Different staining methods

1. STAINING TECHNIQUES
Dr.Sreenadh H

2. INTRODUCTION
• Bacteria – contains clear protoplasm
• Their refractive index is closer to the medium in which they grow
• Difficult to see using ordinary microscope
• So we use special methods of illumination like dark field,phase
contrast,fluorescence,silver impregnation and staining

3. • Staining– important in observation and identification of microbes
• It is done to :-
• Demonstrate microorganisms & human cells in a specimen
• Demonstrate structural details of microbes such as capsule, spores,
flagella,intracellular lipids and polysaccharides, nuclear materials, viral
inclusion bodies, elementary bodies of Rickettsiae and intracellular granules
• To differentiate organisms based on certain properties
• As constituents of culture media as indicators(eg: Chromogenic agar) or
selective inhibitors(eg: Eosin Methylene Blue Agar for GNB)
• Stains used are synthetic coal tar dyes made from aromatic
hydrocarbons

5. • Dye – used as a colouring material in textile industry and
for general purpose.
They are crude compounds.
• Stain - used to colour biological entities under
microscope.
They are purified compounds.

6. CLASSIFICATION
• NATURAL
• Haematoxylin(from logwood tree), crocin(from Saffron crocus).
• SYNTHETIC
• Acidic – Eosin, acid fuchsin, picric acid, nigrosin, malachite green
• Basic – Crystal violet, methyl violet, gentian violet, basic fuchsin, methylene blue
• Neutral – Giemsa (MB + Eosin)

7. • Microbial surface – phosphates and bicarbonate ions – negatively charged
• BASIC STAIN – Coloured cation and colourless anion
attracted to negatively charged surface
-- stain internal components like proteins and nucleic acid as well
Eg: crystal violet,methylene blue, safranin,basic fuchsin
• ACIDIC STAIN – Coloured anion and colourless cation
stains background
Eg: nigrosine,picric acid,eosin,acid fuchsin,india ink
• NEUTRAL STAIN – stain nucleic acids and cytoplasm
Eg:Giesma stain

8. Vital Stains
• Differentiate living cells from dead cells
• Living cells donot take up dye(intact plasma membrane)
• Dead cells take up dye
• Viability of tissue assessed by measuring the % of cells that donot take up the
dye

9. • SUPRAVITAL STAINING – Staining the tissues that are removed from an
organism during life or immediately after death.
• Crystal violet
• Methyl violet
• Janus green B
• Brilliant cresyl blue
• Nile blue
• Toluidine blue
• Janus Green B - Basic dye – stain Mitochondria
O2 present – indicator turns blue
O2 absent – indicator turns pink

10. • INTRAVITAL STAINING – Process of staining the tissues by injecting stain
into the body in the living state and removing that tissue after staining.
Eg: Hoechst stain(nuclear stain)
Isolectin (cytoplasm stain)
Maleimide (matrix stain)
Methylene blue

11. Methylene Blue Intravital Staining

12. SIMPLE STAINING
• Single stain is used
• So same colour to all structures
• For bacterial morphology, arrangement and nature of cells
• For bipolar staining, demonstration of granules
• Eg:- Methylene blue, Dillute carbol fuchsin.

13. DIFFERENTIAL STAINING
• Distinguishing bacteria into different groups based on the structural differences
among different bacterial species/genus
• Application of 2 stains separately to heat fixed smears

14. SPECIAL STAINING
• Demonstration of flagella, spores, capsules, metachromatic granules,
Colonies of Mycoplasma
• Demonstrating Spirochetes, Rickettsiae, Parasites, Fungus, Viral inclusion
bodies, Intracellular lipid, Cell Polysaccharides.

15. MORDANTS
• Facilitates staining
• An organism can be stained when a chemical is added before(pre
mordanting/onchrome), during(meta mordanting/metachrome) or after(post
mordanting/afterchrome) the application of stain
• Chemical used – MORDANTS
• Basic mordants react with acidic dyes and vice versa
• Eg:- Iodine in Gram’s staining(as afterchrome)
Phenol in Ziehl Neelsen staining(as onchrome)

16. ACCENTUATION
• Making a reaction rapid, intensive and selective by adding a compound
which doesn’t react with stain but make it mature/ripened.
• Eg:- KOH in Loeffler’s methylene blue
K2CO3 in Polychrome Methylene blue
PROGRESSIVE STAINING
• Several stains applied in a sequence for specified time without use of
decolouriser.
• Eg:- Giemsa stain

17. REGRESSIVE STAINING/DIFFERENTIAL STAINING
• All constituents of a smear are stained followed by selective removal of
excess stain by a process called differentiation or decolourisation.
• For basic stain, acidic decolouriser is used
• Eg:- Gram’s staining,Ziehl Neelsen staining.

18. Points to be considered in making staining solutions
• Stains should bear certification label of biological stain
• All dyes should be weighed accurately
• Dyes – ground in mortar and diluent to be added in small amounts till all dyes
dissolved
• Stain to be filtered before use. If allowed to stand for >24hrs – not needed
• If precipitation occurred – not to be used
• Stock solution can be kept for a long time.So diluted solution should be made
in small quantities
• Label the concentration of dye and preparation date
• Keep stains in glass stoppered bottles away from sunlight.

20. FIXATION
• After air drying– heat fixation
• Passing slide rapidly 2- 3 times through the flame till its just hot on the back
of the hand
• Methanol/ ethanol fixation is more bactericidal(used in Dry India Ink
staining,Giemsa staining)
• Cell bodies are better preserved esp pus cells
• Better method for intracellular diplococci like Gonococci and Meningococci
• Other fixatives
• 40gm/L KMnO4(potassium permanganate) for Bacillus anthracis in Polychrome MB Stain.
• Formaldehyde vapour for mycobacteria
• Such smears stain poorly
• Purpose – preserve microorganism and to prevent smear from being washed
away

21. SIMPLE STAINING
• LOEFFLER’S METHYLENE BLUE
• Most useful
• Characteristic morphology of Polymorphs and Lymphocytes
• Content – Saturated solution of Methylene blue in alcohol – 300ml
KOH, 0.01% in water – 1000ml
• Stain smear with MB for 3 mins
• Do not readily overstain

23. • POLYCHROME METHYLENE BLUE
• Slow ripening(>12 months) of Loeffler’s Methylene blue
• Kept half filled in bottles – intermittent shaking for aeration – slow
oxidation – Violet compound – Polychrome properties
• Quick ripening – 1%K2CO3.
• Acidic structure – Red purple
• Eg:- McFadyean reaction for Anthrax
• Gurr’s polychrome methylene blue McFadyean stain
(readymade stain)

24. • McFadyean reaction
• Fairly thick smear of blood, exudate or tissue fluid
• Dry in air
• Fix imperfectly by quickly passing through flames 3 times
• Stain with Polychrome methylene blue for 30 sec
• Wash and dry
• Irregular pink purple capsular material,surrounding the bacilli and chain of
bacilli -- diagnostic

25. • In-vitro Mcfadyean reaction
• Convenient and inexpensive when large no of isolates have to be tested
• Inoculate material from a 24 hr colony or subculture into 2-3 ml of sterile
heparinised or defibrinated horse or sheep blood.
• Incubate at 6-8 hrs at 37°C
• Thick smear made
• Fix and stain it as before
• Blue stained bacilli with pink purple capsular material

27. • BUFFERED METHYLENE BLUE
• It is for stool examination for Trophozoites.
• Shows nuclear details
• Nucleus and inclusion bodies – dark blue eg: Heinz bodies in G6PD deficiency
• Cytoplasm light blue
• Methylene blue 0.06% in an acetate buffer at pH 3.6

28. Heinz bodies in Buffered methylene blue

29. Borax Methylene Blue
• Similar to Polychrome methylene blue
• Methylene Blue - 20gm
• Borax - 50gm
• Water -1000ml
• Used in spectrophotometric studies
• Used as a counter stain in Modified Leifson’s method for flagellar staining.

30. • Dilute Carbol Fuchsin
• Diluting ZN stain with 10-20 times its volume of water
• Stain the smear for 10-25 sec
• Overstaining to be avoided as it may stain protoplasm along with nuclei and
bacteria.

32. DIFFERENTIAL STAINING
•GRAM‘S STAINING
• It is the most common staining method used in Microbiology
• It differentiates bacteria into 2 groups
• GRAM POSITIVE
• GRAM NEGATIVE

33. HISTORY
• Devised by Danish Physicist –
HANS CHRISTIAN JOACHIM
GRAM
• In 1883
• Published his work in 1884
• Studied on lung tissues of
patients who had died of
pneumonia– distinguished
Klebsiella from Streptococcus
pneumoniae.

34. • CARL WEIGERT
• German pathologist from
Frankfurt
• Added final step of staining with
• Carmalum solution

35. • Original formulation used
• Aniline gentian violet
• Lugol’s iodine(iodine 1gm + KI 2gm + H2O 300ml)
• Absolute alcohol
• Bismarck brown

36. • Culture of Gram positive species of aged culture – may contain Gram negative
species – not vice versa
• Gram positive may appear Gram negative – in antibiotic treated cases– due
to cell wall damage.
• Tissue cells, leukocytes, and debris in inflammatory exudates – Gram
negative.

38. MECHANISM

39. Modification Violet dye Iodine Decolourizer Counter stain
KOPELOFF-
BEERMAN
1922 –General use
(modificationof
Burke’s)
Sol A – Methyl violet + DW
Sol B – NaHCO3 +DW
30V A+8V B
5 min
Iodine + 4%
NaOH +DW
2 min
Acetone
2-3 sec
Basic fuchsin
30 sec
BURKE’S
(General use)
1% aqueous MV + 5drops of 5%
NaHCO3
3min
Wash with
Iodine + KI +
DW
Cover with
Fresh Iodine
1-2 min
Acetone
1-2 sec
Safranin or
Neutral red
>/ 10 sec
JENSEN’S
(Gonococci &
Meningococci)
0.5% Methyl violet
30 sec
Lugol’s Iodine
30 sec
95- 100 % alcohol
30 sec
Neutral red
2 min
WEIGERT’S
(for tissues first
counter stain with
Carmalum solu: for
10min
Carbol gentian violet
2-3 min
Gram’s iodine
1 min
Aniline xylol
Till stain ceases to
remove (examine in
low power for nuclei
of pus cells,should
be pale violet.
Dil carbol
fuchsin
10-25 sec

40. Modification Violet dye Iodine Decolourizer Counter stain
Hucker’s
1920
Aniline + AA +
Gentian violet
45 sec
I+ KI
1 min
Aniline oil + Xylol +
Alchohol
(95%)
Until no more violet
comes
Bismarck Brown +
Alchohol (95%)
45 sec
Claudius
1897
(for Fungus in
tissue section)
1% aqueous Methyl
violet
1-2 min
Chloroform or
aniline + 0.1 %
picric acid
Until no more violet
comes
wash with half
saturated solution of
picric acid (0.6%) then
cover with fresh picric
acid for
1-2 min then rinse with
xylol and mount in
Canada balsam
Preston &
Morrell’s
1962
Ammonium oxalate
CV
30 sec
Lugol’s iodine
(1%)
30 sec
Iodine acetone
decolouriser
30 sec
Dilute carbol fuchsin
30 sec

41. Modification Violet Dye Iodine Decolourizer Counter Stain
Brown and Brenn
(for tissues
containing
Nocardia,
Actinomyces,
Microsporidia)
Crystal violet
1 min
Lugol’s Iodine
1 min
Aetone
2-3 sec
Basic Fuchsin
3min
Again decolourize
by dipping into
acetone- 2 dips
then
Dip into Picric acid
acetone mixture
until salmon colour
then again
decolourize with
acetone.Air dry and
dip into xylene and
put coverslip.

44. QUICK GRAM STAIN FOR SINGLE SLIDES
• Held the slide with forceps.
• Flood with crystal violet or methyl violet and allow to act for 5secs
• Tip off stain then flood the slide with iodine solution for 5secs
• Tip off iodine then flood with acetone for 2secs
• Flood with Basic fuchsin for 5secs.

45. GRAM METHOD FOR MULTIPLE SLIDES
• Place the slides side by side in a staining rack over the sink
• Cover each slide with methyl violet for 30secs
• Wash then cover each slide with lugol’s iodine 1% for 30secs
• Wash then cover each slide with iodine acetone for 30secs
• Wash then cover each slide with basic fuchsin for 30secs.

46. • Reporting of Gram’s staining
• Intra/extrcellular
• Shape – cocci/bacilli/coccobacilli
• Colour – Gram positive or Gram negative
• Arrangement – scattered/clusters/pairs/chains
• Accompanying cells – Pus cells/ epithelial cells/ yeast cells
• Disadvantages
• Thick smear– gram negative organism appears as gram positive
• Integrity of cell wall disrupted

47. • GP anaerobes easily decolourise
• Overdecolourisation with acetone
• To confirm Gram staining results
• KOH test
• 2 drops of 3% KOH on slide – emulsify a loopful of colony – loop
raised 1-2 cm from surface
• GN – Cell wall broken –release of DNA – mucoid solution –
STRING formation
• L alanine para nitroanilide
• GN bacteria – aminopeptidase enzyme – hydrolyses above said
substrate – nitroalanine – Yellow coloured
• Commercial disk available – added to broth –disk to yellow colour

49. ZIEHL NEELSEN STAINING
• Modification of Ehrlich’s(1882) original method
• With Aniline Gentian violet and strong HNO3
• Ordinary aniline dye – does not penetrate the bacilli
• As Mycobacterial cell wall contains Mycolic acid and arabinogalactan
moeities
• So powerful staining solutions containing Phenol or application of heat –
facilitate penetration of dye
• Upon staining– bacilli withstand decolourisation for a considerable time

51. • REAGENTS
• ZN carbol fuchsin
• Basic fuchsin 5g
• Phenol 25g
• 95%/100% alcohol 50ml
• Distilled water 500ml
• H2SO4(20%)
• Conc H2SO4 (98%) 250ml
• Distilled water 1 litre
• Alcohol 95%

52. • Acid alcohol decolouriser
• Conc. HCl 75ml(3%)
• Methylated spirit 2425ml
• Methylene blue( Loeffler’s Methylene Blue)
• Distilled water 100ml
• KOH 0.1 ml
• Saturated solution of MB 30ml

53. • Malachite green
• Malachite green 5g
• Distilled water 500ml
• 1% stock solution – diluted to make working solution
1% stock solution 40ml +
Distilled water 360ml
• Pale green background –is formed which is used for method which uses deep blue green
filter for easy recognition of tubercle bacilli

54. • Use of alcohol for secondary decolourisation
• 95% alcohol used
• Tubercle bacilli – Acid and alcohol fast
• Advantages :- quicker decolourisation
Margins and underside of slide more
cleaner
• Certain bacilli acid fast but not alcohol fast – smegma
bacillus – can be confused with tubercle bacilli in urine
• Acid alcohol as decolouriser
• 3% HCl in 95% alcohol
• Less corrosive and more convenient to make
• Expensive

55. • Prepare smear, dry it and heat fix
• Stain with filtered concentrated carbol fuchsin
• Heat untill steam arises x 5 min with intermittent heating
• Donot allow stain to dry
• Wash both sides of slides
• Decolourise with 20% H2SO4( Yellowish brown colour) x 1 min
• Wash several times and repeat the steps for atleast 10 min
• Methylene blue or Malachite green counterstain for 3 minutes
• If tissue or background appear red, no adequate decolourisation.

58. • PRECAUTIONS
• Should not be done in staining jars
• As AFB from positive smear may get detached to stain/decolouriser and adhere to
subsequent negative smears
• Use fresh, clean blotting paper to avoid False positive results
• If large sheets used – AFB from positive smears adhere to blotting paper and get
transferred to negative smears
• Use fresh distilled water to prepare reagents
• As tap water may contain AFB

59. • ZN FOR WEAKLY ACID FAST ORGANISM
• M leprae – 5% H2SO4
• Actinomycetes, nocardia and Mycobacterium – 1% H2SO4
• Brucella differential staining(for Brucella abortus)
• Stain with dilute carbol fuchsin without heating for 15 minutes.
• Decolourise with 0.5% acetic acid for 15 sec
• Counterstain with Loeffler’s methylene blue for 1 min
• Spores – 0.25 – 0.5%
Excellent results – if 2% HNO3 in absolute alcohol used as
decolourizer

60. • 1% H2SO4 – Cryptosporidium parvum
Cyclospora cayetanensis
Cystoisospora belli
Microsporidia
Eggs of T.saginata
Hooklets of hydatid cyst
Miracidium of S.mansoni
• 0.5 -1% H2SO4 – sperm head
Legionella micdadei

61. • KINYOUN’S COLD ACID FAST STAINING
• No heating
• Phenol concentration in CF is increased
• Carbol fuchsin stain – 5 min
• Basic fuchsin 4g Solution A
• Ethyl alcohol(95%) 20ml
• Distilled water 100ml Solution B
• Liquid phenol 8 g
• Decolouriser – 3-5 secs
• 3ml conc HCl + 97 ml of 95% ethanol
• Counterstain – 3 min
• Methylene blue

63. Modification Dye Decolouriser Counter stain
Muller Chermock
( in Veterinary
use)
Carbol fuchsin +
tergitol
2 min
No heating
Acid – alcohol
Till colourless
Methylene blue
5- 15 sec
Cooper
(in tissues)
Cooper Carbol
fuchsin
3-5 min
5% HNO3 in 95%
alcohol
Cooper Brilliant
Green
>30 sec
Gabbett Carbol fuchsin
10 min
No heating
Here 2 and 3rd
step is done as
single step
Gabbet’s MB
(it contains
H2SO4)
2 min

64. Gabbett’s Methylene Blue
Methylene Blue – 1gm
H2SO4 - 20ml
Absolute alchohol - 30ml
Distilled H2O - 1000ml

65. SPECIAL STAINING
STAINING OF GRANULES
• Volutin/metachromatic granules – accumulation of inorganic
polyphosphates
• Well developed granules – as round refractile bodies – in unstained wet
preparations
• Basic dyes – stain more strongly
• Toluidene blue/methylene blue – metachromatic (reddish-purple)
• Corynebacterium diphtheriae – volutin staining – (young cultures)18-24 hr old
colonies on BA or serum medium

66. • METHYLENE BLUE
• Stain the dry heat fixed smear with
Methylene blue for 1-2 min
• Granules -- deep blue
• Body -- lighter blue

67. • ORIGINAL ALBERT’S METHOD
• Stain
• Toluidene blue 1.5g
• Methyl green 2g
• Glacial acetic acid 10ml
• Alcohol(95%) 20ml
• Distilled water 1 litre
• Dissolve dyes in alcohol – add to H2O and acetic acid – filter after 24hrs.
• Albert’s iodine
• KI 3g
• Iodine 2g
• Distilled water 300ml

68. • Dry and heat fix smears
• Stain with Albert’s stain for 5minutes
• Drain,but do not wash
• Use Albert’s iodine for 1 min
• Wash
• Dry and examine
• Granules –dark green to black
• Body – light green

69. • ALBERT-LAYBOURN METHOD
• Laybourn modification (1924)
• Malachite green instead of methyl green
• Staining solution – Malachite green instead of methyl green
• Albert’s iodine
• KI 9g
• Iodine 6g
• Distilled water 900ml

70. • Cover slide with Albert’s stain x
3-5 min
• Wash with water
• Cover the slide with Albert’s
iodine x 1min
• Wash
• Granules – bluish black
• Protoplasm – green
• Other organisms – light green

71. • NEISSER’S STAIN
• Stain
• Methylene blue 1g
• Absolute alcohol 50 ml
• Glacial acetic acid 50 ml
• Distilled water 1000ml
• Neisser’s Bismark brown
• Bismark brown Y 0.2g
• Distilled water 100ml

72. • Stain with Neisser’s stain x 30-
60 sec
• Wash
• Bismark brown x 10sec
• Wash
• Granules – Bluish black
• Protoplasm -- Brown

73. Modified Neisser’s Method
• Stain with Neisser’s MB for 3min
• Wash off and cover with Iodine solu: (Kopeloff & Beerman’s Iodine) and
for 1min
• Counterstain with Neutral Red for 3min
• Deep blue Granules and Pink protoplasm.

75. • PONDER’S STAIN
• Stain
• Toluidene blue 0.02gm
• Acetic acid 1ml
• Alcohol 2ml
• Distilled water 100ml
• Dissolve dyes in alcohol
• Mix acid with water
• Mix both

76. • Stain with Ponder’s stain for 3min
• Wash with dilute iodine solution
• Wash with water
• Counterstain with Neutral red x 3min
• Wash
• Granules – reddish purple
• Protoplasm – light blue

77. • GOHAR STAINING
• Stain smear with methylene blue x 5min
• Decolourise with 1:1000 H2SO4
• Rapidly wash
• Counterstain with Eosin 1% in distilled water x 30sec
• Wash and dry
• Granules – black
• Body -- yellow

78. • LJUBINSKY STAINING
• Stain
• Crystal violet 0.25gm
• Glacial acetic acid 5ml
• Distilled water 95ml
• Stain with Ljubinsky stain x 2min
• Wash
• Counterstain with Bismark brown x 30 sec
• Wash
• Granules – deep purple
• Bodies -- Brown

79. STAIN GRANULES PROTOPLASM
Methylene blue Deep blue Lighter blue
Original Albert’s Dark green to black Light green
Albert-Laybourn Bluish black Green
Neisser’s Bluish black Brown
Gohar Black Yellow
Ljubinsky Deep purple Brown
Ponder’s Reddish purple Light blue

80. CAPSULAR STAINING
• Capsules of organism in animal tissue, blood, serous fluids and pus – stained
by common stains such as Gram’s stain, Basic fuchsin, Polychrome
methylene blue.
• Artificial media – special stains –Relief or negative staining
eg: Wet India Ink method
Dry films by India ink, Nigrosin, Eosin.
• Best method – India ink wet film

81. • BURRI’S INDIAN INK WET FILM METHOD
• Adv :- capsules don’t shrink as they are not dried or heat fixed
• India ink – must be dense,homogenous , very finely granular and free from
contaminants with capsulated and non-capsulated bacteria

82. • Wipe a slide
• Add a large loopful of India ink
• Emulsify small portion of solid culture/loopful of liquid culture
• Place clean grit free coverslip on ink drop and press it down through a sheet
of blotting paper – thin and pale film
• Observe under OIF – highly refractile bacterium
Between refracile surface and dark background – clear
space – capsule.
• Loose slime – irregular strands lighter than ink – gradually disperse from
bacteria and dissolve in the ink.

84. • Phase contrast microscope –ideal for wet India ink films where bodies of
bacteria appear dark and in clear contrast to bright capsular zone
• DRY INDIA INK STAINING(BUTT et al)
• Loopful of 6% glucose in H20 – placed on one end of a clean slide
• Small portion of colony mixed with it
• Loopful of india ink added to it
• Mixture spread to make a thin film
• Fix with Methanol or undiluted Leishman stain
• Drain and warm over flame
• Pour Methyl violet and keep for 1-2 min
• Wash, dry and examine under OIF

85. CAPSULE STAINING BY NIGROSIN
• To a loopful of CSF or a light aqueous or saline suspension of growth
from an agar culture
• To this add a loopful of Nigrosin stain
Ingredients: Nigrosin 10gm
Formalin 0.5ml(0.5%)
DW 100ml
• Mix well and cover with a cover slip
• Examine under high power and then oif.

87. RELIEF STAINING WITH EOSIN
Staining solution: 10% H2O soluble Eosin 4parts + Serum 1part + Crystal of
thymol
Allow the mixture to stand at room temp for several days, centrifuge and store
the supernatant.
• On a slide with 1mm diameter mix 1 drop of exudate/fluid culture/suspension
in broth with 1 drop of ZN Carbol fuchsin and allow to stain for 30secs.
• Then add 1 drop of eosin solution and leave for 1min.
• Spread the film with another glass slide.
• Examine under oif after drying(do not heat).
• Red background and organism with unstained capsule.

88. • HISS’S CAPSULAR STAINING
• Capsule of B.anthracis – Polyprotein – doesn’t bind to protein stains like
crystal violet
• To demonstrate boundary of capsule – bacteria suspended in Proteinaceous
fluid like milk and serum
• Stain
• 5ml saturated alcoholic solution of Gentian violet in 95 ml distilled water

89. • Mix material to be stained with equal amount of serum
• Make smear and dry
• Fix in flame/ 1:10 dil commercial formalin
• Apply Hiss stain and heat till steaming for few seconds
• Wash with 20% watery solution of CuSO4
• Dry in vertical position
• Bacterial bodies, cells and background – Purple
• Capsule -- colourless

91. • MACNEAL CAPSULAR STAINING
• Stain
• Eosin Y, certified 1gm
• Methylene blue, certified 1gm
• Methylene violet 0.2gm
• Methanol, neutral & acetone free 1000ml
• Mix and heat to 500 C x 30 min – keep at 37 0 C x 2days -- filter

92. • Make a thin smear
• Dry in air
• Stain with MacNeal tetrachrome stain x 3-5min
• Wash and dry
• Capsule – colourless and pale

93. • LAWSON CAPSULAR STAINING
• Mordant
• 5% solution of Phosphomolybdic acid
• Stain
• Wright stain 10ml
• Glycerin 5ml
• Add glycerin to wright stain just before use.

94. • Prepare direct smear from culture
• Dry in air
• Add mordant x 30 sec
• Wash with methyl alcohol
• Cover with 20 drops stain x 2 min
• Add 30 drops of distilled water, mix stain in slide with water and allow to act x
10-20 sec
• Rinse with distilled water
• Drain and examine
• Capsules – colourless and pale

95. STAINING OF SPORES
• Best observed in unstained wet films under phase contrast m/s – large,
refractile, oval/spherical bodies within bacteria or free
• Ordinary stains – Body deeply stained, spore unstained
• Modified ZN – Spore red, bacteria blue

96. ACID FAST STAIN FOR SPORES
• Films are made in usual manner and fixed by air drying
• Stain with ZN Carbol Fuchsin for 3-5 min
• Wash with H2O
• Treat with 0.25% / 0.5% H2SO4 for several minutes
• Another better and convenient method of decolourisation is by dipping the
slide in 2% solution of HNO3 & Absolute Alchohol and washing immediately.
• Counterstain with 1% aqueous MB for 3min
• Spore stain bright red and protoplasm blue

97. • An alternative to this is counter staining the slide with Nigrosin(spreading the
in the smear with another slide) instead of MB.
• Bright red spores, unstained protoplasm and dark background.

99. • MALACHITE GREEN STAIN
• Schaeffer and Fulton method, modified by Ashby, 1938
• Films are dried and fixed with minimal flaming
• Slide kept over a beaker of boiling water with film uppermost
• Large droplets condenses on underside of slide
• Flood with 5% aqueous Malachite green x 1min while the water continues to
boil
• Wash in cold water
• Treat with 0.5% safranine or 0.05% basic fuchsin x 30 sec
• Wash and dry
• Spores – green , bacilli – red , lipid granules- unstained

101. • DORNER SPORE STAINING
• DORNER’S SOLUTION
• Nigrosin 10gm
• Distilled water 100ml
• Boil for 30min in an Erlenmeyer flask(conical flask)
• Add preservative 0.5ml formalin(40%)
• Filter twice
• Stopper tubes with aluminium foiled corks
• To stain background

102. • Heavy suspension of organism in 3-4
drops of distilled water in test tubes.
• Stain with 3-4 drops of ZN carbol
fuchsin in test tubes.
• Boil the tube in water bath for 10-15
min.
• Loopful of preparation on slide and heat
fix, add a drop of Dorner solution.
• Make a thin smear
• Exmine under oif
• Spores – red
• Vegetative cells – unstained
• Background -- grey

103. • WIRTZ – CONKLIN SPORE STAINING
• Make a thin smear and heat fix
• Stain with 5% aqueous malachite green and steam for 3 -6 min
• Wash with distilled water
• Counterstain either with 0.5% aqueous safranine or 0.5% aqueous
mercurochrome x 30-60 sec
• Dry and examine under oif
• Spores – green
• Body of bacteria – pink
• Useful to detect early germinating spores within 30 min of incubation in a
germinating environment.

104. • ABBOTT SPORE STAINING
• Make a thin smear and flame fix
• Stain with methylene blue and heat to boil, but not continuosly x 1min
• Wash
• Cover with 1 part saturated alcoholic eosin Y solution and 9 parts distilled
water x 5-10 min
• Wash and blot
• Spore – blue
• Body -- pink

106. SPORE STAINING SPORES VEGETATIVE
CELLS/BACTERIA
Ordinary Stain Unstained Colour according to stain
Modified ZN Red Blue
Malachite Green Green Red
Dorner’s Staining Red Unstained
Wirtz-Conklin Staining Green Pink
Abbott Spore Staining Blue Pink

107. FLAGELLAR STAINING
• Flagella is 0.02μm thick
• Best demonstrated using electron microscopy by negatively stained films with
phosphotungstic acid.
• To be resolvable for light microscope – must be thickened at least 10x –
superficial deposition of special stain procured by action of mordant usually
tannic acid.

108. • MODIFIED LEIFSON’S METHOD
• Good results depend on cleaning and flaming of slides
• Method of cleaning slides
• Clean slide with absolute alcohol wiping with fine cotton
• Put in conc.H2SO4 saturated with potassium dichromate solution for several
days at RT.
• Do not touch even the ends of slide with bare hands
• Using forceps – transfer slides to a clean Coplin jar – for rinsing, drying and
storing– don’t overcrowd
• Rinse with Tap water followed by distilled water
• Drain and dry in the air with jar inverted on clean blotting paper
• Store with the jar closed

109. • Before use – flame the slide for few seconds
• Place it on a clean metal rack and cool
• For quick use – store slides in solution for 1 hr @ 900 C in a strong metal
container with sand while heating
• Dichromate – H2SO4 cleaning solution
• 63 gms of Na/K dichromate by heating in 35ml of water
• Cool and add conc H2SO4 to 1 litre

110. • Precautions
• Use rubber glove
• Splash – wash with water
• Residual acid – Na2CO3 solution – wash off with H2O
• Never pour water to acid – boiling and spurting
• Add acid to water slowly in repeated small volumes

111. • PROCEDURE
• Fix broth culture or saline suspension of organism in test tube in 1-2 %
formalin
• Sediment bacilli by centrifuging at 2000-3000rpm
• Discard supernatant and gently resuspend bacilli in distilled water by rotating
tube in opposite direction rolling in between palms of hand
• Again centrifuge and resuspend in distilled water
• Final suspension – slightly cloudy
• Place a loopful onto slide – spread over an area of 1-2 cm diameter
• Dry in air at room temp– donot fix

112. • Stain
• Tannic acid 10g
• NaCl 5g
• Basic fuchsin 4g
• Thouroughly mix powdered ingredients in a mortar
• Store in stoppered container
• 1.9 g of powdered mix + 33ml of 95% ethanol
• When mostly dissolved – add DW to make 100ml
• Adjust pH – using NaOH/HCl
• Store in stoppered bottle in refrigerator @ 3-5 0 C

113. • Staining of smear
• Slide placed horizontally on levelled staining rack
• Pippette 1ml stain onto slide
• Leave at RT
• Several smears should be stained for different times – 6,8,10,12 mins –
inorder to choose best one(apparent thickness of flagella increase with
duration of stain)
• Rinse by gently placing under a slowly running water
• Don’t pour of the stain before rinsing
• Conterstain with Borax methylene blue for 30min – colour protoplast
• Wash with water, dry in air, examine under OI

115. • WET MOUNT FLAGELLAR STAIN
• Mordant solution
• Phenol 5% w/v aqueous solution 10ml
• Tannic acid 2g
• Aluminium potassium sulphate,saturated aqueous solution: 10ml
• Ryu’s stain
• Crystal violet, saturated ethanolic solution(12g/100ml) : 10ml
• Mordant solution 100ml
• Filter the stain and keep in syringe with 0.22μ pore size porous membrane b/w
syringe and needle and needle stuck in rubber stopper – prevent drying
• Remain stable for several weeks at ambient temperature

116. • Procedure
• Grow bacteria on non inhibitory medium(eg: blood agar) for 16-24 hrs
• Touch loopful of water onto edge of colony and let motile bacteria swim into it
• Transfer that loopful to a loopful of water on a slide – faintly turbid – cover
with coverslip
• Bacterial suspension made with minimal agitation only – flagella would detach
• After 5-10 min, apply 2 drops of Ryu’s stain to edge of coverslip and leave to
diffuse
• Examine after 5-15 min in oif

118. • SILVER STAIN FOR FLAGELLA
• Solution A (Mordant/ prestain)
• Saturated aqueous aluminium phosphate 25ml
• 10% aqueous tannic acid 50ml
• 5% aqueous ferric chloride 5ml
• Mixture is black – stored in dark at 50 C
• Solution B (silver stain)
• 5% silver nitrate and conc NH4OH – 2-4 ml
• Prepare by slowly adding conc NH4OH to silver nitrate
• Brown ppt – more stain added – dissolves
• Stop just as solution clears
• Reverse – add 5% AgNO3 one drop at a time untill solution becomes faintly cloudy
• Store at 50C in dark

119. • Procedure
• Smear prepared from 18-24 hr old culture from TSA slant
• Light suspension of organism in 3ml DW
• Large loop – spread on the slide
• Slides – staining rack
• Flood with solution A for 4min
• Wash
• Flood with solution B
• Heat till steam rises for 4min
• Examine under OI

121. • GRAY STAIN FOR FLAGELLA
• Solution A (mordant)
• Potash alum saturated solution of 20% aqueous tannic acid
• Mercuric chloride saturated solution
• Solution B
• Methyl alcohol, basic fuchsin in DW ( ZN Carbol Fuchsin)
• Mix alcoholic carbol fuchsin with mordant each time before use

122. • Procedure
• Place 1 drop of DW on a clean slide
• Select part of young colony and touch gently on drops of water
• Gently rotate
• Air dry
• Donot fix
• Add mordant
• Keep for 10min
• Wash with DW
• Add ZN Carbol fuchsin for 10 min
• Wash and dry – examine under OI

124. • RHODE’S METHOD
• Flood smear with iron tannate mordant
• Incubate at room temperature for 3-5 min
• Rinse with water
• Flood the slide with hot ammoniacal AgNO3 solution
• Leave to act for 3-5 min
• Rinse thoroughly with water
• Blot dry

125. STAINING TO DEMONSTRATE CELL WALL
• Bouin’s fluid
• Saturated aqueous solution of Picric acid, Formalin and glacial acetic acid
• Make impression preparation fixed in Bouin’s fluid
• Mordant with 5- 10% tannic acid for 20-30 min
• Stain with crystal violet 0.02% for 1min
• Cell wall – purple
• Bouin’s fluid – This fixative is useful for the investigation of viral inclusion
bodies.

126. STAINING FOR COLONIES/DIENE’S STAIN
• To differentiate Mycoplasma colony from an artefact or other bacterial colonies on a plate
• Stain
• Methylene blue 2.5gm
• Azure II 1.25gm
• Maltose 10gm
• Na2CO3 0.25gm
• Distilled water 100ml
• Flood agar containing Mycoplasma with 1ml of Diene’s stain working solution.
• Immediately rinse agar surface with DW to remove stain.
• Decolourise with 1ml 95% ethanol for 1min.
• Wash with DW and allow to dry.
• Observe the medium for colonies under low and then high power.
• Dark blue if Mycoplasma colonies present.

128. STAINING TECHNIQUES
Dr.Sreenadh H

129. STAINING OF INTRACELLULAR LIPIDS
1, Burdon’s (1946) Method
• Sudan black stain
Sudan black B powder 0.3gm
70% Ethanol – 100ml
Shake thoroughly at intervals, stand overnight, keep in
stoppered bottles.

130. Procedure
• Make film, dry in air and fix by flaming.
• Cover slide with Sudan black for 15min
• Drain off excess stain,blot,dry in air
• Rinse thoroughly with xylene and again blot dry
• Counter stain with Safranine or Dilute carbol fuchsin for
• 5-10min
• Lipid inclusion granules are stained blue black or blue grey while bacterial
cytoplasm is stained light pink.

132. 2, LIPID/SPORE STAIN(Holbrook & Anderson in 1980)
• This method combines Burdon’s Staining for lipids and Ashby’s staining
for spores.
Procedure
• Prepare film from centre of a 1day old colony or from edge of a 2 day old
colony.
• Air dry and heat fix.
• Stain with Malachite green for 2min while the slide is held a little above the
surface of boiling H2O in a beaker.
• Wash, stain with Sudan black B in ethanol for 15min.
• Wash with xylene for 5secs and blot dry.

133. • Counterstain with 0.5% safranine for 20secs.
• Wash with water and examine.
Lipid stain Black
Spore stain Green
Cytoplasm stain Red
Useful in studying morphology of Bacillus cereus and related
Bacillus species.

134. STAINING FOR INDIVIDUAL BACILLI
• For Yersinia Pestis (WAYSON STAIN)
• Rapid method
• Yersinia pestis ( bipolar staining)
• Stain with methylene blue in 95% ethanol and 5% phenol x 10-20 min
• Wash and observe under OI
• Blue coccobacilli with pink ends

136. • For Spirochetes
• Larger ones (Borrelia) – ordinary stains
Gm –ve, Leishman’s stain, Geimsa’s stain.
• Smaller ones (Treponemes and leptospires) – Unstained wet film under dark
ground microscope – Bright appearance and motile.
• Permanent preparation – silver impregnation method

138. SILVER IMPREGNATION METHODS
1, FONTANA’S METHOD FOR FILMS
• Fixative
• Acetic acid 1ml
• 40% formaldehyde 20ml
• Distilled water 100ml
• Mordant
• Phenol 1g
• Tannic acid 5g
• Distilled water 100ml
• Ammoniacal AgNO3
• 10% NH3
• 0.5% AgNO3
• Distilled water

139. • Treat film 3 times with fixative – 30 sec each
• Wash with absolute alcohol and allow to act for 3 min
• Drain off excess alcohol, burn off remaining until film is dry
• Pour on mordant, heat till steam rises, allow to act for 30 sec
• Wash with distilled water and dry
• Treat with Ammoniacal AgNO3, heat till steam rises x 30sec
• Film becomes brown
• Wash well, dry and mount in Canada balsam

140. Brownish black spirochetes in Brownish yellow background

141. 2, MODIFIED BECKER’S METHOD FOR FILMS
• The fixative and mordant are as in Fotana’s method.
• Stain
Basic fuchsin in saturated alchohol solu: +
Shunk’s mordant B +
Ethanol + Aniline oil
DW

142. Procedure
• Filter stain and reagents into jars for use
• Make film dry in air
• Place in fixative for 1-3 min
• Wash with H2O for 30secs
• Treat with mordant for 3-5min
• Wash with H2O for 30secs
• Place in staining solu: for 3-5min
• Wash with water and dry.
• Spirochetes are Pink in colour.

143. 3, LEVADITTI’S METHOD FOR TISSUES
Procedure
• Fix the tissue in formalin for 24hrs
• Wash the tissue with H2O for 1hr and place it in alchohol for 24hrs.
• Place tissue in a solu: of AgNO3 + Pyridine for 2hrs
• Heat it to 50°C for 4-6hrs
• Wash with 10% pyridine solu:

144. • Transfer to Reducing fluid containing
Formalin +
Acetone +
Pyridine.
• Keep the tissue in this fluid for 2days in RT in the dark.
• Wash well with H2O and dehydrate with Alchohol.
• Embedded in Paraffin
• During examination remove the paraffin using xylene and
mount in Canada balsam.

146. THE ROMANOWSKY STAINS
• The original Romanowsky stain was made by dissolving Eosin & Zinc
free Ripened Methylene Blue in Methyl alchohol.
• On combining these 2 stains a precipitate is formed which is dried and
dissolved in pure Methanol.
• The methanol should be at a pH of 6.5.
• They impart a reddish purple colour to the chromatin of malaria and
other parasites.
• Various modifications of Romanowsky stain are present which vary
according to the method used for ripening and the relative proportions of
Eosin & Methylene Blue.

147. MODIFICATIONS OF ROMANOWSKY STAIN
1, Giemsa’s Stain
• Used for staining blood smears for Malarial parasites, Pathogenic
Spirochaetes and Mouse or Rat blood for Trypanosomes.
• This consists of a no: of compounds made by mixing different proportions of
Methylene blue & Eosin.
• Azure A Eosinate
• Azure B Eosinate
• Methylene Blue Eosinate.

148. • 2, Leishman’s Stain- for Malaria, Trypanosomes,
• 3, Wright’s Stain -
• 4, Jenner’s Stain – for Cytological examination of blood

149. STAINING PARASITES IN BLOOD FILMS
• BLOOD SMEARS
• Thin smears
• Allows optimal assessment of the morphology of any parasitic forms
• Fresh blood obtained by a finger prick or from an EDTA-anticoagulated
specimen collected by venepuncture

150. • Blood from Finger Prick
• Puncture the bulb of the finger or lobe of ear(after wiping the area
with alchohol and allow to dry) -- carefully wipe away the first
drop of blood.

151. • Place a clean glass slide on a horizontal
surface.Place a small drop of blood on the surface
of the glass slide about 1 cm from one end.
• With a second glass slide held at an angle of 45
degrees to the glass with the blood drop, back into
the drop of blood, hesitate momentarily while the
blood extends to both margins of the spreader
slide, then quickly and steadily advance the angled
slide to the opposite end.
• The result is a thin, feathered film that becomes
progressively thinner

152. • Limitation
• Parasitic forms may be missed in light infections.
• Smears -- within 1 hour after venepuncture.
• The morphology of parasitic forms and the erythrocytes become atypical after
that time from direct action of the anticoagulant.

153. CHARACTERISTICS OF A GOOD THIN FILM
• The surface of film should be even and uniform
• The margins of the film should not extend to the sides of the slide.
• The “Tails” end near about the centre of the slide.
• It consists of a single layer of RBCs.

154. • Thick Blood Film
• Relatively large volume of blood providing a better opportunity to recover
parasitic forms in light infections. Red blood cells are lysed -- detect parasitic
forms against a more transparent background.
• Fresh blood obtained by a finger prick or an EDTA-anticoagulated specimen
collected by venepuncture.
• Heparin or sodium citrate anticoagulated blood samples -- trypomastigotes or
microfilaria is suspected

155. • Blood from Finger Prick
• Touch a large drop of blood to the surface of the middle of a clean glass slide.
• Spread with a needle or with corner of another slide to form an area of half
inch square.
• The thickness should be such that newsprint can barely be seen through the
slide.
• If it is too thick, spread it with a circular motion using an applicator.
• If too thin, a second slide must be prepared.
• The spread of blood should be even, without ridges, smudges, or streaks

156. • Species identification of malarial parasites--difficult to
impossible
• Smears must be prepared from anticoagulated blood
within 1 hour after venipuncture.
• The morphology of parasitic forms and the erythrocytes
become atypical after that time from direct action of the
anticoagulant

159. • For thin film we use Giemsa’s or Leishman’s stain.
• For thick film we use Giemsa’s, Leishman’s or Field’s stain.
• For thick film we should dehaemoglobinise the smear before staining with
Giemsa’s or Leishman’s stain.
• Dehaemoglonisation is done by flooding the slide with Glacial acetic acid and
Tartaric acid mixture till the film becomes greyish white in colour.

160. FIELD’S STAIN
• Method used for thick films without fixation and dehaemoglobinisation.
• Consisits of solu: A and solu: B
• Solution A: Methylene blue 0.8gm
Azure I 0.5gm
Na2HPO4 5gm
KH2PO4 6.25gm
Distilled water 500ml

161. • Solution B
Eosin 1gm
Na2HPO4 5gm
KH2PO4 6.25gm
Distilled water 500ml
Stains are kept in staining jar with upto 3 inch
filled.

162. Procedure
• The thick film is placed in solu A for 1-2 secs
• Immediately rinsed with water
• Then place it in solu B for 1 sec
• Again rinse with water
• Allow to dry in vertical position.

163. FIELD’S STAIN SHOWING SIGNET RING CELLS (THEILERIASPECIES)

164. • GIEMSA STAIN
• Consists of number of compounds made by mixing different proprtions of
Methylene blue and eosin
• Eg:- Azure I, Azure II, and Azure II eosin
• Preparation of Giemsa
• Azure B eosinate – Methylene blue, conc H2SO4, Potassium dichromate, 5% eosin,
NaHCO3
• Azure A eosinate -- More of potassium dichromate
• Methylene blue eosinate
• Solvent – Methanol + glycerol
pH 7 – adjusted with PO4 buffer

165. • Grind Azur I(Azure B), II(Azure A) and II eosinate separately into fine powder
in 3 clean mortars
• Weigh
• 500mg Azur B +
• 100 mg Azur A +
• 400mg MB eosinate and
• 200mg methylene blue
• Decant mixed powder on to surface of 200ml solvent gradually.
• Keep at 50-60 0 C shaking intermittently for 2-3 days
• Smear fixed in methanol or ethanol x 3min

166. • RAPID METHOD
• Fix films by methanol for 3min.
• Stain with a mixture of 1 part stain in 10 parts buffer solution x 1 hour
• Wash with buffer solution x 30 sec
• Blot and air dry
• Excellent results with thin films for Malaria – Schuffner’s dots well defined
• Trypanosomes also well demonstrated

168. RAPID METHOD FOR SPIROCHAETES
• Fix films by heating or with Ethanol.
• Cover the film with freshly prepared solu: of Giemsa’s Stain.
• Warm till heat rises and allow to cool for 15secs.
• Pour of the stain and repeat this process of staining 4-5 times.
• Dry, mount and examine.

170. • SLOW METHOD
• For staining organisms which are difficult to be stained
eg: Certain pathogenic Spirochaetes.
• Allow diluted stain to act for a considerable period
• Stain causes a fine ppt – this should not get deposited on film

171. • Fix film in methanol x 3min
• Mix 1ml stain with 20ml buffer solution of pH 7 in a petri dish
• Place a piece of thin glass rod in the petri dish and slides.
• The fixed slide is kept with film downwards in the stain with one end resting
on the rod
• Leave to stain x 16-24 hrs
• Wash in buffer solution
• Air dry, mount and examine
• Reddish purple colour to chromatin of malarial and other parasites

172. • Used for
• Microfilaria, trypanosoma, Malaria, Leishmania
• Viral inclusions in herpetic vesicles
• Corneal scraping from suspected case of Chlamydia trachomatis
• Monolayer of infected cell culture such as McCoy cells of invitro Chlamydia culture
• Toxoplasma infection in brain tissue

176. Adachi’s Modification of Slow Method
• Used for staining the Flagella of Spirillum minus and for delicate spirochaetes.
Procedure
• Fix the smear with osmic acid vapour for 30-60secs.
(osmic acid vapour produced by heating solu: containing osmic acid 1gm, DW
100ml, 10drops of 1% Mercuric chloride)
• Stain overnight in Dilute Giemsa’s solu: containing 1% K2CO3.

178. • LEISHMAN STAIN
• Staining Protozoa in blood film
• 0.15 g Leishman’s powder in 100 ml methanol
• Dry unfixed films are used
• Undiluted stain – poured on slide – allowed to act for 1 min
• Add double volume of distilled water to the slide, mix alternatively using a
pipette
• Allow diluted stain to act for 12 mins
• Flood slide with distilled water – allow preparation to differentiate until film
becomes bright pink usually within 30secs. Blot and air dry.

179. • Distilled water -- should be neutral – or else colour of granules of WBC
changes and may look pathological.
A buffer solution with pH 7 can also be used instead of DW
• Na2HPO4 (anhydrous) – 5.4 g
• KH2PO4 -- 4.7 g
• Add 1g of above mixture to 2l of distilled water
• If little acidic pH of 6.8 needed
• Na2PO4 -- 4.5 g
• KH2PO4 -- 5.9g
• Add 1g to 2L of distilled water.

181. Demonstrating Schuffner’s dots in Benign Tertian Malaria
• Here we stain thin blood film with Giemsa stain f/b Leishman’s stain(By
Dinscombe in 1945).
Procedure
• Fix thin blood film with Leishman’s stain for 15-60secs.
• Dilute with twice volume of buffer solu: and allow to stand for 15min.
• Wash with dilute Giemsa’s stain and allow to stand for 30min.
• Wash with buffer solu: blot and dry.

183. JASWANT SINGH-BHATTACHARJI STAIN
• Rapid Romanowsky’s method of staining malarial parasites.
• Consists of Solu: 1 and Solu: 2
Solu: 1 Methylene Blue 0.5gm
Potassium Dichromate 0.5gm
H2SO4(1%) 3ml
KOH(1%) 10ml
DW 500ml

184. • Solu: 2
Eosin 1gm
Tap water 500ml
Solutions kept in staining jars.
For Thick film the procedure is:
• Immerse the slide in solu:1 for 10secs
• Wash for 2 secs in jar containing water adjusted to a pH 6.2-6.6 by addition of 5%
acetic or citric acid.
• Stain with solu:2 for 1sec
• Wash in the pH adjusted water for 5secs
• Immerse in solu:1 again for 10sec
• Wash in the same way for 10secs
• Dry and examine.

185. STAINING FOR PARASITES IN PUS AND STOOL
• WET MOUNT STAINING
• Pus and Stool
• With different types of I2 solutions – Lugol’s and D’Antonio’s I2
• D’Antoni’s Iodine
• KI 1g
• Powdered I2 crystals 1.5g
• DW 100ml
• Cyst – golden yellow
• Glycogen – brown
• Nuclei – pale refractile

187. • WET SALINE MOUNT
• Chromatoidal bodies visible

188. • NAIR’S BUFFERED METHYLENE BLUE
• MB(0.06%) in an acetate buffer at pH 3.6
• Nuclear details
• Cytoplasm– pale blue
• Nucleus – dark blue

190. • PERMANENT STAINED SMEARS
• Detection and correct identification of Intestinal protozoa
• Preparation of fresh material
• Schaudinn’s solution
• Absolute alcohol 100ml
• Saturated HgCl2 solution 200ml
• Glacial acetic acid 15 ml
• Prepare smear
• Fix it with Shaudinn’s solution x 30 min
• Time can be shortened to 5min by heating to 600C
• Allow slides to dry for several hours @ 350C or overnight @ RT

191. • If liquid specimen – 3 or 4 drops of PVA(Polyvinyl Alchohol) with 1 or 2 drops
of faecal material directly on to slide
• PVA PRESERVED MATERIAL
• Stool specimen added to PVA x 30 min
• After fixation – sample thoroughly mixed
• Small amount – on a paper towel – to absorb extra PVA.
• Eg :- Wheatley trichrome staining
Iron hematoxylin staining
Toluidene blue staining
GMS

192. • TRICHROME STAINING
• Originally developed by Gomori for tissue differentiation
• Adapted by Wheatly for intestinal parasites
• Ingredients
• Chromotope 2R
• Light green SF
• Phosphotungstic acid
• Glacial acetic acid
• Distilled water
• Principle
• Internal elements of cysts and trophozoites – best visualised with a stain that enhances
morphological features
• 90% alcohol -- decolouriser

193. • Cytoplasm – blue
green to purple
• Nuclei – red or purple
• Helminthic eggs and
larvae – dark purple or
red
• Background – pale
green

194. Iron Hematoxylin Stain
• Fix wet smears in Schaudinn’s fluid for 5min
• Wash films in 50% alchohol and apply Gram’s Iodine for 2min to remove
Mercury salt
• Wash with water
• Stain with Iron Hematoxylin for 10-20min.
• Wash films in water
• Counterstain with van Geison’s Stain for 15-30secs.
• Pass through alchohol, clear with xylol and mount in balsam.

195. • Iron Hematoxylin Stain:
(a)Haematoxylin 1gm
Absolute alchohol 100ml
(b) Liquor ferri perchloride 4ml
Concentrated HCl 1ml
DW 100ml
Mix equal parts of (a) & (b)
• van Gieson’s Stain :
Saturated aq solu: of Acid Fuchsin
Saturated aq solu: of Picric Acid

196. MODIFICATIONS OF IRON HAEMATOXYLIN STAIN
• Spencer and Monroe short haematoxylin stain
• No decolourisation
• Tonkins and Miller
• Use of phosphotungstic acid as decolourizer.
• Iron hematoxylin with carbol fuchsin
• Intestinal protozoa
• AF organisms
• Used with Sodium acetate & formalin fixed stool specimen
• Background and organism – grey blue to black
• Cellular inclusions and nuclei – Darker than CP

197. • DOBELL’S METHOD
• Similar to Original method except the following:
• Mordant for 10min with Ammonium Molybdate instead of Gram’s Iodine.
• Counter Stain is not used.

199. • TOLUIDENE BLUE
• Solution A
• Sulfation reagent – 1N NaOH in ether or conc. H2SO4
• Solution B
• Toluidene blue O +
• Toluidene blue D+
• Conc HCl
• Absolute alcohol
• Smear is made with Mucoid part of sputum or centrifuged bronchial lavage
• Fix in absolute alcohol x 1 min
• Slide placed in sulfation reagent x 5 min

200. • Solution B x 3 min
• Dehydrate with isopropyl alcohol
• Clear with xylene
• Principle :- Thick cell wall of parasites and some fungi – after sulfation –
retains Toluidene Blue O stain
• Pneumocystis – reddish blue / purple
• Cysts – Crescent shaped/ punched
• Background -- blue

201. Pneumocystis Pneumonia

202. • JENNER GIEMSA STAINING
• Oocysts of Cryptosporidium in faeces
• Blue spherical bodies containing few eosinophilic granules

203. FUNGAL STAINING
Wet Preparation
1. KOH Wet Mount
2. India Ink Stain
3. Nigrosin Stain
4. Lactophenol Cotton Blue Stain
5. PHOL Stain
6. Neutral Red Stain
7. Diazonium Blue B Stain

204. Differential Stains
• Gram’s Stain
• Modified Acid Fast Stain
• Hematoxylin And Eosin Stain
• May-Grunwald Giemsa Stain
• Diff-Quik Stain
• Periodic Acid-Schiff Stain

205. • Gridley’s Fungal Stain
• Gomori’s Methenamine Silver Stain
• Mayer’s Mucicarmine Stain
• Masson-Fontana Stain
• Schmorl’s Melanin Stain
• Toluidine Blue O Stain

206. Wet Preparations
• KOH Wet Mount
• Clearing of specimens – fungi more readily visible
• Advantage – rapid detection of fungal elements
• Disadvantage – background artifacts – confusing

207. Ingredients:
• KOH 10gm
• Glycerol 10ml
• DW 80ml
1. Slide KOH
• Place epidermal scales,skin scrapings,nail scrappings, homogenized biopsy
tissue on clean glass slide.
• Pour drop of KOH(10%) on specimen and place cover slip
• Gently warm over flame for cleansing
• If DMSO is used no need of heating

208. 2. Tube KOH
• Used for biopsy specimen which take longer time for dissolution.
• Specimen is dissolved in KOH(10%) in a test tube and kept in incubator at
37°C overnight and examine by slide mount.

209. India Ink Stain
• Negative staining for encapsulated fungi like Cryptococcus species.
• Ingredients
India Ink: 150ml
Merthiolate: 3ml
Tween80 : 0.1ml
• A drop of specimen such as CSF and an equal volu: of India Ink mixed on
glass slide.
• Cover slip is put and examine under oif.

210. Nigrosin Stain
• Negative staining similar to India Ink.
Ingredients
Nigrosin granules: 10gm
Formalin(10%): 100ml
• Better than India Ink as it can last for >1 yr with no carbon particles
• Rest of technique similar to India Ink Staining.

211. • LACTOPHENOL COTTON BLUE
• Phenol crystals 20g
• Lactic acid 20 ml
• Glycerol 40ml
• Distilled water 20ml
• Cotton blue 0.075g
• Dissolve phenol crystals in the liquids by gentle warming and then add the dye
• Lactic acid – Preserves fungal morphology
• Glycerol -- Increases viscosity and prevents drying
• Phenol -- Disinfectant
• Cotton Blue stains outer wall of fungus.

212. • Used to stain fungal isolates grown in culture
• Place one drop of stain on a slide – add small amount of culture on it
• Tease with 2 teasing needles
• Place cover slip
• Gently press
• Seal edges to keep it for long periods with nail polish
• Advantage :-
• Can study micro and macroconidia, chlamydospore, hyphae
• Help to speciate fungi.

214. LPCB WITH PVA
• LPCB preparation can be permanently preserved if PVA is used
• Here instead of cotton blue safranine can also be used

215. • PHOL STAINING
• Similar to LPCB Staining.
• Pal, Hasegawa, Ono, Lee
• Formalin instead of phenol
• Methylene blue instead of cotton blue
• For Fungus and Prototheca(Green Algae).

216. PHOL Staining of Prototheca

217. Narayan Staining
• Similar to India Ink
• Glycerine instead of phenol
• Methylene Blue instead of Cotton Blue
• Dimethyl sulfoxide as cleansing agent.

218. • NEUTRAL RED STAINING
• Useful and easily applicable
• Evaluation of vitality of fungal elements
• Water soluble dye – passes through intact plasma membrane – stored in
lysosome of viable cells
• So when cell memb: or lysosome are damaged stain uptake ceases.
• So used as a vital stain for Dermatophytes and Candida.

220. Diazonium Blue B Stain (DBB Stain)
• Used for differentiating the heterogenous genera,
basidiomycetous Trichosporon and
ascomycetous Geotrichum.
• Positive reaction is indicated if dark red or violet red colour develops within 2
mins.
• Trichosporon form pink to violet pigments with DBB
• While Geotrichum is DBB negative.

221. Differential Stains For Fungus
Gram’s Stain
• Modifications for fungus
Hucker’s modification
Claudius Modification
Brown and Brenn Modification
Modified Acid-Fast Stain
Kinyoun’s Cold Acid Fast Staining can be used for Pichia
anomala, Saccharomyces cerevisiae, Microsporidium etc.

223. Hematoxylin and Eosin Stain
• Nucleus and Fungi are stained by Hematoxylin(Shades of Blue)
• Tissue fibres and Cytoplasm stained by Eosin(Shades of Red & Pink)
• Stain
Hematoxylin Stain
Hematoxylin - 3gm
Ethanol - 125ml
K or NH3 Alum -160gm
Iodine – 1gm
DW – 2000ml

224. Eosin 1% aq solu:
Differentiator – HCl(1%) in Ethanol
Bluing agent - Scott’s Tap Water
NaHCO3 - 2gm
MgSO4 – 20gm
Water – 2L

225. Procedure
• Bring sections to water
• Stain wiyh Hematoxylin solu:
• Wash with H2O apply Differentiator
• Wash with H2O apply Bluing agent for 3-5min
• Wash with H2O and Counterstain with Eosin solu: for 30secs
• Wash, dehydrate with alchohol and mount the specimen.
• Fungi- Blue to Purple

226. PulmonaryAspergillosis

227. May-Grunwald Giemsa Stain
Commonly used to demonstrate intracellular yeast like Histoplasma capsulatum
and other fungi.
Stain
• May-Grunwald solu:
May-Grunwald dye - 1gm
Methanol - 300ml
• Giemsa Solu:
Giemsa dye – 1gm
Methanol – 85ml
Glycerol – 54ml
Working solu: made by diluting with water.

228. Procedure
• Fix smears in methanol for 5-10min
• Stain in diluted MG solu: for 10min
• Without washing transfer the slides to diluted Giemsa solu: and allow to stand
for 15min.
• Wash and keep water in slides for 1-2min for differentiation.
• Air dry and examine.
Nucleus – Purple
Cytoplasm – Blue
RBC - Pink

230. Diff - Quik Stain
This is a dip stain.So fixative & staining solu: are dispensed in coplin jars.
Solutions required:
1. Fixative – Fast Green in Methanol
2. Stain solu: 1 - Eosin G in PO4 buffer
3. Stain solu: 2 – Thiazine dye in PO4 buffer

231. Procedure
• Allow smears to dry
• Dip slide in fixative 5 times for 1 sec each, allow
excess to drain after each dip.
• Dip slide in stain 1 solu: 5 times for 1sec each,
allow excess to drain.
• Dip slide in stain 2 solu: 5 times for 1sec each,
allow excess to drain
• Wash with distilled water.
• Blot and dry
• Examine under low power and then oif.
• Yeast cells stain Blue.

233. PERIODIC ACID - SCHIFF STAIN
• Principle
Based on Feulgen reaction wherein hydrolysis of Polysaccharides with HCl
liberates Aldehydes which recolour Schiff reagent.The polysaccharides of fungi
are oxidized to aldehyde by Periodic acid giving a magenta colour.
• Proteins and nucleic acid are unstained

234. • Advantage – stain fungal elements well – hyphae and moulds can be well
distinguished
• Disadvantage – Nocardia species don’t stain well
Many components of tissue having
polysaccharides are also stained to magenta along with fungi.
This stains only Live Fungi

235. Solutions required
• Periodic acid 1% aqueous solu:
• Schiff’s Reagent:
Basic fuchsin – 1gm
Na metabisulfite - 2gm
Conc HCl – 2ml
Activated Charcoal – 2gm
DW – 200ml

236. Procedure
• Bring sections to water.
• Oxidize with Periodic acid solu: for 5min
• Rinse well in DW
• Treat with Schiff’s reagent for 15min
• Wash in running tap water for 5-10min to intensify the stain
• Stain nuclei with Hematoxylin.
• Dehydrate, mount and examine.
PAS +ve substance – Magenta
Nucleus - Blue

238. Gridley’s Fungal Stain
This is like PAS staining but Chromic acid is used as the Oxidizing agent.
Aldehydes produced recolour Schiff’s reagent and give fungus a Red colour

239. Procedure
• Treat sections with Chromic acid 4% aq solu: for 1hr.
• Wash and treat with Schiff’s reagent for 15min
• Treat with Sulfurous acid for 6min
• Wash and treat with Aldehyde Fuchsin for 20-30min
• Wash with 50% alchohol and then with water
• Stain with Tartrazine solu: for 30secs
• Wash with Cellosolve
• Dehydrate, mount and examine
Fungus – Purple
Background - Yellow

241. CALCOFLOUR WHITE FLOURESCENT STAINING
• Advantage – can be mixed with KOH
Detects fungi rapidly d/t flourescence
used to study morphology
• Disadvantage – requires use of flourescent m/s
vaginal secretions are difficult to interpret
• Stock solution A
• Calcoflour white 1.9g
• Distilled water 100ml
• Stock solution B
• Evan’s blue 0.05g
• Distilled water 100ml

242. • Working solution
• Solution A 1ml
• Solution B 9ml
• Tissue and sputum mixed with a drop of working solution and one drop of 20%
KOH
• Allow for maceration
• Place coverslip
• Flourescent m/s with UV light or one with blue light (400-450 nm)
• Fungal cell wall – bright green or blue white

244. • Cellulose in fungal cell wall – bind to stain
• Enhances the visibility on tissue or other specimens

245. • GOMORI’S METHANAMINE SILVER STAIN
• Principle : - depends upon the availability of free
aldehyde groups for reduction of alkaline methanamine
silver nitrate complex to metallic silver
• Free aldehyde – liberated from chromic acid treatment of
cell wall polysaccharide
Fungi stained black(both live and dead).
It also stains Actinomycetes unlike PAS stain
Mucopolysaccharide dark grey, Cytoplasm old rose and
Tissue stain pale green.
• Aq Chromium trioxide solution, 5%
• CrO3 5gm
• DW 100ml

246. • Methanamine silver nitrate stock solution
• 5% AgNO3 5ml
• 3% Methanamine 100ml
• Shake to dissolve white ppt – clear solution – stable @ 40C
Working Solution
Stock solution 25ml
5% Borax solu: 2ml
FeCl3 0.1%
Na metabisulfite 1% aq solu:
Na thiosulfate 5% aq solu:

247. Stock Light – Green:
Light green SF(yellow) 0.2gm
Glacial acetic acid 0.2 ml
DW 100ml
Working Light Green solu: Dilute this to 5times DW

248. Procedure:
• Treat section with CrO3 solu: for 1hr wash with H2O
• Bleach in metabisulfite solu: for 1min
• Wash with water
• Working methanamine silver solu: is preheated to 56°C for 1hr using Coplin
jar.
• Treat section in this solu: and examine after 10-20min. Until fungi are
blackened treat with this solu: repeatedly.
• Wash with DW and tone in 0.1% aq FeCl3 for 3min
• Wash in water and fix in Na Thiosulfite for 5min.
• Wash again

249. • Counterstain in Light green for 1-2min
• Wash in water
• Dehydrate, clear and mount as desired.
Cellulose, chitin, fungi including Pneumocystis jirovecii, amoebae some
mucins, melanin, glycogen, and starch stain Black in a green background.

250. • Fungi – black
• Mucin – dark grey
• Background – pale green

251. Mayer’s Mucicarmine Stain
• Used for staining Crytococcus and Rhinosporidium species
• Cryptococcus stains deep rose red, nuclei black and tissue yellow.
• In Rhinoporidiosis sporangium and endospore are stained by mucicarmine
stain.

252. Staining solu:
• Grind 1gm carmine and place in large conical flask
• Add 100ml of 50% alchohol and mix.
• Add 1gm Al(OH)3
• Mix and add 0.5gm anhydrous Al(Cl)3
• Mix and boil gently for 2.5min
• Cool and filter.
• Store at 4°C

253. Procedure:
• Treat section with alum hematoxylin solu:
• Differentiate in acid alchohol solu: and apply bluing agent (as in Hematoxylin
and Eosin stain)
• Stain with mucicarmine solu: for 20min
• Wash in water, dehydrate clear and mount.

255. Masson- Fontana Stain(MF Stain)
• Used to identify Melanin containing fungi
• Melanin possesses the ability to bind silver from silver solu: and reduce it
directly to metallic silver without the use of an external reducer.
• Melanin containing fungi are called Phaeoid fungi and are now an emerging
significant pathogen.

256. Staining solu:
• Aq AgNO3 +
• Strong NH3 +
• DW +
• Aq Na thiosulfate
• Aq Neutral red

257. Procedure:
• Treat section with Ammoniacal silver solu: in dark container.(Coplin jar painted
black) for 20-30min at 56°C
• Wash with water
• Treat with 0.5% Na Thiosulfate for 2min
• Wash and counterstain with neutral red solu: for 3-5min
• Wash, dehydrate and mount.
Melanin, Argentaffin granules, Chromaffin and some Lipofuscin pigment
stain black
Nuclei stain red

259. Schmorl’s Melanin Stain
• It is a melanin based stain like(MF Stain)
• Here Ferricyanide is converted to Ferrocyanide which is again converted into
insoluble Prussian Blue in the presence of ferric ions.
• Here Melanin contents are stained Bluish - green.

260. Staining Solution
• FeCl3 (1% aq solu:)
• Potassium Ferricyanide (1% aq solu:)
• DW
Procedure:
• Immerse section in coplin jar containing stain solu: upto 5min
• Wash with water
• Treat with 1% acetic acid for 5min
• Counterstain with nuclear fast red for 5min
• Wash in water, dehydrate, mount.

261. • Melanin stain shades of Blue colour
• Nuclei – Pinkish red
• Cytoplasm – Pale pink

263. Toluidine Blue O Stain
• Used for rapid detection of Pneumocystis jirovecii in lung biopsy, imprint
smears and BAL fluid.
• It stains the cysts as reddish blue or dark purple against a light background.
• Trophozoites are not detectable

264. Procedure:
• Place air dried slide in sulfation reagent(Glacial acetic acid + H2SO4) for
10min
• Wash with cold water for 5min
• Drain and place in Toluidine Blue O for 3min
• Wash with Ethanol and then Xylene
• Dry and observe under oif
Pneumocystis jirovecci cysts - Purple
Tissue remanants - Blue
Disadvantage -Trophozoite – Not detectable

266. STAINING FOR RICKETTSIAE AND CHLAMYDIAE
• MACCHIAVELLO STAIN
• Extracellular and intracellular rickettsiae and elementory bodies of psittacosis
• Constituents
1. Basic fuchsin 0.5%
2. Citric acid, 0.5%
3. Methylene blue, 1%
• Procedure
• Air dry smears – heat fix
• Basic fuchsin x 4min – drain , wash
• Dip in citric acid immediately wash in tap water
• Stain with 1% methylene blue x few secs

267. • Rickettsiae – red staining coccoid forms
• Psittacosis elementary bodies – red , found clusters in macrophages
• Early dvlptl forms of psittacosis – blue large coccoid bodies – initial bodies

268. • CASTENADA’S METHOD
• Solution A
• 1% potassium phosphate
• 25% aqueous NaPO4
• 1 ml formalin
• Solution B
• Methanol
• Methylene blue
• Mix 20 ml solution A + 0.5 ml of solution B + 1ml formalin

269. • Solution C
• 0.2 % aqueous safranine
• 0.1% acetic acid
• Thin film
• Air dry
• Cover with stain – A +B
• Drain (Don’t wash)
• Counter stain with safranine O x 1-4 sec
• Rickettsiae – blue
• Polymorphs -- red

271. STAINING FOR VIRAL INCLUSION BODIES
• HERZBERG VICTORIA BLUE STAIN
• For variola, vaccinia, HSV and varicella
• Stain
• 3ml Victoria blue in 100 ml DW – age solution in a brown bottle for 14 days
• Air dry x 24 hrs – wash in DW x 10 min
• Dry at 37 0 C x 1hr
• Stain
• 5min for vaccinia
• 20 min varicella
• 30 min HSV
• Rinse in DW, dry and examine

272. • MAY-GRUNWALD-GIEMSA STAIN
• For tissue culture
• Stain
• May-Grunwald stain -- 0.25 gm dye in 1000ml absolute methyl alcohol– age for 1 month
prior to use
• Giemsa solution – 1 gm Giemsa in 66 ml of glycerol by heating at 55-60 0 C x 1.5 hrs
• Add 66 ml methyl alcohol
• Wash tissue culture with HBSS(Hanks’ balanced salt solu) x 15min
• Fix in absolute methyl alcohol x 5 min
• Stain with May-Grunwald x 10 min
• Stain in dil.Giemsa x 20min

273. • Dehydrate in 2 changes of acetone – do not allow slide to dry
• Clear by rinsing in 3 changes of acetone – xylol (2:1), 3 changes of acetone-
xylol (1:2) and fresh xylol x 10 min
• Mount in balsam and examine
• RNA – blue
• DNA– red purple

275. • METHYL GREEN – PYRONINE STAIN
• Differentiate DNA from RNA
• Methyl green – specific for DNA – green
• Pyronine – specific for RNA – red
• Stain
• Methyl green – 0.5 gm in 50ml DW with 0.5% phenol
• Pyronin Y – 0.5gm in 50ml DW with 0.5% phenol
• Mix both – then add 95% ethanol 2.5 ml and glycerin 20ml
• FAA(formalin-acetic acid-alchohol) fixative
• 95% ethyl alcohol 90ml
• Glacial acetic acid 5ml
• Neutral formalin 5ml

276. • Fix tissue in cold FAA fixative x 1 hr
• Rinse in 50% alcohol and in dw
• Stain in Methyl green-pyronin solution x 20-25 min
• Wash in DW
• Dehydrate with acetone x 3sec
• Clear in 2 changes of xylol
• Mount and examine

278. • ACRIDINE ORANGE STAIN
• Rapid, simple – for determining the type of nucleic acid
• DNA – flouresce yellow green
• RNA – reddish orange
• Carnoy fixative
• McIlvaine buffer , pH 3.8
• Acridine orange stain
• Cell monolayer in PO4 buffer, 6.8
• Fix coverslip in Carnoy fixative x 20min
• Rinse for 2 min in 3 changes of McIlvaine PO4 buffer
• Stain with 0.5% acridine orange x 30 min

279. • Wash for 30min in 3 changes of buffer
• Mount in buffer and seal with nail polish

280. • HEMATOXYLIN EOSIN STAIN
• Paraffin embedded and sectioned tissues
• Differentiate Coxsackie A and B
• Solutions
• Alum hematoxylin solution – 0.6 gm of H, 5 gm of ammonium alum and 0.1 gm of NaOH in
70 ml DW + 2ml GAA and 20 ml glycerol – filter and dilue to 1:20
• NaHCO3 solution – 1 gm in 100ml dw
• Eosin Y solution – 0.5 gm in 100ml dw
• Rinse with 3 changes of HBSS
• Fix in neutral buffered formalin x 30 min – rinse in DW
• Stain with 1:20 dilution of alum hematoxylin sol x 10 min

281. • Rinse in tw
• Treat with 1% NaHCO3 untill cells assume blue colour
• Rinse in dw
• Dehydrate rapidly with 2 changes of acetone
• Rinse in 3 changes of acetone-xylol (2:1), 3 changes of acetone-xylol(1:2) and
xylol
• Clear in fresh xylol x 10 min
• Mount and examine

282. • MODIFIED GIEMSA
• Urinary sediments for intranuclear cytomegalic inclusions
• Fixative
• Equal parts of ethyl ether and absolute ethyl alcohol
• Stain
• 2% solution of Giemsa in 100ml PO4 buffer (7.2)
• Centrifuge urine – 2500 rpm x 15 min
• Sediment – clean slide coated with 25% bovine albumin
• Fix x 10 min
• Stain x 1 hr
• Wash in buffer x 3 min
• Examine

283. • SELLER’STECHNIQUE
• Rabies – demo of intra cytolasmic inclusion bodies – neuronal cells – Negri
bodies
• Negri bodies – acidophilic – 1- 20 internal basophilic granules
• Matrix stains pink
• Internal granules– blue to deep blue

284. • Solutions
• MB stock – 1 gm MB in 100 ml absolute methyl alcohol
• BF(basic F)stock – 1 gm BF in 100 ml Absolute methyl alcohol – refrigerator at 2- 4 0 C
• Seller stain – 2 parts MB + 1 part BF – keep tightly stopperd to prevent alcohol evaporation
– check stain periodically with rabies positive tissue
• Amputate head – take to lab as soon as possible – operator should wear
Rubber gloves
• Place head on a board – remove skull cap by sawing – remove brain
• Make an incision straight down on one side of hemisphere into lateral ventricle

285. • At post part of lat ventricle– incision deep into cortex – Ammon’s horn – Negri
bodies +
• Prepare smears – spreading tissue b/w 2 slides – smears from B/L Ammon’s
horn, cortical areas of cerebrum and cerebellum
• Tissue still moist – dip in Seller stain x 3 sec
• Rinse in TW
• Dry without blotting -- examine

287. MANN’S METHYL-BLUE EOSIN STAIN
• Contains 1% aqueous solu: of methyl blue, eosin and DW.
• Fix in Bouin’s solu: or Zenker’s fluid
• Stain for 12hrs in incubator at 37⁰ C
• Rinse with water
• Add 70% alchohol and Orange G solu: to differentiate
• Dehydrate and mount in Canada balsam