MORPHOLOGICAL AND TINKTORIAL PROPERTIES OF BACTERIA - E. L. Zaytseva - 2015

CHAPTER III. PREPARATION OF STAINS AND MICROBIAL PREPARATIONS

Biological staining is used to clearly visualize the outlines of microorganisms. Special staining techniques make it possible to identify the Morphology and Structure of microbial Cells. Microbiology employs various Dyes, which are classified according to their ability to stain specific Structural components of microbial cells.

Groups of stains:

1. Basic (or nuclear) - selectively stain The Nucleus and basophilic structures of bacterial cells, presumably due to the acidic nature of nuclear material: neutral red, basic fuchsin, crystal violet, hematoxylin, malachite green, methylene green, vesuvin, chrysoidine, induline, etc.

2. Acidic (or cytoplasmic) - predominantly stain the Cytoplasm, and less frequently Cell walls: acid fuchsin, eosin, erythrosin, Congo red, picric acid, nigrosin (Indian ink), etc.

3. Neutral - selectively stain individual cytoplasmic components: Sudan III, Nile blue, etc.

4. Fluorochromes - a group of dyes capable of fluorescence under specific excitation wavelengths: auramine, coryphosphine, primuline, acridine orange, etc.

COMPOSITION AND PREPARATION OF STAINS FOR BACTERIAL SMEARS

Preparation of saturated alcoholic dye solutions. The required amounts of dye powders are placed into bottles, mixed with 96% ethanol, and incubated in an incubator at 37°C for 18–24 hours with intermittent shaking. During this period, a significant portion of the dye dissolves, leaving a small residue at the bottom of the bottle, which indicates that the solution is saturated. Saturated solutions can be stored for long periods in dark Glass bottles.

Preparation of aqueous-alcoholic dye solutions. Aqueous-alcoholic solutions are prepared from saturated alcoholic solutions. To do this, a saturated alcoholic solution of the dye is mixed in various proportions (depending on the dye) with distilled Water.

Crystal Violet

A. Crystal violet - saturated alcoholic solution.

Crystal violet - 10 g.

Ethanol 96% - 100 ml.

Crystal violet can be substituted with gentian violet or methyl violet.

B. Crystal violet - aqueous-alcoholic solution.

1 ml of saturated alcoholic solution is diluted with 10 ml of distilled water.

C. Crystal violet - carbol solution.

Crystal violet - 1 g.

Crystalline carbolic acid - 2 g.

Ethanol 96% - 10 ml.

Distilled water - 100 ml.

Carbol solutions of crystal, gentian, or methyl violet are prepared similarly to the first formulation of Ziehl's carbol fuchsin (described below), or by mixing 10 ml of saturated alcoholic dye solution with 100 ml of carbolic acid. The resulting mixture is thoroughly shaken and filtered through filter paper. Solutions of crystal, methyl, and gentian violet are unstable, and preparing them in large quantities for long-term storage is not recommended.

D. Crystal violet acetate.

Crystal violet (saturated solution) — 0.25 ml.

Acetic acid 5% — 100 ml.

The prepared mixture is thoroughly shaken and filtered through filter paper. A saturated solution of methyl violet can be used instead of crystal violet. The stain is unstable and should be stored for no more than a month in a dark glass bottle.

Fuchsin

A. Fuchsin — saturated alcoholic solution.

Basic fuchsin — 8–9 g.

Ethanol 96% — 100 ml.

B. Fuchsin — hydroalcoholic solution.

1 ml of saturated alcoholic fuchsin solution is diluted with 10 ml of distilled water.

C. Ziehl's carbol fuchsin (Ziehl, 1882).

Basic fuchsin — 1 g.

Ethanol 96% — 10 ml.

Crystalline carbolic acid — 5 g.

Glycerol — 0.5 ml.

Distilled water — 100 ml.

Option 1. Grind 1 g of basic crystal fuchsin in a porcelain mortar with 5 g of crystalline carbolic acid and a few drops of glycerol. While grinding, add alcohol in small portions. After the stain is thoroughly ground, add 100 ml of distilled water in small portions with constant stirring. Filter the staining solution through a wet filter paper. Ziehl's fuchsin is very stable and keeps quite well for a long time in a dark glass bottle with a ground-glass stopper.

Option 2. Add 90 ml of 5% carbolic acid to 10 ml of a saturated alcoholic solution of basic fuchsin (pour the carbolic acid into the stain, not the other way around). Shake the mixture vigorously for a few minutes, filter through filter paper, and pour into a storage bottle. In its pure form, this stain is used exclusively for staining acid-fast Bacteria and spores.

D. Pfeiffer's hydroalcoholic fuchsin (Pfeiffer's solution).

Ziehl's carbol fuchsin — 1 ml.

Distilled water — 9 ml.

Add 9 parts of distilled water to 1 part of Ziehl's carbol fuchsin and shake the solution vigorously for a few minutes. The solution is highly unstable and must be prepared in small quantities immediately before use. It should not be kept beyond the following day.

Methylene blue

A. Methylene blue — saturated alcoholic solution.

Methylene blue — 8–9 g.

Ethanol 96% — 100 ml.

B. Methylene blue – alcohol-aqueous solution.

1 ml of saturated alcoholic solution of methylene blue is diluted with 30 ml of distilled water.

C. Loeffler's methylene blue.

Saturated 8% alcoholic solution of methylene blue – 30 ml. Distilled water – 100 ml.

1% potassium hydroxide (KOH) solution – 1 ml.

The saturated 8% solution of methylene blue is filtered through filter paper, the potassium hydroxide solution is added, and the mixture is diluted with distilled water. The dye solution can be stored in a bottle for a long time without altering its properties. During storage, its staining capacity as well as its ability to produce metachromatic staining of nucleic compounds increase due to The formation of azures.

D. Neisser's blue.

Two stock solutions are prepared separately.

Solution No. 1:

Methylene blue – 0.1 g.

96% ethyl alcohol – 2 ml.

Glacial acetic acid – 5 ml.

Distilled water – 100 ml.

Solution No. 2:

Crystal violet – 1 g.

96% ethyl alcohol – 10 ml.

Distilled water – 300 ml.

Immediately before staining volutin granules, mix 2 PARTS OF THE first solution with 1 part of the second solution.

Toluidine blue acetate

Toluidine blue – 0.25 – 0.5 g.

Glacial acetic acid – 2 ml.

96% ethyl alcohol – 5 ml.

Distilled water – up to 100 ml.

Chrysoidine – aqueous solution

Chrysoidine powder – 1 g.

Distilled water - 150 ml.

The weighed amount of dye is placed in a funnel lined with filter paper, followed by The addition of boiling water.

Vesuvin (Brown Stain)

Vesuvin powder - 2 g.

Ethyl alcohol 96% - 60 ml.

Distilled water - 40 ml.

Mix all ingredients, heat to a boil over low heat, and after cooling, filter through paper.

Safranin

Safranin - 2.0 g.

Distilled water - 100 ml.

Weigh the specified amount of safranin, transfer it to a funnel with filter paper, and pour boiling water over it.

Alcoholic Solution of Brilliant Green

Brilliant green - 0.5 g.

Ethyl alcohol 96% - 100 ml.

The stain should be stored for extended periods in a dark glass bottle in a dark place, sealed with a ground-glass or rubber stopper.

Romanowsky-Giemsa Stain

Azure I - 3.772 g.

Eosin - 2.165 g.

Methylene blue (medicinal grade) - 1.563 g.

Methanol (analytical grade) - 750 ml.

Glycerol (analytical grade) - 256 ml.

The Romanowsky-Giemsa staining mixture, based on Romanowsky-Wright stain, is dissolved in powder form (commercial dye) in a mixture of equal volumes of methyl alcohol and glycerol (800 mg of dye per 100 ml of solvent). Since the dye dissolves poorly, it is best to first triturate it with the solvent at a rate of 300 mg per 100 ml, and then gradually add the remaining dye while stirring until the desired concentration is reached.

Preparing the stain can sometimes take several days. It is essential to use chemically pure methyl alcohol and glycerol as Solvents, as impurities degrade the staining properties. Methyl alcohol can be substituted with 95% ethyl alcohol. Store the prepared staining mixture in a dry, cool place in a tightly sealed container.

The Romanowsky-Giemsa stain is a mixture of azure (an organic dye derived from methylene blue), eosin, and methylene blue. It is commercially available ready-made. Immediately before use, add 10 drops of the commercial Romanowsky-Giemsa stain to 10 ml of distilled water (pH 7.0–7.2).

India Ink for Burri Staining Technique

Black mascara - 5 ml.

Distilled water - 15 ml.

Add 3 parts of distilled water to 1 part of black mascara. Pour the diluted mascara into a centrifuge tube and centrifuge for 15-20 minutes at 2000-3000 rpm to sediment large mascara particles. Decant the supernatant layer of mascara and sterilize in an autoclave at 0.5 atm for 30 minutes. Instead of sterilization, a few drops of formalin can be added to the mascara. The mascara prepared in this way is kept for two weeks until all large particles settle to the bottom. To prepare the specimen, carefully take only the upper layer of the settled liquid.

G. P. Kalina Reagent

0.5% potassium iodide solution - 96 ml.

5% alcoholic solution of basic fuchsin - 2 ml.

5% alcoholic iodine solution - 2 ml.

The 0.5% potassium iodide solution is prepared by heating in a water bath. After the potassium iodide sample is completely dissolved in 96% ethanol, the latter is combined with the fuchsin and iodine solutions. The mixture is stored in a dark glass bottle with a ground-glass stopper in a refrigerator.

Lugol's Solution

Crystalline iodine - 1 g.

Potassium iodide - 2 g.

Distilled water - 300 ml.

Dissolve the potassium iodide in 5-10 ml of distilled water, then add the crystalline iodine, leave for several hours until it is completely dissolved, and then add the remaining amount of water.

Kozlov Stain (for detecting granular forms of tuberculosis mycobacteria)

Gentian violet - 1 g.

Crystalline phenol - 6 g.

Glycerin - 40 ml.

Distilled water - 100 ml.

Grind the gentian violet and phenol in a mortar, gradually adding glycerin and distilled water. Before use, dilute the stock solution with 3 parts of distilled water and filter.

Raskina Stain for Staining Volutin Granules

Ziehl's carbol fuchsin - 4 ml.

Saturated methylene blue - 4 ml.

Glacial acetic acid - 5 ml.

96% ethyl alcohol - 95 ml.

Distilled water - up to 200 ml.

DYE SOLUTIONS FOR PREPARING STAINING PAPERS

Crystal violet or methyl violet for Gram staining

Crystal violet - 1 g.

Ethyl alcohol 96% - 100 ml.

Glycerin - 5 ml.

Basic fuchsin

Fuchsin - 2 g.

Ethyl alcohol 96% - 100 ml.

Glycerin - 1 ml.

Ziehl's carbol fuchsin

Add 1-2 ml of glycerin to 100 ml of the prepared stain.

Methylene blue

Methylene blue - 1 g.

Ethyl alcohol 96% - 100 ml.

Glycerin - 1 ml.

PREPARATION OF SMEARS FOR STAINING

To prepare a smear from microorganisms grown in a liquid medium, take the test material from the tube using a bacteriological loop.

Pay attention to proper hand positioning: hold the tube containing the Microbial growth in meat peptone broth (MPB) with your left hand, and the bacteriological loop with your right hand.

Materials:

1) Microscope slide;

2) test tube with a bacterial culture grown in a liquid nutrient medium;

3) bacteriological loop or sterile Pasteur pipette;

4) alcohol burner.

Preparation technique:

1. Sterilize the loop in the flame of the alcohol burner until red-hot.

2. Remove the cotton plug from the test tube with the right hand (using the little finger), and flame the rim of the tube over a spirit lamp.

3. Insert the inoculation loop into the test tube containing nutrient broth, cool the loop against the inner glass surface, and then pick up 1 drop of the culture.

4. Flame the rim of the test tube and the plug, close the tube, and place it in the test tube rack (while holding the loop with the microbial material in the right hand).

5. Take a grease-free glass slide—flamed and cooled—between the thumb and index finger of your left hand.

6. Using the bacteriological loop, apply the material onto the slide, spreading it evenly in circular motions to form a circle 8-10 mm in diameter.

7. Sterilize the loop in the flame of the spirit lamp until it glows red.

8. Air-dry the smear and then fix it by flaming or gentle heating, passing the slide through the spirit lamp flame three times, and allow it to cool.

Smears should contain sufficient material for examination without being too thick.

Preparation of a smear from a microbial culture grown on a solid medium (meat peptone Agar - MPA).

Materials:

1) microscope glass slide;

2) distilled water or isotonic sodium chloride solution;

3) a test tube or Petri dish with a bacterial culture grown on a solid nutrient medium;

4) bacteriological loop;

5) spirit lamp.

Technique:

1. Place a drop of water or saline solution onto a degreased glass slide.

2. Carefully pick up some material from The surface of the medium using the bacteriological loop (ensure the test tube with the culture is positioned correctly, with the slanted surface facing upward).

3. Flame the rim of the test tube, plug it, place it in the rack, and retain the bacteriological loop with the culture in your hand.

4. Transfer the material from the loop into the drop of water on the slide, spread it evenly, and outline the smear area on the underside of the slide with an oval or square.

5. Air-dry the preparation.

6. Burn off any excess microbial material remaining on the loop in the spirit lamp flame.

7. Fix the smear by flaming—passing the slide through the spirit lamp flame three times—and allow it to cool.

Preparation of a smear from clinical material. Preparation of a Blood smear. Place a drop of blood near one end of a thoroughly degreased glass slide by touching the narrow edge of the slide directly to a finger puncture. Place a second spreader slide—which must be narrower than the glass slide—at a 45° angle to the first slide and bring it into contact with the drop of blood. Once the blood spreads along the edge of the spreader slide, make a smooth, sweeping motion from right to left, distributing the blood in an even, thin layer across the surface. A correctly prepared smear has a light pink color and a uniform thickness throughout.

Preparation of a "thick drop" smear. Place a drop of blood in the center of a glass slide using a Pasteur pipette, or Touch the slide directly to the drop of blood emerging from the finger. Spread the blood on the slide with a bacteriological loop to a diameter of 8-10 mm. Leave the slide in a horizontal position until the blood dries. The blood in a "thick drop" is distributed unevenly, forming ragged edges.

Preparation of a smear from liquid pathological material (urine, CEREBROSPINAL FLUID). A small drop of the test fluid is placed onto a glass slide and spread in circular motions using a bacteriological loop to form a uniform round layer 8-10 mm in diameter.

Preparation of a smear from viscous material (sputum, pus). Sputum or pus is transferred onto a glass slide near its narrow edge and covered with another slide. The slides are gently pressed together to spread the material between them. Then, the free ends of the slides are gripped with the thumb and index fingers of both hands and pulled apart horizontally in opposite directions, ensuring that the slides remain tightly pressed against each other during the motion.

Preparation of a smear from Organs and Tissues. When examining an organ with a soft consistency, its surface is cauterized with heated forceps tips (for sterilization), an incision is made at that site, and a small piece of tissue is excised from the depth using pointed scissors. This piece is placed between two glass slides, and the subsequent Procedure is identical to that for preparing a smear from pus or sputum.

If the tissue of the organ is dense, a scraping is taken from the depth of the incision using a scalpel. The resulting material is spread in a thin layer over the surface of a slide using a scalpel or a bacteriological loop.

To prepare an impression smear (touch imprint): a small piece of tissue excised from the center of an organ is grasped with forceps, and its cut surface is pressed against a glass slide several times in succession (forming a series of impression smears).

Preparation of a smear from native cerebrospinal fluid. Native CSF is centrifuged at 3000 rpm for 5-10 minutes. A drop of CSF is collected from the sediment and placed onto a degreased glass slide. The smear is air-dried at room Temperature or in an incubator at 37°C.

FIXATION OF SMEARS

Fixation is necessary to inactivate bacteria, attach (fix) them to the slide, prevent cellular autolysis, and enhance stain uptake.

Methods of smear fixation are divided into Physical and Chemical. The physical method is based on The Effect of high temperature on the microbial cell, whereas the chemical method relies on chemical agents that cause coagulation of cytoplasmic Proteins.

The most common method of bacterial fixation is heat fixation. This is performed by passing the slide with the smear side up through the flame of an alcohol lamp or gas burner three times. The slide should remain in the flame for no more than 3 seconds. To check whether the fixation is adequate, the non-smear side of the glass slide is applied to the back of the left hand. When properly fixed, the glass should feel hot without causing a burn sensation. Afterward, the bacterial smears are stained using various methods. This method is suitable when working with pure microbial cultures. While heat fixation satisfactorily preserves the overall bacterial shape, cell dimensions are less well-preserved due to cell shrinkage. As for cellular inclusions and other structures, heat fixation is suitable for volutin granules and endogenous spores (for rough examination). Heat fixation must not be used for flagella, protoplasm, nuclei, blood smears, or organ imprints.

Chemical fixation is used to reveal and differentiate structural elements such as flagella, protoplasm, nucleoids, and cell walls, as well as when working with mixed material (blood, organ imprints, mucosal scrapings, pus, sputum).

The following agents are used as fixatives:

✵ methyl alcohol (fix for 5 minutes);

✵ 96% ethyl alcohol (fix for 10-20 minutes);

✵ Nikiforov’s mixture: equal volumes of alcohol and ether (fix for 10-20 minutes);

✵ alcohol-formalin: a mixture of 5 ml of undiluted formalin and 95 ml of 96% ethyl alcohol (fix for 15 minutes);

✵ Bouin’s fluid (Bouin, 1898): undiluted formalin 10 ml, glacial acetic acid 2 ml, saturated aqueous picric acid solution 30 ml (fix for 5-10 minutes);

✵ acetone (fix for 5 minutes);

✵ Carnoy’s fluid (Carnoy) — glacial acetic acid 10 ml, chloroform 30 ml, and 96% alcohol 60 ml (fix for 10-15 minutes). Carnoy’s fluid is an excellent fixative, particularly for studying the Fine Structure of bacteria. Carnoy’s fluid must not be used for fixing blood smears and internal organ imprints, as it causes complete hemolysis of erythrocytes.

There are two methods of chemical smear fixation:

1. The glass slide with the dried smear is immersed in a jar containing the fixative, air-dried, and examined under a microscope.

2. Drops of the fixative are applied to horizontally placed glass slides with smears and allowed to dry.

The fixation time depends on the specific fixative agents used.