Periodic acid-Schiff (PAS) is a staining technique for demonstrating the carbohydrates and fungal cell wall components. PAS can detect the presence of glycogen, polysaccharides, and mucin in the tissue that is either formalin-fixed, paraffin-embedded, or frozen tissue sections. PAS is performed in the laboratory for histological studies.

Principle of Periodic acid- Schiff (PAS) Staining

The periodic acid of the stain reacts with carbohydrates in an oxidative process. In this process, the polysaccharide and the periodic acid reaction form an oxidized compound- aldehyde. Now, the aldehyde reacts with the Schiff reagent, which gives the purple-magenta color. Similarly, the appearance of the pink color suggests the presence of intracellular or extracellular mucin. In contrast, using hematoxylin or methyl green as counter-stain helps in staining the nuclei. Likewise, a light green colored counter stain is preferred to demonstrate the fungal organisms.

Solutions and Reagents

0.5% Periodic Acid Solution

• Periodic acid crystals-0.5 g
• Distilled water-100 ml

To prepare 0.5% periodic acid solution mix 0.5 g periodic acid crystals in 100 ml distilled water.

Schiff’s reagent

Dissolve 5 g basic fuchsin in 900 ml boiled water. Once it is cools to 50°C  add 100 ml 1M HCl to the mixture. Again add 10 g of K₂S₂O₅ once the mixture cools down to 25°C. After completely mixing, shake the solution for 3 minutes and let it incubate for 24 hours in a dark room.

After incubation, add 5 g of activated charcoal to the mixture. Then shake the solution for 3 minutes and filter. The refiltration and retreatment of the solution are necessary if the solution is not crystal clear. Store the solution at 4°C in the foil-covered bottle. It is suitable to use the solution for 2-3 weeks if stored properly.

For testing the purity of the prepared Schiff’s reagent:

Pour 10 ml of 10% formalin into a beaker, then add a few drops of the prepared Schiff’s reagent. Then change in color is interpreted as 

• The red-purple color means it is a good Schiff’s reagent.
• Deep blue-purple color means poor Schiff’s reagent (delayed reaction).

Mayer’s Hematoxylin

• Aluminum potassium sulfate (alum)- 50 gram
• Distilled water- 1000 ml
• Hematoxylin- 1 gram
• Sodium iodate- 0.2 gram
• Glacial acetic acid- 20 ml

Dissolve alum in distilled water. Add hematoxylin when alum is completely dissolved. When hematoxylin is completely dissolved, add sodium iodate and acetic acid. Then boil and cool it.

Procedure for Periodic Acid-Schiff (PAS) Staining

1. Firstly remove the paraffin from the tissue sections by washing in distilled water.
2. Then, place the tissue in 0.5% periodic acid solution for 5 minutes. It oxidizes the tissue.
3. After that, rinse the tissue properly in distilled water.
4. Then, cover it with Schiff’s reagent for 5-15 minutes which turns into light pink.
5. After that, wash the stain for 5 minutes using lukewarm water, which turns it into dark pink.
6. Then counter-stain the tissue using Mayer’s Hematoxylin for 1 minute.
7. After that, wash it with running tap water for 5 minutes and rinse using distilled water.
8. Finally, dehydrate the slide, place the coverslip and mount it using synthetic mounting media.

Results interpretation for Periodic acid-Schiff (PAS) Staining

1. Glycogen, mucin and some basement membranes-Red/ purple
2. Fungi-Red/ purple
3. Background-Blue

Applications of Periodic Acid-Schiff (PAS) Staining

• The periodic acid-Schiff (PAS) staining is used to detect glycogen deposits in the liver. This test is useful when a person is suspected of the glycogen storage disease.

Glycogen storage disease (GSD) is a genetic disease in which the body cannot produce enzymes to break the complex sugar glycogen into simpler forms. It affects the different parts of the body like the liver, muscles etc.

• It demonstrates the glycogen granules in the bladder, kidney, ovary, pancreas, and lung tumors.
• It is used to visualize the basement membrane present in the various tissues in the body. Likewise, PAS demonstrates the thickness of the glomerular basement membrane for detecting abnormality in renal tissues.
• PAS stain is used in the diagnosis of glandular carcinomas (adenocarcinoma).
• PAS identifies Candida albicans, Aspergillus fumigatus, and Cryptococcus neoformans infections in the tissue samples.
• PAS staining detects the neutral mucins in the gastrointestinal tract and some epithelial mucins. 
• PAS can be used to study the amorphous or granular globules of the pulmonary alveolar proteinosis.

Pulmonary alveolar proteinosis is a rare disorder which causes the air sacs in the lungs to become clogged with surfactant.

• PAS is used to study the skin’s eosinophilic globoid bodies or Kamino bodies.

References

1. (IHC World, 2011)IHC World. (2011). PAS (Periodic Acid Schiff) Staining Protocol. IHC World. http://www.ihcworld.com/_protocols/special_stains/pas.htm
2. Stain, Periodic Acid Schiff Test – Test Results, Normal Range, Cost And More. Lybrate. (2022). Retrieved 21 June 2022, from https://www.lybrate.com/lab-test/stain-periodic-acid-schiff.
3. (2022). Retrieved 21 June 2022, from https://www.labce.com/spg949466_periodic_acid_schiff_pas_diagnostic_applications.aspx. 

Giemsa stain is a type of Romanowsky stain named after Gustav Giemsa, a German chemist who created a dye solution. It was primarily designed for the demonstration of malarial parasites in blood smears, but it is also employed in histology for routine examination of blood smears.

Principle of Giemsa Stain

Giemsa stain is a differential stain and contains a mixture of azure, methylene blue, and eosin dye. It is specific for the phosphate groups of DNA and attaches itself to where there are high amounts of adenine-thymine bonding.

Azure and eosin are acidic dye that variably stains the basic components of the cells like the cytoplasm, granules, etc.

Methylene blue acts as the basic dye, which stains the acidic components, especially the nucleus of the cell.

Methanol act as a fixative as well as a cellular stain. The fixative does not allow a further change in the cells and makes them adhere to the glass slide.

Preparation of Giemsa Stain

Giemsa is the most commonly used stain for staining blood films for malaria diagnosis. It is available commercially as a ready-to-use product, but the quality varies according to the source. By following simple rules, laboratories can prepare a stock solution of Giemsa stain using Giemsa stain powder, thus ensuring the use of consistent, high-quality stain.

Composition

The essential ingredients of Giemsa stain are the same; however, dilutions can be made depending on their use.

Ingredients	Gm/L
Giemsa powder	7.6
Glycerol	500 ml
Methanol	500 ml

Supplies, Materials, and equipment  

1. Giemsa powder or stain, 7.6 g (preferably Biological Stain Commission grade, to ensure a very good product of standard quality;
2. absolute methanol, pure, high-grade, acetone-free, 500 mL;
3. glycerol, high-grade, pure, 500 mL;
4. methanol-cleaned solid glass beads, 3-5 mm in diameter, 50-100 pieces;
5. a spatula or measuring spoon;
6. weighing paper;
7. a graduated cylinder;
8. a glass or plastic funnel;
9. a screw-capped, dark or amber glass bottle, clean and dry, 500-ml capacity (If not available, a chemically clean, dry, clear hard glass or polyethylene bottle of suitable size may be used, but should be wrapped in dark paper);
10. an analytical balance capable of weighing to 0.01 g; and
11. a shaker, if available.

Note:

• The person preparing the Giemsa stain should follow universal precautions, including the use of relevant personal protective equipment (PPEs) such as gloves, safety glasses, and a laboratory gown.
• Avoid contact and inhalation of methanol and Giemsa stain. Methanol and Giemsa stain are inflammable and highly toxic if inhaled or swallowed. Keep both chemicals in a locked cabinet or cupboard when they are not in use.

Preparation of Giemsa Stock Solution

Working Solution of Giemsa Stain

Working solution of Giemsa stain should be freshly prepared from Giemsa stock solution. Depending upon the method of staining used to stain malaria blood films, the Giemsa working solution is either 10% (for the rapid method) or 3% (for the slow method).

A rapid method is used in outpatient clinics and busy laboratories where a quick diagnosis is essential for patient management, whereas a slow method is used for staining a large number of slides collected during epidemiological or field.

Rapid (10% working solution) method

1. Commonest method for staining 1-15 slides at a time.
2. Used in outpatient clinics and busy laboratories
3. Efficient method but costly (as more stain is consumed)

Slow (3% working solution) method

1. Used for staining a larger number of slides (>20)
2. Ideal for staining blood films collected during cross-sectional or epidemiological surveys, field research, or for preparing batches of slides for teaching
3. Time-consuming method, so less appropriate when a quick result is needed
4. Less expensive compared to the rapid method as it requires much less stain.

Materials and Supplies

• Giemsa stain, transferred and filtered from the stock solution into a 25-or 50-ml bottle;
• buffered water, pH 7.2;
• a beaker or tube, clean, 5-10-ml capacity;
• a Pasteur pipette and
• Whatman filter paper, grade #1.

Preparation of Giemsa Working Solution

Prepare either 10% or 3% Giemsa working solution, depending on your need. About 3 mL of stain is required for each slide with a blood film.

1. Place 90 mL of prepared buffered water, pH 7.2, into a clean beaker or tube.
2. Filter the Giemsa stock solution through paper Whatman #1 and transfer it to a 25 to 50 mL container.
3. Add 10 mL of Giemsa stock solution using a clean, dry pipette. Do not take the aliquot from the large bottle containing the Giemsa stock solution to avoid contaminating it.
4. Prepare the Giemsa working solution just before staining the blood film(s), and use it within 15 minutes of preparation. Discard any unused stain.

To prepare 3% Giemsa working solution, follow the procedure mentioned above, but mix 97 mL of buffered water with 3 mL of Giemsa stock solution.

Staining of the Slides

For Thin blood smears

1. Fix air-dried film in absolute methanol by dipping the film briefly (two dips) in a Coplin jar containing absolute methanol.
2. Remove and let air dry.
3. Stain with a working solution of Giemsa stain
4. Wash by briefly dipping the slide in and out of a Coplin jar of buffered water (one or two dips).
   Note: Excessive washing will decolorize the film.
5. Let air dry in a vertical position. Observe under the microscope first at 40X and then using an oil immersion lens

For Thick blood smears

1. Allow the film to air dry thoroughly for several hours or overnight. Do not dry films in an incubator or by heat, because this will fix the blood and interfere with the lysing of the RBCs.
   Note: If a rapid diagnosis of malaria is needed, thick films can be made slightly thinner than usual, allowed to dry for 1 hour, and then stained.
2. DO NOT FIX.
3. Stain with diluted Giemsa stain
4. Wash by placing the film in buffered water for 3 to 5 min.
5. Let air dry in a vertical position, observe under the microscope at 40X, and then use an oil immersion lens.

For Chlamydia trachomatis

Follow the aforementioned steps with the dilute stain of 1:40 dilution (add 0.5 ml stock Giemsa solution to 19.5 ml buffered water) and leave the stain for 90-120 minutes.

Observation

On microscopic observation, cell organelles, bacteria, and parasites are distinguished based on their morphology and color;

Cell Components	The color observed after staining
Red blood cells	Mauve-pink
Neutrophils	Reddish purple nuclei with pink cytoplasm
Eosinophils	Purple nuclei, faintly pink cytoplasm, and red to orange granules.
Basophils	Purple nuclei, blue coarse granules.
Lymphocytes	Dark blue nucleus with light blue cytoplasm.
Monocytes	Pink cytoplasm with a purple color nucleus.
Platelets	Violet to purple color granules.
Nuclei of host cells	Dark purple
Nuclei of WBCs	Dark purple
The cytoplasm of host cells	Pale blue
The cytoplasm of white cells	Pale blue or grey-blue
Melanin granules	Black green
Bacteria	Pale or dark blue
Chlamydia trachomatis inclusion bodies	Blue-mauve to dark purple depending on the stage of development
Borrelia spirochetes	Mauve-purple
Yersinina pestis coccobacilli	Blue with dark stained ends (bipolar staining)
Malaria parasite	Malaria parasites have a red or pink nucleus and blue cytoplasm. If P. vivax is seen, the Schüffner dots are seen as an even carpet of pink dots in the cytoplasm of red blood cells. If P. falciparum is observed, Maurer clefts will be seen as unevenly distributed, coarse bodies in the red cell cytoplasm.

Uses of Giemsa Stain

Wright-Giemsa’s stain is commonly used to demonstrate the cellular elements in peripheral blood and bone marrow smears. Giemsa stain is used to obtain differential white blood cell counts. It is also used to differentiate the nuclear and cytoplasmic morphology of the various blood cells like platelets, RBCs, and WBCs.

In Microbiology, Giemsa stain is used for staining inclusion bodies in Chlamydia trachomatis, Borrelia species, and if Wayson’s stain is not available, to stain Yersinia pestis. Giemsa stain also is used to stain Histoplasma capsulatum, Pneumocystis jiroveci, Klebsiella granulomatis, Talaromyces marneffei (formerly called Penicillium marneffei), and occasionally bacterial capsules.

Cytogenetics also uses this stain to stain the chromosomes and identify chromosomal aberrations. It is commonly used for G-banding (Giemsa-Banding)

Parasitology

In microbiology, this stain is most commonly used in parasitology to detect intraerythrocytic (plasmodia, babesiae) and exoerythrocytic (trypanosomes, microfilaria) parasites. It is also used for the detection of intracellular amastigotes of Leishmania species or Trypanosoma cruzi.

Photomicrograph of a Wright-Giemsa-stained peripheral blood smear illustrating several stages of Plasmodium species. 

Giemsa stain is also used for the laboratory diagnosis of Toxoplasmosis. Tachyzoites of Toxoplasma gondii are best seen in needle aspirates, or impression smears stained with Wright-Giemsa. In Giemsa-stained smears characteristics, bow-shaped or crescent-shaped tachyzoites with the central dark-staining nucleus are seen.

The Wright-Giemsa-stained impression smear illustrates a few background macrophages and numerous tiny 2 to 3 amastigotes of Leishmania. These forms are often difficult to differentiate from the yeast cells of Histoplasma capsulatum. Careful observation, however, will reveal that many of these forms have a small, rod-shaped kinetoplast, characteristics of Leishmania amastigotes.

Bacteriology

Wright-Giemsa stain has little use for staining bacteria, but it can be used for the laboratory diagnosis of various obligate intracellular parasites. 

The diagnosis of Chlamydia trachomatis infection can be made if large numbers of chlamydial inclusion bodies are seen in a sample stained by the Giemsa or Gimenez methods.

The laboratory diagnosis of granuloma inguinale relies on the staining of intracellular bacteria in mononuclear cells and observation of “Donovan bodies” in tissue smears or biopsy specimens examined by Giemsa and Wright stains.

Wright-Giemsa stains of peripheral blood smears of people suffering from bubonic plague reveal the characteristics of bipolar staining typical of Yersinia. Note: bipolar staining “closed safety pin” shaped cells.

In people suffering from Carrion’s disease, Bartonella bacilliformis can be seen in the tissues both intra-and extracellularly. On Giemsa-stained blood films, the organism appears blue-to-purple extraerythrocytic and intraerythrocytic bacilli and coccobacilli.

Mycology

Detect the intracellular yeast forms of Histoplasma capsulatum.

Virology

The stain is also helpful for demonstrating specific intracellular viral inclusions. Herpes simplex virus produces multinucleated giant cells with intranuclear inclusions, which can be visualized after staining with Wright’s stain (or Wright-Giemsa stain).

References

1. Stockert, J. C., Blázquez-Castro, A., & Horobin, R. W. (2014). Identifying different types of chromatin using Giemsa staining. Methods in molecular biology (Clifton, N.J.), 1094, 25–38. https://doi.org/10.1007/978-1-62703-706-8_3 
2. Barcia J. J. (2007). The Giemsa stain: its history and applications. International journal of surgical pathology, 15(3), 292–296. https://doi.org/10.1177/1066896907302239

Capsule stain is a type of differential stain which uses acidic and basic dyes to stain the background and bacterial cells, respectively, so that presence of the capsule is easily visualized. The capsule is synthesized in the cytoplasm and secreted outside the cell, where it surrounds the bacterium.

Most capsulated bacteria have a capsule made up of a polysaccharide layer, but some bacteria have a capsule made up of polypeptide or glycoprotein. Capsules are associated with virulence in several microorganisms, including Streptococcus pneumoniae and Neisseria meningitides, because capsules resist phagocytosis.

In capsule staining procedure “we do not heat fix and rinse the smear with water” as heat and water may dislodge capsules from bacteria.

Principle of Capsule Stain

Bacterial capsules are non-ionic, so neither acidic nor basic stains will adhere to their surfaces. Therefore, the best way to visualize them is to stain the background using an acidic stain (e.g., Nigrosine, congo red) and to stain the cell itself using a basic stain (e.g.,crystal violet, safranin, basic fuchsin, and methylene blue).

Various types of methods are available for the demonstration of bacterial capsules. The results (stain of the cells, background, and capsule) depend on the method used. Two commonly used methods are discussed here:

A. India ink method

In this method, two dyes, crystal violet, and India ink are used. The capsule is seen as a clear halo around the microorganism against the black background. This method is used for demonstrating Cryptococcus.

• The background will be dark (color of India ink).
• The bacterial cells will be stained purple (bacterial cells take crystal violet-basic dyes as they are negatively charged).
• The capsule (if present) will appear clear against the dark background (capsule does not take any stain).

Expert opinion: Tasha Sturm, Microbiologist at Cabrillo College,  “I use Nigrosin instead of India ink as Nigrosin gives a more even background and spreads little easier”. Read details in the comment section.

B. Anthony’s stain method

In this type of capsule staining procedure, the primary stain is crystal violet, and all parts of the cell take up the purple crystal violet stain. There is no mordant in the capsule staining procedure. A 20% copper sulfate solution serves a dual role as both the decolorizing agent and counterstain. It decolorizes the capsule by washing out the crystal violet, but will not decolorize the cell. As the copper sulfate decolorizes the capsule, it also counterstains the capsule. Thus, the capsule appears as a faint blue halo around a purple cell.

Materials and Reagents

• Test bacteria: 36-48 hour culture of capsulated bacteria e.g. Klebsiella pneumoniae growing on a slant of EMB agar or culture of other capsulated bacteria and non-capsulated bacteria [Note: Growing Klebsiella pneumoniae in milk-based media (e.g. Skim milk) increase its capsule size, making it easier to visualize.]
• Stain solutions: Depending on the method used (crystal violet, India ink, Nigrosin, copper sulfate, Basic carbol fuschin solution, methylene blue solution, etc).
• Microscopic slides
• Inoculating loop
• Light Microscope with 100x objective lens (oil immersion)
• Immersion oil
• Gas burner
• Tissue paper

Capsule Stain procedure

A. India Ink Method

1. Place a single drop of India ink on a clean microscope slide, adjacent to the frosted edge.
2. Using a flamed loop and sterile technique, remove some Klebsiella pneumoniae from culture tube or plate and mix it into the drop of India ink. Be sure there are no large clumps of organism, but try to avoid spreading the drop.
3. Place the end of another clean microscope slide at an angle to the end of the slide containing the organism. Spread out the drop out into a film. This is done by contacting the drop of India ink with the clean microscope slide and using the capillary action of the dye/ slide to spread the India ink across the smear.
4. Allow the film to air dry (will take 5-7 minutes). DO NOT heat or blot dry!  Heat will melt the capsule!
5. Saturate the slide with crystal violet for 1 minute and rinse slightly & very gently with water. Be cautious water may remove the capsule from the cell.
6. Let the slide air dry for a few minutes. DO NOT blot the slide! Blotting will remove the bacteria from the slide and/or distort the capsule.
7. Observe the slide under oil immersion.

Results: Look for purple cells surrounded by a clear halo on a dark background. The halo is the capsule. You may need to decrease the amount of light in order to make the capsule easier to see.

B. Anthony’s stain method

1. Place a single drop of crystal violet on a clean microscope slide, adjacent to the frosted edge.
2. Using a flamed loop and sterile technique, add three loopful of test bacterium (any capsulated bacteria such as Klebsiella pneumoniae, Streptococcus pneumoniae) from broth culture. If you are adding bacteria from a culture plate make sure that there are no large clumps of the organism, but try to avoid spreading the drop.
3. Place the end of another clean microscope slide at an angle to the end of the slide containing the organism. Spread out the drop out into a film. This is done by contacting the drop of crystal violet with the clean microscope slide and using the capillary action of the dye/ slide to spread the crystal violet across the smear.
4. Allow the film to air dry (will take 5-7 minutes). DO NOT heat or blot dry!  Heat will melt the capsule!
5. Tilt the slide and rinse with 20% copper sulfate solution. DO NOT RINSE WITH WATER! Water will remove the capsule from the cell.
6. Let the slide air dry for a few minutes. DO NOT blot the slide! Blotting will remove the bacteria from the slide and/or distort the capsule.
7. Observe the slide under oil immersion.

Results: Look for purple cells surrounded by a clear or faint blue halo on transparent background. The halo is the capsule. You may need to decrease the amount of light in order to make the capsule easier to see.

Points to remember

• Clean your microscope with lens cleaner, removing all oil from lenses.
• Dispose of staining waste and slides in designated waste containers.
• Be cautious while handling the slide, since the organisms have not been killed.

References and further reading

1. Microbugz . Capsule Stain. https://www.austincc.edu/microbugz/capsule_stain.php   
2. Color Atlas and Textbook of Diagnostic Microbiology, Koneman, 5th edition

The simple stain can be used as a quick and easy way to determine the cell shape, size, and arrangement of bacteria. True to its name, the simple stain is a very simple staining procedure involving a single stain solution. Any basic dye, such as methylene blue, safranin, or crystal violet, can be used to color the bacterial cells.

These stains will readily give up a hydroxide ion or accept a hydrogen ion, which leaves the stain positively charged.  Since most bacterial cells and cytoplasm surface is negatively charged, these positively charged stains adhere readily to the cell surface. After staining,  bacterial cell morphology (shape and arrangement) can be appreciated.

Procedure

Preparation of a smear

1. Using a sterilized inoculating loop, transfer a loopful of liquid suspension containing bacteria to a slide (clean grease-free microscopic slide) or transfer an isolated colony from a culture plate to a slide with a water drop.
2. Disperse the bacteria on the loop in the drop of water on the slide and spread the drop over an area the size of a dime. It should be a thin, even smear.
3. Allow the smear to dry thoroughly.
4. Heat-fix the smear cautiously by passing the underside of the slide through the burner flame two or three times. It fixes the cell in the slide. Do not overheat the slide as it will distort the bacterial cells.

Staining 

1. Cover the smear with methylene blue and allow the dye to remain in the smear for approximately one minute (Staining time is not critical here; somewhere between 30 seconds to 2 minutes should give you an acceptable stain, the longer you leave the dye in it, the darker will be the stain).
2. Using distilled water wash bottle, gently wash off the excess methylene blue from the slide by directing a gentle stream of water over the surface of the slide.
3. Wash off any stain that got on the bottom of the slide as well.
4. Saturate the smear again but this time with Iodine. Iodine will set the stain
5. Wash any excess iodine with gently running tap water. Rinse thoroughly. (You may not get a mention of steps 4 and 5  in some textbooks)
6. Wipe the back of the slide and blot the stained surface with bibulous paper or with a paper towel.
7. Place the stained smear on the microscope stage smear side up and focus the smear using the 10X objective.
8. Choose an area of the smear in which the cells are well spread in a monolayer. Center the area to be studied, apply immersion oil directly to the smear, and focus the smear under oil with the 100X objective.

Results

The bacterial cells usually stain uniformly and the color of the cell depends on the type of dye used. If methylene blue is used, some granules in the interior of the cells of some bacteria may appear more deeply stained than the rest of the cell, which is due to the presence of different chemical substances.

Uses

Diagnostic microbiology laboratory generally does not perform simple staining methods. Differential stainings such as Gram staining and AFB staining are commonly used to identify and differentiate bacterial isolates. Simple staining can be useful in the following circumstances.

1. To differentiate bacteria from yeast cells: When endocervical swab culture is done in blood agar both Staphylococcus spp and yeast cells may give similar-looking colonies in Blood agar (a common error for a new technologist or microbiologist with less experience). Performing the wet mount technique or simple staining from the isolate can be helpful.
2. To presumptively identify the bacterial isolate
   Due to their ubiquitous presence, Bacillus spp may present as a contaminant in the culture plates. In some circumstances (e.g. growth in Blood agar but no growth in MacConkey agar), identifying the shape of the bacteria (rod or cocci) may help to eliminate the isolate as possible contaminants (e.g., Bacillus spp) or further process as a potential pathogen (cocci).

References

1. Moyes, R. B., Reynolds, J., & Breakwell, D. P. (2009). Preliminary staining of bacteria: simple stains. Current protocols in microbiology, Appendix 3, . https://doi.org/10.1002/9780471729259.mca03es15 
2. Tripathi, N., & Sapra, A. (2023). Gram Staining. In StatPearls. StatPearls Publishing. 
3. Dagnall, G. J., & Wilsmore, A. J. (1990). A simple staining method for the identification of chlamydial elementary bodies in the fetal membranes of sheep affected by ovine enzootic abortion. Veterinary microbiology, 21(3), 233–239. https://doi.org/10.1016/0378-1135(90)90034-s

Acridine orange is a fluorescent dye that intercalates or binds with the nucleic acid (either DNA or RNA) present in organisms and fluoresces to emit various colors that help differentiate cellular organelles. This binding results from the electrostatic interactions of acridine molecules between the nucleic acid-base pairs. Due to its metachromatic properties, acridine orange (AO) is commonly used in fluorescence microscopy and flow cytometry analysis of cellular physiology and cell cycle status, including the fluorescent microscopic examination of microorganisms.

Principle

Acridine orange is a cell-permeable, nucleic acid selective fluorescent dye that emits green fluorescence when bound to dsDNA (at 520 ) and red fluorescence when bound to ssDNA or RNA (at 650 nm). Since it is a cationic dye, it also enters acidic compartments such as lysosomes which, in low pH conditions, will emit orange light.

Acridine orange is a carcinogen when absorbed through the skin. Wear gloves when working with this stain.

Staining procedure:

Staining procedures vary according to their use

1. For staining clinical specimens with acridine orange at low pH (Acridine orange acid stain)

• Requirements: acridine orange, glacial acetic acid, distilled water
• Preparation of reagent: 50 mg acridine orange is dissolved in 10 ml of distilled water to prepare a stock solution and stored in the refrigerator.1 ml of acridine orange stock solution and 0.5 ml of glacial acetic acid is added to 50 ml of distilled water to prepare a working solution.
• Staining procedure:

1. Prepare a smear in a clean grease-free slide and allow it to air dry.
2. The slide is then fixed with methanol and dried again.
3. It is then put in a trough with an acridine orange staining working solution (i.e 0.01 percent).
4. After 2 minutes of staining, the slides are washed gently with water, dried, and examined in a fluorescent microscope.

Observance: Bacteria stain orange against a green to a yellow background of human cells and debris.

2. For staining cells for analysis by flow cytometry.
   Requirements: 0.1M Citric Acid (dissolve 1.921g per 100ml distilled water), 0.2M Dibasic Sodium Phosphate  (dissolve 2.839g per 100ml distilled water) ,Triton X-100 (Baker), 0.5M EDTA, Sodium chloride(NaCl), Acridine Orange (Powder) and Sucrose.
3. Preparation of reagents:
   • Stock Buffer I:20mM Citrate-Phosphate, pH 3.0, 0.1mM EDTA, 0.2M Sucrose, 0.1% Triton X-100
     (To 125ml distilled water add 40µl 0.5M EDTA, 26.48ml 0.1M Citric Acid, 6.85ml 0.2M Dibasic Sodium Phosphate, 13.69g Sucrose, 0.2ml Triton X-100 .QS to 200ml and 0.2µ filter. Store at 4°C)
   • Stock Buffer II:10mM Citrate-Phosphate, pH 3.8, 0.1M NaCl (To 150 ml distilled water, add 9.92ml 0.1M Citric Acid, 5.46ml 0.2M Dibasic Sodium Phosphate, 1.7g NaCl. QS to 200ml and 0.2m filter. Store at 4°C)

Staining Procedure

1. Make a 2mg/ml solution of acridine orange in distilled water and dilute to 1:100 in Buffer II
2. Aliquot cells: 105- 106 in 100µl PBS or media.
3. Add Buffer I (0.5ml) at room temp, agitate to suspend.
4. Add Buffer II + AO (0.5ml) at room temp, agitate to suspend.
5. Run on flow cytometer. Excitation 488 nm; dot plot of green fluorescence at 530nm versus red fluorescence >600 nm).

Observation

• Green fluorescence when bound to dsDNA and
• Red fluorescence when bound to ssDNA or RNA.

Applications

Acridine orange stain is used to confirm the presence of bacteria in positive blood cultures when Gram stain results are difficult to interpret or when the presence of bacteria is highly suspected, but none are detected using light microscopy. Other applications of acridine orange stains are listed here;

• For enumerating the microbial load in a sample since acridine orange binds with the nucleic acid of both living and dead bacteria.
• Detection of cell wall-deficient bacteria (e.g., mycoplasmas) grown in cultures. Cell wall deficient bacteria are hard to visualize in Gram stain as they cannot retain Gram stain dyes.
• For differential staining of human cells and prokaryotic cell with a fluorescence microscope. Human cells are stained black to faint green in which bright orange organisms are easily detected.
• Acridine orange is also used in a method referred to as the quantitative buffy coat (QBC), a rapid screening tool for the detection of malaria.
• For analyzing mitochondria and lysosomal content by flow cytometry.
• For visual detection of nucleic acids on agarose and polyacrylamide gels.
• For identifying engulfed apoptotic cells because they will fluoresce upon engulfment.

 Research showed that acridine orange staining is a sensitive, rapid, and reliable method for detecting bacteria in blood cultures early during incubation and can be substituted for blind subcultures. Acridine orange is better than Gram stain in cases with low amounts of organisms.

Limitations

• Acridine orange stain is non-specific. It stains all nucleic acid-containing cells, so it does not discriminate between gram-negative and gram-positive bacteria.

Quality Control

1. Examine the acridine orange staining solution for color and clarity. The solution should be clear, orange, and without evidence of precipitate.
2. Each use time, stain a prepared slide of known bacteria, such as Escherichia coli mixed with staphylococci, and examine for the desired results. Record results and refer out-of-control results to the supervisor.
   1. Gram-negative rods and Gram-positive cocci are fluorescent (orange).
   2. The background is nonfluorescent (green-yellow).

References

• Bailey & Scott’s Diagnostic Microbiology, Forbes, 11th edition
• Review of Medical Microbiology and Immunology, Lange Medical Books, 13th edition