As bacteria consist of clear protoplasmic matter, differing but
slightly in refractive index from the medium in which they are
growing, it is difficult with the ordinary microscope, except
when special methods of illumination are used, to set them in
the unstained condition.
Staining, therefore, is of primary importance for the
recognition of bacteria.
Staining may be simple staining and differential staining.
Basic dyes: Methylene blue, Basic fuchsin, Crystal violet, Safranine,
Malachite green have positively charged groups (usually from
penta-valent nitrogen) and are generally sold as chloride salts.
Basic dyes bind to negatively charged molecules such as nucleic
acids, many proteins and surfaces of bacterial and archeal cells.
Acidic dyes: Eosin, Rose Bengal and Acid fuchsin possess groups
such as carboxyls (-COOH) and phenolic hydroxyls (-OH). Acidic
dyes, in their ionized form, have a negative charge and bind to
positively charged cell structures.
4. SIMPLE STAINING
• These show not only the presence of organisms but also the
nature of the cellular content in exudates.
• A single stain is used.
• Examples are Loeffler’s methylene blue, polychrome
methylene blue, dilute carbol fuchsin.
• Simple staining is of positive staining and negative staining.
6. Negative staining
• Negative staining procedure helps to study the cell shape, cell
breakage, refractable inclusion bodies and spores besides
poly- hydroxy butyrate granules.
• It is useful for those bacteria which are difficult to stain.
• Very slender bacteria like spirochaetes that are not detectable
by simple staining methods can be viewed by negative
Negative staining requires the use of acidic stains such as Indian
ink and nigrosin. The acidic stain with its negatively charged
chromogen will not penetrate the cell because of the negative
charges on the surface of the bacteria. So unstained cells are
diffferentiable against the dark background. Since heat
fixation is not required, the cells are not subjected to the
destaining effect of chemicals and heat, their natural size,
shape and arrangement can be seen by this method. It is
possible to observe bacteria that are difficult to stain.
• A loopful of undiluted Indian ink was placed on one end of clean
• A loopful of inoculum was transferred into the drop of stain.
• By using a second dirt-free slide with smooth edge over the
suspension, it was spread uniformly along the edge.
• The suspension was spread to the end of the slide so as to form a
• The slide was then air dried and observed under oil immersion
• Colorless bacteria are seen against a dark background.
14. GRAM’S STAINING
• Gram Stain is developed in 1884 by the Danish physician
Christian Gram, is the most widely used method in
• It is first and usually the only method employed for the
diagnostic identification of bacteria in clinical specimens.
• Violet dye and the iodine combine to form an insoluble, dark
purple compound in the bacterial protoplasm and cell wall.
• This compound is dissociable in the decolorizer, which
dissolves and removes its two components from the cell.
• But the removal is much slower from Gram-positive than
from the Gram-negative bacteria, so that by correct timing
the former stay dark purple whilst the latter become colorless.
• The difference between the two types of bacteria is that the
Gram positive have thicker and denser peptidoglycan layers
in their cell walls, which makes them less permeable to the
stain than those of the Gram negative bacteria.
• The iodine has a critical role in enhancing this difference.
• It seems to bind temporarily to the peptidoglycan and make it
even less permeable to the dye.
• Step 1- Crystal violet (primary
stain) for 1 minute. Water rinse.
• Step 2- Iodine (mordant) for 1
minute. Water rinse.
• Step 3 – Alcohol (decolorizer)
for 10-30 seconds. Water rinse.
• Step 4 - Safranin (counterstain)
for 30-60 seconds. Water rinse.
• Cells stain purple.
• Cells remain purple.
• Gram-positive cells remain
purple. Gram negative cells
• Gram positive cells remain
purple. Gram-negative cells
23. ACID – FAST STAINING
• This is also known as Ziehl – Neelsen staining.
• This method is a modification of Ehrlich’s (1882) original
method for the differential staining of tubercle bacilli and
other acid-fast bacilli with aniline-gentian violet followed by
strong nitric acid.
• Stain used consists of basic fuchsin, with phenol added.
Acid fast bacteria retain primary stain (carbol fuchsin) even
after washing with a strong acid. It appears red while non-
acid fast bacteria are decolorized on washing with acid and
takes the color of the counter stain (methylene blue). The
property of acid fasting appears due to the presence of
mycolic acid in their cell walls. Mycolic acid is a group of
branched chain hydroxy lipids.
25. It is most commonly used to identify M.tuberculosis and M.leprae,
the pathogen responsible for tuberculosis and leprosy,
respectively. These bacteria have cell walls containing lipids
constructed from mycolic acids, a group of branched chain
hydroxyl fatty acids, which prevent dyes readily binding to the
However, M.tuberculosis and M.leprae can be stained by harsh
procedures such as the Zeihl-Neelson method which uses heat and
phenol to derive basic fuchsin into cells.
Once basic fuchsin has penetrated, M.tuberculosis and M.leprae
are not easily decolorized by acidified alcohol (acid-alcohol) and
thus are said to be acid-fast.
Non acid-fast bacteria are decolorized by acid-alcohol and thus
are stained blue by methylene blue counterstain.
• A clean sterile glass slide was taken.
• A thin uniform smear was prepared in the slide using a
• The smear was allowed to air dry and then heat fixed.
• The smear was flooded with carbol fuchsin and heated until
• Preparation was allowed to stay for 5-7 min. The stain must
not be allowed to evaporate or dry in the slide. Pour more
carbol fuchsin on slide is necessary.
• It is then washed under a low steam of running water.
• The smear was decolorized with 20% sulphuric acid.
• The slide is then washed again under running tap water.
• Counter stain the smear with methylene blue for the 2 min.
• Washed under running tap water.
• Slide was blot dried and examined under microscope.
31. ZN methods for weakly
1. Leprosy bacilli are acid-fast, but usually to lesser degree
than the tubercle bacillus. They are stained in films or
sections in the same way as the tubercle bacillus, except that
5% sulphuric acid is used for decolorization in the place of
20% sulphuric acid or acid-alcohol.
2. Sections of tissues containing ‘clubs’ formed by
actinomycetes, mycobacteria and nocardiae can be stained
by ZN stain and decolorized with 1% sulphuric acid to
demonstrate the acid-fastness of the clubs.
32. ZN methods for weakly acid-
3. Brucella differential stain. Brucella abortus in infected tissue
or exudate may be distinguished from the latter by its
weakly acid-fast reaction. Stain with dilute (1-in-10) carbol
fuchsin, without heating, for 15 min. Decolorize with 0.5%
acetic acid solution for 15 seconds, wash thoroughly with tap
water and counter stain with Loeffler’s methyene blue for 1
• Volutin granules are a type of cytoplasmic inclusion bodies
found in many bacteria as well as in some fungi, algae,
• These granules are composed mainly of polyphosphate, RNA
• These granules are found most prominent in old cultures
before starvation occurs.
• The method of volutin granule staining is known as ALBERT-
Albert’s stain contains cationic dyes like toludine blue and
Due to the highly acidic nature of the granules, they can be
selectively stained by acidified basic dyes.
The toludine blue preferentially stain volutin granules while
malachite green stains the cytoplasm.
Later due to application of Albert’s iodine, the dye molecule are
fixed by precipitation.
Well developed granules of volutin (polyphosphate) may be seen
in unstained wet preparations as round refractile bodies within
the bacterial cytoplasm
• A thin uniform smear of culture was made. It was air dried and
• Lower the slide with Albert’s stain A and allowed to react for 3-5
• The slide was then washed under running tap water.
• Flood the slide with Albert’s Iodine and allowed to react about 1
• Slide was then washed and blot dried.
• The slide was observed under oil immersion objective of a
38. SPORE STAINING
• The morphology of bacterial endospores is best observed in
unstained wet films under the phase contrast microscope,
where they appear as large, refractile, oval or spherical bodies
within a bacterial mother cells or else from the bacteria.
• If spore-bearing organisms are stained with ordinary dyes , or
by Gram’s stain , the body of the bacillus is deeply colored ,
whereas the spore is unstained and appears as clear area in
• This is the way in which spores are most commonly observed.
39. Spore staining
• If desired , however , it is possible by vigorous staining
procedures to introduce dye into the substance of the spore.
• When thus stained , the spores tends to retain the dye after
treatment with decolorizing agents, and in this respect
behaves similarly to the tubercle bacillus, but is more weakly
• The spores are thick walled structures and very resistant to
physical and chemical agents.
• The spores have a capacity to survive for long periods even in
unfavourable environmental conditions.
• The heat resistance by spores is due to the high content
calcium- dipicolinic acid.
• The spores are differentially stained using special procedures
that help dye to penetrate the spore wall.
• An aqueous primary stain, malachite green is applied and
steamed to enhance the penetration of the impermeable spore
• Once stained the endospore does not readily decolorize even with
the application of decolorizer and they appear, but the cytoplasm
of the cell takes the color of safranine and appears red.
• A modified Ziehl-Neelsen stain in which weak, 0.25% sulphuric
acid is used as decolorizer, yields red spores in blue-stained
bacteria. Lipid granules also stain red, appearing like small
Films are dried and fixed with minimal flaming.
1. Place the slide over a beaker of boiling water , resting it on
the rim with the bacterial film uppermost.
2. When , within several seconds , large droplets have
condensed on the underside of the slide , flood it with 5%
aqueous solution of malachite green and leave to act for 1
min while the water continues to boil.
3. Wash in cold water.
4. Treat with 0.5% safranine or 0.05% basic fuchsin for 30
5. Wash and dry.
This method colors the spores green and the vegetative bacilli
red. Lipid granules are unstained.
46. CAPSULE STAINING
Some bacteria secrete a prominent slimy or gummy material on their
surface usually polysaccharide make up the capsule. It is not
essential for life, but may serve as a reserve food. It provides
protection against dehydration and also against phagocytosis. The
chemical composition of capsule varies according to organism but
usually consist of polysaccharides e.g., glucose, galactose, amino
Capsule are colorless and have low refractive index and so are
difficult to observe without a special staining technique. Moreover
the capsule is non-toxic, hence cannot be stained in the usual
manner. Techniques like negative staining can be used to
demonstrate the capsule.
Congo red is a negative stain. It stains the back ground leaving
the capsule unstained. Acid fuschin fixes congo red and
reacts with it to give a blue back ground. Acid fuschin also
stains the organism pink, leaving the capsule colorless.
• Placed a drop of Manevali’s solution I on a clear glass slide.
• Sterilised the nichrome loop, cooled it and just touched to the
growth of organism on the slant.
• Mixed the culture on loop with a drop of Manevali’s solution I
on the slide.
• The drop was spread into a thin film.
• The film was allowed to dry completely.
• The smear was covered with Manevali’s solution II and
allowed to react for 1 min.
• The excess stain was discarded and dried in air.
• Observe under oil immersion objective. 48
50. FLAGELLAR STAINING
Flagellar staining provides taxonomically valuable information
about the presence and distribution pattern of flagella on
prokaryotic cells. Bacterial and archaeal flagella are fine,
threadlike organelles of locomotion that are so slender (about
10 to 30 nm in diameter) they can only seen directly using
electron microscope. To observe bacterial flagella with the
light microscope, their thickness is increased by coating them
with mordants such as tannic acid and potassium alum, and
then staining with pararosalineor basic fuchsin.
• Grow bacteria for 16 – 24 hrs on a non-inhibitory medium ,
e.g. tryptic soy agar or blood agar.
• Touch a loopful of water onto the edge of a colony and let
motile bacteria swim into it.
• Then transfer the loopful into a loopful of water on a slide to
get a faintly turbid suspension and cover it with a cover-slip.
• The bacterial suspension is thus prepared with a minimum of
agitation , which would detach the flagella.
• After 5-10 min , when many bacteria have attached to the
surfaces of the slide and cover-slip , apply two drops of Ryu’s
stain to the edge of the cover-slip and leave the stain to diffuse
into the film.
• Examine with the microscope after standing 5-15 min at
55. PERIODIC ACID-SCHIFF (PAS)
METHOD FOR FUNGI IN
The polysaccharide constituents of bacteria and fungi are
oxidized by periodate to form polyaldehydes which yield red-
colored compounds with Schiff’s fuchsin-sulphite; the
proteins and nucleic acids remain uncolored. The method
may be used to reveal fungal elements in sections of infected
animal tissue; the fungi stain red, while the tissue material,
except glycogen and mucin, fails to take the stain.
• Bring sections to distilled water.
• Treat for 5 min with a freshly prepared 1% solution of
periodic acid in water.
• Wash in running tap water for 15 min and rinse in distilled
• Stain with fuchsin-sulphite for 15 min.
• Wash two or three times with sulphite wash solution. Wash
• Wash in running tap water for 5 min and rinse in distilled
• Counterstain with dilute aqueous malachite green or with
0.1 % light green in 90 % alcohol for 1 min.
• Dehydrate rapidly in absolute alcohol , clear in xylene and
mount in Canada balsam.
59. LEISHMAN’S STAIN
Dry unfixed films are used. The stain is first used undiluted ,
and the methyl alcohol fixes the film.
The stain is then diluted with distilled water, and the staining
properly carried out.
• Pour the undiluted stain on the unfixed film and allow it to
act for 1 min.
• By means of a pipette and rubber teat, add double the volume
of distilled water to the slide, mixing the fluids alternately
sucking them in the pipette and expelling them.
60. Leishman’s stain
• Allow the diluted stain to act for 12 min.
• Flood the slide gently with distilled water , allowing the
preparation to differentiate in the distilled water until the
film appears bright pink – usually about 30 seconds.
• Remove the excess water with blotting paper and dry in air.
62. GIEMSA STAIN
This consists of a number of compounds made by mixing
different proportions of methylene blue and eosin. These have
been designated Azur I, Azur II and Azur II-eosin. It is an
advantage of Giemsa’s stain over Leishman’s that fixation is
by alcohol instead of by undiluted stain which, particularly in
the tropics, may deposit precipitate on the film.
Rapid and slow methods are preferred for staining and multiple
films of malaria blood and tissue aspirates.
1. Fix films in methyl alcohol for 3 min.
2. Stain in a mixture of 1 part stain and 10 parts buffer soution
pH 7.0 for 1 hr.
3. Wash with buffer solution, allowing preparation to
differentiate for about 30 seconds.
4. Blot and allow to dry in air.
This method of staining gives excellent results with thin blood
films for malaria parasites, Schuffner’s dots being well
defined. Trypanosomes are also well demonstrated.
65. STAINING FOR
The large spirochaetes borreliae stain by ordinary method including
Gram’s (giving a negative reaction), Leishman’s and Giemsa’s. The
smaller ones, e.g, treponemes and leptospires, are too thin to be
demonstrated by ordinary stains. They are best observed in
unstained wet films under the dark-ground microscope where
their bright appearance and motility draw attention to them.
If a permanent preparation of small spirochaetes is required, use
may be made of a silver impregnation method which artificially
thickness them with a deposit of silver. The classical method are
Fontana’s (for films) and Levaditi (for sections). Faulkener and
Lille’s modified silver methods has been recommended for
demonstrating leptospires in sections of tissue.
• Treat the film three times, 30 seconds each, with a fixative.
• Wash off the fixative with absolute alcohol and allow the
alcohol to act for 3 min.
• Drain off the excess alcohol and carefully burn off the
remainder until the film is dry.
• Pour on the mordant, heating till steam rises, and allow it to
act for 30 seconds.
• Wash well in distilled water and again dry the slide.
• Treat with ammoniated silver nitrate, heating till steam rises,
for 30 seconds, when the film becomes brown in color.
• Wash well in distilled water, dry and mount in Canada
It is essential that the specimen be mounted in balsam under a
cover-slip before examination, as some immersion oils cause
the film to fade at once.
The spirochaetes are stained brownish-black on a brownish-
69. LACTOPHENOL COTTON
Lactophenol cotton blue stain is used for making a semi
permanent microscopic preparation of fungi. It stains the
fungal cytoplasm and provides a light blue background
against which walls of the hyphae can readily be seen. It
contains four constituents, phenol serves as fungicidal agent,
lactic acid acts as a clearing agent, cotton blue stains the
cytoplasm of fungus and glycerine which gives the semi
permanent preparation. A permanent preparation is made
by incorporating polyvinyl alcohol in place of glycerine.
A drop of lactophenol cotton blue was placed on a clean glass
A small part of fungal colony was taken from Potato Dextrose
Agar (PDA) plate using a sterile needle and it was placed on the
drop of lactophenol cotton blue that had been taken on the glass
slide. The fungal elements were spread with sterile needle.
A coverslip was placed over the specimen taking care to avoid the
formation of air bubbles.
Excess stain was wiped off. The slide was kept for 5-10 min and
was observed first under 10X then under 45X objective.