The document provides instructions for performing bacterial staining techniques. It describes how to make smears from clinical samples and the importance of proper smear preparation for staining and microscopy. It then explains the basic principles and types of bacterial staining, including simple, differential, and special staining. Gram staining and Ziehl-Neelsen staining are differential techniques discussed in detail, outlining the staining processes, reagents used, and bacterial groups that each method can identify based on staining characteristics. Proper staining allows clear visualization of bacterial morphology and differentiation of types under the microscope.

1. BACTERIAL
STAINING
Learning objectives
At the end of this chapter, the student will be able to:
– Describe smear preparation for bacterial examination
– Define stains and bacterial staining.
– Describe the uses of staining solutions.
– List the types of staining methods that are used to
stain bacteria.
– List the different types of bacterial staining
techniques.
– Discuss the principles of Gram’s stain and Ziehl
- Nelseen stain.
– Describe the factors that contribute for false Gram’s
stain and Zihel -Nelseen stain results.
– Prepare the Gram’s stain and Zihel -Nelseen
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2. PREPARATION OF SMEARS
– Making smear is a precondition to facilitate
staining and further observation of
microorganisms under microscope.
– If smears are to provide reliable information
they must be prepared, labeled and fixed
correctly prior to being stained.
– Smears should be spread evenly covering an
area of about 15 – 20 mm diameter on a slide
– Whenever smears are made every precaution
and aseptic technique should be followed
strictly!
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3. PREPARATION OF SMEARS
Making smear is a precondition to facilitate staining and
further observation of microorganisms under
microscope.
Materials for bacteriological Smear
preparation
• Microscope slide
• Swab-for making smear taking sample from
body parts or culture media
• Applicator stick- for taking sample from &
make smear on slide
• Lead pencil – (not grease pencil or ink) – for
lebeling
• Heat or alcohol – for fixation
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4. PREPARATION OF
SMEAR
S…
• Purulent body fluids( contain pus /dead WBC)- using a
sterile wire loop, make a thin preparation.
• Non purulent body fluids - centrifuge the fluid and
make a smear from a drop of the well mixed
sediment.
• Culture – Emulsify a colony in sterile distilled water and
make a thin preparation on a slide
• Sputum- Use a clean stick to transfer and spread
purulent material on a slide. Soak the stick in a phenol
or hypochlorite disinfectant before discarding it.
• Swabs- Roll the swab on a slide. Rolling the swab
avoids damaging the pus cells.
• Feaces (stool) –???? .
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5. Method of collecting and making skin smear of
M.leprae
• The site sampled should be the edge of a leprosy
lesion.
1. Explain the procedure to the patient
2.Fit a new scalpel blade (surgical blade) in its
scalpel holder.
• Sterilize the blade by wiping it carefully with a
piece of absorbent cotton wool soaked in 70%
v/v ethanol and flaming it for 2-3 seconds in the
flame of a sprit lamp. Allow the blade to cool.
3. Wearing gloves, cleanse the area from where the
smear is to be taken, using a cotton wool swab,
moistened with 70% v/v ethanol. Allow the area to
dry.
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6. PREPARATION OF SMEARS…
4. Pinch the skin tightly between the thumb and index finger
until it becomes pale due to loss of blood.
NB. The area should be kept bloodless.
5. Using the sterile blade, make a small cut through the skin
surface, about 5mm long and deep enough in to the
dermis (2- 3mm) where the bacteria will be found,
continue to hold the skin tightly.
6. Turn the scalpel blade until it is at a right angle to the cut,
using the blunt edge of the blade, scrape firmly two or
three times along the edges and bottom of the cut to
collect a sample of tissues juice and cell (not blood)
7. Transfer the sample to a slide. Make a small circular
smear, covering evenly an area measuring 5 – 7 mm
in diameter.
8. Cover the cut with small dressing
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7. The thickness of the smear should allow to read a text when
placed under the smear.
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8. PREPARATION OF SMEARS…
Drying and fixing smears
• After making a smear, leave the slide in a safe
place for the smear to air – dry protected from
direct sun light.
• When a smear requires urgent staining, it
can be dried quickly using gentle heat.
• Smears are fixed by heat, alcohol, or
occasionally by other chemicals.
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9. PREPARATION OF SMEARS…
Heat Fixation
- This is widely used but can damage organisms and alter their
staining reactions especially when excessive heat is used.
- Heat fixation also damages leukocytes and is therefore unsuitable
for fixing smears which may contain intracellular organisms such as
N. gonorrhoeae and N. meningitides.
• When used, heat fixation must be carried out with care.
1. Allow the smear to air – dry completely
2. Rapidly pass the slide, smear upper most, three times through
the flame of a sprit lamp or pilot flame of a Bunsen burner.
NB: do not heat smears excessively
3. Allow the smear to cool before staining.
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10. Alcohol Fixation
• is far less damaging to microorganisms than heat.
Cells, especially pus cells, are also well
preserved.
• Therefore recommended for fixing smears when
looking for Gram negative intracellular diplococci
• It is more bactericidal than heat. (E.g. M.
tuberculosis is rapidly killed in sputum smears
after applying 70% v/v alcohol).
• When use:
– Allow the smear to air dry completely
– fix with one or two drops of absolute methanol or
ethanol
– Leave the alcohol on the smear for a minimum
of 2 minutes or until the alcohol evaporates.
PREPARATION OF SMEARS…
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11. Staining
Precautions that should be taken when staining
smears
• Do not immerse slides in containers of stain
because this can lead to contamination of stain and
transfer of organisms from one smear to another.
• Do not attempt to stain a smear that is too thick.
• When washing smears of CSF sediment and
specimens which can be easily washed from a
slide, direct the water from a wash bottle on the
back of the slide, not directly on the smear.
• After staining, place the slides at an angle in a
draining rack for the smears to air dry. Do not blot
smears to dry with filter or blotting paper.
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12. STAINING IN
BACTERIOLOGY
• PRINCIPLE OF STAINING
– Even with the microscope, bacteria are
difficult to see unless they are treated in a
way that increases contrast between the
organisms and their background.
– The most common method to increase
contrast is to stain part or all of the microbe.
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13. Definition
Stains (Dyes) are coloured chemical
compounds that are used to selectively give
(impart) colour to the colourless structures of
bacteria or other cells.
 Bacterial staining is the process of imparting
colour to the colourless structures (cell wall,
spore, etc.) of the bacteria in order to make it
visible under the microscope.
STAINING
…
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14. STAINING
…
Uses of staining
1.To observe the morphology, size and
arrangement of bacteria
2.To differentiate one group of bacteria from the
other group.
Staining reactions are made possible because
of the Physical phenomena of capillary
osmosis, solubility,
adsorption, and absorption of stains or dyes by
cells of microbes.
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15.  Basic principle:
The cellular components of mammalian as
well as microbial cell are different.
For example the nuclei of cell is negatively
charged because of the presence of acidic
component (DNA) hence it combines with
positively charged compounds, (basic dyes).
and
the cytoplasm parts of a cell is generally
positively charged therefore combines with
negatively charged compounds (acidic dyes).
STAINING
…
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16. STAINING …
Why are stains not taken up by every microbe?
Factors controlling selectivity of microbial cells
are:
1. Number and affinity of binding sites
2. Rate of reagent uptake
3. Rate of reaction
4.Rate of reagent loss (differentiation or
regressive staining)
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17. STAINING …
Type of stain in Microbiology
There are three kinds of stains
1. Basic stains
2. Acidic stains
3. Neutral stains
NB: This classification is not based on the pH of
stains
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18. STAINING
…
A. Basic Stains
 Are stains in which the colouring substance is contained
in the base part of the stain and the acidic part is
colourless.
 The bacterial cells can easily stain with basic dyes
e.g. Methylene blue stain, Safranin, Gentian
violet, Carbolfuchsin etc.
B. Acidic Stains
 Are stains in which the colouring substance is contained
in the acidic part of the stain and the base part is
colourless.
 Acidic dyes are mainly
used in histology laboratory but not commonly used
in microbiology laboratory ; e.g. eosin.
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19. STAINING
…
C. Neutral Stains
 Are stains in which the acidic and basic
components of stains are coloured.
 Neutral dyes stain both nucleic acid and
cytoplasm. these stains are commonly used in
haematology laboratory. e.g. Giemsa’s stain,
Wright’s stain.
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20. Type of staining methods
1. Simple staining method
2. Differential staining
method
3. Special staining method
STAINING
…
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21. 1. Simple staining: type of staining method in which
only a single dye is used. There are two types
–
a. Positive staining: The bacteria or its
parts are stained by the dye. e.g. Methylene blue
stain, Crystal violent stain
Procedure
1. Make a smear and label it
STAINING
…
2. Allow the smear to dry in air
3. Fix the smear over a flame
4. Apply a few drops of positive simple
stain
5. Wash off the stain with water
6. Air -dry and examine under microscope
Result-all the bacterial surface is stained.
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22. b. Negative staining-
• The dye stains the background and the bacteria
component remain unstained. e.g. as nigrosin, India
ink, or eosin
• Procedure
– apply a small amount of bacteria to one end of a
clean microscope slide
– Add 1 to 2 loops of nigrosin, India ink, or eosin solution
to the bacteria and mix thoroughly.
– Spread the mixture over the slide using a second slide.
– Allow the smear to air dry. Do not heat-fix!
– With the low-power objective, find an area of the smear
that is of the optimal thickness for observation.
– Use the oil immersion lens to observe
STAINING
…
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23. STAINING
…
2. Differential staining method
• A method in which multiple stains (dye) are
used to distinguish different group of bacteria.
e.g. Gram’s stain, Ziehl-Neelson stain.
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24. A. Gram stain
• This method was developed by the Danish
bacteriologist Christian Gram in 1984.
• The most commonly used for the direct examination of
specimens and bacterial colonies because it has a broad
staining spectrum
Basic concepts:
• Most bacteria are differentiated by their gram reaction due to
differences in their cell wall structure.
• The surface of bacterial cell has got a negative charge due to
the presence of polysaccharides and lipids (PG) this has
made the surface of the bacteria to have affinity to
cationic or basic dyes
24
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25. Gram stain
(cont’d)
Principle
• The cells are first fixed to slide by heat or alcohol
and stained with a basic dye (e.g. crystal
violate), which is taken up in similar amounts by
all bacteria.
• The slides are then
treated with an Gram’s iodine to
fix (mordant) the crystal violet stain on Gram
positive bacteria, decolorized with acetone alcohol,
and finally counter stained
with Safranin.
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26. Gram stain
(cont’d)
 Gram positive bacteria stain dark purple with
crystal violet (CV) and are not decolorized by
acetone or ethanol.
 Gram negative bacteria stain red because after
being stained with CV they are decolorized by
decolorizer and take up red counter stain
(Neutral red, Safranin or diluted carbolfuchsin).
 The reason for the retention of the primary stain
(CV) by the Gram positive bacteria after
decolorization is due to the presence of more
acidic protoplasm (PG layer) of these organisms
which bind to the basic dye.
26
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27. Gram stain
(cont’d)
 Gram positive bacteria
• Staphylococcus,
• Streptococcus,
• Clostridium
• Bacillus
• Corynebacterium … etc
• Gram negative bacteria
• Neisseria spp.
• Haemophilus spp.
• Enterobacteriaceae …etc.
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28. Gram stain
(cont’d)
Required reagents
• Crystal violet
• Gram’s Iodine
• Acetone-Alcohol or 95%
Alcohol
• Safranin or Neutral red
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29. Gram’s stain reagents
preparation 1.Crystal violet
stain
– Crystal violet ………………………….
20g
– Ammonium oxalate ……………………
9g
– Ethanol or methanol absolute ………..
95ml
– Distilled water ……………………….. to
1 litre
Gram stain
(cont’d)
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30. 3. Acetone -alcohol
To make 1 litre
– Acetone …………………………… 500ml
– Ethanol or methanol absolute ….. 475 ml
– Distilled water ……………….……. 25ml
• N.B: some workers prefer to use acetone by
itself or ethanol 95% v/v as decolorizing
solution.
• A mixture of acetone and alcohol is
recommended because it decolorizes more
rapidly than ethanol 95% v/v and is less likely
to over decolorize smears than acetone with
out alcohol added.
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31. Gram stain
(cont’d)
2. Gram’s iodine
– Potassium iodide …………………..
20 g
– Iodine …………………….…………
10 g
–Distilled water ………………………
to 1 litre Should be stored in a brown
bottle
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32. 4. Safranin
1. Prepare a stock solution
 SafraninO 2.5g
 Ethanol (95%) 100ml
Mix until all the Safranin is dissolved. Transfer the
solution to a glass – stoppered bottle.
Label the bottle ( Safranin stock solution) and write the
date.
2. Prepare a working solution in a glass stoppered bottle
 Stock solution 10ml
 Distilled water ----------------------------------------------
90ml
Label the bottle ( Safranin working solution) and write the
date
Gram stain
(cont’d)
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33. Gram stain (cont’d)
5.Neutral red; 1g/l
(w/v) To make 1 litre
– Neutral red ……………. 1g
– Distilled water ………… 1 litre
N.B: Neutral red is selected as the counter stain
because it stains well gonococci and
meningococci. Safranin can also be used.
The dilute carbolfuchsin (1 in 10) is
recommended for staining Vincent’s
organisms, Yersinia, Haemaophilus,
campylobacter, and vibrio species.
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34. Gram stain (cont’d)
Procedure of Gram’s Stain
1. Fix the dried smear with heat by gently passing it over
sprit lamp or Bunsen burner.
Note: When the smear is for the detection of gonococci
or meningococci, it should be fixed with methanol for 2
minutes
2. Cover the fixed smear with crystal violet stain for 30
– 60 seconds
3. Rapidly wash off the stain with clean water
4. Tip of all the water, and cover the smear with Gram’s
iodine for 30 – 60 seconds
5. Wash off the iodine with clean water
6. Decolorize rapidly with acetone-alcohol for 30 seconds &
wash immediately with clean water.
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35. 7.Cover the smear with Neutral red or Safranin
for 2 minutes
8. Wash off the stain with clean water.
9.Wipe the back of the slide clean, and place it
in a draining rack for the smear to air dry
10.Examine the smear microscopically, first with the
40x objective to check the staining and to see the
distribution of material, and then with oil
immersion objective to report the bacteria and
cells.
Gram stain
(cont’d)
35
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36. Gram positive Gram negative
Step 1: Crystal
Violet
Step 2: Gram’s
Iodine
36
Step 3:
Decolorization
(Aceton-
Alcohol)
Step 4: Safranin
Red
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37. Results
Gram positive bacteria
…..……………Purple(blue)
Yeast cells
……………………………………Dark purple
Gram negative bacteria …….…………Pale
to red
Nuclei of pus cell
…….………….………Red
Epithelia cells
Gram stain
(cont’d)
37
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38. Gram stain (cont’d)
Reporting of Gram’s stained smear
The report should include the following:
1. Number of bacteria present whether
many, moderate, few or scanty
2. Gram reaction of the bacteria whether
Gram positive or Gram negative
3. Morphology and arrangement of the
bacteria whether cocci, diplococci,
streptococci, rods, or coccobacili; also
whether the organisms are intracellular
4. Presence and number of pus cells.
5. Presence of yeast cells and epithelia
cells. 38
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39. Gram stain
(cont’d)
AbateA. 39
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40. Gram stain
(cont’d)
Figure: Gram positive
an
d Gram negative
bacteria
40
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41. Gram stain
(cont’d)
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42. Gram stain
(cont’d)
42
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43. Factors which contribute to false
Gram’s staining result
I. False Gram positive reaction
• Preparation of too tick smear – the stain will
not be fully washed.
• Presence of sediment in crystal violet - this
can be avoided by filtering the stain before
use.
• Decolourizing time - when insufficient
decolourization time is used or ( when not
fully decolorized).
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44. GRAM’S STAIN …
II. False Gram Negative reactions
– Making smear from old culture.
– Cell wall damage due to antibiotic.
– Excessive heat fixation of smear.
– Over decolourization of smear (decolourization for longer
time).
– Use of an iodine solution which is too old (i.e. yellow
instead of brown in colour) (its mordant effect will be
decreased).
Solution for the above problem
• Always check new batches of stain and reagent for correct
staining reaction using a smear containing known gram
positive and gram negative organism as a control.
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45. Gram stain
(cont’d)
45
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46. Ziehl-Neelsen technique
• Developed by Paul Ehrlich in1882, and modified
by Ziehl and Neelson
• Ziehl-Neelson stain is used for staining
Mycobacteria which are hardly stained by Gram‘s
staining method.
• Once the Mycobacteria is stained with primary stain
it can not be decolorized with acid, so named as
acid-fast bacteria.
• The high lipid content of Mycobacteria accounts for
unusual properties of the organism.
E.g Relative impermeability to stains, acid
fastness, unusual resistance to killing by acid and
alkali
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47. Ziehl-Neelsen (cont’d)
Principle
 Sputum smear is heat –fixed, flooded with a
solution of carbolfusin (a mixture of basic
fuschin and phenol) and heated until steam
rises.
 The heating will facilitate penetration
(entrance) of the primary stain into the
bacterium.
 After washing with water, the slide is covered with
3% HCl (decolourizer). Then washed with water
and flooded with methylene blue ( M.
tuberculosis) and malachite green (M. leprae).
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48. Ziehl-Neelsen (cont’d)
Materials for AFB staining
• Materials
– Wire loop or applicator stick
– Microscope slide
– Marking pen
– Bunsen burner or sprit
– Staining racks (staining rods)
– Slide rack
• Stain: Carbolfuchsin, 3% Acid alcohol,
Methylene blue (malachite green)
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49. Ziehl Neelson stain reagents
preparation
A. Carbolfuchsin
- Solution A (saturated solution of basic
fuchsin) Basic fuchsin ………….. 3gm
Ethanol …………………..100ml
- Solution B ( phenol aqueous solution, 50g/L (
5%) Phenol ……………………..... 10gm
Distilled water ……………… 200ml
- Mix 10 ml of solution A with 90ml of solution
B.
-Transfer resulting mixture to a glass
stoppered amber bottle and label.
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50. Ziehl-Neelsen (cont’d)
B. 3% Acid –alcohol (decolorizer)
– Hydrochloric acid (conc) ……………
3ml
– Ethanol
………………………………..97ml
– Label the bottle.
N.B It is recommended to use 25 %
H2SO4
a
s decolorizing solution
• Content per liter:
– sulfuric acid (minimum 95%)--------250
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51. Ziehl-Neelsen (cont’d)
Preparation:
1. Measure 750ml of water into a 2-liter flask
2. Measure 250mL of concentrated sulphuric
acid in a cylinder
3. Pour it slowly into the flask containing
the water, directing the flow of acid
gently along the inner side of the flask
4. Stop and swirl flask regularly as a lot of
heat is generated until all acid is added
5. Mix well and allow to cool before use
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52. Ziehl-Neelsen (cont’d)
C. Methylene Blue Stain
– Methylene blue chloride .............................. 0.5g
– Distilled water......................................... 100ml
D. Malachite green
– Malachite green …………………. 0.5g
– Distilled water ……………………. 100ml
• Using a pestle and mortar, grind the malachite green
crystal to a powder. Dissolve the grind powder in 100ml
distilled water and store in a dark brown bottle.
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53. Ziehl-Neelsen (cont’d)
Hot and cold Ziehl Nelson technique
• In the hot Zn technique the phenol – carbolfuchsin
stain is heated to enable the dye to penetrate the
waxy Mycobacterial cell wall.
• Techniques that do no heat the stain are referred to
as ‘cold’ techniques. In these, a penetration of the
stain is usually achieved by increasing the
concentration of basic fuchsin and phenol.
• Comparison between the ‘hot’ and ‘cold’ method has
shown that both M. leprae and M. tuberculosis stain
less well by the cold method.
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54. Ziehl-Neelsen (cont’d)
Difference between Acid fastness of
Mycobacterium species
• M. tuberculosis and M. ulcerance are strongly acid
fast, when staining specimens for these species, a
3% v/v acid solution or 25%H2SO4 is used to
decolorize the smear.
• M. lepreae is only weakly acid fast. A 1% v/v acid
or 5% H2SO4 decolorizing solution is
therefore used for M. lepreae smears and also
different staining and decolorizing times.
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55. Ziehl-Neelsen (cont’d)
Procedure
1. Use blood specked and/or purulent sputum (do not use saliva)
and a variety of specimens (eg. body fluid, tissue, pus, CSF,
urine etc.) from EPTB can be used can be for smear
preparation
2. Heat fix the dried smear,
– Alcohol fixation is recommended when the smear has
not been prepared from sodium Hypochlorite (bleach)
– Heat fixation of untreated sputum will not kill M.
tuberculosis where as alcohol – fixation is
bactericidal.
3. Cover the smear with carbolfuchsin stain
4. Heat the stain until vapor just begins to rise. Do not over heat
– Allow the heated stain to remain on the slide for 5 minutes.
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56. 5. Wash off the stain with clean water
6. Cover the smear with 3% acid alcohol for 5 minutes or
until the smear is sufficiently decolorized. i.e. pale
pink.
7. Wash well with clean water
8. Cover the smear with methylene blue for 1 – 2
minutes.
9. Wash off the stain with clean water.
10. Wipe the back of the slide clean, and place it in a
draining rack for the smear to air dry (do not blot
dry)
11. Examine the smear microscopically, using the
100x oil immersion object.
12. Examine a minimum of 300 fields systematically
before the smear is reported as negative.
Ziehl-Neelsen (cont’d)
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57. Ziehl-Neelsen (cont’d)
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58. Results of Ziehl-Neelson staining
• AFB............. Red, straight or slightly curved
rods, occurring single or in a small groups
• Cells......................................... Blue
• Background Material ……….. Blue
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59. Ziehl-Neelsen (cont’d)
REPORTING OF SPUTUM SMEAR
• When any definite red bacilli are seen report
the smear as AFB positive and give an
indication of the number of bacteria present
as follows:
When
1- 9 AFB /100 fields --------------- exact
number
10 – 100 AFB/100 fields ------------------ +
1 – 10 AFB/field --------------------------- ++
> 10 AFB/field ------------------ -----------+++
(source : Manual basic techniques for a
health lab WHO,2003)
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60. REPORTING OF SPUTUM SMEAR
N.B: AFB means number of acid fast bacilli seen.
• when very few AFB are seen E.g When only
one or
two AFB are seen, request further
specimen for
Ziehl-Neelsen (cont’d)
examination.
• When no AFB are seen after examining 300
fields report the smear as ‘ No AFB seen’ do
not report as “Negative” because organisms
may be present but not seen in those field
examined.
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61. Sensitivity of direct microscopy
• 5,000 to 10,000 tubercle bacilli /ml of sputum are
required
• The sensitivity can further be improved by examination
of more than one smear from a patient.
• So up to three specimens (spot– morning- spot)
are required
NB: Studies have shown that centrifugation of bleach
treated sputum increased the sensitivity of direct
microscopy from 43.4% to 76.3%, increasing the
number of smear-positive patients detected. Sensitivity
is significantly increased in HIV/AIDS patients.
Ziehl-Neelsen (cont’d)
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62. Sodium hypochlorite
centrifugation
technique
• To concentrate AFB
1. Transfer 1-2 ml of sputum to a screw -
cap container of 15m-20 ml capacity
2. Add an equal volume of concentrated
sodium hypochlorite (bleach) solution
and mix well.
3. Leave at room temperature for 10-15
minutes, shake at intervals to break down
the mucus in the sputum.
4. Add about 8 ml of distilled water, and mix
well.
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63. 5. Centrifuge at 3000g for 15 minutes.
N.B: When centrifugation is not possible, leave the
NaOCl treated sputum to sediment overnight.
6. Using a glass Pasteur pipette, remove and
discard the supernatant fluid. Mix the sediment.
7. Transfer a drop of the well mixed sediment to a
clean scratch free glass slide. Spread the
sediment to make a thin preparation and allow to
air dry.
8. Heat fix the smear and stain it using Ziehl
Neelsen technique and examine microscopically.
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64. Factors which bring false positive results
• Utilization of slides that have been positive.
These should be discarded.
• Using scratched slides on which deposits of
stain may look like bacilli
• Using unfiltered fuchsin which may contain
crystals .
• Carelessness in heating the fuchsin, allowing it
to dry and crystallize on the smear
• Inadequate decolorizing of the smear.
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65. Fluorescence staining for
Mycobacteria
• Fluorescence microscopy uses illumination
from either a quartz halogen lamp or a high
pressure mercury vapor lamp.
• The advantage of fluorescence microscopy is
that a low magnification (20x, 40x) is used to
scan smears, allowing a much larger of the
smear to be seen and resulting in more rapid
examination.
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66. Fluorescence staining (cont’d)
Reagents:
• Solution1 (Auramine O)
– Auramine--------------------------------------------------0.1gm
– 95% Ethanol----------------------------------------------10 ml
– Dissolve
• Solution 2 (Phenol)
– Phenol crystal--------------------------------------3.0 gm
– Distilled water-------------------------------------87ml
– Dissolve
– Mix solution 1 and 2 and store in amber bottle away
from light and heat.
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67. Fluorescence staining (cont’d)
Decolorizing solution
3% acid alcohol
Counter stain
Either Potassium permanganate or acridine orange can be used
• Potassium permanganate
Potassium permanganate (KMnO4) ------------------0.5g Distilled
water 100ml
Dissolve and store in amber bottle
• Acridine orange
Acridine orange--------------------------------------------0.01g
Anhydrous dibasic sodium phosphate (NA2HPO4) -0.01g Distilled
water-----------------------------------------------100ml
Dissolve sodium phosphate in distilled water. Add Acridine orange.
Store in amber bottle.
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68. Fluorescence staining (cont’d)
Procedure
1.Place the dried labeled smear on
staining rack
2.Flood entire smear with Auramine O and
allow to stain for 15min ensuring that
staining solution remains on smear
3.Rinse with distilled water and drain. Tap
water contains chlorine which may
interfere with fluorescence.
4.Decolorize with 3% acid alcohol for two
minutes
5.Rinse with distilled water and drain
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69. 6. Flood the smear with Potassium
permanganate or acridine orange and allow to
counter stain for two minutes.
– Time is critical with potassium
permanganate because counter staining
for longer time may quench the
fluorescence of acid fast bacilli.
7. Rinse with distilled water and drain
8. Allow the smear to air dry. Do not blot. Read as
soon as possible.
– Examine fluorescence stained smear
within 24 hours as the fluorescence may
Fluorescence staining (cont’d)
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70. Fluorescence staining
(cont’d)
Figure: Fluorescent stained Mycobacteria
under fluorescence microscope
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71. 3. Special Staining
method
• These are stains, which are used to stain
capsules and spores.
There are two types of special staining
methods
A.Capsule Staining method
B.Spore staining method
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72. A. Capsule staining method
• This technique is used for showing the
presence of capsules around bacteria
Required reagents
• Crystal Violet ………………………..10g/l (1%
w/v) stain
• Copper sulphate……………………200g/l
(20%w/v)
Special
Staining …
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73. Procedure
1. Fix the direct smear using alcohol. Do not heat fix .
Capsules stick well to glass, and heat may destroy the
capsule
Note: when preparing the smear, mix the organism
suspension with a drop of normal serum this will help to
show the capsules
2 Cover the smear with crystal violet stain and heat gently
until the steam just begins to raise, leave to stain for 1
minute.
3 Wash off the stain with the copper sulfate solution
N.B don’t use water
4 Wipe the back of the slide clean and air dry
5 Examine the smear microscopically first with the 40x,
then with the 100 x oil immersion objective to look for
Special
Staining …
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74. Result
• Bacterial cell dark
purple
• Capsule outline ---------------------------------- Pale
blue
Special
Staining …
Fig.: Capsule stain of Enterobacter aerogenes
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75. B. Spore staining
method
Materials needed:
– Dye kit (malachite green,
Safranin)
– Staining rack
– Hot plate
– Paper towel (cut the size of the
slide)
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76. Special Staining …
 It is based on the binding of the malachite green
and the permeability of the spore vs. cell wall.
– The steaming helps the malachite green to
permeate the spore wall.
• The primary dye malachite green is a relatively
weakly binding dye to the cell wall and spore
wall.
– In fact, if washed well with water, the dye
comes right out of the cell wall.
– That is why there does not needa
decolourizer in this stain.
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77. Procedure
1.Make a smear, air-dry and heat-fix.
2.Put a beaker of water on the hot plate and boil
until steam is coming up from the water. Then
turn the hot plate down so that the water is
barely boiling.
3.Place the wire stain rack over the beaker which
now has steam coming up from the boiled
water.
4.Cut a small piece of paper towel and place it
on top of the smear on the slide.
Special
Staining …
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78. 5. Flood the smear with the primary dye, malachite green, and
leave for 5 minutes. Keep the paper towel moist with the
malachite green. DO NOT let the dye dry on the towel.
6. Remove and discard the small paper towel piece.
7. Wash really WELL with water and move the slide and wire
rack
Special
Staining …
from the boiling water to the regular stain tray
8. Place the smear in the stain jar or flood the smear
with the counter stain dye, Safranin, and leave for 1
minute.
9. Wash off the excess safranin with water, blot dry, and
observe using oil immersion microscopy. With this
endospore staining procedure, endospores will stain green
while vegetative bacteria will stain red
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79. Special
Staining …
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80. The cold method spore stain
• Without heat you have to really rough up the
spore wall to get in the dye.
• Heat fix the smear by running the slide through
the flame about 20 times, and leave malachite
green on for 20 minutes during the stain
process.
Special
Staining …
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81. Special
Staining …
Interpretation
• Spores will be a light green:
• Vegetative cell walls will pick up the counter
stain safranin and become red(Red)
Notice: the position of the spore could be --
terminal, central, or subterminal and this may be
helpful in the identification of the unknown
species of bacteria.
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