This document discusses various special staining methods used in bacteriology. It describes acid-fast staining including Ziehl-Neelsen technique. Other staining methods discussed are fluorochrome staining for acid-fast bacilli, spore staining including Schaeffer-Fulton method, capsule staining using positive and negative techniques, flagella staining using Ryu's stain, lipid staining with Burdon's and Holbrook & Anderson methods, and Romanowsky staining techniques like Giemsa stain. The document provides detailed procedures for these special staining methods used to identify different bacterial structures under the microscope.

1. SPECIAL STAINING METHODS IN
BACTERIOLOGY
Dr.P.B.PRAVEENKUMAR
FIRST YEAR POST GRADUATE
DEPARTMENT OF MICROBIOLOGY
THANJAVUR MEDICAL COLLEGE

2. ACID FAST STAINING
• First discovered by PAUL EHRLICH
(1882)
• Modified by ZIEHL-NEELSON
• EHRLICH’S METHOD: Staining was
done using aniline-gentian violet
followed by strong nitric acid
• As the ordinary aniline dyes do not
readily penetrate the tubercle
bacillus this method was modified
• Most widely useful staining for acid
fast bacilli is the ZIEHL-NEELSON
technique

3. PRINCIPLE
• Acid fastness is attributed to the
presence of large quantities of
unsaponifiable wax fraction called
mycolic acid in the cell wall .
• Degree of acid fastness varies in different
bacteria which depends on the content
of mycolic acid
• In this method application of heat helps
the dye to penetrate the tubercle bacillus
• Once stained the tubercle bacilli resist
decolourizing action of acid-alcohol
• Other organisms that get decolourized
are considered non acid fast

4. COMPONENTS OF AFS
• Primary and mordant Staining: strong carbol
fuchsin
• Decolourize with acid alcohol: 3% HCL &
25%H2SO4 or 95% ETHANOL
• Counter stain: methylene blue/Malachite
green
• Acid fast cells-red
• Non acid fast cells-blue

5. PROCEDURE (HOT METHOD)
• Heat fix the smear by gently passing over the
flame(coagulation of proteinaceous material)
• Cover with strong carbol fuchsin (carbol fuchsin +
phenol) for 5 mins
• Allow it to stain for 5 mins. During this period heat the
slide intermittently, don’t allow to dry
• Rinse with water
• Cover with 25% H2SO4 for 10 minutes for
decolourization
• Wash the slide with water
• Cover the slide with methylene blue for 15-20secs
• Wash ,air dry & examine

6. INTERPRETATION
• Mycobacterium TB
appears as long slender,
straight/beaded red
color acid fast bacilli
• Other non acid fast
organisms and
background take up
counter stain and
appear blue

7. MODIFICATIONS
1. KINYOUN’S modification: Cold method where the intermittent heating
is not required.
2. Acid alcohol can be used as decolourizer
3. Malachite green can be used as counter stain
4. Concentration of H2SO4 depends on the acid fastness of the organism
5. Brucella abortus – weakly acid fast (0.5% acetic acid – decolourizer)

8. FLUROCHROME STAINING
• Stain containing a fluorescent dye and
examined under fluorescent microscope after
staining
• AURAMINE – RODAMINE is an example where
AFB appear yellow against a black background
• More rapid, sensitive & accurate compared to
ZIEHL-NEELSON method in identifying AFB ,
because of higher cost & technical complexity
it is less feasible

9. PROCEDURE
• Thin sputum smear that is dried & heat
fixed is taken
• Smear is covered with Auramine phenol
for 10mins
• Wash off with water ,add 1% acid
alcohol and leave to decolorize for 5
minutes
• Wash with water, cover the smear with
0.1% potassium permanganate counter
stain for 15secs
• Wash, air dry examine by fluorescence
microscopy. Tubercle bacilli appears as
yellow luminous rods in dark field

10. STAINING OF VOLUTIN CONTAINING
GRANULES
• Volutin granules also called babes ernst /
metachromatic granules / polar bodies are well
developed granules within the bacterial cytoplasm
• They appear as round refractile bodies in unstained
wet preparations
• They are made up of polyphospate
PRINCIPLE: Granules exhibit metachromatic property
whereas the remaining bacteria take up the contrasting
counter stain
• STAINS USED : ALBERT’S STAIN , NEISSER’S OR PUCH’S
STAIN, PONDER’S STAIN.

11. STAINING OF VOLUTIN CONTAINING
GRANULES (cont.)
• With toludine blue/methylene blue the granules stain metachromatically
and appear reddish purple
ALBERT’S STAIN: Granules in diphtheria bacilli exhibit metachromasia and hence
appear bluish black when stained with toluidine blue present in ALBERTS STAIN
I REAGENT ,while the diphtheria bacillus appears green due to methyl green in
the reagent
ALBERT-LAYBOURN METHOD: (Instead of methyl green, malachite green used)

12. ALBERT STAIN
PROCEDURE:
1. Prepare the film, air dry, heat fix
2. Cover the film with ALBERT’S
STAIN I for 3-5 mins
3. Wash with water and blot dry
4. Add ALBERT’S STAIN II ( ALBERTS
IODINE) for 1 min
5. Wash and blot dry
GRANULES-BLUE BLACK
BACILLI-GREEN

13. SPORE STAINING
• Spores are highly resistant and metabolically inactive forms
• Morphology of bacterial endospores is best observed in
unstained wet films under phase contrast microscope where
they appear as large, refractile, oval bodies
• Different staining methods are available for staining spores
1. Schaeffer & fulton method- malachite green
2. Modified Ziehl-Neelson stain-0.25% H2SO4 (appears red
with blue bacilli), lipid granules also appear red
3. Toluidine blue stain
4. Grams stain- Appears as a clear area within stained bacilli
5. Moeller stain

14. SCHAEFFER FULTON METHOD
MODIFIED BY ASHBY
• Films are air dried and heat fixed
• Place the slide with the smear over a beaker of boiling
water, resting it on the rim with bacterial film on the upper
side
• When within several seconds large droplets have
condensed on the underside of the slide ,flood the smear
with 5% aqueous solution of malachite green and allow to
act for 1min while the water continuous to boil
• Wash the slide with cool water
• Then cover it with 0.5% safranine or 0.05% basic fuschin for
30 seconds
• Rinse it with water and observe under microscope

15. SPORE STAINING (cont.)
• Spores take up malachite green where as the
decolourized vegetative cell takes up the counter stain
and appears red. Lipid granules remain unstained

16. CAPSULE STAINING
• Capsules are the protective substances secreted by the
bacteria around the cell which prevents them against
penetration by toxins
• Capsule staining is more difficult than other types of
differential staining because capsular materials are
water soluble and may be removed easily by vigorous
washing Also called as SLIME LAYER/LOOSE SLIME
• Bacterial smears should not be heated as resultant cell
shrinkage may create a clear zone around the organism
that may be mistaken for a capsule

17. CAPSULE STAINING (cont.)
• Capsule is non ionic , so that the dyes commonly
used will not bind to it. Two dyes, one acidic and
one basic are used to stain the background and
cell wall respectively
METHODS:
1. POSITIVE STAINING: Capsule is stained
2. NEGATIVE STAINING: background is stained
3. McFadyean’s reaction
The best method for staining capsules on bacteria
from cultures is the wet film India ink method

18. POSITIVE STAINING
• Prepare a smear from capsulated
bacteria and allow it to air dry.
SHOULD NEVER BE HEAT FIXED
• Flood the smear with crystal
violet . Allow it to stand for 5-7
minutes
• Wash the smear with 20%CuSO4
solution and blot dry
• Observation: capsule is seen as
light blue in contrast to the deep
purple color of cell

19. NEGATIVE STAINING
(WET FILM METHOD)
• Take a clean glass slide
• Place a large loopful of undiluted India ink on the
slide
• Then add a small quantity of liquid bacterial
culture to India ink & emulsify
• Take a cover slip and place on the India ink &
press it down
• Observe under high power
• Capsule appears as clear halo around the
organism against black background

20. NEGATIVE STAINING
(DRY FILM METHOD)
• Nigrosin, modification of India ink 2% mercurochrome can
be used
• Take a clean glass slide
• Place a small drop of nigrosin close to one end of the slide
• Using a sterile loop, loopful of broth culture is mixed with
nigrosin
• With the edge of second slide make a thin smear
• Air dry and observe
• Capsule appears as clear halo around the organism against
black background
• Occasionally shrinkage spaces may form & give false
appearance as capsules

21. DEMONSTRATION OF FLAGELLA
• Flagella are best demonstrated with electron
microscope /metal shadowed films/films made
with phosphotungstic acid because of their
extreme thinness
• Special staining methods are used by which
flagella is thickened ten folds by superficial
deposition of stain produced by action of
mordant
• Easier and most reliable method for staining
flagella in wet mounts was described by
Heimbrook et al (1989)

22. WET MOUNT FLAGELLAR STAIN
PROCEDURE
• Grow bacteria for 16-24 hrs on a non-inhibitory
medium, eg: tryptic soy agar/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 and cover with cover slip
• After 5-10 minutes apply two drops of Ryu’s stain
and leave it to diffuse into the film
• Examine after 5-15mins

23. RYU’S STAIN
• Ryu’s stain:
1. Crystal violet in
saturated ethanol
solution
2. Mordant solution(tannic
acid, 5%phenol,
Aluminium phosphate)

24. STAINING OF LIPIDS
BURDON’S METHOD:
• Prepare a film, air dry & heat fix
• Cover the entire slide with SUDAN
BLACK stain for 15mins
• Drain excess stain & air dry
• Rinse thoroughly with xylene and blot
dry
• Counter stain with 0.5% safranine or
dilute carbol fuschin for 5-10 seconds,
rinse with water & air dry
Lipid granules-blue black
Bacterial cytoplasm-light pink

25. LIPID STAINING (cont.)
HOLBROOK & ANDERSON METHOD: (Mainly for bacillus species)
• Differential staining technique which combines BURDONS stain for lipid &
ASHBY’S stain for spores
• Prepare a film, air dry & heat fix
• Stain with 5% malachite green for 2 minutes with the slide placed over
boiling water, wash, blot dry.
• Cover the entire slide with SUDAN BLACK stain for 15mins
• Drain excess stain & air dry
• Rinse thoroughly with xylene for 5 secs and blot dry
• Counterstain with 0.5% safranine or dilute carbol fuchsin for 5-10 seconds,
rinse with water & air dry
Lipid granules-blue black
Spores- green
Bacterial cytoplasm-red

26. STAINING OF SPIROCHAETES
• Larger spirochaetes,eg., Borreliae can be seen by
ordinary methods like gram’s stain, leishman’s,
giemsa’s stain
• Smaller spirochetes are too thin hence best
observed in unstained wet films under dark
ground microscopy
• Silver impregnation techniques help for
permanent preparation from film/sections
• FONTANA’S METHOD(film), LEVADITI (tissue
sections)

27. FONTANA’S METHOD
• Treat the film 3 times, 30 seconds each time with fixative
(acetic acid, formalin, distilled water)
• Wash the fixative with absolute alcohol & allow the alcohol to
act for 3 minutes
• Drain the excess alcohol and burn off the remainder till it
becomes dry
• Pour on the mordant (phenol,tannic acid,distilled water) heat
till steam arises ,allow to stand for 30 seconds
• Wash with distilled water and air dry
• Treat with ammoniated silver nitrate heat till steam arises ,
allow to stand for 30 seconds till the film turns brown
• Mount under cover slip & examine
Spirochetes- brown black on brown yellow background

29. ROMANOWSKY STAINING
• Neutral stains
• Original stain was made by dissolving the compound
formed by interaction of watery solutions of eosin and
zinc free methylene blue in methyl alcohol
• IMPARTS REDDISH PURPLE color to the chromatin of
malaria
• Various modifications were formed later which are
easy to use
• Giemsa’s stain, Leishman’s, Wrights, Jenners stain
• Each modifications of romanowsky stain varies
according to the ripening and relative proportion of
eosin and methylene blue

31. GIEMSA STAIN (TO SEE SPIROCHAETES)
• Consists of Azur I, Azur II and Azur II-eosin.
(Lillie’ preparation)
• Fix the film in methyl alcohol for 3 minutes.
• Pour 1 ml stain with 20 ml buffer solution in Petri
dish.
• Place a piece of thin glass rod in dish.
• Place the fixed slide, film side downwards in dish
with one end resting on the rod. (Sufficient stain)
• Over night incubation, wash it, dry and examine.

32. ACRIDINE ORANGE STAINING
• Acridine orange R has a
marked affinity for nucleic
acids
• When cells stained with it
are viewed by UV rays, the
RNA components fluoresce
orange-red and DNA
yellow-green
• Useful in studying growth
of viruses in host cells and
for revealing scanty
bacteria in wet films.

33. ACRIDINE ORANGE STAINING (cont.)
• A drop of 0.01% acridine orange in centre of
slide
• Then add one drop of specimen in the slide.
• Apply cover slip and focus it using UV
fluorescence microscope. (40X objevctive)

34. BIPOLAR STAINING
WAYSON STAINING CARBOL THIONINE STAINING
• Solution A (Basic fuchsin,
methylene blue, ethyl alcohol)
• Solution B (Phenol 5%)
• Stain the smear after filtration
using both solutions one by
one.
• Thionine stock solution
• Phenol
• Distilled water
• Filter and use this stain

35. THANK YOU