore STRUCTURE OF THE peptidoglycan in Gram-positive bacteria prevents the dye from washing out when the bacterial smear is treated with alcohol.

The cell wall of gram-negative bacteria contains 1-2 layers of peptidoglycan, and its thickness is much smaller (10-20 nm) than that of gram-positive microorganisms. The cell wall of gram-negative bacteria includes an outer membrane linked by a bimolecular lipid layer over the surface peptidoglycan layer. The outer membrane has a mosaic structure consisting of phospholipid, polysaccharide, and protein molecules. Outer Membrane Proteins—"porins"—surround hydrophilic pores through which water and other molecules with a mass of up to 800 Da pass.

When treated with alcohol, gram-negative bacteria, due to an insufficient amount of peptidoglycans in the cell

wall, lose the iodine-crystal violet complex, become decolorized, and then acquire a contrasting color from counterstains.

The cell component that enters into a ternary complex with the dye and mordant is magnesium ribonucleate. When a suspension of gram-positive microbial cells is treated with Bile salts, magnesium ribonucleate is extracted, and the cell becomes gram-negative. Adding the extracted magnesium ribonucleate back to such gram-negative cells restores their gram-positivity.

Peculiarities of preparing the smear for Gram staining. Nowadays, in The process of preparing smears for the Gram stain, all those methodological details that were previously considered important are usually ignored. Such neglect in many cases leads to indistinct or even erroneous results.

Bacterial concentration. To obtain reliable staining results, the smear must be thin. In thick smears where cells are clustered (in "dumps"), dye precipitation frequently occurs, and the decolorization of gram-negative organisms is more difficult. Conversely, gram-positive organisms in clustered areas decolorize significantly faster than single cells. Bacteria in the smear should be spaced as far apart as possible from one another.

Drying the smear. As mentioned above, a positive Gram reaction is associated with the presence of magnesium ribonucleate in the cell. The extraction of this compound deprives the cell of its gram-positivity. Saline solution is among the extractants capable of extracting Nucleic Acids and their salts from the cell. Preparing a bacterial suspension from solid-medium cultures in saline and prolonged drying can lead to the partial extraction of magnesium ribonucleate from the cells and their partial conversion into gram-negative forms. Prolonged storage of a bacterial suspension in saline or distilled water partially or completely converts gram-positive cells into gram-negative ones. Even prolonged drying on a glass slide of a drop of suspension prepared in saline can lead to a partial loss of resistance to decolorization. Therefore, a minimal amount of liquid culture or cell suspension from a solid culture should be applied to the slide with a loop and spread in as thin a layer as possible to maximize the drying speed of the smear. Rapid drying by heating over a flame is not recommended, as it promotes the extraction of magnesium ribonucleate from the cell.

Spreading the suspension on the slide. Trauma to gram-positive bacterial cells can lead to a loss of their ability to resist decolorization. Too vigorous rubbing of the suspension with a loop on the slide during smear preparation can lead to a partial change in their Gram-staining reaction.

Fixation. A smear for Gram staining should be fixed only over a burner/alcohol lamp flame. The Use of other fixation methods, in particular with methyl alcohol, an ethyl alcohol-ether mixture, etc., can produce a distorted picture. However, with heat fixation, the intensity of heating the smear and the intensity of decolorization of gram-positive bacteria are in direct proportion—the more the gram-positive cells are subjected to thermal exposure, the more gram-negative individuals are found among them. Therefore, fixation over a burner flame should be cautious and gentle.

The Staining procedure of the smear should be initiated only after it has completely cooled down. It is not recommended to pour the dye solution onto a smear that is still hot after fixation.

Factors influencing Gram stain results. By exposing the culture to various substances or cultivating it in media mixed with these substances, one can achieve The conversion of gram-negative microbes into gram-positive ones and vice versa.

In addition to creating such unusual living conditions, fluctuations in the Gram reaction can also occur under physiological conditions. Each bacterial species has a specific maximum culture age at which the characteristic Gram reaction of that species is most clearly expressed. Young cultures of gram-positive bacteria are more resistant to decolorization than old ones, although there are indications that 48-hour cultures are more resistant than 24-hour ones. In the most critical cases, especially when determining the Gram reaction of a newly described microbe, it is recommended to prepare Gram-stain smears from cultures of three ages: 6-8-hour, 12-24-hour, and 48-hour.

Even the Location On the surface of the Agar slant can affect Gram stain results. A smear prepared from the lawn scraped off the upper, already drying area of the agar may yield completely different results than one prepared from the lower, more moist part of the agar.

Materials:

1) glass microscope slide;

2) carbol-gentian violet or crystal violet solution (1 g of dye, 10 ml of 96% alcohol, 2 g of crystalline carbolic acid, 100 ml of distilled water, or 10 ml of 4% alcohol dye solution and 100 ml of 2% carbolic acid);

3) Lugol's solution (Gram's modification): 2 g of potassium iodide, 10 ml of distilled water, 1 g of iodine crystals. The mixture is left to stand for a day, after which 300 ml of distilled water is added;

4) 96% ethyl alcohol;

5) aqueous solution of fuchsin;

6) water for rinsing the smear;

7) bacteriological loop or sterile Pasteur pipette;

8) alcohol burner.

Staining technique:

1. The fixed smear is stained with carbol-gentian violet solution for 1-2 minutes.

2. The excess dye is drained off without rinsing the smear with water, and Lugol's solution is poured on for 1-2 minutes until the smear turns black.

3. Pour off the Lugol's solution and decolorize the stained smear with 96% alcohol (dip the slide into a beaker of alcohol several times until purple streaks stop coming off). Decolorization should take no longer than 20–30 seconds.

4. Rinse the slide with water.

5. Counterstain the smear with an aqueous fuchsin solution for 1–2 minutes.

6. Pour off the stain, rinse the slide with water, let it dry, and examine under an oil immersion microscope.

Interpretation of Results. Gram-positive bacteria stain dark violet or blue, while Gram-negative bacteria stain red or pink.

Gram staining as modified by A.I. Sinev. This method utilizes small pieces of filter paper impregnated with a 1% alcoholic solution of crystal violet and dried. To prepare them, a 1–2% solution of the dye (in 96% ethanol) is poured over sheets of filter paper laid out on a glass surface. Once dry, the stained paper is cut into 2x4 cm pieces and stored in dark glass jars with ground-glass stoppers. Filter paper prepared in this manner retains its staining properties for a very long time.

Staining technique:

1. Place a strip (or square) of filter paper impregnated with 1% alcoholic crystal violet solution onto the fixed smear.

2. Add a few drops of water and stain for 1–2 minutes.

3. Remove the filter paper strip.

4. Flood the smear with Lugol's solution for 1 minute (until the stain turns black).

5. Pour off the Lugol's solution.

6. Rinse the smear in 96% alcohol for 30 seconds to 1 minute (until the dye stops bleeding).

7. Rinse with water.

8. Counterstain the decolorized elements and cells with diluted Pfeiffer's fuchsin applied via a piece of filter paper moistened with water; press it against the smear and leave for 30 seconds to 1 minute.

9. After staining, thoroughly rinse the slide with water and dry it.

10. Examine using an oil immersion system.

Interpretation of results. Under the microscope, Gram-positive bacteria appear blue-violet, and Gram-negative bacteria appear pink-red.

Gram staining as modified by C.N. Atkins. (Atkins K.N., 1920). This method is used to enhance the staining of Gram-positive bacteria, although the dye composition differs slightly. Smears stained by the Atkins method are more resistant to decolorization because the mordant holds the gentian violet much more securely. This is particularly important for bacteria that are highly sensitive to decolorization (Streptococcus pneumoniae, Bacillus spp.).

Alcoholic gentian violet solution:

✵ Crystal gentian violet (85–90%) – 20 g;

✵ 96% ethanol – 200 ml;

✵ Ammonium oxalate [(NH4JC2O4] – 8 g;

✵ Distilled water – 800 ml.

Dissolve the crystal violet by grinding it in a mortar while adding alcohol, and dissolve the oxalate in water; mix both solutions and filter the prepared stain after 24 hours.

Iodine solution:

✵ Crystalline iodine (I2) – 20 g;

✵ Sodium hydroxide (1 N solution) – 100 mL;

✵ Distilled water – 900 mL.

Dissolve the iodine in the alkali and add water. Store at room Temperature in a dark glass container. Pure acetone is used for decolorization.

Safranin solution:

✵ Safranin O (2.5% solution in 95% ethyl alcohol) – 25 mL;

✵ Distilled water – 75 mL.

First, prepare a 2.5% alcoholic solution of safranin, then mix it with water in the specified proportion.

Materials:

1) glass slide;

2) alcoholic solution of gentian violet;

3) iodine solution;

4) pure acetone;

5) safranin solution;

6) water for rinsing the specimen;

7) bacteriological loop or sterile Pasteur pipette;

8) spirit lamp.

Staining technique:

1. Stain the fixed smear with an alcoholic solution of gentian violet for 1–2 minutes.

2. Pour off the excess stain without rinsing the specimen with water, and apply the iodine solution for 1–2 minutes until the specimen turns black.

3. Drain the iodine solution and decolorize the stained smear with pure acetone (dip the specimen into a beaker of acetone several times). Decolorization takes no more than 20–30 seconds.

4. Rinse the specimen with water.

5. Counterstain the smear with safranin solution for 1–2 minutes.

6. Pour off the stain, rinse the specimen with water, blot dry, and examine under an oil immersion microscope.

The Atkins method offers no advantages for staining Gram-negative microorganisms; moreover, the detection of such microorganisms can be hindered in certain specimens, such as Blood smears.

G.P. Kalina staining method. Professor G.P. Kalina developed a gentle staining procedure specifically tailored for Neisseria group microorganisms, as they tend to decolorize unevenly with alcohol during the Gram stain.

Materials:

1) glass slide;

2) G.P. Kalina reagent;

3) 0.5% brilliant green solution;

4) physiological sodium chloride solution;

5) water for rinsing the preparation;

6) 5-10% aqueous fuchsin solution;

7) 30% ethyl alcohol;

8) bacteriological loop or sterile Pasteur pipette;

9) spirit lamp.

Staining procedure:

1. Suspend the test culture in a small drop of physiological saline on a glass slide.

2. Mix the suspension with one drop of 0.5% alcoholic brilliant green solution and spread the smear evenly into a thin layer.

3. Air-dry and heat-fix over the flame of a spirit lamp.

4. Apply Kalina's reagent to the fixed smear for 1.5-2 min.

5. Pour off the stain, rinse with water, and then decolorize with 30% ethyl alcohol until the dye stops shedding.

6. Rinse again with water.

7. Counterstain the preparation with a 10% aqueous fuchsin solution for 2 min.

8. Rinse the smear with water, blot dry with filter paper, and examine under a microscope.

Interpretation of results. Under microscopy, Gram-positive bacteria appear green-black or violet, while Gram-negative bacteria appear pink-red.

Romanovsky-Giemsa staining method. A universal cytological staining technique used in light microscopy for Protozoa, bacteria, rickettsia, chlamydia, spirochetes, and various Cellular Structures and Tissues (including blood). It was proposed in 1904 by Gustav Giemsa (Giemsa).

Principle of the method. In the original version, the stain is named "Giemsasche Lözung für die Romanowsky färbung" (Giemsa's solution for Romanovsky staining). The Romanovsky-Giemsa stain consists of methylene blue, eosin, and azure, allowing it to stain microorganism elements and Blood Cells in distinct colors.

Materials:

1) glass slide;

2) distilled water;

3) methyl alcohol or 95% ethyl alcohol for smear fixation;

4) bacteriological loop or sterile Pasteur pipette;

5) spirit lamp;

6) standard Romanovsky-Giemsa stain solution.

Staining procedure:

1. Smears fixed in methyl alcohol for 3 min are air-dried and stained with a dye solution prepared immediately before use (add 10 drops of commercial Romanovsky-Giemsa stain to 10 ml of distilled water).

2. After an hour, the stain is drained, the slide is washed with water, air-dried, and examined under oil immersion.

Evaluation of results. Bacteria stain purple-red, Cell Cytoplasm stains light blue, and nuclei stain purple-red.

When staining protozoa, their cytoplasm turns blue, and nuclei turn reddish-purple.

Zdrodovsky staining method. A cytological method for staining rickettsiae proposed by Zdrodovsky.

Materials:

1) glass slide;

2) distilled water;

3) diluted Ziehl's fuchsin (add 10–15 drops of Ziehl's fuchsin to 10 ml of distilled water);

4) 0.5% citric acid solution or 0.01% Hydrochloric acid solution;

5) methylene blue;

6) bacteriological loop;

7) spirit lamp.

Staining procedure:

1. Stain the smear with diluted Ziehl's fuchsin (10–15 drops per 10 ml of distilled water) for 5 minutes.

2. Wash with water.

3. Treat the smear with 0.5% citric acid solution or 0.01% hydrochloric acid solution.

4. Wash with water.

5. Stain with methylene blue for 1 min.

6. Rinse with water and air-dry the smear.

7. Examine using oil immersion microscopy.

Results evaluation. Rickettsiae appear red, the cytoplasm of host cells is light blue, and nuclei are dark blue.

Rapid METHOD FOR DETERMINING the Gram type of microorganisms. This method is based on the lysis of Gram-negative bacterial cells in an alkaline environment (3% KOH solution) and the release of free DNA. The formation of mucus indicates that KOH disrupts the Introduction/37.html">Bacterial cell wall, liberating the DNA (viscous thread formation).

Materials:

1) glass slide;

2) 3% KOH solution;

3) bacteriological loop;

4) spirit lamp.

Staining procedure:

1. Place a drop of 3% KOH solution and one loopful of a 24-hour agar culture on a glass slide, and mix thoroughly.

2. After 5-10 seconds, slowly lift the bacteriological loop to a height of 2-3 cm.

Results evaluation. Within 5-7 seconds, Gram-negative cultures form a viscous thread 1-2 cm long when the loop is pulled upward; if no mucus forms, the test culture is Gram-positive.

STAINING ACID-FAST BACTERIA

Ziehl-Neelsen staining method. This technique for detecting acid-fast bacteria was first proposed in 1882–1883 by the German physicians Franz Ziehl and Friedrich Neelsen. It is designed to differentiate acid-fast bacteria (such as the causative agents of tuberculosis, leprosy, certain actinomycetes, and dormant endospores) from non-acid-fast microorganisms.

Principle of the method. The cell wall and cytoplasm of acid-fast bacteria contain a high concentration of Lipids, Waxes, and hydroxy acids with 50-100 carbon atoms, which render the cell wall impermeable to crystal violet and other common dyes. Consequently, these microorganisms resist staining with diluted dye solutions. To facilitate dye penetration into the cells, concentrated staining solutions are heated, detergents are added, and staining times are extended. Once the dye successfully penetrates the cell, it cannot be decolorized by standard acid or alcohol Solvents.

Materials:

1) glass slide;

2) Ziehl's carbol fuchsin;

3) 5% sulfuric acid solution (H2SO₄);

4) wash water;

5) aqueous methylene blue solution (Loeffler's methylene blue);

6) 95% ethanol;

7) bacteriological loop or sterile Pasteur pipette;

8) spirit lamp.

Staining technique:

1. Place a small piece of filter paper, not exceeding the size of the glass slide, onto the fixed smear.

2. Pour Ziehl's carbol fuchsin (primary stain) and gently heat the smear over a burner/spirit lamp for 3-5 minutes until vapors appear, then leave the stain until the preparation cools down slightly.

3. Remove the paper with fuchsin and rinse the slide with water.

4. Decolorize the stained preparation with a 5% sulfuric acid solution (differentiating agent) for 3-5 s, or with a mixture of 10 parts of alcohol and 1 part of hydrochloric acid for 1-2 min, or with 96% ethanol containing 3% hydrochloric acid (HCl) by volume, by repeatedly dipping the glass slide with the smear into a beaker of acid alcohol until the smear appears pale pink.

5. Thoroughly rinse the preparation with water.

6. Rinse it with 96% ethanol (this step is frequently omitted in laboratory practice).

7. Rinse again with water.

8. Counterstain for 3-5 min with an aqueous solution of Loeffler's methylene blue (secondary stain).

9. Rinse the preparation with water, dry, and examine under a microscope.

Interpretation of results. Acid-fast microorganisms stain ruby-red, while non-acid-fast ones stain blue.

It is convenient to use A.I. Sinev's modification: filter paper is impregnated with an alcoholic stain solution (2% alcoholic fuchsin solution), then dried, cut into strips matching the size of a glass slide, and stored for future use. Apply a few drops of distilled water to the fixed slide, place the pre-stained paper on top, heat until vapors rise, and then proceed as in the previous method starting from the end of the first step.

Ziehl-Neelsen staining of urine smears using Huze's modification. Along with acid-fast M. tuberculosis, urine may contain acid-fast smegma bacilli (M. smegmatis), which differ from the former by being susceptible to alcohol decolorization. Huze proposed using this feature to differentiate them. Huze's modification follows the standard Ziehl-Neelsen procedure, but after treating the smear with cold 5% H₂S0₄ and rinsing with water, an additional Treatment with ethanol (20 s) is applied.

Materials:

1) glass slide;

2) Ziehl's carbol fuchsin;

3) 5% sulfuric acid solution;

4) water for rinsing the preparation;

5) Loeffler's methylene blue solution;

6) 95% ethanol;

7) bacteriological loop or sterile Pasteur pipette;

8) spirit lamp.

Staining technique:

1. Stain the smear with Ziehl's carbol fuchsin (primary stain) with heating for 3-5 minutes, or using fuchsin-stained paper until vapors appear, without bringing the stain to a boil.

2. Cool the slide, remove the paper, drain off the stain, and rinse with water.

3. Decolorize with a 5% sulfuric acid solution (acting as a differentiating agent) for 1–2 minutes.

3. Drain off the residual acid, rinse the slide with water, and treat with ethanol for 20 s.

4. Counterstain for 3–5 minutes with Löffler's aqueous methylene blue solution (counterstain).

5. Rinse the slide with water, blot dry, and examine under a microscope.

Evaluation of results. M. tuberculosis are ethanol-resistant and do not decolorize, retaining a red color. M. smagmatis lose this stain upon exposure to alcohol, but turn blue when counterstained with methylene blue.

Bunge–Trautenroth staining. This method is used to stain urine sediment when differentiating between M. tuberculosis and M. smagmatis.

Materials:

1) microscope slide;

2) absolute alcohol;

3) carbol fuchsin;

4) 1N chromic acid (H₂CrO₄);

5) 10% sulfuric acid solution;

6) water for rinsing the slide;

7) saturated methylene blue solution;

8) bacteriological loop or sterile Pasteur pipette

9) spirit lamp.

Staining procedure:

1. The slide with the prepared smear is immersed in absolute alcohol for 3 hours to extract Fatty acids.

2. It is then immersed in chromic acid for 15 minutes.

3. Rinse the slide with water.

4. Stain with carbol fuchsin while heating.

5. Decolorize with 10% sulfuric acid for 3 minutes.

6. Counterstain for 5 minutes with a saturated alcoholic solution of methylene blue.

Evaluation of results. Acid-fast M. tuberculosis bacteria appear crimson, while less acid-fast M. smagmatis appear blue.

Semenovich-Martynovsky staining method. This staining technique is intended for acid- and alcohol-fast microorganisms.

Materials:

1) glass slide;

2) Ziehl's carbol fuchsin;

3) Loeffler's methylene blue solution;

4) water for washing the smear;

5) bacteriological loop or sterile Pasteur pipette;

6) alcohol burner.

Staining procedure:

1. The smear is stained with diluted (1:3) Ziehl's carbol fuchsin for 2 min.

2. The slide is washed with water.

3. It is stained with Loeffler's methylene blue solution for 3–5 min.

4. Wash with water, dry, and examine under a microscope.

Evaluation of results. Acid- and alcohol-fast microorganisms stain red, while all other microbes stain blue.

Baumgarten staining method. This method is used to differentiate leprosy bacilli (M. leprae) from M. tuberculosis.

Materials:

1) glass slide;

2) saturated alcoholic solution of fuchsin;

3) 95% ethyl alcohol;

4) distilled water;

5) hydrochloric acid (HCl);

6) diluted methylene blue;

7) water for washing the smear;

8) bacteriological loop or sterile Pasteur pipette;

9) alcohol burner.

Staining technique:

1. Stain the smear in a fuchsin solution (5 drops of saturated alcoholic fuchsin solution and 5 mL of distilled water) without heating for 5–7 min.

2. Then decolorize in a mixture of 10 parts alcohol and 1 part hydrochloric acid for 15 s.

3. Rinse the slide with water.

4. Counterstain with diluted methylene blue.

5. Rinse with water, air-dry, and examine under a microscope.

Result evaluation: M. leprae stain pinkish-red, while M. tuberculosis appear blue.

Much's staining method. This method is used to detect the granular form of tubercle bacilli (Much's granules).

Materials:

1) glass slide;

2) saturated alcoholic solution of methyl violet;

3) 2% aqueous solution of carbolic acid;

4) Lugol's solution;

5) 5% nitric acid solution;

6) 3% hydrochloric acid solution;

7) 95% ethanol solution;

8) acetone;

9) Pfeiffer's fuchsin;

10) water for rinsing the preparation;

11) bacteriological loop or sterile Pasteur pipette;

12) alcohol burner.

Staining technique:

1. Keep the smear for 24 h in a mixture of 10 mL of saturated alcoholic methyl violet solution and 100 mL of a 2% aqueous carbolic acid solution.

2. Next, treat with Lugol's solution for 1–2 min.

3. Drain off Lugol's solution and apply a 5% nitric acid solution to the smear for 1 min.

4. Next, apply a 3% hydrochloric acid solution for 10 s.

5. Drain the solution and immerse the slide in a mixture consisting of equal volumes of alcohol and acetone.

6. Rinse the specimen with water and counterstain with Pfeiffer's fuchsin.

7. Rinse the specimen with water, air dry, and examine under a microscope.

Results evaluation. Acid-fast bacteria stain purple.

Staining of granular forms of M. tuberculosis by Kozlov's method.

Materials:

1) microscope slide;

2) Kozlov's mixture;

3) Lugol's iodine solution;

4) 0.1% safranin solution;

5) water for rinsing the specimen;

6) bacteriological loop or sterile Pasteur pipette;

7) spirit lamp.

Staining procedure:

1. Immerse the fixed smear in a jar containing Kozlov's mixture for 45 min.

2. Rinse the smear with tap water.

3. Apply Lugol's solution to the specimen for 20 s.

4. Rinse with tap water for 2-3 s.

5. Counterstain with 0.1% safranin solution for 1-2 min.

6. Rinse the specimen with tap water, air dry, and examine under a microscope.

Results evaluation. Acid-fast bacteria stain purple, while other microorganisms appear raspberry-red. M. tuberculosis shows prominent granulation.

DETECTION OF THE MICROBIAL CELL WALL

Standard staining techniques do not allow visualization of the microbial cell wall. Gram or Ziehl-Neelsen staining does not differentiate the cell wall from the bacterial body. To achieve contrast, the smear requires a special pretreatment prior to staining. The specimen is prepared from a highly diluted microbial suspension so that there is sufficient free space between the bacterial cells.

Gutstein's staining method.

Materials:

1) glass slide;

2) Bouin's fluid;

3) 10% tannin solution;

4) water;

5) 0.02% hydro-alcoholic solution of crystal violet;

6) bacteriological loop;

7) alcohol burner.

Staining technique:

1. Air-dry the prepared smear.

2. Fix in Bouin's fluid for 3 min.

3. Mordant with a 10% (w/v) aqueous solution of tannin for 25 min.

4. Rinse with tap water.

5. Stain with a 0.02% hydro-alcoholic solution of crystal violet for 30–60 sec.

6. Drain off the excess stain and, without rinsing with water, air-dry the slide and examine under a microscope.

Evaluation of results. Cell walls appear as long, thin dark-violet or black filaments outlining the cell bodies, which feature white or light-blue cytoplasm.

DETECTION OF THE CELL ENVELOPE

Peshkov staining method.

Materials:

1) glass slide;

2) 25% NaCl solution;

3) 10% tannin solution;

4) water for rinsing smears;

5) Pfeiffer's fuchsin;

6) bacteriological loop;

7) spirit lamp.

Staining technique:

1. Add 1-2 loops of Yeast to 1-2 ml (hypertonic solution for water extraction from cells) - 30-40 minutes.

2. Prepare a smear, air-dry, and fix it over a spirit lamp flame or in alcohol-formalin.

3. Cover the fixed specimen with a 10% tannic acid solution (mordant).

4. Pour off the tannin and rinse carefully with water.

5. Stain the specimen with Pfeiffer's fuchsin.

6. Rinse the specimen thoroughly with water, air-dry, and examine under oil immersion.

Evaluation of results: bacteria are red.

BACTERIAL SPORE STAINING

Spore formation involves the Condensation and ISOLATION OF A specific region of the vegetative cell cytoplasm, followed by The Development of a round or oval body inside the bacterium. This body is encased in a thick, multi-layered coat rich in lipids, calcium, and dipicolinic acid. Because the dense spore coat is impermeable to water, it is extremely difficult to stain; consequently, standard staining techniques make spores appear as unstained vacuoles within the cell.

Once the spore fully matures, the vegetative portion of the cell may undergo lysis. Among pathogenic microbes, The ability to form spores is restricted to rod-shaped Gram-positive bacteria.

Spores are stained using specialized methods that involve preliminary heating of the spore along with the use of mordants (acids or alkalis). Mordants loosen the spore coat, facilitating the penetration of the dye. Unlike the vegetative body of the microbial cell, which is decolorized by acid treatment, stained spores exhibit acid-fastness.

Oerszkow's staining method. The Oerszkow method is similar to the Ziehl-Neelsen technique, but it employs a hydrochloric acid solution as a mordant to loosen the spore coat, which normally resists dyes. Following mordanting with hydrochloric acid and heating for 2-3 minutes, the smear is fixed and stained using the Ziehl-Neelsen procedure: the specimen is stained with a primary dye, decolorized with acid, and counterstained with a contrasting color. Spores strongly retain carbol fuchsin and stain red, while the bacterial cytoplasm is decolorized with 5% sulfuric acid and turns blue after counterstaining with methylene blue.

Materials:

1) glass slide;

2) 0.5% hydrochloric acid solution;

3) water for rinsing smears;

4) Ziehl's carbol fuchsin;

5) 5% sulfuric acid solution;

6) methylene blue;

7) bacteriological loop;

8) spirit lamp.

Staining technique:

1. Apply a few drops of 0.5% hydrochloric acid (HCl) solution to a dried, unfixed smear (prepared thick and positioned at the edge of the slide) and heat gently over a burner/spirit lamp flame for 1-2 minutes until boiling, then pour off the remaining acid.

2. The preparation (cooled) is washed with water, dried, and fixed over the flame of a burner/alcohol lamp.

3. The smear is stained with Ziehl's carbol fuchsin (primary stain) with heating until vapor appears.

4. Decolorize with a 5% sulfuric acid solution (differentiating agent) for a few seconds.

5. Wash with water.

6. Counterstain for 3-5 min with Loeffler's methylene blue (secondary stain), dry, and examine using oil immersion microscopy.

Evaluation of results. Bacterial spores stain red, the cytoplasm turns blue, and the vegetative bodies of the microbial cells become light blue.

Peshkov's staining method. M. A. Peshkov proposed staining spores with Loeffler's methylene blue, employing boiling to alter the resistance of the cell wall. Both spores and cytoplasm are stained upon heating. Washing the preparation with water leads to the decolorization of the cytoplasm, whereas the spore firmly retains the stain.

Materials:

1) microscope slide;

2) Loeffler's methylene blue;

3) water for washing smears;

4) 0.5% aqueous solution of neutral red;

5) bacteriological loop;

6) alcohol lamp.

Staining technique:

1. The prepared smear is fixed over the flame of an alcohol lamp.

2. Loeffler's methylene blue is applied to the fixed smear and heated to boiling over the flame of an alcohol burner for 20-30 seconds.

3. The cooled preparation is washed with water and counterstained with a 0.5% neutral red solution for 30-60 seconds.

4. Wash again with distilled water and air-dry.

5. Examine using oil immersion microscopy.

Evaluation of results. Spores stain light blue or dark blue. The protoplasm of the bacterial vegetative body is pink or red.

Schaeffer and Fulton staining method.

Materials:

1) microscope slide;

2) 5% aqueous solution of malachite green;

3) wash water for smears;

4) 0.5% safranin solution;

5) bacteriological loop;

6) spirit lamp.

Staining technique:

1. Apply a 5% aqueous solution of malachite green to the fixed smear and heat 3–4 times until vapors appear (3–6 min).

2. Rinse with water.

3. Counterstain with a 0.5% aqueous safranin solution for 30 s.

4. Rinse with water and air-dry the slide.

5. Examine under oil immersion.

Results evaluation. Bacterial spores stain green, and vegetative cells stain red.

Wirtz-Conklin staining method.

Materials:

1) microscope slide;

2) 5% aqueous solution of malachite green;

3) wash water for smears;

4) 0.5% aqueous safranin solution;

5) bacteriological loop;

6) spirit lamp.

Staining technique:

1. Fix the slide over a spirit lamp flame.

2. Stain the fixed smear with a 5% aqueous solution of malachite green, heating it 3–4 times until vapors appear (3–6 min).

3. Rinse the smear with water.

4. Counterstain the slide with a 0.5% aqueous safranin solution.

Results evaluation. The bacterial body stains red, and the spores stain green.

The negative staining method for spore detection is used in the quantitative assessment of sporulation by counting the number of spores and vegetative cells under various bacterial growth conditions.

Materials:

1) microscope slide;

2) methylene blue or fuchsin;

3) bacteriological loop;

4) spirit burner.

Staining procedure:

1. Prepare a thin smear of sporulating bacterial cells on a microscope slide, air dry, and heat-fix over a flame.

2. Apply methylene blue for 3–5 minutes or fuchsin for 1–3 minutes.

3. Gently air dry the preparation.

4. Examine under oil immersion.

Evaluation of results. The vegetative bacterial cells take up the stain, whereas the spores—possessing a multilayered, impermeable coat—do not. They appear as spherical or oval structures located either inside or outside the bacterial cells, depending on the stage of sporulation.

METHODS FOR DETECTING MICROBIAL CAPSULES

Certain species of bacteria produce a mucous substance that concentrates around the microbial cell body to form a capsule. The chemical composition of capsules varies among different bacteria; therefore, they cannot be visualized using a single universal staining method. Furthermore, capsules are easily deformed during staining, and capsular material binds dyes poorly, washing away readily during slide preparation. With conventional staining techniques, capsules remain colorless. This allows for the use of simple staining methods to identify encapsulated microorganisms in smears prepared from Organs and tissue fluids. Special staining techniques are employed to color the capsular substance.

Duguid's staining method. The simplest and most effective method for staining capsules.

Materials:

1) microscope slides and coverslips;

2) India ink;

3) filter/blotting paper;

4) bacteriological loop;

5) spirit burner.

Staining procedure:

1. Using a loop, place a drop of India ink on a clean microscope slide and mix it with the culture.

2. Place a coverslip over a portion of the resulting bacterial mixture.

3. Firmly press the coverslip against the slide using blotting paper until a thin brown layer of liquid appears between them. Remove any excess liquid.

4. Examine the preparation under high magnification using dry or oil immersion objectives.

Evaluation of results. Capsules appear as transparent zones around the microorganisms against a brownish-black background.

The Burri staining method is a negative staining technique: the background is stained while the microorganisms themselves remain unstained. This involves using dyes that do not penetrate bacteria, such as India ink. Alternatively, a 10% nigrosin solution, 20% collargol solution, or 3% Congo red solution can be used instead of India ink.

Materials:

1) microscope slide;

2) India ink;

3) bacteriological loop;

4) spirit lamp.

Staining procedure:

1. Place a drop of India ink in the center of a microscope slide.

2. Emulsify a loopful of broth culture or a small portion of a colony taken from a solid nutrient medium in the drop.

3. Spread the liquid evenly along the slide using the edge of a spreader slide.

4. Air-dry the smear and, without fixing, examine it microscopically using an immersion system.

Evaluation of results. The background of the slide is stained dark smoky grey, while the microbial cells and capsules do not take up the India ink and remain colorless.

The Burri-Gins staining method is used to stain encapsulated bacteria and is based on the principle that the capsule does not absorb dyes. The capsule is revealed by negative contrast of the background using the Burri technique.

Materials:

1) microscope slide;

2) India ink;

3) Nikiforov's mixture (ethyl alcohol, etc.);

4) water for washing smears;

5) Ziehl's carbol fuchsin;

6) bacteriological loop;

7) spirit lamp.

Staining procedure:

1. Mix the black India ink with the culture, prepare a smear in the manner of a blood smear, and allow it to dry.

2. Next, the specimen is chemically fixed using Nikiforov's mixture or other suitable fixatives.

3. Wash with water.

4. Stain the microbial cells with Ziehl's carbol fuchsin diluted 1:3 for 3–5 minutes.

5. Rinse the slide with water, air-dry, and examine under a microscope using an oil immersion system.

Evaluation of results. Colorless capsules are clearly visible against the dark background of the slide, enclosing red to bright crimson bacterial cells. Occasionally, small colorless zones—known as false capsules—may appear around stained bacteria that do not actually form capsules. These artifacts result from improper drying or smearing fixation.

Johne's staining method is a specialized technique for staining bacterial capsules.

Materials:

1) glass slide;

2) distilled water;

3) 2% aqueous solution of gentian violet or methyl violet;

4) 1–2% acetic acid;

5) bacteriological loop;

6) spirit lamp.

Staining procedure:

1. Apply a 2% aqueous solution of gentian violet (or methyl violet) with a pipette onto the prepared, dried, and fixed smear, and gently heat for 1–2 minutes, then rinse the slide with water.

2. Apply 1–2% acetic acid for 5–10 seconds, then rinse thoroughly with water.

3. Examine under a microscope directly in water.

Negative staining technique.

Materials:

1) microscope slides and coverslips;

2) India ink;

3) fuchsin;

4) bacteriological loop;

5) spirit lamp.

Staining procedure:

1. A small amount of cells from a solid medium is placed on a microscope slide in a drop of diluted fuchsin.

2. Mix with a drop of India ink.

3. Cover with a coverslip and observe using a 40x objective.

Evaluation of results. Against the dark background of the specimen, the colorless capsules surrounding the pink-stained microbial cells are clearly visible.

Methods for Staining BACTERIAL NUCLEOID

Staining of the bacterial nucleoid by the Feulgen nucleal reaction. A prominent place among various methods for staining nuclear material is held by the technique based on the specific reaction of sulfurous fuchsin with the aldehyde groups of deoxyribonucleic acid, proposed in 1924 by Feulgen and Rossenbeck.

Principle of the method. Deoxyribonucleic acid contains thymine among its nitrogenous bases and also incorporates four residues of the pentose sugar deoxyribose. Subjecting nuclear Nucleoproteins to mild acid Hydrolysis causes the Cleavage of purine and pyrimidine bases, thereby exposing deoxyribose molecules. During hydrolysis, deoxyribose is converted into β-hydroxylevulinic aldehyde, which reacts with the sulfurous component of colorless sulfurous fuchsin (Schiff's reagent), releasing free fuchsin and thus producing a specific reddish-purple coloration.

Materials:

1) microscope slide;

2) Carnoy's fluid;

3) 80% ethanol;

4) 1 N НСl;

5) sulfurous fuchsin (Schiff's reagent);

6) distilled water;

7) bacteriological loop;

8) spirit lamp.

Staining procedure:

1. Prepare a smear from a bacterial or yeast culture.

2. Air-dry and fix in Carnoy's fluid for 7 min.

3. Rinse with 80% ethanol.

4. Treat with 1 N НСl preheated to 60°C for 7 min, then transfer the slide to cold 1 N НСl for 1-2 min.

5. Treat with sulfurous fuchsin (Schiff's reagent) for 3-4 hours.

6. Wash with water, dry, and examine under oil immersion.

Evaluation of results. The nuclear material stains purple.

Staining of nuclear material using the Romanowsky-Giemsa method. This technique is primarily used for the microscopic examination of organ impression smears and blood smears. The Romanowsky-Giemsa stain consists of methylene blue, eosin, and azure, which allows it to stain different structures of microorganisms and blood cells in distinct colors. Before applying the stain to the specimen, it must be diluted 10-to 20-fold with water (pH 7.0-7.2). The optimal dilution varies depending on the dye batch and the pH of the water; therefore, it is recommended to test each new batch of stain and water using blood smears.

Materials:

1) glass microscope slide;

2) Carnoy's fluid or Nikiforov's fixative;

3) commercial Giemsa stain solution;

4) distilled water;

5) bacteriological loop;

6) spirit lamp.

Staining technique:

1. Air-dry the prepared slide.

2. Fix with Carnoy's fluid or Nikiforov's fixative.

3. Air-dry the fixed smears and stain them with working Giemsa solution (add 10 drops of commercial Giemsa stain to 10 ml of distilled water) by immersing the slide in a coplin jar containing the stain.

4. After 1 h, decant the stain and rinse the slide with distilled water.

5. Air-dry the finished slide and examine it under a microscope.

Evaluation of results. The protoplasm of protozoan tissue formed elements stains light blue, cell nuclei stain violet-red, and microbial cell bodies acquire a violet-red color.

Blood elements stain as follows: erythrocytes pink, leukocyte nuclei violet, cytoplasm blue, basophilic granules blue, eosinophilic granules red, and neutrophilic granules lilac.

Pikarsky method.

Materials:

1) glass microscope slide;

2) 1N HCl solution;

3) commercial Giemsa stain solution;

4) distilled water;

5) bacteriological loop;

6) spirit lamp.

Staining technique:

1. Treat the slide with 1N HCl solution while heating to 60°C for 7 min.

2. Rinse the slide with water and proceed with Romanowsky-Giemsa staining.

Interpretation of results. Nuclear elements appear dark red, and the cytoplasm is pink.

METHODS FOR STAINING BACTERIAL INCLUSIONS

Under specific growth conditions, the cytoplasm of many bacteria accumulates metabolic inclusion bodies, such as Glycogen and polysaccharide granules, poly-β-hydroxybutyric acid, lipid droplets, sulfur deposits, various crystals, and volutin (polyphosphate).

Staining of volutin granules. Volutin accumulates when nutrients are abundant in the environment, serving as an intracellular reserve for Nutrition and METABOLISM/26.html">Energy Metabolism. Volutin granules exhibit metachromasy and are readily visualized using specialized staining techniques. The term «volutin» was coined by Meyer, who first discovered it in Spirillum volutans. Volutin is a phosphorus- and nitrogen-containing reserve substance derived from nucleic

acids. A hallmark property of volutin is its metachromasy—the ability to stain a different color than the dye itself (for instance, methylene blue stains volutin violet-red).

1. Demonstration of volutin granules using methylene blue.

A method for the metachromatic detection of volutin granules in corynebacteria and nucleic acid aggregates in bacteria.

Materials:

1) microscope slide;

2) aqueous-alcoholic methylene blue solution;

3) water for rinsing smears;

4) bacteriological loop;

5) spirit lamp.

Staining procedure:

1. Apply 1–2 drops of the aqueous-alcoholic methylene blue solution to a fixed smear and stain for 3–5 minutes.

2. Pour off the stain, rinse with water, and air-dry.

3. Examine under oil immersion.

Interpretation of results. The smear shows pale blue cells with deep blue, almost black volutin granules (depending on the staining intensity).

2. Demonstration of volutin granules using acetic methyl violet.

Used to detect volutin granules in the Laboratory Diagnosis of diphtheria. Due to a high concentration of metaphosphates and other phosphorus compounds, volutin granules stain much more intensely than the cytoplasm when treated with acetic methyl violet.

Materials:

1) microscope slide;

2) acetic methyl violet solution (methyl violet, gentian violet, or crystal violet – 0.25 g, 5% acetic acid solution – 100 mL);

3) water for rinsing smears;

4) bacteriological loop;

5) spirit lamp.

Staining technique:

1. Apply 1-2 drops of acetic acid solution of methyl/gentian violet to a prepared fixed smear. Stain for 5-10 minutes.

2. Pour off the stain, rinse with water, and air dry.

3. Examine under an oil immersion microscope.

Results evaluation. In the smear, the cytoplasm of diphtheria corynebacteria stains light lilac, while volutin granules stain dark violet.

3. Neisser's staining method is used to detect volutin granules. The smear is stained with acetic methylene blue, during which a chemical reaction occurs between the dye and volutin. When rinsed with water, the cell body becomes decolorized and is subsequently counterstained with vesuvin.

Materials:

1) microscope slide;

2) Neisser's acetic methylene blue;

3) Lugol's iodine solution;

4) water for washing smears;

5) vesuvin (chrysoidine) solution;

6) bacteriological loop;

7) spirit lamp.

Staining technique:

1. Prepare a smear, air dry, and heat fix.

2. Flood the smear with 1-2 drops of Neisser's acetic methylene blue and stain for 1-2 minutes.

3. Drain off the stain, apply Lugol's solution for 1 min.

4. Rinse with water, blot dry with filter paper.

5. Apply vesuvin (chrysoidine) solution to stain the bacterial cell body for 1-3 min.

6. Wash off with water, dry, and examine microscopically.

Results evaluation. In the smear, the bacterial cytoplasm stains yellowish-brown, and volutin granules stain from blue-green to blue-black.

4. Raskina's staining method.

Materials:

1) microscope slide;

2) Ziehl's carbol fuchsin;

3) Raskina's stain;

4) water for washing smears;

5) bacteriological loop;

6) spirit lamp.

Staining procedure:

1. Pour Raskina's stain onto an air-dried, unfixed smear.

2. Gently heat the slide over a flame until the alcohol ignites.

3. Rinse with water, blot dry, and examine under oil immersion.

Results evaluation. Bacteria stain light red, while volutin granules appear black-blue (dark cherry).

5. Meyer's staining method.

Materials:

1) microscope slide;

2) Loeffler's methylene blue;

3) 1% sulfuric acid solution;

4) 40% aqueous KOH solution;

5) 0.25% solution of light green or chrysoidine;

6) water for rinsing preparations;

7) bacteriological loop;

8) spirit lamp.

Staining procedure:

1. Prepare two duplicate smears and fix them over a spirit lamp flame.

2. Stain the smears with Loeffler's methylene blue for 10 minutes.

3. Next, one slide is immersed for 5 min in a 1% sulfuric acid solution (slide No. 1), and the other for the same duration in a 40% aqueous KOH solution (slide No. 2).

4. Without washing, the slides are blotted with filter paper and counterstained with a 0.25% solution of light green or chrysoidine.

5. They are then rinsed with water and dried.

Evaluation of results. On slide No. 1, the bacterial cytoplasm is stained yellow-brown or green, while volutin inclusions appear cherry-red.

On slide No. 2, the volutin granules are decolorized and appear as empty, unstained spots against the background of faintly stained bacteria.

6. Omeliansky's staining method.

Materials:

1) glass slide;

2) Ziehl's fuchsin;

3) 1% sulfuric acid solution (H₂SO₄);

4) methylene blue (1:40);

5) water for washing the slides;

6) bacteriological loop;

7) alcohol burner.

Staining technique:

1. The fixed smear is stained with Ziehl's fuchsin solution for 30-60 sec.

2. The slide is washed with water and blotted with filter paper.

3. Apply a 1% H₂SO₄ solution to the slide for 20-30 seconds.

4. The slide is washed with water and blotted with filter paper.

5. Counterstain the smear with methylene blue for 15-30 seconds.

6. Rinse it with water and blot with filter paper.

7. Examine the slide under an oil immersion system with a x90 objective.

Evaluation of results. Volutin granules are red, and the cells are blue.

Glycogen staining. Glycogen is a carbohydrate and animal starch (polysaccharide). It frequently accumulates in the cells of Yeasts and bacilli. It is known that glycogen occurs in bacteria more often than starch. Reserve Polysaccharides are utilized by microorganisms as sources of carbon and energy.

Materials:

1) glass slides and cover slips;

2) iodine solution in potassium iodide:

✵ iodine — 7 g;

✵ potassium iodide — 20 g;

✵ distilled water — 100–300 ml;

3) filter paper;

4) bacteriological loop or sterile Pasteur pipette;

5) spirit lamp.

Staining procedure:

1. Place a small drop of the microorganism suspension on a clean glass slide, add an equal drop of iodine-potassium iodide solution (7 g of iodine and 20 g of potassium iodide in 100–300 ml of distilled water), and let stand for 2–3 minutes.

2. Cover with a cover slip and remove excess liquid using filter paper.

3. Examine the preparation under oil immersion (place a drop of oil onto the cover slip) using a x40 objective.

Interpretation of results. The cell cytoplasm stains light yellow, while glycogen granules stain reddish-brown.

Granulose staining. Granulose is a carbohydrate, a starch-like substance. It accumulates in large quantities within the cells of butyric acid bacteria (Clostridium acetobutyricum) prior to sporulation.

Materials:

1) glass slides and cover slips;

2) Lugol's iodine solution;

3) filter paper;

4) bacteriological loop or sterile Pasteur pipette;

5) spirit lamp.

Staining procedure:

1. Add a small drop of Lugol's solution to a small drop of the butyric acid bacteria suspension.

2. Cover with a cover slip and blot away excess liquid with filter paper.

3. Apply a drop of oil to the cover slip and examine using an immersion system.

Fat staining. Fats are present in the cells of virtually all microorganisms, accumulating in particularly large amounts as the culture ages.

Materials:

1) Sudan III (0.1 g of Sudan III dissolved in 200 ml of 90° alcohol);

2) 40% formalin solution;

3) methylene blue solution (1:40);

4) glass slides and coverslips;

5) bacteriological loop or sterile Pasteur pipette;

6) alcohol burner.

Staining technique:

1. Place a small drop of 40% formalin solution onto a glass slide.

2. Using the loop, introduce the microbial culture into the drop. Formalin kills the cell and loosens its cell wall.

3. After 5 minutes, add a small drop of methylene blue (1:40) to the same drop, and 10 minutes later, add a drop of Sudan III (a fat-soluble dye and indicator of lipid-like substances).

4. Cover the resulting combined drop with a coverslip and remove excess liquid using filter paper.

5. Examine the preparation under oil immersion.

Results evaluation. The cell cytoplasm stains blue, while fat inclusions stain pinkish-orange.

Staining of poly-β-hydroxybutyric acid. Poly-β-hydroxybutyric acid serves as a reserve lipid-like substance in many bacteria (e.g., members of the genus Pseudomonas). Poly-β-hydroxybutyric acid is an excellent source of carbon and energy.

Materials:

1) glass slide;

2) 0.3% solution of Sudan Black B prepared in Ethylene glycol;

3) xylene;

4) 0.5% aqueous safranin solution;

5) water for washing preparations;

6) bacteriological loop;

7) alcohol burner.

Staining technique:

1. Immerse a heat-fixed smear on a glass slide into a 0.3% solution of Sudan Black B prepared in ethylene glycol. Stain for 5–15 minutes (staining time is determined empirically).

2. Air-dry the preparation.

3. Immerse the microscope slide into xylene several times, then blot the preparation dry using absorbent material.

4. Stain the preparation with a 0.5% aqueous safranin solution for 5-10 s.

5. Wash with water, air-dry, and examine under a microscope.

Results evaluation. Poly-β-hydroxybutyric acid inclusions appear as blackish-blue droplets, while the bacterial cytoplasm stains pink.

Staining of polyphosphate granules.

Materials:

1) microscope slide;

2) Loeffler's methylene blue solution or 1% toluidine blue solution;

3) water for washing the smears;

4) bacteriological loop;

5) spirit lamp.

Staining procedure:

1. Heat-fix the smear on the microscope slide.

2. Apply Loeffler's methylene blue solution or 1% toluidine blue solution to the preparation for 10-30 s.

3. Rinse the preparation with water.

4. Air-dry and examine under a microscope.

Results evaluation. When stained with methylene blue, polyphosphate granules appear as blue or violet spheres, and the cytoplasm stains light blue. Toluidine blue stains metachromatically, with granules appearing red against a light-blue cytoplasmic background.

DETECTION OF FLAGELLA IN MOTILE MICROORGANISMS

Bacterial flagella are difficult to stain; therefore, mordants are used to increase their apparent thickness and facilitate staining. When preparing slides for flagella detection, it is essential to use exceptionally clean, grease-free glass slides. It is recommended to use new coverslips that have been pre-boiled for 10 min in chromic acid cleaning mixture. After boiling, the slides are washed for 5 min in a dilute sodium hydroxide (NаОН) solution, thoroughly rinsed with water, rinsed with alcohol, and stored in jars containing Nikiforov's mixture. The slides are removed from the jars using clean forceps, and the residual Nikiforov's mixture is removed by briefly flaming the glass. The procedure requires young microbial cultures (12-16-hour cultures) grown at the optimum temperature for the given species.

Morosov's staining method is designed for detecting Viruses via silver impregnation, as well as for revealing the Fine Structure of bacteria (including rickettsiae and spirochetes), flagella, argyrophilic and argentophilic inclusions, and granules.

Principle of the method. Treatment of the smears with tannin preparations loosens The structure of bacterial flagella, making them bulkier and more accessible to silver nitrate. Subsequently, treating the preparation with ammoniacal silver solution causes the salt to be reduced within the test object, selectively staining specific cellular structures in brownish-black.

Materials:

1) microscope slide;

2) distilled water;

3) precipitation tube;

4) 1% formalin solution;

5) bacteriological loop;

6) spirit lamp;

7) Reagents:

✵ Reagent No. 1: 1 ml of glacial acetic acid, 2 ml of formalin, 100 ml of distilled water.

✵ Reagent No. 2: 5 g of tannin, 1 ml of liquid carbolic acid, 100 ml of distilled water.

✵ Reagent No. 3: Dissolve 5 g of crystalline silver nitrate in 100 ml of distilled water, pour 20 ml into another vessel, and add ammonia solution dropwise to the remaining 80 ml of the silver solution until the precipitate formed dissolves and a slight opalescence remains. If too much ammonia is added, use the 20 ml of silver solution set aside earlier, adding it dropwise until the required faint opalescence is achieved.

To stain the specimen, dilute the silver solution with distilled water at a 1:10 ratio.

Staining procedure:

1. Using a flamed and cooled loop, gently Touch The surface of the colonies or lawn growth of the microbial culture to avoid mechanical damage to the flagella.

2. Transfer the collected material into a precipitation tube containing 0.1–0.2 ml of a 1% formalin solution at the bottom. Leave the loop undisturbed for a few minutes. During this time, some of the microbes will transfer from the loop into the solution. Repeat this procedure 2–3 times until the liquid shows a faint opalescence.

3. Place the prepared suspension in an incubator at 37°C for 1–2 hours to ensure an even DISTRIBUTION OF MICROORGANISMS in the liquid medium.

4. Next, add 1–2 drops of the formalin-fixed bacterial suspension to a tube containing 1.5–2 ml of distilled water. After 10–15 minutes, once the bacteria are relatively evenly distributed throughout the liquid volume, apply 5–6 drops of the resulting suspension onto a microscope slide without touching the slide with the loop.

5. Air-dry the drops without smearing them.

6. For better mordanting, treat the slides with Reagent No. 1 (glacial acetic acid with formalin) for 1 minute.

7. Pour off the excess mordant and wash the smear with water.

8. After the smear has partially dried, apply Reagent No. 2 to the specimen and gently heat over a low flame until vapors appear (1 minute).

9. Rinse thoroughly with water (1–2 minutes).

10. Apply Reagent No. 3 to the air-dried slide and let it act until the smear turns dark brown.

11. Rinse thoroughly with water.

12. Dry and examine under an oil immersion microscope.

Evaluation of results. The bacterial cell bodies stain brownish-black (coal-black), while the flagella acquire various shades of brown and are clearly visible against a pale yellow background.

Gray's staining method is based on the artificial enlargement of flagella through the application of a mordant.

Materials:

1) glass slide;

2) water for washing smears;

3) Ziehl's carbol fuchsin;

4) bacteriological loop;

5) spirit lamp;

6) reagents:

✵ Reagent No. 1: saturated aqueous solution of potassium alum - 5 ml, 20% aqueous solution of tannin - 2 ml, saturated aqueous solution of mercuric chloride - 2 ml.

✵ Reagent No. 2: saturated alcoholic solution of basic fuchsin - 0.4 ml.

The mordant, consisting of solutions No. 1 and No. 2, is prepared 24 hours before use.

Staining technique:

1. Pour the mordant over the preparation for 8-10 min.

2. Wash with water and stain with Ziehl's carbol fuchsin for 5 min.

3. Wash the smear with water.

4. Air dry and examine under a microscope.

Evaluation of results. Flagella stain red.

Schenk's staining method.

Materials:

1) glass slide;

2) water for washing smears;

3) Loeffler's methylene blue or 1% alcoholic safranin solution;

4) bacteriological loop;

5) spirit lamp;

6) reagents:

✵ Reagent No. 1: 30 ml of saturated aqueous tannin solution and 10 ml of 5% aqueous iron(III) chloride solution.

✵ Reagent No. 2: 1 ml of aniline and 4 ml of 95% ethyl alcohol.

Staining technique:

1. The mordant, consisting of solutions No. 1 (8 drops) and No. 2 (1 drop), is applied to the smear.

2. The mordant is drained off.

3. The smear is stained with Loeffler's methylene blue or a 1% alcoholic solution of safranin.

CHLAMYDIA STAINING

Materials:

1) microscope slide;

2) tap water;

3) methyl alcohol;

4) 96% ethyl alcohol;

5) 10% Lugol's iodine solution;

6) bacteriological loop;

7) spirit lamp.

Staining technique:

1. The smears are fixed for 10 minutes with methyl alcohol, followed by 20 minutes with 96% ethyl alcohol.

2. After fixation, a 10% Lugol's solution is applied to the specimen and stained for 5 minutes.

3. The stain is drained off, and the micropreparation is rinsed with water.

4. It is air-dried and examined under oil immersion (eyepiece х7, х10, objective х100).