Bacterial Staining Methods
Staining Methods
Bacterial Staining
Simple Staining
(Monochrome staining)
Solution required
Loeffler' methylene blue
|
Methylene blue
chloride |
0.3 g |
|
95% Ethanol |
30 ml |
|
0.1 % KOH |
100 ml |
Dissolve methlylene blue
chloride in ethanol. Add 0.1% KOH. Filter the solution before use. Store at
room temperature.
OR
a.
1% Crystal violet
OR
b.
Carbol fuchsin
(see acid fast staining). Dilute 10 times before use.
Procedure
1. Prepare the smear and heat fix
2. Treat the smear with 5-7
drops of staining solution.
3. Allow the smear to react as
follows
Loeffler's methylene blue for 120 to 150 seconds.
OR
Per cent crystal violet for 60 to 120 seconds.
OR
Diluted carbol fuchsin for 15 to 30 seconds.
4. Pour off the staining
solution and wash the slide by gentle
5. Flow of tap water.
6. Dry the slide in air.
7. Examine under oil immersion lens.
Bacteria in Milk
(Breed's smear count method)
Solution Required
A. Xylene or chloroform.
B. 95% ethyl alcohol.
C. Breed's methylene blue.
|
Methylene blue chloride |
0.3 g |
|
95% Ethanol |
30 ml |
|
0.1 % KOH |
100 ml |
Dissolve methylene blue
chloride in ethanol. Then add above solution to phenol in water. Mix
D. 90% alcohol.
Procedure
1.
Mark a clean
slide with glass marking pencil to make one centimetre square.
2.
Place 0.01 ml of milk sample in the centre of the square.
3.
Spread the sampfe
with inoculating needle to form uniform smear covering the square.
4.
Heat fix the
smear and treat the slide with solution a for about 1 minute.
5.
Treat the smear
with solution b for 3 minutes.
6.
Treat the smear
with solution c for 2 minutes.
7.
Wash the smear
with solution d till smear appears faintly blue. (approx 30 seconds)
8.
Dry in air and
examine under oil immersion lens.
Result: Bacteria appear blue in colour.
Staining of Azotobacter Cysts
Solution Required
a. Staining solution
|
Glacial acetic acid |
8.5 ml |
|
Sodium sulphate (anhydrous) |
3.25 g |
|
Neutral red |
200.0 g |
|
Light green S.F. yellowish |
200.0 mg |
|
Ethanol |
50.0 ml |
After 15 minutes of incubation remove amorphous precipitate by filtering through
0.5 pm membrane filter.
Procedure
1.
Suspend the growth of the bacteria in the solution a for
wet mount preparation.
2. Observe under oil immersion lens.
Result
1. Vegetative cens appear light yellowish green.
2.
The early stage
of encystment shows dark green cytoplasm.
3.
Cyst: Intine
appears colourless
4.
Exine appears
brownish red.
5. Cytoplasm appears green.
Staining of Actinomycetes
Solutions required
a)
Absolute methanol
b)
Hucker's crystal
violet (see grams staining)
Procedure
1.
Treat the growth
on the coverslip with few drops of solution a for 15 minutes.
2.
Wash with tap
water and blot dry.
3.
Stain with
solution b for 1 minute.
4.
Wash with tap
water, dry in air.
5.
Observe under oil
immersion lens.
Results Mycelium
and spores appears violet in colour.
Note Slide
cultures must first be dried by placing them over boiling water for about 5
minutes until agar has dried.
Staining of Actinomycetes
Solutions required
a. Staining solution
|
Bismark brown stain (0.1%
w/v) |
40 ml |
|
Toluidine blue stain (0.1%
w/v) |
40 ml |
|
Saturated ammonium sulphate
solution Mix together |
20 ml |
Procedure
1.
Grow the culture
of actinomycetes on the surface of sterile cellophane placed on placed on
solidified nutrient agar medium.
2.
Remove the
cellophane bearing growth from the agar surface.
3.
Treat the growth
for 2 minutes with the solutions a.
4.
Wash the slide
with tap water.
5.
Air dry and
observe under oil immersion lens.
Result: Vegetative mycelium appears light yellow and the spores blue.
Staining of Mycoplasmas Colony
Dine's method
Solution required
|
Methylene blue |
2.5 g |
|
Azure II |
1.25 g |
|
Sodium carbonate |
0.25 g |
|
Benzoic acid |
0.20 g |
|
Maltose |
10.0 g |
|
Distilled water to |
100 ml |
Procedure
1.
Flood the plate
containing suspected colonies of mycoplasmas with 1:9 dilued solutions in
distilled water.
2.
Remove the stain.
3.
Examine the plate
under low power microscope.
Result: The colony appears granular, royal blue to greenish blue in colour.
Staining with cresyl-fast violet
Solution required
a. Cresyl-fast violet solution
Stock solation
|
Cresyl - fast
violet |
1.0 g |
|
Distilled water to (pH-3.7, adjusted with Glacial acetic
acid 1-5 drops/100 ml) 100 ml
|
|
Allow the solution to
ripen for 48 hours.
Working solution
|
Stock solution |
20.0 ml |
|
Sodium chloride |
0.05 g |
|
Maltose |
7.0 g |
Mix
sodium chloride in stock solution. Filter. Add maltose.
Procedure
Similar to that of Dienes method.
Result: Colonies of mycoplasmas appear red to purple.
Staining method for Bruce11 Abortus
Solutions required
a. Carbol
fuchsin 10 times diluted (see acid fast staining ZNCF method.)
b. 0.5
acetic acide.
c. Loeffler's methylene blue (see monochrome staining).
Procedure
a.
Prepare
the smear do not heat fix.
b.
Treat
the smear for 15 min with solution a.
c.
Drain
the solution a treat with solution b for 1'2-20 seconds.
d.
Wash
with water and treat with solution c for 1-2
min.
e.
Wash
with water, dry and observe under oil immersion lens.
Results: Brucella abortus appears red in colour. Other organisms appear blue in colour.
Acid Fast Staining
1. Ziehl-Neelsen method
Solutions required
a. Carbol fuchsin (Ziehl-Neelsen)
|
Basic fuchsin |
1.0 g |
|
95% ethanol |
10 ml |
|
5% phenol |
100 ml |
Dissolve
basic fuchsin in ethanol then mix with phenol. Allow this solution to ripen for
1-2 weeks.
a)
Sulphuric
acid (20% solution)
b)
Leffler's
methylene blue (see monochrome staining)
OR
1% Malachite green in water.
Procedure
1.
Prepare
a smear and heat fix.
2.
Treat
the smear with solution a and heat the slide by gentle flame for five minutes.
(The stain must not be allowed to evaporate and dry on the slide.)
3.
Allow
the slide to cool.
4.
Wash
with water.
5.
Treat
with solution b till red cnlnl~rn o longer comes out (usually for 90 secs.)
6.
Wash
with water.
7.
Treat
the smear with solution c for 20 to 30 seconds.
8. Wash, air dry and examine under oil immersion lens.
Results: Acid fast cells
stain bright red, while non-acid fast are stained green or blue colour
according to solution c used.
Note:
1.
Following
decolourisers are usually used for different
organisms.
2.
5
per cent sulphuric acid for M. leprae.
3.
3
per cent HCl in 95% ethanol for M. tuberculosis.
4. 1 per cent sulphuric acid to demonstrate acid fast clubs of Actinomyces and Nocardia. When malachite green is used as a counterstain (solution c), use deep green filter in the light source for microscopic observasion.
2.
Method of Gross
Solutions required
a. Basic fuchsin with Tween 80
|
Basic
fuchsin chloride |
2.0
g |
|
Phenol |
6.0 ml |
|
95%
ethanol |
12.5
ml |
|
distilled water to |
150.0
ml |
Dissolve basic fuchsin chloride in
phenol at 80°C. Add 95% ethyl alcohol by stirring. Make the final volume to 150
ml with distilled water. Allow it to ripen for 1-2 weeks. Filter before use and
add Tween 80.
b. 3%
HCl in ethanol.
c. Loeffler's methylene blue (see monochrome staining).
Procedure
1.
Prepare
the smear and heat fix.
2.
Treat
the smear with solution a for 5-10 min.
3.
Wash
the smear with water.
4.
Wash
the smear with solution b till red colour not longer comes out (usually 120
seconds).
5.
Wash
the slide with water.
6.
Treat
the slide with solution c for 3 min.
7.
Wash,
air dry and examine under oil immersion lens.
Results: Acid fast organisms appear red. Non acid fast organisms and background appear blue in colour.
3. Method of Trauant et al.
Solutions required
a. Staining solutions
|
Auramine "0" |
0.3 g |
|
Phenol |
3.0 g |
|
Distilled water |
97.0 ml |
Dissolve the phenol in water with gentle heat. Add the auramine slowly. Shake vigorously until dissolved, filter and store in dark stopped bottle.
b. Traunt's decolourizer.
|
NaCl |
0.5 g |
|
HCl |
0.5 ml |
|
75% Ethyl alcohol to |
100 ml |
c. Potassium
permanganate solution (1 to 1000) aqueous. Procedure
Procedure
1.
Prepare
the smear and heat fix.
2.
Treat
it with solution a for 15 minutes.
3.
Wash
the smear with water and treat the smear with solution b for 5 minutes.
4.
Wash
with water then treat the smear with solution c for 30 seconds.
5.
Wash
air dry & examine under 8 mm dry objective &a high power eye piece (20x).
Result: Tubercle bacilli appear luminescent yellow coloured. Background appears dark.
Negative Staining
Solution required
a. Nigrosin solution
|
Nigrison (G.T Gurr) |
10 g |
|
Distilled water to |
1000ml |
Dissolve nigrosin in warm distilled
water (require an hour) & filter. Add formalin
0.5 % (i.e., formaldehyde 0.19 %) as a
preservative.
OR
2 % congored solution.
OR
India ink.
Procedure
Take
a loopful bacterial suspension and a drop of solution a at one end of clean
glass slide. Mix.
Spread
this mixture as a film using the another slide.
Allow
it to air dry and examine under high power and oil immersion lens.
Result: Bacteria appear colourless with dark background, blue black with
nigrosin, red with congored and blue with India ink.
Note: Film should not be too thick or too thin.
Capsule staining
A. Positive staining methods
1. Method of Hiss (modification of Anthony)
Solution required
1.
1%
Crystal violet.
2.
20%
CuSO4.5H2O (aqueous solution)
Procedure:
1.
Prepare
a smear. Do not heat fm.
2.
Treat
the slide with solution a for 2 min.
3.
Remove
the solution a by washing with solution b.
4.
Dry
and examine under oil immersion lens.
Result: cells appear dark purple, capsules appear pale blue.
2. Method of Moller
Solutions required
a. Moller's fixations (see fixatives)
b. Moller's crystal violet
solution
|
Crystal violet chloride |
0.5 g |
|
95% ethyl alcohol |
10 ml |
|
Distilled water to |
100 ml |
Dissolve crystal violet chloride in 95% ethyl alcohol. Then add distilled water.
c. Saturated aqueous CuS04.5H20 solution
Procedure
1. Prepare the smear and treat
the smear with solution a for 15 seconds. Pour off the solution a and dry the smear.
2.
Treat the smear with solution b for 2 minutes. Pour off the solution.
3. Treat the smear with solution c for 10 seconds
4. Dry the smear and examine under oil immersion lens.
Result: Capsules appear
light purpule violet. Bacterial cells appear dark violet.
B. Negative Stain methods
1. Method of Howie an Kirkpatrick (Releif staining)
Solution required
a. Staning solution
10 per cent water soluble eosin,
'Yellowish' or
'bluish' or erythrosine in distilled water 40 ml Serum (human, rabbit, sheep or ox heated at 56OC for thirty minutes) 100 ml Crystal of thymol
Mix well. Allow the mixture to stand at room temperature for several days. Centrifuge and store the supernatant fluid at room temperature.
Procedure
1.
Take
one drop of suspension and one drop of solution a at Y one end of slide. Mix
well. Wait for 1 minute.
2.
Spread
this mixture with the help of another slide as a thin
3. Dry without heating and examine under oil immersion lens.
Result: Background and bacteria appear red while
capsule appear stained or lightly stained.
2. Method of Maneval's
Solution required
a.
1% congored.
b.
Maneval's stain I
|
5% aqueous solution of phenol |
30.0 ml |
|
20% acetic acid |
10.0 ml |
|
30% FeC12.6H2O |
4.0 ml |
|
1% acid
fuchsin |
2.0 ml |
Mix thoroughly.
Procedure
1. Take
one drop of bacterial suspension and one drop of solution a at one end of
slide. Mix. Spread over a slide as a thin film with the help of another slide.
2. Air
dry the smear.
3. Treat
the smear with solution b for 1 minutes.
4. Wash with distilled water, air dry and examine under oil immersion lens. Dissolve the nigrosin in warm distilled. water, add the formalin and filter.
b. Loeffler's alkaline methylene blue. (see monochrome staining).
Procedure
1. Take
one looful of culture on a clean slide.
2. Add
one looful of freshly filtered solution a.
3. Mix,
allow to dry in air and fix with gentle heat.
4. Treat
the smear with solution b for 30s.
5. Rinse rapidly in water, air dry and observe under oil immersion lens.
Result: Bacterial cell appear blue, capsule appear colourless unstained against a dark grey background of nigrosin.
Note: Safranin may be used in place of the methylene blue.
(solution b)
Dry India ink film method
Solutions required:
a.
6%
glucose solution in water.
b.
8%
aqueous solution of nigrosin
OR
Indian
ink.
a.
Methanol
or Leishman stain.
b. Methyl violet stain (1% aqueous solution).
Procedure:
1.
Take
one drop of bacterial suspension and one drop of solution a at one end of
slide.
2.
Add
one drop of solution b to it.
3.
Mix
and spread the mixture over a slide as a thin film with the help of another
slide.
4.
Dry
the smear in air.
5.
Treat
the smear by solution c. Pour off immediately, dry in air.
6.
Treat
the smear with solution d for 1-2 minutes.
7. Wash with water, dry and examine under oil immersion lens.
Result: Capsules appear colourless, cells appear purpule. Background appear grey-
India ink preparation (Wet film)
Solution required:
a. Indian ink
Procedure:
1. Place one loopful of solution a on a perfectly clean glass slide.
2. Emulsify a small portion of solid bacterial culture in the drop of
solution a or mix in a loopful of liquid culture.
3. Cover the mixture with a clean cover glass and press the
4.
latter
down firmly, to form a very thin film.
5. Seal the edges of the cover glass with paraffin wax or other suitable medium.
6. Examine the smear under oil-immersion objective.
Result: Bacteria highly refractile, surrounded by a clear zone against a dark grey background of ink particles. Noncapsulated bacteria do not show this clear zone.
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