In this article we will discuss about the staining procedure used for detecting bacteria.
1. Simple Staining Procedure:
When a single staining-reagent is used and all cells and their structures stain in the same manner, the procedure is called simple staining procedure.
This procedure is of two types – positive and negative (Fig. 17.5). In positive staining, the stain (e.g., methylene blue) is basic (cationic) having positive charge and attaches to the surface of object that is negatively charged.
In negative staining, the stain (e.g., India ink, nigrosin) is acidic (anionic) having negative charge and is repelled by the object that is negatively charged, and thus fills the spaces between the objects resulting in indirect staining of the object.
2. Differential Staining Procedure:
When more than one staining reagents are used and specific objects (e.g., specific microorganisms and/or particular structure of a microorganism) exhibit different staining reactions readily distinguishable, the procedure is called differential staining.
The most widely used differential staining in microbiology are Gram-staining and acid-fast staining. Acid-fast staining is especially useful in identifying Mycobacterium tuberculosis, the causative agent of tuberculosis.
A. Separation of Microbes into Groups:
(i) Gram-Staining:
A Danish scholar Christian Gram in 1884 devised a differential staining procedure which differentiates bacteria into gram-positive and gram-negative. This procedure is called Gram- staining technique. This technique has experienced numerous modifications from time to time and proves to be valuable for staining smears of pure cultures of bacteria.
Procedure:
The staining procedure (Fig. 17.6) is as follows:
I. A thin film of young culture (smear) is fixed on a clean slide.
II. The smear is stained for one minute with ammonium oxalate crystal violet. This stain sometimes yields over stained preparations in which certain gram-negative organisms (e.g. Gonococcus) also retain the stain. If this trouble is encountered, lesser amount of crystal violet should be used.
III. The slide is washed in tap water for not more than 2 seconds to remove excess stain.
IV. The slide is immersed for one minute in Lugol’s iodine solution. The bacteria become deeply stained and appear deep purple in colour due to crystal violet-iodine-complex formation.
V. The slide is washed in tap water and blot-dried.
VI. The slide is gently agitated for 30 seconds in 95% ethyl alcohol and blot-dried, gram-negative bacteria lose their stain in this step (i.e. decolourize). However, the gram- positive ones retain deep purple colour.
VII. The slide is now counterstained for 10 seconds in the safranin solution.
VIII. The slide is washed in tap water, dried and examined.
Results:
Gram-positive bacteria: deep purple (blue); gram-negative bacteria: pink (red).
Mechanism:
Although different explanations have been given to answer why bacteria respond differently to the Gram-stain, it seems likely that the answer is related to the physical nature of their cell walls as when cell wall is removed from gram-positive bacteria, they become gram-negative. The peptidoglycan appears to act as a permeability barrier preventing loss of crystal violet-iodine-complex.
When gram-positive bacteria are treated with destaining agent (alcohol), the alcohol is thought to dehydrate the thick layer of peptidoglycan resulting in shrinkage of pores of peptidoglycan. Shrinkage of peptidoglycan pores prevents crystal violet-iodine-complex from escaping and the bacteria remain deep purple.
In contrast, peptidoglycan is very thin in gram-negative bacteria, not as highly crossed-linked as is in gram-positive ones, and has larger pores. Alcohol, therefore, readily penetrates the lipid-rich outer layer of the cell wall and extracts enough lipid thus increasing the porosity further.
For these reasons, alcohol more readily removes the deep purple crystal violet-complex from gram-negative bacteria and the latter become decolourized.
Precautions:
1. Fresh and young culture (less than 24 hours old) should always be used to avoid misleading results because old cultures of gram-positive bacteria tend to decolourize more rapidly and show gramnegativeness.
2. Smears should be thin and uniform to avoid over populated bacteria.
3. During heat-fixing, excessive heating should be avoided.
4. Over decolourization should be avoided.
(ii) Acid-Fast Staining:
The acid-fast stain is a differential stain developed first by Paul Ehrlich in 1882 and later on modified by Ziehl- Neelsen, and is in use even today by microbiologists. Majority of the bacteria are stained with simple stain and Gram-stain but certain bacteria do not do so because they have waxy components of the cell wall, hence their cell wall has limited permeability.
Such bacteria belong to genera like Mycobacterium and Nocardia; and are stained by acid-fast stain; the latter is used to identify Mycobacterium tuberculosis and M. leprae, the pathogens responsible for tuberculosis and laprosy, respectively.
The acid-fastness property of these bacteria is correlated with high lipid contents, which makes them difficult to stain. Hence for staining of these bacteria heating with strong dye is required. Once the acid-fast bacteria are stained, it is difficult to decolourize them even with acid and alcohol. Moreover, acid-fast staining serves also as good identification tool for a number of harmless saprophytic bacteria.
Reagent Preparation:
The reagents are prepared as given below:
Carbol Fuchsin Stain:
Basic fuchsin – 0.3 g
Ethanol (95%) – 10.0 ml
Phenol (heat melted crystals) – 5.0 ml
The basic fuchsin is dissolved in ethanol and phenol is dissolved in water.
These two are mixed and kept for several days before filter and use:
Decolourising solvent (acid-alcohol):
Ethanol (95%) – 97.0 ml
Hydrochloric acid (conc.) – 3.0 ml
Counter stain:
Methylene blue chloride – 0.3 g
Distilled water – 100.0 ml
Procedure:
The staining procedure (Fig. 17.7) is as follows:
I. A thin film of young culture (smear) is heat-fixed and air-dried on a clean slide.
II. The smear is now flooded with carbol fuchsin.
III. The slide is steamed over boiling water for 3-5 minutes More stain is added time to time to prevent smear from becoming dry.
IV. Slide is cooled and washed with distilled water until no colour appears from the smear.
V. Smear is decolourized with decolourising solvent (acid-alcohol) for 15-20 seconds. Some bacterial cells appear red (faint pink) in colour, while others decolourize. Slides arc washed with distilled water.
VI. Smears are now counterstained with methylene blue for 1-2 minutes and washed with distilled water.
VII. Slides are blot-dried with bibulous paper and examined directly under oil-immersion.
Results:
Acid-fast bacteria appear red.
Non-acid-fast bacteria appear blue.
Mechanism:
Acid-fast staining helps classifying bacteria into two groups: acid-fast and non-acid-fast. Acid fast bacteria, particularly those in the genus Mycobacterium, do not bind simple stains but when stained by heating with a mixture of carbol fuchsin (basic fuchsin + phenol) they retain carbol fuchsin (the primary stain) after washing even with strong acid.
Once basic fuchsin penetrates with the aid of heat and phenol, cells of acid-fast bacteria are not easily decolourized by an aid-alcohol wash and hence remain red.
This is due to the quite high lipid content of cell walls of acid-fast bacteria; in particular, mycolic acid (a group of branched chain hydroxy lipids) appears responsible for acid-fastness. The nonacid-fast bacteria get decolourized after washing with acid-alcohol; they retain the counterstain methylene blue hence appear blue.
Precautions:
1. If necessary, more carbol fuchsin stain should be added to avoid its evaporation and dryness.
2. Carbol fuchsin stain should be prevented from heating to avoid “messy” preparations.
3. Over decolourization of the smear should be avoided.
B. Visualization of Various Structures:
(i) Endospore Staining:
Bacteria in the genera Bacillus and Clostridium form an exceptionally resistant structure capable of surviving for long periods in an unfavourable environment. This dormant structure is called an endospore since it develops within the cell. Endospore morphology and location vary with species and often are valuable in identification. Endospores are not easily stained well by most dyes.
Considerable amount of heating is required in order to make the stain penetrate the spore-coat, a thick wall primarily responsible for endospore resistance. But once stained, they strongly resist decolourization. This property is the basis of endospore staining techniques.
However, there are two staining procedures used by microbiologists to stain the endospores. These methods are the Schaeffer-Fulton method and Dorner method. For convenience, Schaeffer-Fulton method is given here.
Procedure:
The staining procedure of endospore (Fig. 17.8) by Schaeffer-Fulton method is as follows:
i. A thin film of young culture (smear) of endosporous bacteria is fixed on a clean slide.
ii. The smear is heat-fixed on to the slide by gentle warming.
iii. Smear is covered with the solution of malachite green which is a very strong stain that can penetrate the spore-coat of an endospore.
iv. The slide is kept on a suitable stand and heated with steam from below for 5 minutes. If the stain dries up during heating, more stain is added to the smear from time to time as per requirement.
v. The slide is washed gently under tap water.
vi. The slide is counter-stained with safrain for about 30 seconds.
vii. It is then washed with distilled water and dried with blotter.
viii. The smear is observed under oil immersion.
Results:
The endospore appears green while rest of the cell appears red.
Mechanism:
Endospores are extremely resistant due to their thick wall, the spore coat. The spore coat does not take the stain easily. Malachite green, however, penetrates the spore coat of endospore after considerable heating. Once stained, the endospore does not decolourizes easily hence appears green even after washing. In contrast, the counter stain fails entering the endospore but stains rest of the cell content that appears red.
(ii) Flagella Staining:
Many bacteria are motile due to the presence of flagella that originate in the cytoplasm and project out from the cell wall. Bacterial flagella are fine, threadlike organelles that are so slender (about 10 to 30 nm in diameter) that they can only be observed directly using electron microscope.
To observe flagella with the light microscope, the thickness of flagella is increased by coating them with mordants like tannic acid and potassium alum, and they are stained with pararosaniline (Leifson’s method) or basic fuchsin (Gray’s method).
Flagella staining procedures provide taxonomically valuable informations (e.g., presence, distribution pattern, number) which are used in the identification and classification of bacteria.
However, Gray’s method of flagella staining is as follows:
Reagent preparation:
The reagents are prepared as given below:
Solution A (Mordant):
Tannic acid (20% aqueous solution) – 2.0 ml
Potassium alum (saturated aqueous solution) – 5.0 ml
Mercuric chloride (saturated aqueous solution) – 2.0 ml
Solution B (stain):
Basic fuchsin (3%) in 95% alcohol – 0.4 ml
Solutions A and B are prepared by mixing their ingredients less than 24 hour before using and are stored in coloured bottles. Both solutions separately may be kept indefinitely.
Procedure:
I. Bacteria are grown on a suitable broth. If it is a solid medium, small quantity of 1% peptone is added in which the bacteria swim.
II. Medium is removed by centrifugation to obtain the pellet.
III. The pellet is suspended in 10% formaline to produce a light, faint turbidity, and for the prevention of flagella.
IV. The loopful of suspension is transferred on to a new greese-free slide and spread to prepare a thin film of young culture (smear).
V. The smear is flooded with freshly filtered mordant (solution A) and allowed to act 8-10 minutes.
VI. The smear is washed with a gentle stream of distilled water.
VII. The smear is flooded with freshly filtered basic fuchsine stain (solution B) and allowed to stand 5 minutes without heating.
VIII. The smear is gently washed with distilled water and air-dried.
IX. The slide is examined directly under oil-immersion.
Results:
Flagella stained red are observed.
Precautions:
1. Especially cleaned greese-free slides should be used.
2. Smears should not be heat-fixed.
3. Smears should not be blot-dried.
4. Reduced illumination should always be used in the microscope to observe flagella clearly.
(iii) Capsule Staining:
Bacterial capsules are more easily confused with artifacts than any other structure pertaining to the organisms. Inasmuch as capsules sometimes show merely as unstained areas around the bacterial cells, there is a temptation to call any such surrounding area a capsule; very often, however, they merely represent the tendency of a lightly stained surrounding medium to retract from bacterial cells on drying.
For this reason, the best way to demonstrate capsules is actually to stain them by some procedure which differentiates them from the bacterial cells itself. Though there are several methods of staining to accomplish this, the procedure of Anthony’s method is much simpler. E.E. Anthony devised this method for capsule staining in 1931.
However, Anthony’s method for capsule staining is the following:
Reagents:
The reagents used in the procedure are the following:
Crystal violet aqueous (85% dye content) – The stain
Copper sulfate aqueous (20%; CuSO4.5H2O) – Decolourizing agent as well as counter stain Distilled water.
Procedure:
The staining procedure (Fig. 17.9) is as follows:
I. Two loopfuls of young culture of bacteria (e.g., 36-48 hour milk cultures of Klebsiella pneumoniae) is put on a clean glass slide.
II. A heavy smear is prepared and air-dried.
III. Smear is flooded with crystal violet stain for 2 minutes.
IV. The stain is washed off with copper sulfate (20%)
V. The stain is washed off with copper sulfate (20%).
VI. Copper sulfate is drained and the smear is gently blot-dried.
VII. The slide is examined directly under oil-immersion.
Results:
The bacterial cells appear dark blue, encircled by blue violet coloured capsule indicating that the bacteria are capsulated.
Mechanism:
Crystal violet aqueous (85% dye content) is the primary stain, and on its application both the capsular material and the bacterial cell wall take the colour of the stain and appear dark blue. But the capsule being non-ionic fails to absorb the primary stain, while the cell wall being ionic absorbs it.
Copper sulfate aqueous (20%) acts both as a decolourizing agent as well as a counter stain. On application, the copper sulfate first decolourizes, the capsule by removing crystal violet and then imparts blue violet colour to it. As a result, the capsule finally appears blue violet in contrast to the dark blue colour of the bacterial cell.
Precautions:
1. Heavy smear should be prepared to counter the removal of bacterial cells during washing.
2. Smear should not be heat-fixed because the heating results in shinkage which may create a clear zone surrounding the cell that is an artifact and that can be mistaken for the capsule.
3. Washing off the smear with water should be avoided because the capsular materials are water soluble and may be dislodged and removed with water.