Laboratory Manual
on
Microbial Morphology and Taxonomy
Surajit Mandai
S. K. Tomar
Rameshwar Singh
Dairy Microbiology Division
National Dairy Research Institute (ICAR)
(Deemed University)
Karnal-132001, Haryana (India)
Laboratory Manual
on
Microbial Morphology and Taxonomy
Surajit MandaI
S. K. Tomar
Rameshwar Singh
Dairy Microbiology Division
National Dairy Research Institute
(leAR)
(Deemed University)
Karnal - 132001, Baryana (India)
Published by:
Director
National Dairy Research Institute (Deemed Universi lY)
Kamal-132001, Haryana, India
Tel: 91-0184-2252800; Fax: 91-01842250042
First edition: March 2012
NDRI Publication No_ 7712012
Price: Rs_ 100/·
Pri"ted at:
Intech Printers & Publishers
# 353, Mugha\ Canal, Kamal - 13200 1
Tel. 0184-4043541 , 3292951
PREFACE
Microbes are microscopic organisms that comprises either a single cell (unicellular), cell
clusters, or no cell at all (acellular). Bacteria are often viewed as the cause of diseases in
humans and animals. Some bacteria are useful, for example certain bacteria aids in
digestion. Bacteria make up the base of the food web in many environments. Bacteria are
of such immense importance because of their extreme flexibility, capacity for rapid
growth and reproduction, and great age. Bacteria cannot be visualized by naked eye and
hence, observed through microscopes. A bacterium has a cell envelope which includes a
I~yered cell wall and external surface adherents. The appendages of cell wall include
flagellae-the organs of locomotion and fimbriae which help in adhesion of bacteria.
Internally the bacterium has loose arrangement of DNA, i.e. nuclear apparatus surrounded
by an amorphous cytoplasm which contains ribosomes. Mesosomes and inclusiongranules
are otloer structures present in bacterium. Microorganisms have diverse morphological
features such as staining properties (e.g. Gram positive or negative, spore formation,
capsules etc), shape (cocci, bacilli etc.), size and arrangements (cells in paris, tetrads,
chains, clusters etc), presence of specialized structures (flagella, fimbriae) etc. Therefore,
morphological f~atures and ultra-structures are important for identification, grouping, and
classification microorganisms.
The emphasis has been given on the process development of introductory understanding
of techniques for conducting experiment, analysis the data and interprets the results. A
great effort has been made to confer sufficient scientific understanding of each exercise,
materials required for the experiment, experimental protocol for smooth conducting of the
practical.
The authors sincerely thank Dr. A.K. Srivastava, Director and Vice-Chancellor, National
Dairy Research Institute (Deemed University), for entrusting and granted the permission
for preparing the laboratory manual. The authors also convey the gratitude .to Dr. S.L.
Goswarni (IDR), Dr. G.R. Patil (IDA) and all other colleagues of Dairy Microbiology
Division for their support and guidance. Authors thank ICAR for providing funds for
manual preparation under the scheme for strengthening and development of agricultural
education.
Authors look forward to the acceptance of manual amongst students, scholars and
teachers.
S. K. Tomar
Surajlt MandaI
Rameshwar Singh
"I'
Content
Pg. no.
Exercise I
Simple staining for visualization of bacteria under microscope
I
Exercise 2
Gram staining for differentiation of bacteria based on cell wall
6
Exercise 3
Examination of shape and arrangement of bacteria
12
Exercise 4
Measurement of size of microorganisms by micrometry
16
Exercise 5
Examination of bacterial spores
25
Exercise 6
Staining of acid fast bacteria
30
Exercise 7
Staining of bacterial cell wall
34
Exercise 8
Staining of bacterial capsules
37
Exercise 9
Staining of bacterial flagella
42
Exercise 10
Staining of bacterial inclusion! storage bodies
49
Exercise II
Staining of bacterial nucleoids
55
Exercise 12
Examination of bacterial motility
58
Exercise \3
Preparation of bacterial protoplasts
60
Exercise 14
Preparation of bacterial spheroplasts
64
Exercise IS
Study of morphology of yeasts
66
Exercise 16
Study of morphology of moulds
69
Exercise 17
Detection and enumeration of bacteriophages in cheese whey
71
Exercise 18
Application of computer software in bacterial identification
73
Exercise 19
Examination of ultra-structure of microbial cell using Scanning
Electron Microscopy (SEM)
78
Exercise 20
Examination of ultra-structure of microbial cell using Transmission
Electron Microscopy (TEM)
82
Viva voice
85
References
86
I
Exercise 1- Simple stainh ·
Introduction
.( visualization of bacteria under microscope
Microorganisms are too small (generally in micrometer range size) to be seen by our un-
aided eyes. Therefore, to see these microorganisms, microscope is required, which
magnifies the microorganisms to some enlarged size to be seell by our eye.
Microorganisms contain ..... 99% water and thus the optical density or refractive index of
microbial cells is nearly equal to the surrounding medium. i.e. water. Thus, eveD under
microscope, the size of microorganisms are enlarged to visible range, it is difficult to
differentiate microorganisms from the surrounding aqueous medium. To improve the
visibility, the optical density of microorganisms is enhanced by applying stains or dyes.
Simple stains used to increase color contrast in cells. They are generally able to react with
all types of bacteria. Aniline dyes, such as methylene blue, crystal violet, basic fuchsin,
eosin Y, and safranin 0 , are examples of simple stains. These are basic dyes, as opposed
to acidic dyes, and bind with the acidic portions of cells. The dye particles bind to the
microbial surfaces and thus increase the optical density as compared to surrounding
medium for visualization under microscope. Optical density of microorganisms can be
enhanced by using positive staining (staining of bacteria) or negative staining (staining of
surrounding medium). Bacteria are routinely stained with different dyes (positive staining
by methylene blue, cry tal violet etc) in order to reveal different properties and to
enhance contrast for vi wing with conventional bright field microscopy. In negative
staining. smears are pro uced by mixing material with acidic dyes (such as nigrosin). The
acidic dyes have negati e charge and are repelled by the negatively charged cell walls.
Cells remain unstained gainst a dark background. Microscopic slide of given bacterial
culture is prepared by imple staining protocol and observe under IOOx (oil immersion)
objective to see the mO$hological spape and arrangement of bacteria.
I
Bacterial cultures: dne bacillus and onc coccus cultures
l
Materials
•
• Simple stain soluti n: Methylene bluel crystal violet solution (positive staining) or
nigrosin solution ( egative staining)
• Microscopic slides
• Inoculating loop
• Immersion oil
I
\
• Microscope (10M objective lens)
• Gas burner
• Tissue papers
Protocol
a) Positive staining
I. Prepare two smears from broth culture of Lae/dfaeillus spp. and Lae/oeoeeus spp. on
clean and grease free microscopic slides and heat fix over the gas flame.
2. Flood the smear with the methylene blue or crystal violet solution for 1-2 min.
I
3. Gently wash the slide with water to remove the excess dye.
4. Blot dry followed by drying over the gas flame of the stained smear on the slide and
examine under the microscope (IOOx, oil immersion objective).
5. Draw and identify the shape and arrangement of the bacterial cells in the circles
below.
b) Negative staining
I. Take two clean and grease free microscopic slides.
2. Transfer two loopful of broth culture and one loopful of nigrosin solution on the slide
and mix thoroughly with the help of inoculating loop.
3. Spread unifonnly the mixture of bacterial culture and nigrosin dye to get a thin and
transparent smear on the slide.
4. Allow to air dry of the smear and examin e under the microscope ((IOOx, oil
immersion objectives
5. Draw and identify the shape and arrangement of the bacterial cells in the circles
below.
Observations
Results
Inferences
1. Ziebls Carbol-fucbsln
Solution A: Basic fuchsin solution 10 ml:
Basic fuchsin dye
0.3 g
Ethyl alcohol (95%)
IOml
Solution B: 5% carbolic acid solution
100ml
Phenol
5g
Distilled water
95 ml
Mix solutions A and B
2. Ammonium Oxalate Crystal Violet (Hucker's)
Solution A:
Crystal violet dye
0.2 g
Ethyl alcohol (95%)
20 ml
Solution B:
Ammonium oxalate
0.8 g
Distilled water
80 ml
Mix solutions A and B
3. Crystal Violet in Dilute Alcohol
Crystal violet dye
2g
Ethyl alcohol (95%)
20ml
Distilled water
80 mI
4. Loeffler's Alkaline Methylene Blue
Solution A: Metbylene blue solution in alcohol 30 ml
Methylene blue dye
0.3 g
Ethyl alcohol (95%)
30 mI
Solution B: Sol KOH in distilled water (1:10,000) 100 ml
Dilute KOH (0.0 I % by weight)
100 ml
Mix solutions A and B
5. Methylene Blue in Dilute Alcohol
Methylene blue dye
0.3 g
Ethyl alcohol (95%)
30 mI
Distilled water
100m!
3
6. Carbol Rose Bengal
Rose bengal dye
Ig
Phenol (5% aqueous solution)
100 ml
CaCh
0.01-0.03 g
(The amount ofCaCl, added determines the intensity of staining.)
7. Kinyoun's Carbol Fuchsin
Basic fuchsin dye
4g
Phenol crystals
8g
Ethyl alcohol (95%)
20ml
Distilled water
100 ml
This fonnula is preferred in some quarters to the Ziehl carbol fuchsin. It is attributed to
Kinyoun, but the reference to its original publication has not been located.
8. Carbol Crystal Violet
Solution A:
Gentian violet solution
10 ml
Crystal violet dye
0.4 g
Phenol (1% aqu sol.)
100mI
Ethyl alcohol (95%)
10 mI
Solution B:
Phenol
Ig
Distilled water
100 ml
Mix solutions A and B
Staining protocol
Prepare bacterial smears on microscopic slides and heat-fix. Stain for 5-60
water, dry and examine under microscope.
Results
The results depend on which of the above staining fluids is selected.
NEGATIVE STAINING OF BACTERIA
1. Dorner's Nigrosin Solution
Nigrosin, water soluble (nigrosin B)
109
Distilled water
100 mI
4
S,
wash with
Immerse in boiling water bath for 30 min, and then add as preservative fonnalin 0.5 ml.
Filter twice through double filter paper and store in serological test tubes, about 5 ml to
the tube. This staining solution is used for the negative demonstration of bacteria.
Staining protocol
1. Mix a loopful of the bacterial suspension on the slide with an equal amount of the
staining solution (If prepared from growth on solid media, the suspension must not be
too heavy.)
2. Allow the mixture to dry in the air, and examine under microscope.
Results
Unstained ceUs in a background which is an even dark gray if the preparation is well
made.
2. Benian. Congo Red
Congo red dye
2g
Distilled water
100ml
Staining protocol:
1. Place a drop of the above staining fluid, on a slide ..
2., Mix culture with ,the drop and spread out into a rather thick
, film,
3. After film has dried, wash with I % He!.
4. Dry and examine under oil immersion objective (lOOx) of compound microscope.
Results
Bacterial cells are unstain.e d in a blue background. Good results are not to be expected
from broth cultures or from cultures in salt solutions unless the cells are first removed by
centrifuging.
5
Exercise 2 - Gram staining for differentiation of bacteria based on cell wall
Introduction
Bacteria are routinely stained with different dyes in order to reveal different properties
and to enhance contrast for viewing with conventional bright field microscopy. The Gram
stain is routinely used as an initial procedure in the identification of an unknown bacterial
species. Bacteria bear a slight net negative charge and usually bind positively charged
dyes such as methylene blue and crystal violet. A species can he classified as Gram
positive, Gram negative, or Gram variable depending on the ability if cells to retain the
blue dye. Gram negative bacteria do not retain the dark blue color, but can be
counterstained a light red so that they can he seen in bright field microscopy. Gram
staining is called a differential staining. In Gram staining bacteria fixed to a slide are
treated with a basic dye that binds electrostatically to the negatively charged cells. Next,
the preparation is treated with a mordant such as iodine to form an insoluble dye-iodine
complex. The slide is then washed with alcohol to solubilize and remove the dye-mordant
from Gram negative cells but not Gram positive ones. Differential extraction of the dyemordant by the decolorizing agent is the critical step that distinguishes the bacteria. A
counterstain, safranin, is applied in the final step. Cells that have been decolorized will
take up the second basic dye whereas those already stained with the first dye will not. The
thinner cell wall of Gram negative bacteria would be readily penetrated by the
decolorizer. Since many Gram positive bacteria tend to become Gram negative with age,
the Gram stain should be used with overnight cultures. Sample from the edge of a colony,
where cells are actively growing. Gram (+) cells have thicker cell walls - more
peptidoglycan and teichoic acid. Gram (-) cells have Lipopolysaccharide (Endotoxin) in
the outer membrane of their cell wans. Gram (+) cells are generally more sensitive to
those antibiotics which interfere with cell wall biochemistry like penicillin, vancomycin
and cephalosporin. Gram (+) cells are more sensitive to lysozyme - a peptidoglycan
digesting enzyme.
Materials
• Bacterial cultures: one Gram's positive and one Gram 's negative bacteria
• Gram' staining kit:
a) Gram's crystal violet: 1% aqueous crystal violet dye; Hucker's crystal violet - 2 g
crystal violet 90% dye content, 20 m1 ethyl alcohol, 0.8 g ammonium oxalate, 80
ml distilled water
b) Gram's iodine: 1 g iodine, 2 g potassium iodide, 300 ml distilled water
c) Gram's safranin: 4 g safran in powder, 200 ml ahydrous ethanol, 800 ml distilled
water
d) Gram's decolorizer: 25% acetone, 75% isopropyl alcohol
• Microscopic slides
• Inoculating loop
• Immersion oil
6
• Microscope with l00x objective lens
• Gas burner
• Tissue papers
Protocol
1. Prepare two smears from broth culture of Lactobacillus spp. and Escherichia coli on
clean and grease free microscopic slides and heat fix over the gas flame.
2. Cover the smear completely with a few drops of a solution of crystal violet· (purple
basic dye).
3. After 30-60 sec, rinse the smear with water by squirting the slide above the smear and
letting the water wash over it until the water runs clear.
4. Cover the smear with iodine solution (the mordant) and after 60 sec, rinse the smear
with water to remove the excess iodine.
5. Wash the smear with a few drops of Gram's decoloriser (isopropanol-acetone mixture
or similar solvent) until the wash solution runs colorless and rinse with water to
remove excess decoloriser.
6. Apply the aqueous safranin for 30-60 sec and rinse the excess dye with water.
7.
Dry the smear with blotting paper to remove excess water.
8. Blot dry followed by drying over the gas flame of the stained smear on the slide and
examine under the microscope (IOOx, oil immersion objective).
9. Draw and identify the bacterial cells (Gram positive or negative).
Observations
7
Results
Inferences
8
OTHER METHODS
There are numerous modifications of the Gram's stain. The Hucker's modification is
valuable for staining smears of pure cultures; Kopeloff and Beerman for preparations of
body discharges such as gonorrhoeal pus, also for pure cultures of strongly acid-fonning
organisms. The latter is itself a variation of the modification by Burke.
1. Hucker's Modification
Primary slain: Ammonium Oxalate Crystal Violet (Hucker's)
Solution A:
Crystal violet dye
0.2 g
Ethyl alcohol (95 %)
20ml
Solution B:
Arrunonium oxalate
O.S g
Distilled water
SOml
Mix solutions A and B
Gram's iodine: Gram's Modification of Lugol's Solution
Iodine
Ig
KI
2g
Distilled water
300ml
Counter staiD:
Safranin 0 (2.5 % solution in 95 % ethyl alcohol)
IOmI
Distilled water
lOOml
Staining protocol
1. Stain smears 1 min with ammonium oxalate crystal violet.
2. Wash in tap water for not more than 2 sec.
3. Immerse 1 min in iodine solution.
4. Wash in tap water, and blot dry.
5. Decolorize 30 sec with gentle agitation, in 95 per cent ethyl alcohol. Blot dry.
6. Counterstain 10 sec in the above safranin solution.
7. Wash in tap water.
8. Dry, and examine.
Results
Gram-positive organisms, blue; gram-negative organisms, red.
2. Burke and
Kopeloff~Beerman
Modifications
Alkaline Gentian Violet
Solution A:
Gentian or crystal violet
I g
9
Distilled water
100 ml
Solution B:
NaHCO,
Ig
20 ml
Distilled water
Burke's IodiDe Solution
Iodine
Ig
KI
2g
Distilled water
100mi
Kopeloff and Beerman's Iodine Solution
Iodine
2g
Normal NaOH (40.0 I g per liter)
lOml
After the iodine is dissolved, make up to 100 mt with distilled water.
Burke' s Counterstain
Safranin 0 dye
2g
Distilled water
100ml
Kopeloff and Beerman's Counter stain
Basic fuchsin dye
0.1 g
Distilled water
100 ml
Staining Protocol
I. Dry thinly spread films in the air without heat.
2. Flood with solution A; mix on the slide with 2·3 drops (or more, depending on size of
flooded area) of solution B, and allow to stand 2-3 min.
3. Kopelotf and Beennan mix the two solutions in advance, 1.5 ml of solution A to 0.4
ml of solution B, and allow to stay on slide 5 min or more.
4. Rinse with either of the above iodine solutions. (The committee indicates no
preference between the two; some workers prefer one, some the other.)
5. Cover with fresh iodine solution, and let stand 2 min or longer.
6. Rinse with tap water; then blot water from surface of smear, without drying.
(Kopeloff and Beennan omit the washing.) The amount of drying is important in this
step. One must get rid of all free water but not allow the cells to dry.
7. Follow the blotting very quickly with decolorization in ether and acetone (1 vol of
ether to 1-3 vol of acetone), adding to the slide drop by drop until practically no color
comes off in the drippings (usually less than 10 sec). In this step the speed of
decolorization can be varied by varying the ratio of ether to acetone; the more
acetone, the more rapid the process. It is sometimes desirable to slow down the
process by using a ratio of I: I .
8. Dry in the air.
10
9. Counter-stain 5-10 s in one of the above given counter-stains. Burke's (i.e., safranin)
is preferred.
10. Wash in tap water.
11 . Dry, and examine under oil immersion objective (lOOx) of the microscope.
Results
Gram-positive organisms are blue; gram-negative organisms are red. This technique is
advantage of not giving false positives due to vacuolar bodies that resist decolorization by
other gram-staining procedures.
Interpretation of the Gram Stain
Proper cares is essential for interpretation of gram stain. Organisms are generally
described as gram-positive or gram-negative. Many organisms are gram-variable. Hence,
interpretation of gram reaction of an unknown organism on the basis of a single test is not
possible. Hence, repeated procedures with different age cultures should use more than onc
staining technique in order to detenninc the constancy of the organism toward the stain.
Gram-positive organisms can be made gram-negative by treatment with ribonuclease and
that the gram-positive reaction can be restored subsequently by treatment with
magnesium ribonucleate. Some organisms have granules which resist decolorization and
which may cause misinterpretation. Therefore, gram stain does not always give a clearcut reaction and the results must be interpreted carefully.
II
Exercise 3 - Examination of shape and arrangement of bacteria
Introduction
The microorganisms are of varying in shapes and arrangements. Different shaped
microorganisms are occurred, such as spherical or coccus shaped, cylindrical or rod or
bacillus shaped and spiral or belical shaped. Following binary fission, the newly formed
bacteria spontaneously assume (or, end up with) a characteristic shape. This shape varies
significantly among species of bacteria and may be employed in the course of isolate
identification. Two major shape classes are coccus and bacillus, though there are many
variations on these themes, additional distantly related shapes, as well as many variations
on the degree to which cells separate or interact following their division.
Most bacteria come in one of three basic shapes: coccus, rod or bacillus, and spiral.
a) Cocci, singular - coccus, from the Latin coccinus (scarlet) and derived from the Greek
kokkos (berry) are any microorganism (usually bacteria), whose overall shape is spherical
or nearly spherical. The cocci are spherical or oval bacteria having one of several distinct
arrangements based on their planes of division. An average coccus is about 0.5-1.0
micrometer in diameter. Spherical shaped bacteria can be classified based on their
arrangements as follows:
I) Monococcus: Single coccus/ spherical cells, e.g. Micrococcus spp.
2) Diplococcus: cocci occur as pair. A diplococcus (plural diplococci) is a round
bacterium (a coccus) that typically occurs in pairs of two joined cells. The cell divides
in one plane. Cocci that remain in pairs after they divide, e.g. Lactococcus lactis ssp.
cremoris, Streptococcus pneumoniae, Moraxella catarrhalis, Neisseria gonorrhoeae
and Neisseria menjngitidis.
3) Tetards/ tetra cocci: coccus occurs as combination of four cells along the diagonal of a
square. Division in two planes produces a tetrad arrangement. Cocci that fail to
separate after the division, but instead remain in groups of four forming squares, e.g.
Pediococcus spp., Micrococcus luteus.
4) Sarcina: Eight cocci occur in cubical shape. Division in three planes produces a sarcina
arrangement. Cocci that fail to separate after the division, but instead remain in groups
of eight fonning cubes, e.g. Sarcina spp.
5) Streptococci: cocci occur in chains. Cocci that fail to separate after the division, but
instead remain in chains of cells, e.g. Streptococcus thermuphilus, Enterococcus
faecalis, Streptococcus pyognes.
6) Staphylococci: cocci occur in irregular, often grape bunch like arrangement due to the
division in random planes. Cocci that fail to separate after the division, but instead
remain in amorphous sheets or clumps, e.g. Staphylococcus aureus.
b) Bacilli, rod-shaped bacteria, divide in one plane producing a bacillus, streptobacillus
or coccobacillus arrangement. Variations on rod-shaped bacteria - rod, tapered rod, staff,
cigar, oval and curved. Basically, bacilli are longer than they are wide and lack extreme
curvature. Bacilli typically divide only across their short axis. An average bacillus is 0.5·
12
1.0 micrometer wide by 1.0-4.0 micrometer long. Cy lindricallrodslbaci lli shaped bacteria
are classified based on arrangements as follows:
..
•
.......
diplococcus
<OC<Us
a
streptococcus
tetrad
sarcina
Figure la: Arrangements of cocci shaped bacteria
Source: http://student.ccbemd.eduicourseslbioI41Ilecguidelunitl /shapeJu] coCcus.hlm]
.
•
t
..,.,. ., '" -
*•
. ""
-.
A
.'
•
•
1-
•
Figure I b: Staphylococcus aureus
-
.I>
•
,
;~
-• \.. ... • ...••-, •
•
•
.,
•,. ,
•
10 ""
Source: http://student.ccbemd.eduicourseSlbioI4 IlIecguidelun itl /shape/staphnegimage.html
binary fiuion
•
bacjlus
streptobaCMlus
coccobaciHus
Figure 2: Arrangement of bacil li shaped bacteria
Source: hnp:l/sludenl.ccbcmd.edulcourses/bio 141 ltecguidelunit I/shapelu I rod.hlml
13
1) Bacillus: single bacilli, e.g. Escherichia coli.
2) Diplobacilli: Ba~iIli that fail to separate after the division and remain in paired celis,
e.g. Bacillus megaterum.
3) Streptobacillus: Bacilli arranged in chains, e.g. Bacillus cereus
4) Coccobacillus: oval and similar to coccus. A short bacillus, nearly looks like a coccus.
A coccobacillus (plural coccobacilli) is a type of rod-shaped bacteria. The word
coccobacillus reflects an intermediate shape between coccus spherical bacillus
(elongated). Coccobacilli rods are so short and wide that they resemble cocci, e.g.
Bordetella pertussis, Haemophilus injluenzae, Coxiella burnetti, AcinelObacte,. strains
may grow on solid media as coccobacilli.
c) Spiral shaped: Spirals come in one of three forms, a vibrio, a spirillus, or a spirochete.
Spirals range in size from I micrometer to over 100 micrometer lengths.
I) Vibrio: a curved or comma shaped. e.g. Vibrio cholera e
2) Spirillum: a thick, rigid spiral.
3) Spirochete: a thin, flexible spiral. e.g. Leptospira spp., Treponema pallidllm
Simple staining (single stain, positive or negative) stain is used to study the
morphological shape and arrangement of microorganisms. Simply, microscopic slide of
given bacterial culture is prepared by simple staining protocol and observe under lOOx
(oil immersion) objective to see the morphological shape and arrangement of bacteria.
Materials
• Bacterial cultures: one rod shaped and one cocci shaped culture
• Simple stain solution: Methylene bluel crystal violet stain solution (positive staining)
or nigrosin stain solution (negative staining)
• Microscopic slides
• Inoculating loop
• Immersion oil
• Microscope with lOOx objective lens
• Gas burner
• Tissue papers
Protocol
c) Positive staIning
1. Prepare two smears from broth culture of Lactobacillus spp. and Lactococcus spp. on
clean and grease free microscopic slides and heat fix over the gas flame.
2. Flood the smear with the methylene blue or crystal violet stain solution for \-2 min.
3. Gently wash the slide with water to remove the excess dye.
4. Blot dry followed by drying over the gas flame of the stained smear on the slide and
examine under the microscope (IOOx, oil immersion objective).
5. Draw and identify the shape and arrangement of the bacterial cells in the circles
below.
14
d) Negative staining
1. Take two clean and grease free microscopic slides.
2. Transfer two loopful of broth culture and one loopful of nigrosin solution on the slide
and mix thoroughly with the help of inoculating loop.
3. Spread unifonnly the mixture of bacterial culture and nigrosin dye to get a thin and
transparent smear on the slide.
4. Allow to air dry of the smear and examine under the microscope «(IOOx, oil
immersion objectives
5. Draw and identify the shape and arrangement of the bacterial cells in the circles
below.
Observations
Resulls
Inferences
15
Exercise 4 - Measurement of size of microorganisms by micrometry
Introduction
Bacteria range in size from approximately as small as the largest viruses to large enough
for single cells to hf' visihl e hy the naked eye. That is. from about 0.1 to about 600 11m
over a single dimension. Bacteria vary in size as much as in shape. The small est (e.g.,
some members of the genus Mycoplasma) are about 100 to 200 nm in diameter,
approximately the size of the largest viruses (poxviruses). Escherichia coli, a bacillus of
about average size, is 1.1 to 1.5 11m wide by 2.0 to 6.0 11m long. A few become fairl y
large; some spirochetes occasionally reach 500 11m in length, and the Cyanobacterium
oscil/atoria is about 7 11m in diameter (the same diameter as a red blood cell). Recently a
huge bacterium has been di scovered in the intestine of the brown surgeonfish, Acall/hunts
nigrofusclls. Epulopisdumjishelsoni grows as large as 600 11m by SO 11m, a little smaller
than a printed hyphen. It is now clear that a few bacteria are much larger than the average
eucaryotic cell.
Because of their small size, bacteria have a large surface-la-volume ratio. For examp le.
spherical bacteria with a diameter of2 11m have a surface area of about 12 Ilm2 (4.1t.d2/4)
and a volume is about 4 Ilm3 (4/3.1t.d 3/S). Their surface-to-volume ratio is 12:4, or 3: I. In
contrast, eukaryotic cells with a diamter of 20 Ilm have a surface area of about 1200 11m2
and a volume of about 4000 Ilm3. Their surface-to-volume ratio is 1200:4000, or 0.3: I,
only one-tenth as great. The large surface-to-volume ratio of bacteria means that no
internal part of the cell is very far from the surface and that nutrients can easily and
quickly reach all parts of the cell. The large surface to volume ratio seen in bacteria is one
reason that procaryotes are so successful despite their relatively simple morphologies.
An average bacillus is 0.5-1 .0 micrometer wide by 1.0-4.0 micrometer long. An average
coccus is about 0.5-1.0 micrometer in diameter. The size of bacillus shaped bacterium is
expressed as L x B (Length x breadth) and for coccus shaped bacterium is expressed as
micrometer of diameter (0).
Rod shaped
Coccus shaped
Figure 3: Size expression of cocci and rods
Materials
•
Bacterial cultures (Lactobacillus spp., LaCIOCOCCliS spp.)
16
•
Simple stain solution: Methylene bluel crystal violet stain solution (positive staining)
•
Microscopic slides
•
Stage micrometer
•
Ocular micrometer
•
Inoculating loop
•
Immersion oil
•
Microscope with 100x objective lens
•
Gas burner
• Tis~ue papers
Protocol
a) Calibration of ocular micrometer with stage micrometer
I. Stage Micrometer: Stage micrometer looks like a microscopic slide having a
micrometer scale on it. Each division of stage mi crometer scale is 10 micrometer
or 0.01 mm.
2. Ocular micrometer: This looks like a circular glass disc with a graduated scale
of 100 divisions. The distance between each two divisions is constant and is
calibrated using stage micrometer.
0.01 MM
a) Stage micrometer
Figure 4: Stage micrometer and ocular micrometer
17
b) Ocular micrometer
When the stage micrometer is placed under oil immers ion objective (lOOx) of a
compound microscope, the microscopic fiel d is looked as fo llows:
Eyepiece ( IOx)
Body tube
Objeclive lens (I OOx)
Slagc micromeler
Condenser lens
Stage divisions
Microscopic field .
Figure 5: Microscopic field after insertion of stage micrometer under IOOx objective
18
The ocular micrometer is set in ocular lens system and after insertion the microscopic
field is looked as follows:
Eyepiece (lOx)
Ocular micrometer
f-'I--. Body tube
\..._T
Objective lens (IOOx)
L_____S----\~ Condenser lens
to
20
30
.
so
.
1\1
"
90
I IIIII
/
Oi;ular divisions
---/--+
Microscopic field
Figure 6: Microscopic field after insertion of ocular micrometer in eyepiece (lOx).
19
3. Calibration protocol
The each division length of ocular micrometer is calibrated using the stage micrometer.
r:/.~'1-----. Eycp!e'Ce (lO:-:}
¥:::::~----. Ocular micrometer
Body tube
/!:::=::[J::=~:::::::::::':;: Objective lens (IOOx)
L.:::::~_~'::::=:::~_~/ ,~'r--+
Siage micrometer
\ - - - - - - \ - -.... Condenser lens
Points of coincide of ocular
and
divisions
70
W
---...".4
Microscopic field
Figure 7: Microscopic field after insertion of stage micrometer under objecvie (IOOx) and
ocular micrometer in eyepiece ( lOx)
20
The stage micrometer and ocular micrometer are inserted into their respective positions in
a compound bright field microscope. The microscopic field is looked as follows (Figure
7), where both stage and ocular micrometer scales are observed in same microscopic
filed. The ocular micrometer's division scale can be adjusted by rotating the eyepiece to
align the ocular and stage micrometer division scales in parallel. Now, by adjusting the
stage micrometer, the scale of division of both micrometers can be coincided. The
number of ocular divisions is coinciding with the number of stage micrometer divisions
are counted and length of each ocular division is calculated as follows:
Let, y (56) ocular micrometer divisions cover the x (5) stage micrometer divisions.
Therefore,
y (56) ocular divisions = x (5) stage division = I Ox5 = 50 micrometers
One ocular division = 10xly = 50/56 = 0.9 micrometer.
Thus, each ocular division is calibrated as 10xly = 0.9 micrometer with respect to the
particular microscope (fixed ocular and fixed objective). The calibrated value varies from
microscope to microscope, objective to objective and ocular 10 ocular and their
combinations also.
When, the ocular micrometer is inserted in the microscope each division observed in the
microscopic filed under loox objective is 10xly = 0.9 micrometer in length.
Now the system is ready for measurement of microorganisms ' sizes. Simple stained
microscopic slides of the culture is inserted under oil immersion objective and adjusted to
visualize the microbial cells. After this, the ocular micrometer is adjusted by rotating the
ocular lens system and alignment of cell is adjusting by moving the slide on stage. The
number of ocular division(s) covers the cell (along with diameter for coccus or length and
width for bacillus) is counted. The size of microbial cell is calculated by multiplying the
number of ocular division(s) with calibration factor (lOxly = 0.9 micrometer i.e. the
length of each division) and expressed respectively for coccus and bacillus shaped cell.
b) Preparation of stained bacterial slide
1. Prepare two smears from broth culture of Lactobacillus spp. and Laclococcus spp. on
clean and grease free microscopic slides and heat fix over the gas flame.
2. Flood the smear with the methylene blue or crystal violet stain solution and keep for
1-2 min.
3. Gently wash the slide with water to remove the excess dye.
4. Blot dry followed by drying over the gas flame of the stained smear on the slide and
examine under the microscope (100x, oil immersion objective).
c) Measurement of bacterial size
I. The stained bacterial slide is visualized under 100x objevtive and the ocular
micrometer is inserted in the eyepiece (lOx), which has already been calibrated using
stage micrometer (locular division = 0.9 micrometer).
2. After visualization, the bacterial cell is align to the ocular micrometer division to find
how many ocular division(s) are covered by the cell alongwith diameter (for cocci)
21
and along with length and breadth (for bacilli). From this, the dimeter for cocci or
length and breadth for bacilli are calculated as follows:
3. From Figure 8a, it is observed that one rod covers 1 and 2 ocular divisions along with
breadth and length, respectively. Therefore, L= 2·0.9=1.8 micrometer and B = 1*0.9
= 0.9 micrometer and expressed as 1.8 x 0.9 micrometer.
4. From Figure 8b, it is observed that 3 cocci cover 9 ocular divisions along with
diameter. Therefore, diameter of one coccus, D = 9·0.9/3=2.7 micrometer and
expressed as 2.7 micrometer diameter.
Observations
Results
Inferences
22
f't-:l?t----.
Eyep;ece (lOx)
Ocular micrometer
--+--\;--. Body lube
L-~
Objective lens (I OOx)
Stained slide
L ___--=:::,---"""'I~ Condenser lens
,
, , /.
,20
,,I~
10
I
'
-
--
I
j~
1111 1 II
-, ,
Ocular divisions
I
,
--
---r.... Microscopic field
I
Figure Sa: Microscopic field after insertion of stained bacterial slide under objecvie
(IOOx) and ocular micrometer in eyepiece (lOx)
23
r=t-In----+
Eyepiece (lOx)
F:~---+
-
+-1,----
Ocular micrometer
Body tube
\....---
Objective lens (IOOl()
L __-.::.:s-----', Condenser lens
...
.....
• ••
•
•.....
... ...
- .....•
...
~
Ocular divisions
•
...
...
---++ Microscopic field
Figure 8b: Microscopic fie ld after insertion of stained bacterial slide under objecvic
( IOOx) and ocu lar micrometer in eyepiece (lOx)
24
,
Exercise 5 - Examination of bacterial spores
Introduction
An endospore is a donnant, tough, and non-reproductive structure produced by Grampositive bacteria from the Firmicute phylum, which forms when a bacterium produces a
thick internal wall that encloses its DNA and part of its cytoplasm. Endospores (or simply
spores) fonn within (hence, eoda-) special vegetative cells called sporangia in response to
adverse changes in the environment (Figure 9). The original cell replicates its genet ic
material, and one copy of this becomes surrounded by a tough coating. The outcr cell then
disintegrates, releas ing the spore which is now well protected against a variety of
extremes of heat and cold, radiation, and an absence of nutrients, water, air etc. The
primary function of most endosporcs is to ensure the survival of a bacterium through
periods of environmental stress. These are therefore resistant to ultraviol et and gamma
radiation, desiccation, lysozyme, temperature, starvation, and chemical disinfectants.
Endospores are commonly found in so il and water. where they may survive for long
periods of time. Some bacteria produce exospores or cysts instead.
Figure 9: Schematic diagram of cross section of an endospore
Source: www.bmb.leeds.ac.uklmbiologyluglugteachlnewde ...
Bacteria produce a single endospore internally. The spore is often surrounded by a thin
covering known as the exosporium, which overlies the spore coat. The spore coat is
impenneable to many toxic molecules and may also contain enzymes that are involved in
germination. The cortex lies beneath the spore coat and consists of peptidoglycan. The
core wall lies beneath the eortex and surrounds the protoplast or core of the endospore.
The core has nonnal cell structures, such as DNA and ribosomes, but is metabolically
inactive. Up to 15% of the dry weight of the endospore consists of ca lcium dipicolinate
within the core, which is thought to stabilize the DNA. Dipicolinic acid could be
responsible for the heat resistance of the spore, and calcium may aid in resistance to heat
and oxidizing agents. The position of the endospore differs among bacterial species and is
useful in identification. The main types within the cell are tenninal, sub-teoninal and
centrally placed endospores. Terminal endospores are seen at the poles of cells, whereas
central endospores are more or less in the middle. Sub-terminal endospores are those
between these two extremes, usually seen far enough towards the poles but close enough
25
to the center so as not to be considered either tenninal or central. Lateral endospores are
seen occasionally. Examples of bacteria having terminal endospores include Clostridium
tetani, the pathogen which causes the disease tetanus. Bacteria having a centrally placed
endospore include Bacillus cereus, and those having a sub-terminal endospore include
Bacillus subtiUs. Sometimes the endospore can be so large the cell can be distended
around the endospore, this is typical of Clostridium tetani. Visualizing of endospores
under the light microscope can be difficult due to the impermeability of the endospore
wall to dyes and stains. While the rest of a bacterial cell may stain, the endospore is left
colourless. Because the spore is protected by a thick, tough coveri ng, it is difficult to
stain. To force the stain into the spore body, the thermal energy in stream is used and then
coumer-stains the vegetative cell components that are attached to the spore with a dye of
a contrasting colour. Therefore, a spore stain contains spores alone, spores inside
vegetative cells and vegetative cells without spores formed in them. To combat this, a
special stain technique called a Moeller stain is used. That allows the endospore to show
up as red, while the rest of the cell stains blue. Another staining technique for endospores
is the Schaeffer-Fulton staining, which stains endospores green and bacterial bodies red .
Due to high hydrophobic nature of spore coat and cortex, hydrophillic chemicals
including water cannot penetrate across it. The dyes (weak ly positive), generally used to
stain bacterial spores, are hydrophillic in nature and hence, cannot penetrate and stain the
spores. Thus, the protocol use to stain the vegetative cells is not suitable to stain the
spores as the dye is washed away from the spores during the removal of excess dye from
the smear on the slide with water but attached to the vegetative cells. The fact that the
moist heat can loosen or weak and penetrate the highly hydrophobic spore coat and thus,
helps in spore germination followed by outgrowth to fonn the vegetative cells, which is
called the heat induction for gennination and outgrowth of donnant spores. The idea led
to the development of moist heat induced dye penetration and staining of bacterial spores,
in which the spore smear on the glass slide is covered with dye solution and heated using
steam. The heat treatment helps in loosening the impervious spore coat and penetration of
dye to stain the spore. After cooling the stained smear, nonnal washing to remove the free
excess dye with water dose not remove the stain penetrated in spore and thus spores are
appeared as per the dye used. However, the vegetative portion looses the stain due to the
weakly positive nature of the dye. The vegetative cells or vegetative portion of the
sporulated cells are differentiated by using suitable counter stain.
Materials
•
Bacterial cultures: spore fanner - Bacillus spp. and non-spore former - Lactobacillus
spp.
•
Stain solutions: Malachite green solution (0.5%), Safranine solution (0.5%)
•
Microscopic slides
•
Blotting paper
•
•
Beaker
Inoculating loop
26
•
•
Immersion oil
Microscope with 100x objective lens (oil immersion)
•
Gas burner
• Tissue paper
Protocol
I. Prepare two smears from the colony of Bacillus spp. and Lactobacillus spp.
2. Put a piece of blotting paper of the size of the smears, but small enough so it doesn't
hang over the edges of the slide.
3. Place the slide over a beaker with water and heat the beaker to boil the water using
gas flame while holding on a tripod.
4. Flood the paper on smear with the malachite green stain solution and boil for 5 min
and keep the paper wet with the stain.
5. After 5 min, remove the slide and lift off the paper with your loop and discard.
6. Gently wash the slide with water.
7. Flood the smear with the counter-stain safranin to stain the vegetati ve cells.
8. After one minute, wash off the safranin with water, blot the slide dry and examine
under the microscope ( lOOx oil immersion objective).
9. Draw and identiry the spores and the vegetative cells in the circles below. Descri be
the characteristics (location, form, size etc.) of the spores.
Observations
Results
Inferences
27
OTHER SPORE STAINING METHODS
A) Dorner's Method
I. Zlehls Carhol-fuchsln
Solution A: Basic fuchsin solution 10 ml:
Basic fuchsin dye
0.3 g
Ethyl alcohol (95 %)
IOml
Solution B: 5 % sol carbolic acid 100 ml
Phenol
5g
Distilled water
95 ml
Mix solutions A and B
2. Dorner's Nigrosin Solution
Nigrosin, water soluble (nigrosin B Griibler recommended by Domer; American
nigrosins certified by Commission on Standardization of Biological Stains ordinari ly
satisfactory) 10 g
Distilled water 100 mt
Immerse in boiling water bath for 30 min, and then add as preservative fonnalin 0.5 ml
Filter twice through double filter paper and store in serological test tubes, about 5 mt to
the tube.
Staining protocol
1. Make a heavy suspension of the organism in 2-3 drops of distilled water in a small
test tube.
2. Add equal quantity of freshly filtered Ziehl's carbol fuchsin .
3. Allow the mixture to stand in a boiling water bath 10 min or more.
4. On a cover slip or slide mix one loopfut of the stained preparation with one loopful of
Domer's nigrosin solution.
5. Smear as thinly as possible and do not dry too slowly.
If even backgrounds for exhibiting or photographing arc required, especially in the case
of slime·producing bacteria, the following procedure is recommended:
1. Make the suspension in 0.5 ml of nutrient broth or water.
2. Add I ml of 10% gelatin solution.
3. Add I ml of carbol fuchsin, and stain as in steps I and 2 above.
4. Wash out the colloids with warm tap water, with the help of centrifuge or
sedimentation.
5. Mix with nigrosin, and proceed as above.
Results
Spores are red; vegetative cells are unstained; background is gray.
28
8) Dorner's Method - Snyder's Modification
Staining protocol
Prepare a dried smear on a slide, and cover with a small piece of blotting paper.
Saturate blotting paper with freshly filtered Zieh's carbol fuchsin.
Allow to steam 5-10 min, keeping paper moist by adding more staining fluid.
Decolorize instantaneously with 95% ethyl alcohol (but omit this step if the organisms
do not hold color well).
5. Wash with tap water.
6. Apply a drop of saturated aqueous nigrosin (Domer's fluid), and spread evenly.
7. Allow slide to dry quickly with gentle heat, without prior washing .
Results
I.
2.
3.
4.
Same as with Domer' s method, but this modification proves appl icable to some bacteria
(e.g., Bacillus subtilis) that are difficult to stain by Domer's technique.
C) Conklin's Modification of Wirtz Method
Staining protocol
1. Make smears as usual and fix by heat.
2. Flood slide with 5% aqueous malachite green, and steam for 10 min, keeping slide
flooded by addition of fresh staining fluid .
3. Wash 30 sec in running water.
4. Counter stain I min with 5% aqueous mercurochrome.
5. Wash in running water.
6. Blot dry, and examine.
Results
Spores are green; rest of cell is red.
Trouble is sometimes experienced with.the green fading after the slides have stood a fe w
days. Apparently this is the result of an alkaline reaction and can be prevented by treating
the slides in acid before making the smears. The alkalinity may be due to an invisible film
of soap or washing powder.
D) Bartholomew and Mittwer's "Cold" Method
Heat is not necessary in making an acid-fast stain; it is proving that it may also be
eliminated from spore staining, in which a very similar principle is involved. The
following modification of the Wirtz method by Bartholomew and Mittwer is a good
illustration.
Staining protocol
1. Fix the smear by passing through a flame 20 times.
2. Stain to min with saturated aqueous malachite green (i.e., about 7.6%), without heat.
3. Rinse with tap water for about 10 sec.
4. Stain 15 sec in 0.25% aqueous safranin.
5. Rinse, blot, and dry.
Results - Spores are green; rest of cell is red.
29
Exercise 6 - Staining of acid fast bacteria
Introduction
Acid-fastness is a physical property of some bacteria referring to their resi stance to
decolorization by acids during staining procedures. Acid-fast organisms are difficult to
characterize using standard microbiological techniques (e.g. Gram stain - if you gram
stained an AFB the result would be an abnormal gram positive organism, which would
indicate further testing), though they can be stained using concentrated dyes, particularly
when the staining process is combined with heat. Once stained, these organisms resist the
dilute acid andlor ethanol-based de-colorization procedures common in many staining
protocols- hence the name acid.Jast. The high mycolic acid contcnt of certain bacterial
cell walls, like those of Mycobacteria, is responsible for the staining pattern of poor
absorption followed by high retention. The most common staining technique used to
identify acid-fast bacteria is the Ziehl-Neelsen stain, in which the acid fast bacilli are
stained bright red and stand out clearly against a blue background. Some bacteria may
also be partially acid-fast. This staining method is differential stain used to identify
members of the genera Mycobacterium and Nocardia. These bacteria, and a few others,
have cells walls that contain a waxy, lipoidal material called mycolic acid. These bacteria
retain this primary stain even after treatment with a decolorizing agent called acid alcohol. Bacteria, which are not decolorized by acid-alcohol are said to be acid-fast
Bacteria which are readily decolorized by acid-alcohol are call nonacid-fast.
Materials
•
Bacterial cultures: spore former - non-acid fast - Lactobacillus spp., acid fast Mycobacterium spp,
•
Stain solutions: Basic carbol fuschin solution, Methylene blue solution (0,5%)
•
Microscopic slides
•
Blotting paper
•
Beaker
•
Inoculating loop
•
Immersion oil
•
Microscope with 100x objective lens (oil immersion)
•
Gas burner
•
Tissue paper
Protocol
I. Prepare two smears from the colony of Lactobacillus spp. and Mycobacterium spp. on
a clean and grease free microscopic slide.
2. Place a small piece of bibulous paper over the smear and saturate the paper with
carbolfuchsin.
30
3. Heat the slide gently over the bunsen burner for 5 minutes. Be sure to keep the
bibulous paper
4. Saturated with carbolfuchsin during heating; if the slide is steaming. you're okay; if it
stops steaming, add more carbolfuchsin.
5. Rinse the slide gently with water and di spose of the used bibulous paper in the trash.
6. Deco lorize the slide with acid-alcohol until the rinsate runs clear.
7. Rinse the slide gently with water.
8. Counterstain with methylene blue for 2 minutes.
9. Rinse the slide gently with water.
10. Carefull y blot the slide dry with bibulous paper.
II . Observe the slide under the microscope, using proper microscope technique.
12, Acid-fast cells will stain fuchsia (red). Non-acid- fast cell s will stain blue.
Observations
Results
Inferences
OTHER METHODS
A) Ziehl-Neelsen Method
I . Ziehls Carbol-fuchsin
Solution A: Basic fuchsin solution 10 ml
Basic fuchsin dye
0.3 g
Ethyl alcohol (95 %)
10 ml
Solution B: 5 % sol carbolic acid 100 rnl
Phenol
5g
Distilled water
95 ml
Mix solutions A and B
2. Methylene Blue in Dilute Alcohol
Methylene blue dye
0.3 g
Ethyl alcohol (95 %)
30 ml
Distilled water
100 ml
0) Gross' Cold Method
An effort has been made to eliminate the necessity of applying heat during the fuchsin
staining so as to simplify the technique and to avoid "messy" preparations. Such
procedures seem to have justified themselves and can be recommended for pure culture
work.
Basic fuchsin solution:
Add 25 ml of a stock 4% alcoholic basic fuchsin solution to 75 ml of 6% aqueous phenol.
To this add 3-4 drops ofTergitol No.7 and stir thoroughly.
Methylene blue solution:
Add 30 ml of a stock 1.5% alcoholic methylene blue solution to 100 ml of 0.0 I % aqueous
KOH.
Staining protocol
1. Stain 5-10 min, without heating. in the above basic fuchsin solution.
2. Rinse in wann water.
3. Agitate for 30-60 sec in acid alcohol (3 ml of cone Hel in 97 ml ethyl alcohol).
4. Rinse with cold water.
5. Counterstain 3-5 min in the above methylene blue solution.
C) Fluorescence Method
Although this method is not of special importance in pure culture work, special mention
should be made of it because of the amount of attention now given to it in diagnostic
work. Its real advantage is that it can he used with relatively low magnification, and the
large fields that can he examined assure positive diagnoses in cases where the numbers of
tubercle organisms are few .
32
Solution A:
Auramine 0 (90 % dye content)
0.1 g
Ethyl alcohol (70 %)
lOOml
Solution B:
Liquefied phenol
3 ml
Con. HCI
0.5 ml
Distilled water
97 ml
NaCI
0.5 g
Staining protocol
1. Stain dried smears 2-3 min in solution A.
2. Wash in tap water.
3. De-stain 3-5 min in solution B. freshly prepared.
4. Dry, and examine under a monocular microscope, using 8 mm dry objective and a 20
X ocular; illumination should be a low-voltage, high-amperage microscope lamp,
supplied with a blue (ultraviolet-transmitting) filter. a complementary yellow filter
having been provided for the ocular.
Results
Acid-fast bacteria, bright yellow, fluorescent; other organisms, not visible; background,
nearly black.
33
Exercise 7 - Staining of bacterial cell wall
Introduction
Cell wall is a rigid structure which gives definite shape to the cell, situated between the
outer Ill.Ost slime or capsule and cytopl asmic m('mbnme. II has got thickness in the range
from 10-25 nm. It comprises of about 10-40% of dry weight of cell. It is involved in
growth and cell division of bacteria, apart from giving protection to the cell . Cell wall of
bacteria contains diaminopimelic acid (DAP), muramic acid and tcichoic acid. These
substances are joined together to give rise to a complex polymeric structure known as
peptidoglycan which provides rigidity to the cell. Peptidoglycans(mucopeptides,
glycopeptides, mUfeins) are the structural elements of almost all bacterial cell walls. They
constitute almost 95% of the cell wall in some Gram positive bacteria and as little as 510% of the cell wall in Gram negative bacteria. Un li ke the Gram positive cell wa ll , the
Gram negative cell wall contains a thin peptidoglycan layer adjacent to the cytoplasmic
membrane. This is responsible for the cell wall 's inabiliry to retain the crystal violet stain
upon decolouri sation with ethanol during Gram staining. In addition to the peptidoglycan
layer, the Gram negative cell wall also contains an outer membrane composed by
phospholipids and lipopolysaccharides, which face into the external environment. As the
Jipopolysaccharides are highly-charged, the Gram negative cell wall has an overall
negative charge.
The cell wall has low affinity for stain and therefore, it is not stained with most of the
staining procedures used to stain cytoplasm. Various techniques for staining the bacterial
cell wall have been developed, of which the tannic acid-crystal violet method is one of the
most widely used. This technique, which requires a very dilute solution of crystal violet,
results in the cell wall and the cross walls of many bacteria being stained rather faintly.
The tannic acid-crystal violet technique also has the disadvantage of being prepared as
temporary water mounts. Phosomolybdic acid-methyl green cell wall stain which
promises to be applicable to some bacteria not effectively stained by the tannic acidcrystal vio let method. This stain also must be prepared as a water mount and does not
give uniform results. A new principle in bacterial cell wall staining involves the use of a
basic dye, such as new fuchsin or crystal violet, and decolorization of the cell contents
through the use of congo red, an acid dye. The Chance cell wall stain appears to be
effective on most of the bacteria and on other forms considered difficult to stain, such as
Nocardia and Streptomyces. This technique, however, has the disadvantage of on ly
faintly staining cross septa. An effective technique for staining the bacterial cell wall has
been developed which utilizes procedures of both the tannic acid-crystal violet and
Chancels fuchsin-congo red methods. By using tannic acid as a mordant and congo red as
a selective decolorizing agent, a 0.5 per cent to 1.0 per cent soluti on of crystal violet can
be used without leaving the cytoplasmic area of the cell colored. This method stains the
outer wall and cross walls very sharply and has the added advantage of yielding
permanent dry mounts. This technique appears to have wide application and shou ld prove
especially useful since cross septa are shown clearly. The procedure has been
satisfactorily used to stain several genera of bacteria with uniformly good results. In cell
34
wall staining techniques mordant is applied, which increases the affinity of cell wall for
stain. It also increases the apparent thickness of cell wall due to deposition of fine
precipitates.
Materials
•
•
•
•
•
•
Cultures: Gram positive and Gram negative bacterial cultures
Microscopic glass slides
5% aqueous tannic acid solution
0.5% crystal violet solution
0.5% aqueous congo red solution
Blotting paper
• Gas flame
Protocol
I. Prepare a smear and fix it by heat.
2. Flood the smear with 5% aqueous solution oftanoic acid for 30 minutes.
3. Rinse the slide with water. Do not dry.
4. Stain the smear with 0.5% aqueous solution of crystal violet chloride for 2-3 minutes.
5. Wash off the stain with water and treat the smear with 0.5% aqueous solution of
congo red for 2-3 minutes.
6. Wash, dry and examine under oil-immersion objective.
7. Cell wall including newly developed cross walls and completed transverse septa will
be stained blue.
Observations
35
Results
Inferences
36
Exercise 8 - Staining of bacterial capsules
Introduction
Many bacteria secrete a slimy, viscous covering called a capsule or glycocalyx. Capsules
are often produced only under specific growth conditions. This is usually composed of
polysaccharide, polypeptide, or both. Most capsules are composed of polysaccharides, but
some are composed of polypeptides. The capsule differs from the slime layer that most
bacterial cells produce in that it is a thick, detectable, discrete layer outside the cell wall.
Some capsules have well-defined bnundaries, and some have fuzzy, trailing edges. The
medium in which the culture is grown as well as the temperature at which it is grown and
the age of the culture will affect capsule formation. Older cultures are more likely to
exhibit capsule production. When performing a capsule stain on your unknown, be sure
the culture you take your sample from is at least five days old.
Even thought not essential for life, capsules probably help bacteria to survive in nature.
Capsules help many pathogenic and normal flora bacteria to initially resist phagocytosis
by the host's. In soil and water, capsules help prevent bacteria from being engulfed by
protozoans. Capsules also help many bacteria to adhere to surfaces and thus resist
flushing. Capsules protect bacteria from the phagocytic action of leukocytes and allow
pathogens to invade the body. If a pathogen loses its ability to form capsules, it can
become avirulent. Capsules are formed by organisms such as Klebsiella pneumoniae.
A capsule is a layer of mucoid material that surrounds the bacterial cell, lying outside the
cell call. When this mucoid layer is thick. well organized, and difficult to remove, it is
called a capsule. If it is thin, difJiLse, and easy to wash oj[, it is called a slime layer
Capsules can easily be seen with a light microscope using a capsule stain. Most capsules
are composed of a network of polysaccharide fibers that have been secreted from the cell,
called the glycocalyx. The amount of capsule produced by a cell depends on the culture
conditions. Growth in high carbon, low nitrogen medium promotes capsule [ormation.
The general Function of a Capsule or slime layer is to: resist desiccation, they contain a
great deal of water, to help the bacterium adhere to surfaces, prevent or inhibit
phagocytosis by a host phagocyte (Streptococcus pneumonia). When it lacks a capsule, it
is destroyed easily and does not cause disease. Gliding bacteria often produce slime
which aids in their motility. The capsule stain employs an acidic stain and a basic stain to
detect capsule production. Bacterial capsules are non-ionic, so neither acidic nor basic
stains will adhere to their surfaces. Therefore, the best way to visualize them is to stain
the background using an acidic stain and to stain the celi itself using a basic stain. We use
India ink and Gram crystal violet. This leaves the capsule as a clear halo surrounding a
purple celi in a field of black.
Materials
•
Bacterial cultures: capsule former and non-capsule fonner
•
Stain solutions: Basic carbnl fuschin solution, Methylene blue solution (0.5%)
•
Microscopic slides
•
Blotting paper
•
Beaker
•
Inoculating loop
•
Immersion oil
•
Microscope with 100x objective lens (oil immersion)
•
Gas burner
•
Tissue paper
Protocol
I. Place a single drop of India ink on the left-hand end of a clean microscope slide.
2. Using a flamed loop and sterile technique, remove some K. pnellmoniae (or the
organism you want to stain) from your slant and mix it into the drop of India ink (Be
sure there are no large clumps of organism, but try to avoid spreadi ng the drop).
3. Place the end of another clean microscope slide at an angle to the end of the slide
containing the organism. Spread out the drop out into a film. This is done by
contacting the drop of India ink with the clean microscope slide and using the
capillary action of the dye! slide to spread the India ink across the smear.
4. Allow the film to air dry (Do not heat or blot dry. Heat will melt the capsule).
5. Saturate the slide with crystal violet for 1 minute.
6. Rinse the slide gently with water.
7. Allow the slide to air dry .
8. Observe the slide under the microscope (100x, oil immersion objective).
9. The background will be dark and the bacterial cells will be stained purple.
10. The capsule will appear clear against the dark background.
11. Note: Be sure your culture is several days old as young, fresh cultures won't have
developed capsules yet.
Observations
38
Results
Inferences
39
OTHER MEDHODS
Bacterial capsules are more easily confused with artifacts than any other structure
pertaining to the organisms. Inasmuch as capsules sometimes show merely as unstained
areas around the cells, there is a temptation to call any such surrounding area a capsule;
very often, however, they merely represent the tendency of a lightly stained surrounding
medium to retract from the cells on drying. For this reason the best way to demonstrate
capsules is actually to stain them by some procedure which differentiates them from the
cell itself. Several of the flagella stains accomplish this, notably those of Bailey and
Leifson, given above. Much simpler is the procedure of Anthony described below. The
Anthony method can be recommended because of both its simplicity and its
dependability. Any of the other methods which follow give satisfactory results. The
student is specially urged, however, not to pronounce any organism capsulated, as a result
of any of these staining procedures, until he has carcfully compared it with other
organisms generally recognized as having capsules.
A) Leifson Method
Leifson's stain:
KAI(SO,), 12H,O, or NH,A I (SO,), 12H,O (saturated aq solution)
20 ml
Tannic acid (20% aq solution)
10 ml
Distilled water
10mi
Ethyl alcohol (95%)
15ml
Basic fuchsin (saturated solution in 95% ethyl alcohol)
3ml
Mix ingredients in order named. Keep in tightly stoppered bottle, and the stain may be
good for a week.
Borax methylene blue solution:
Methylene blue (90% dye content)
0.1 g
Borax
I g
Distilled water
100 ml
Staining schedule:
J. Prepare smears of young cultures, on scrupulously cleaned slides.
2. Flood slides with the above solution, and allow to stand 10 min at room temperature in
warm weather or in an incubator in cold weather.
3. Wash with tap water.
4. Stain S-10 min, without heating, in borax methylene bluc.
S. Wash in tap water.
6. Dry, and examine.
Results
Capsules is red and cells are blue.
40
8) Anthony's Method with Tyler's Modification
Original formula
Crystal violet (85 % dye content)
I g
Distilled water
100 rnl
Tyler's modified formula
Crystal violet (85 % dye con-tent)
0. 1 g
Glacial acetic acid
0 .25 rnl
Distilled water
100mi
Staining protocol
I) Prepare smears, and dry them in the air.
2) Slain 2 min in the above aqueous crystal violet or, according to Tyler, 4-7 min in the
above acetic crystal violet.
3) Wash with 20% aqueous copper sulphate (CuS04.5H 20).
4) Blot dry, and examine.
Results
Capsules are blue violet and cells are dark blue.
41
Exercise 9 - Staining of bacterial flagella
Introduction
Perhaps the most recognizable extracellular bacterial cell structures are flagella. Flagella
are whip-like structures protruding from the bacterial cell wall and are responsible for
bacterial motility (i.e. movement). Bacterial flagella account for upte 2% of the dry
weight ofa cell. Anatomically. a bacterial flagellum consists of three parts a basal body. a
hook and main filament. Flagella are complex structures that are composed of many
different proteins. These include flagellin. which makes up the whip-like tube and a
protein complex that spans the cell wall and cell membrane to form a motor that causes
the flagellum to rotate. This rotation is normally driven by proton motive force and is
found in the body. Its shape is a 20 nanometer-thick hollow tube. It is helical and has a
sharp bend just outside the outer membrane; this "hook" allows the axis of the helix to
point directly away from the celL A shaft runs between the hook and the basal body,
passing through protein rings in the cell's membrane that act as bearings. Gram-positive
organisms have 2 of these basal body rings, one in the peptidoglycan layer and one in the
plasma membrane. Gram-negative organisms have 4 such rings: the L ring associates
with the lipopolysaccharides, the P ring associates with peptidoglycan layer, the M ring is
embedded in the plasma membrane, and the S ring is directly attached to the plasma
membrane. The filament ends with a capping protein. The arrangement of flagella about
the bacterial cell is unique to the species observed. Common forms include i) Peritrichous
- Multiple flagella found at several locations about the cell. ii) Polar - Single flagellum
found at one of the cell poles, iii) Lophotrichous, A tuft of flagella found at one cell pnle
etc.
The flagella stains are notoriously difficult to perform because of
(i) the tendency for stain to deposit on background material and indeed onto the surface of
slide and (ii) the ease witl;l which bacteria shad these delicate appendages, if culture is
not handled carefully. For these reasons special care in cleaning of slide and preparing
bacterial smear is essentiaL (a) Cleaning of slide: Use new, heat resistant slide and
immerse it in absolute alcohol for 10-15 minutes. Clean the slide with dichromic acid
cleaning solution. Wash the slide with distilled water. Before use, pass the slide back and
forth through a Bunsen flame until flame shows yellow colour. Cool the slide and use. (b)
Preparation of Smear: Good results can be obtained by preparing smear in following way.
Take agar slant having, actively growing and young culture (18-20 hours old). Add
carefully about 2-4 ml of sterile distilled water through the wall of tube keeping slant
upwards. Rotate the tube slowly between palms. Incubate the tube at 30°C for 30-40
minutes. Check motility of the suspension by hanging drop preparation. If culture is
motile then proceed further. Using sharp glass marker make four rectangular areas
approximately of equal size. Transfer a small drop from the top of bacterial suspension by
means of capillary pipette to the end of square on the reverse side of the marks. Tilt the
slide to about 70° and allow the drop to run slowly to the other end and remove
immediately the excess with the help of blotting paper. Allow the slide to air dry.
42
Because bacterial flagella are very thin and fragile a special stain (flagella stain) is
prepared that contains a mordant. This mordant allows piling of the stain on the flagella,
increasing the thickness until they become visible. Various arrangements of flagella are
seen on different cells. Bacterial flagella are fine, threadlike organelles of locomotion.
They are slender (about 10 to 30 run in diameter) and can only be seen directly using lhe
electron-microscope. In order to observe them with the light microscope, the thickness of
the flagella are increased by coating them with mordants like tannic acid and potassium
alum, and staining them with basic fuchsin (Gray method), pararosaniline (Leifson
method), silver nitrate (West method), or crystal violet (Difeo's method). Although
flagella staining procedures are difficult to carry out, they often provide information
about the presence and location of flagella, which is of great value in bacterial
identification. Difco's SpotTest Flagella stain employs an alcoholic solution of crystal
violet as the primary stain, and tannic acid and aluminum potassium sulfate as mordants.
As the alcohol evaporates during the staining procedure, the crystal violet forms a
precipitate around the flagella, thereby increasing their apparent size.
Materials
•
Young, IS-hour tryptic soy agar slants of Escherichia coli/ Bacillus cereus
(peritriehously flagellated) and Pseudomonas jluorecens (polarly flagellated)
0
Inoculating loop
0
Glass slides
0
Distilled water
0
Microscope
0
Immersion oil
•
Boiling water bath (250 ml beaker with distilled water, rind stand, wire gauze pad, ali.
Bunsen burner or hot plate)
Protocol
West method:
1. Aseptically transfer the bacterium with an inoculating loop from the turbid liquid at
the bottom of the slant to 3 small drops of distilled water in the center of a clean slide
that has been carefully wiped off with clean lens paper. Gently spread the diluted
bacterial suspension over a 3cm area using the inoculating needle.
2. Let the slide air dry for 15 minutes.
3. Cover the dry smear with solution A (the mordant) for 4 minutes.
4. Rinse thoroughly with distilled water.
5. Place a piece of paper towelling on the smear and soak it with solution 8 (the stain).
Heat the slide in a boiling water bath for 5 minutes in an exhaust hood with the fan
on. Add more stain to keep the slide from drying out.
6. Remove the toweling and rinse off excess solution 8 with di stilled water. Flood the
slide with distilled water and allow it to sit for 1 minute while more silver nitrate
residue floats to the surface.
43
7. Then, rinse gently with water once more and carefully shake excess water off the
slide.
8. Allow the slide to air dry at room temperature
9. Examine the slide with the oil immersion objective. The best specimens will probably
be seen at the edge of the smear where bacteria are less dense.
Procedu re (Difco)
1. Draw a border around the clear portion of a frosted microscope slide with a wax
pencil.
2. Place a drop of distilled water on the slide, approximately I em from the frosted edge.
3. Gently touch a colony of the culture being tested with an inoculating loop and then
lightly touch the drop of water without touching the slide. Do not mix.
4. Tilt the slide at a slight angle to allow the drop preparation to flow to the opposite end
of the slide.
5. Let the slide air.dry at room temperature. Do not heat· fix .
6. Flood the slid with the contents of the Difco SportTest flagella stain ampule.
7. Allow the stain to remain on the slide for approximate ly 4 minutes. (Note: the staining
time may need to be adjusted from 2 to 8 minutes depending on the age of the culture,
the age of the stain. the temperature, and the depth of staining solution over the
culture).
8. Carefully rinse the stain by adding water from a faucet or wash bottle to the slide
while it remains on the staining rack. Do not tip slide before this is done.
9. After rinsing, gently tilt the slide to allow excess water to run otT and let the slide air·
dry at room temperature or place on a slide warmer.
10. Examine the slide microscopically with the oil immersion objective. Begin
examination at thinner areas of the preparation and move toward the centre. Look for
fields which contain several isolated bacteria, rather than fields which contain clumps
of many bacteria. Bacteria and their flagella should stain purple.
Observations
Results
Inferences
44
OTHER METHODS
Staining of bacterial flagella is a difficult technique, and there have been numerous
methods proposed for the purpose. It has long been realized that flagella are actually
below the visual limit in size, but of recent years the electron microscope has given a
definite idea how small they really are - around 0,02-0.03 j..l in diameter. Electron
micrographs, in fact, often show many morc flagella than do stained preparations. Until
the electron microscope, however, has become a routine laboratory instrument, one must
have resort to the principle introduced by Loeffier of mordanting the preparations before
staining to increase the apparent size of the flagella. A second difficulty in staining
flagella is the ease with which bacteria shed these delicate appendages unless the cultures
are properly handled. To prevent this one ordinarily employs speciall y cleaned slides and
specially prepared smears on the slides.
Methods for preparing slides: Ordinary cleaning of glassware is not sufficient for the
purpose. Various methods have been proposed, but the following directions seem to give
as good results as any: Use new slides if possible, preferably of Pyrex glass or simi lar
heatresistant properties. (This is because under the drastic method of cleaning to remove
grease. old slides have a greater tendency to break.) Clean first in a dichromate cleaning
fluid, wash in water. and rinse in 95% alcohol; then wipe with a clean piece of
cheesecloth. (Wiping is not always necessary but is advisable unless fresh alcohol is used
after every few slides.) Pass each side back and forth through a flame for some time,
ordinarily until the appearance of an orange color in the flame; some experience is
necessary before the proper amount of heating can be accurately judged.
Unless heat-resistant slides are used, cool sheds gradually in order to minimize breakage.
An ordinarily satisfactory method of doing this is to place the flamed slides on a metal
plate (flamed side up) standing on a vessel of boiling water and then to remove the flame
under the water so as to allow gradual cooling. (Too rapid cooling may result in breakage,
sometimes as long as 2 weeks after the heating.)
Culture preparation: Young and actively growing cultures (18-22 h) on broth agar
slants is used and confinn the motility by banging drop technique. If motility is observed.
wash off the growth by gentle agitation with 2-3 ml of sterile distilled water. Transfer to a
sterile test tube, and incubate at optimum temperature for 10 min and again check motility
under microscope. Tilt the slide. and allow the drop to run slowly to the other end and air
dye the side in tilted position.
Staining Methods
Good results can be obtained with any of the following methods, especially after
familiarity has been obtained with it. Special recommendation must be given to the last of
the four procedures (modified Bailey method). Although seeming a little more
4S
complicated on first reading, it has been found to give the most unifonnly satisfactory
results in inexperienced bands.
A) Casares-GII Flagella Stain
Mordant
Tannic acid
AICI,-6H,O
109
18 g
ZnCl,
109
Basic fuchsin
I.5g
Alcohol (60%)
40 ml
The solids are dissolved in the alcohol by triturating in a mortar, adding 10 ml of the
alcohol first, and the rest slowly. This alcoholic solution may be kept several years. For
use, mix with an equal quantity of water or dilute with 4 parts of water, filter off
precipitate, and collect filtrate on the slide.
Ziehls Carbol-fuchsin
Solution A: Sat ale sol basic fuchsin iO ml :
Basic fuchsin (90 % dye content)
0.3 g
Ethyl alcohol (95 %)
10 ml
Solution B: 5 % sol carbolic acid 100 ml
Phenol
5g
Distilled water
95 ml
Mix solutions A and B
Staining protocol
1. Prepare smears of young cultures, on scrupulously cleaned slides.
2. Filter mordant onto slide as above directed (preferably using Thatcher's I: I dilution);
allow acting for 60 sec without heating.
3. Wash in tap water.
4. Flood slide with freshly filtered Ziehl's carbol fuchsin, and allow to stand 5 mID
without heating.
5. Wash with tap water.
6. Air-dry, and examine. Sometimes considerable search may be needed before finding a
satisfactorily stained part of the smear.
Results
Fagella well stained (red) in the case of those bacteria (e.g., colon-typhoid group, aerobic
spore fonners) that do not have extremely delicate flagella.
46
B) Gray'. Flagella Stain
Solution A (Mordant):
KAI(SO,),12H,O (saturated aq. solution)
5 ml
Tannic acid (20% aq. solution)
2 ml
(A few drops of chloroform must be added to this if a large quantity is made up)
HgCl, (saturated aq. solution)
2 ml
Solution B
Basic fuchsin (saturated ale solution)
0.4 ml
Mix solutions A and B less than 24 h before using. Both solutions separately may be kept
indefinitely, but deteriorate rapidly after mixing.
Staining protocol
1. Prepare smears from young cultures as above directed.
2. Flood slide with freshly filtered mordant, and allow acting 8-10 min.
3. Wash with a gentle stream of distilled water, and follow steps 4-6 of above schedule
(Casares-Gil method).
Results
Fagella well stained (red) in the case of those bacteria (e.g., colon-typhoid group, aerobic
sporefonners) that do not have extremely delicate flagella.
C) Leifson's Stain
KAI(SO.j,12H,O, or NH,AI(SO.j,-12H,O (saturated aqu solution)
20 ml
Tannic acid (20% aqu solution)
10 ml
Distilled water
IOml
Ethyl alcohol (95%)
IS ml
Basic fuchsin (saturated solution in 95 % ethyl alcohol)
3 ml
Mix ingredients in order named. Keep in tightly stoppered bottle, and the stain may be
good for a week.
Staining protocol
1. Prepare slides as for the preceding methods.
2. Flood slides with the above solution, and allow standing 10 min at room temperature
in wann weather or in an incubator in cold weather.
3. Wash with tap water.
4. Dry and examine.
Results
When no counter-stain is used, same as with the two above procedures; with methylene
blue counter-stain, flagella red, cells blue.
47
BaUey Method Modified by Fisher and Conn
This method is specially recommended for bacteria on which flagella are difficult to stain
(as is frequently the case with soil and water non-spore formers and with plant pathogens)
because of slime production, unusually fine flagella or flagella that are readily lost.
Solution A (Mordant)
Tannic acid (10 % aqu solution)
18 ml
FeCl,-6H,O (6 % aqu solution)
6ml
Solution B
Solution A
3.5 ml
Basic fuchsin (0.5 % in ethyl alcohol)
0.5 ml
He l (concentrated)
0.5 ml
Formalin
2.0ml
Ziehls Carbol-fuchsin
Solution A: Sat ale sol basic fuchsin 10 ml:
Basic fuchsin (90 % dye content)
0.3 g
Ethyl alcohol (95 %)
10 ml
Solution 8: 5 % sol carbolic acid 100 ml
Phenol
5g
Distilled water
95 ml
Mix solutions A and B
Staining protocol
5. Prepare smears of young cultures.
6. Filter the above solution A onto the slide and allow it to remain 33 min without
heating.
7. Pour off solution A, and without washing add solution 8 , also through a filter, and
allow it to stand 7 min without heating.
8. Wash with distilled water.
9. Before the slide dries, cover with Ziehl's carbol fuchsin , allowing it to stand 1 min on
a hot plate heated just enough for steam to be barely given off.
10. Wash in tap water.
II . Dry in the air, and examine.
Results
Similar to the preceding methods, but the background precipitate is usually finer and less
conspIcuoUS, thus interfering less with the demonstration of unusually fine, delicate
flagella.
48
Exercise 10- Staining of bacterial inclusionl storage bodies
Introduction
Most bacteria do not live in environments that contain large amounts of nutrients at all
times. To accommodate these transient levels of nutrients bacteria contain several
different methods of nutrient storage in times of plenty for use in times of want. 1-or
example, many bacteria store excess carbon in the form of polyhydroxyalkanoates or
glycogen. Some microbes store soluble nutrients such as nitrate in vacuoles. Sulfur is
most often stored as elemental (So) granules which can be deposited either intra- or
extracellularly. Sulfur granules are especially common in bacteria that use hydrogen
sulfide as an electron source. A number of STORAGE GRANULES may be present in a
cell depending on its physiology & nutritional environment. These may be STARCH,
FAT, SULFUR, or PHOSPHATE. Bacteria exist in a very competitive environment
where nutrients are usually in SHORT SUPPLY, so they tend to store up extra nutrients
when possible. Most of the above mentioned examples can be viewed using a microscope
and are surrounded by a thin non-unit membrane to separate them from the cytoplasm.
Nutrients and reserves may be stored in the cytoplasm in the form of glycogen, lipids,
polyphosphate, or in some cases, sulfur or nitrogen. Special stain generally use to see
storage granules present inside the bacteria. In case of Corynebacterium there is Storage
granules called Volutin or Babes-Ernst granules or Polar bodies.They are the imparts
diagnostic value in the identification of Corynebacteriun. It is possible to see by Staining
with Special stain like I. Albert's stain, 2.Neisser's stain, 3. Ponders stain.
The poly-B-hyclroxybutyrate stain is used to stain granules present within the confines of
the filamentous bacteria cells. These granules are storage components of the cells giving
indication of the cell's ability to take advantage of an opportunity of "free" easily
absorbable and available low molecular weight carbonaceous compounds present in the
environment. Typically, PHB deposition within a cell occurs under specific operational
conditions: High BOD:N:P ratios, anaerobic conditions present or created within a
treatment process (insufficient DO, dead zones within an aeration basin which create
anaerobic conditions which lead to production of low molecular weight organic acids
such as acetic, propionic, butyric acids due to fennentation).
Intracytoplasmic inclusions can be vacuoles, crystals or storage bodies. Bacteria often
store reserve material in the form of insoluble cytoplasmic granules. Inclusions
accumulate when a cell is grown in the presence of excess nutrients and they are often
observed under laboratory conditions. Various examples of these bodies arc:
Starch/Glycogen granules - blue-greens and enteric bacteria, Poly-B-hydroxybutyratc
granules - Azotobacter and Rhizobium, Nitrogen-reserve granules - blue-greens, Sulphur
inclusions - Thiotrix, Lipid inclusions, Volutin granules - Corynebacterium diphtheria.
The inclusion bodies can be appreciated using phase contrast microscope or using special
stains such as Albert ' s stain (volutin granules) or Sudan black (lipid inclusion).
Materials
•
Solution I: Sudan Black B (IV) - 0.3% w/v in 60% Ethanol
•
Solution 2: Safranin 0 - 0.5% w/v aqueous
49
·~·'h"~')'.j
; il.l':.
)F·
.' J~' ,;~ ~ -!C , ... hll .... ... . ,(1, ... ...di u i
lIitf.1(:dhd,\i granules;
yellow-brown is negative.
Observations
Results
Inferences
50
I
'1 ,
p ll a~e ~;o nt rRs,() :
ye l1ow ~ bro\Vn
,l
dlUe Vl old 13 P,)SltlVC (Gllhcr ..::ntin.; cdl or
!Illn~cclluJar
granules;
is negat ive.
Observations
Results
Inferences
,
1
50
"
Protocol
12. Prepare thin smears on microscope slides and thoroughly air dry . Do not heat fix.
13. Stain 10 minutes with solution 1; add more stain if the slide starts to dry out.
14. Rinse I second with water.
15. Stain 10 seconds with so lution 2; rinse well with water; blot dry.
16. Examine under oil immersion at 1000x magnification with direct illumination (not
phase contrast): Blue-violet is positive (either entire cell or intracellular granules;
yellow-brown is negative.
Observations
Results
Inferences
,
,
j
J
~1
50
i
I
.'
PHD Staining
Test organism: Bacillus thuringiensis
Staining protocol
I. Prepare smears of the organism, air dry and heat fix. Flood entire slide with Sudan
Black B andadd more stain as the dye solvent evaporates. Stain for at least 10
minutes.
2. Pour off excess stain (do not wash) and air dry.
3. Clear slide by dipping in a jar of solvent in the fume hood for 5 sec. Air dry in the
fume hood.
4. Counter stain for I min. with safranin.
5. Wash with water, drain, blot and air dry. Examine with oil immersion objective. Cell
is pink; lipids are dark grey or black.
Fat Droplets Staining
Burdon's Method
Staining solution
Sudan black B (commission certified)
0.3 g
Ethyl alcohol 970%)
100 ml.
After the bulk is dissolved, shake at intervals and allow standing overnight.
Staining protocol
I. Prepare smears as usual from 18 to 24 h cultures, and fix by heat.
2. Flood the entire slide with the above staining solution and allow it to stand undisturbed
at room temperature for 5-15 min. (Exact time is unimportant, as good results are often
obtained after only 1 or 2 min; on the other hand no hann results if the slides stain
until the solution is completely dry.)
3. Drain and blot slide completely dry.
4. Cover with xylene by pouring from a dropping bottle or dipping several times in a
staining jar. Blot till dry.
5. Counter-stain 5-10 sec with 0.5 per cent aqueous safranin, taking care not to over stain.
Note: For acid-fast organisms, Ziehl's carbon fuchsin diluted 1: 10 with distilled water
may be applied for 1-3 min, instead of safranin.
6. Wash in tap water, blot, and dry. Fat droplets blue-black or blue-gray; rest of cell pink.
Metachromatic Granules Staining
Various special procedures have been devised for staining the diphtheria organism in such
characteristic
a manner as to render it distinctive in appearance by differentiation
~\.\BRAl\'r
metachromatic granules.
<1s:"l!>~"6Iv-n.
0 ., Coo
Staining protocol
51
10--
1Vl' .I?/
" ~/I _ I?NA,(
.
""~I
3;;
." -Io-J~~
.
tvletachromatic granules arc dark blue to viol et. Bacteria without slich granules are
stained unifonnly. The picture varies little according to the two methylene blue solutions
are employed. The Loeffler fonnula gives purplish shades of staining because of the
oxidation of methylene blue caused by the alkali. Some users consider the polychrome
effect thus obtained to give better differentiation; others think the meta-chromatic
granules show more sharply with the clear blue of the un-poly-chromed dye.
Albert's Diphtheria Stain
Toluidine blue
0.15 g
Methyl green
0.20 g
Acetic acid (glacial)
I ml
Ethyl alcohol (95 %)
2ml
Distilled water
100 ml
Lugol's Iodine
Iodine
2g
KI
3g
Distilled water
300 ml
Laybourn's Modification
Layboum modified the Albert stain by replacing the methyl green with an equal amount
of malachite green.
Staining protocol
I. Make unifonn thin smears and fix with gentle heat.
2. Stain 5 min in either Albert's staining fluid or Layhoum's modification of it. The latter
is claimed to give deeper staining of both granules and body of the cells without
lessening the contrast between them.
3. Drain without washing.
4. Treat I min in a modified Lugol's iodine.
5. Wash briefly with tap water.
6. Blot dry with filter paper, and examine under oil immersion objective (lOOx).
Results
Metachromatic granules are black; bars of diphtheria cells are dark green to black; hody
of cells is light green.
52
Ljubinsky Stain
Solution A:
Pyoktanin (Merck)
0.25 g
Methyl violet 2B or crystal vio-Iet (85 % dye content)
0.25 g
5 % acetic acid
100 ml
Glacial acetic acid
5 ml
Distilled water
95 ml
Solution B:
Vesuvin
0. 1 g
Bismarck brown Y
0.1 g
Distilled water
100mi
Staining protocol
1. Make smears and fix with gentle heat.
2. Stain 30 sec to 2 min with solution A.
3. Wash under gentle flow of tap water.
4. Stain 30 sec with solution B.
5. Wash with tap water.
6. Dry, and examine under oil immersion objective (lOOx).
Results
Metachromatic granules are dark blue or black; rest of cell is reddish or yellowish.
Neisser's Diphtheria
staini~g
Solution I:
Methylene blue (dye content not specified; probably 90 %)
I g
Alcohol (e.g. , 95 %)
20ml
Acetic acid (glacial)
50ml
Distilled water
1000 ml
Solution 2:
Crystal violet (dye content not specified; probably 85 %)
Ig
Alcohol (e.g., 95 %)
10 ml
Distilled water
300 ml
Solution 3
Chrysoidin
lor 2 g
Hot water
300ml
Mix by agitation till the dye dissolved and filter it.
53
Staining protocol
I. Make smears and fix with gentle heat.
2. Stain for 10 sec in a mixture of 2 parts of solution 1 and I part of Solution 2.
3. Wash with tap water and stain 10 sec in solution 3.
4. Wash briefly in water, or not at all.
5. Blot dry and examine under oil immersion objective (IOOx).
54
Exercise 11- Staining of bacterial nucleoids
Introduction
Unlike eukaryotes, the bacterial chromosome is not enclosed inside of a membrane-bound
nucleus but instead resides inside the bacterial ,cytoplasm. This means that the transfer of
cellular infonnation through the processes of translation, transcription and DNA
replication all occur within the same compartment and can interact with other cytoplasmic
structures, most notably ribosomes. The bacterial chromosome is not packaged using
histones to fann chromatin as in eukaryotes but instead exists as a highly compact
supercoiled structure, the precise nature of which remains unclear. Most bacterial
chromosomes are circular although some examples of linear chromosomes exist. Along
with chromosomal DNA, most bacteria also contain small independent pieces of DNA
called plasmids that often encode for traits that are advantageous but not essential to their
bacterial host. Plasmids can be easily gained or lost by a bacterium and can be transferred
between bacteria as a form of horizontal gene transfer.
The chromosome in bacteria is typically a single, closed circle DNA that is concentrated
in a nucleoid region. It is not membrane bound as in eukaryotes. Some bacteria possess
smaller extrachromosomal pieces of DNA called plasmids. Plasmids replicate
independently of the chromosome and carry genes that are not essential for cell survival
but may give some advantage to an organism. The chromosome is attached to an
invagination of the cytoplasmic membrane called mesosome. Mitotic apparatus and
nuclear membrane are completely lacking. The length of E.coli chromosome is
approximately 1.4 mm but is condensed inside the cell by supercoiling. DNA is mainly
negatively charged hence bind readily to basic dyes. It can be demonstrated by Feulgen
stain or by electron microscopy.
Materials
Stain: Add I drop of Giemsa stain to I ml of Sorensen's buffer of pH 6.9-7.0.
Protocol
Fixation and smearing
I . Incubate petri-dish cultures 2-5 h.
2. Remove a block of agar, and fix it from a few seconds to several hours in the vapor of
2% osmic acid.
3. Make an impression smear on a cover slip or glass slide.
4. Store in 70% alcohol till needed.
Staining procedure:
1. Remove preparation from alcohol, and wash in water.
2. Place for 5-10 min in nonnal HCI at 60°C.
3. Remove, and wash three times in tap water.
4. Stain 1-15 min, at 37°C, in the above diluted Giemsa stain.
S. Mount in water for oil immersion examination. The deeper colors (blue and violet)
tend to be localized in the chromatinic material comprising the nuclear structures.
55
Observations
Results
Inferences
56
OTHER METHODS
Feulgen Method
The method for staining of chromoseomes was described by Robert Feulgen. It is widely
used to identify chromosomal material or DNA. This method relies on acid hydrolysis of
DNA, thus fixation using strong acids should be avoided. This method selectively stains
DNA and under controlled conditions can be used for photomerric determination of DNA
content. Fixed bacterial cells is treated with IN HCI at 60°C in water bath for 10 min.
Acid hydrolysis removes purine bases from DNA, thus unmasking the free aldehyde
groups, which react with schiff's reagent to give purple staining. RNA is not hydrolyzed
thus reaction is DNA specific.
Materials
•
Bacterial culture
•
IN HCI
•
Monobromonophatalin solution
•
Oreein solution
•
Schiff's reagent
•
Acetic acid (100%)
Protocol
I . Prepare thin smear of bacterial culture on clean and dry glass slide.
2. Add 2-3 drops ofmonobromonophatalin and keep for 5 h at room temperature.
3. Decant the solution and flood with 100% acetic acid and keep in refrigerator for 10
mm.
4. Drain the acetic acid and flood with IN HCI at 60°C to liberate the purines.
5. Drain the HCI and add Schiff's base and keep at room temperature until colour
develops.
6. Wash the Schifrs reagent and counter stain with oreein and wash after I min.
7. Examine under microscope (100x objective).
Observations
Results
Inferences
57
Exercise 12- Examination of bacterial motility
Introduction
The microorganisms can move from one place to another by the help of their locomotive
system in response to favorable and adverse environmental conditions. The very common
locomotive system of bacteria is flagellum (pI. flagella) and present in motile rod shaped
bacteria but rare or nil in coccus shaped bacteria. Flagella rotate in clockwise or anti clock
wise and thus induce forward and backward movement of bacterial cells. Flagella are
semi-rigid cylindrical structures that are rotated and function much like the propeller on a
ship. Bacterial species differ in the number and arrangement of flagella on their surface;
some have no flagellum (atrichus), single flagellum (monotrichous), a flagellum at each
end (amphitrichous), clusters of flagella at the poles of the cell (lophotrichous), while
others have flagella distributed over the entire surface of the cell (peritrichous).
The flagellum is a rotating structure driven by a reversible motor at the base that uses the
electrochemical gradient across the membrane for power. This motor drives the motion of
the filament, which acts as a propeller. Many bacteria (such as E. coli) have two distinct
modes of movement: forward movement (swimming) and tumbling. The tumbling allows
them to reorient and makes their movement a three-dimensional random walk. The
flagella of a unique group of bacteria, the spirochaetes, are found between two
membranes in the periplasmic space. They have a distinctive helical body 'that twists
about as it moves. Motile bacteria are attracted or repelled by certain stimuli i,n behaviors
called taxes: these include chemotaxis, phototaxis, energy taxis and magnetotaxis. In one
peculiar group, the myxobacteria, individual bacteria move together to form waves of
cells that then differentiate to form fruiting bodies containing spores. The myxobacteria
move only when on solid surfaces, unlike E. coli which is motile in liquid or solid media.
Flagella consist of a.hollow, rigid cylinder composed of a protein called flagellin, which
forms a filament anchored to the cell by a curved structure called the hook, which is
attached to the basal body. Flagellae are, in effect, rotary motors comprising a number of
proteinaceous rings embedded in the cell wall. These molecular motors are powered by
the phosporylation cascade responsible for generating energy within the cell. In action,
the filament rotates at speeds from 200 to more than 1,000 revolutions per second, drivi.ng
the rotation of the flagellum. The organization of these structures is quite different from
that of eukaryotic flagella. The direction of rotation determines the movement of the cell.
Anticlockwise rotation of monotrichious polar flagella thrusts the cell forward with the
flagellum trailing behind. Peritrichous cells operate in the same way. Bacteria can sense
nutrient molecules such as sugars or amino acids and move towards them - a process is
known as chemotaxis. Additionally, they can also move away from harmful substances
such as waste products and in response to temperature, light, gravity, etc. This apparently
intelligent behaviour is achieved by changes in the frequency of tumbles. When moving
towards a favourable stimulus or away from an unfavourable one, the frequency of
tumbles is low, thus the cells move towards or away from the stimulus as appropriate.
Bacteria are very motile during logarithmic phase of growth. The motility of bacteria can
be studied by a) hanging drop technique and b) soft agar diffusion method
58
Materials
A. Hanging drop method of motility
• Special microscope slide with a depression (cavity slide)
• Cover slip
• Micropscope
• Immersion oil
• Actively growing bacterial culture (e.g. Escherichia coli, motile by peritrichus
flagella)
B. Semi-solid agar method
• The agar medium is prepared with the agar content of 0.2%. The medium is put into
test tubes.
Protocol
A. Hanging drop method of motility
I. Take clean and dry cavity slide and cover slip.
2. Place one drop of the culture onto the cover slip and put Vaseline at four comer of the
cover slip.
3. Put the cover slip with culture drop carefully and invert it on the cavity slide (grooved
microscope slide) so that a hanging drop on the cover slip inside the cavity can be
obtained gently.
4. Put a drop of immersion oil on cover slip (just on the culture drop) and visualize
under oi l immersion (lOOx) objective of the microscope. Better visualization can be
achieved by using phase contrast microscope.
5. Motility is characterized by fast unidirectional movement as compared to the
Brownian motion whereby the cells move round in one particular point and can be
observed at periphery of the bright microscopic field.
B. Semi-solid agar method
I. Inoculation is-done by stabbing the medium at the centre.
2. The inoculated medium is incubated at appropriate temperature for 24 h.
3. Motility is detected by observing turbidity at the line of inoculation.
Observations
Results
Inferences
59
Exercise 13- Preparation of bacterial protoplasts
Introduction
Protoplasts are the cells of which cell walls are removed and cytoplasmic membrane is
the outermost layer in such cells. More generally protoplast refers to that unit of biology
which is composed of a cell's nucleus and the surround ing protoplasmic materials.
Protopiasts can be used to study membrane biology, including the uptake of
macromolecules and viruses. Protoplasts are widely used for DNA transformation (for
making genetically modified organisms), since the cell wall would otherwise block the
passage of DNA into the cell. ProtoplaSls may also be used for plant breeding, using a
technique called protoplast fusion.
Protoplast can be obtained by specific lytic enzymes to remove cell wall. Bacterial cell
walls are made of peptidoglycan, a polysaccharide backbone consisting of alternating Nacetyl muramic acid (NAM) and N-acetylglucosamine (NAG) residues in equal amounts.
Cell wall of some Gram positive bacteria is completely dissolved by lysozyme. as this
enzyme attacks the bonds between GA and MA. Therefore, the protoplasts can be made
by degrading cell walls lysozyme+EDTA. During and subsequent to digestion of the cell
wall, the protoplast becomes very sensitive to osmotic stress. This means cell wall
digestion and protoplast storage must be done in an isotonic solution to prevent rupture of
the plasma membrane. Bacterial protoplasts are believed to be cellular units that have
been deprived of their rigid cell wall. Accordingly, they are distinguished by their
spherical shape (in bacillifonn species) and their sensitivity to cytolysis in hypotonic
media. Protoplast of gram-positive bacteria is prepared by treating the cells by lysozyme
in presence of EDTA. A method has now been found for the efficient production of
protoplasts from enteric bacteria such as Escherichia coli and Salmonella typhimurium.
The technique consists essentially of the exposure of growing cells to a medium
containing penicillin, sucrose, and Mg++.
Preparation of protoplasts of Gram's positive bacteria
Materials
• Overnight grown bacterial culture (gram positive bacteria e.g. Lactobacillus spp. or
Bacillus spp.)
• TBS buffer
• Penassay broth (gil it): Tryptone - 5g, yeast extract - l.5g, beef extract - 1.5g, sodium
chloride -3 .5g, dextrose - Jg, potassium phosphate dibasic - 3.68g, potassium
phosphate monobasic - J .32g. pH Adjusted to 7.0±0.2.
• Sucrose solution
• Lysozyme
• EDTA
• Phase contrast microscope
60
Protocol
Method I
1. Take overnight grown bacterial culture and harvest the cells by centrifugation and
wash twice.
2. Suspend the bacterial cells in medium containing 0.5 M sucrose (TBS butTer) and
incubate at 37°C for 20 in a shaker incubator.
3. Add lysozyme into the cell suspension to get final enzyme concentration of 50
microgram! mt and incubate for ;mother 20 min in similar conditions.
4. Add EDTA and incubate for 20-30 min.
5. Separate the cells/protoplast by centrifugation and suspend in 200 micro-liter of TBS
buffer and prepare smear and observe under phase contrast microscopy.
Method II
1. Grow the bacterial cells upto late log phase.
2. Harvest the cells by centrifugation at 10000 rpm for IO min at 4°C.
3. Wash the cells once with double-strength Penassay broth containing 0.5 M-sucrose,
20 mM-MgCI" 20 mM maleic acid and NaOH, pH 7.0±0.2 (neutral SMMP medium).
4. Suspend the cells in SMMP medium in concentrated fonn.
5. Add 0.01 volume of 1% (w/v) lysozyme solution to the suspension.
6. Monitor the protoplast formation at 37°C microscopically.
7. Recover protoplasts by centrifugation at 1000 g for 30 min at 10°C and washed twice
with the SMMP medium and observe under microscopy.
Preparation of protop lasts of Gram's negative bacteria
Preparation of E. coli protoplasts by lysozyme
1. Grow E. coli cells overnight in BHt broth at 37°C with shaking.
2. Harvest the cells by centrifugation at room temperature from 10 ml culture.
3. Suspend the cells in distilled water, and harvest by re-centrifugation.
4. Suspend cell pellet III 5 m1 of cold 0.5 M sucrose
tris(hydroxymethyl)aminomethane (tris) chloride buffer (PH 8.0±0.2).
to
0.03
M
5. Carry out all the subsequent operations at O°C.
6. Add 4 micromoles of sodium EDTA (PH 8.0±0.2) and 15 to 25 mIcrogram of
crystalline egg white lysozyme to the cell suspension.
7. Centrifuge a sample of the suspension and suspend the pellet in water to follow the
action of the lysozyme. (If protoplast formation is completed, the pellet will be
converted to a sticky viscid white material which will be subsequently solubilised on
stirring. If the pellet is suspended in 0.5 M sucrose, however, lysis did not occur.
Perform all experiments as soon as protoplast formation is complete (10 to 30 min
after addition of lysozyme), since, on more prolonged exposure to lysozyme, even at
zero degrees, the protopiasts underwent further changes.).
61
8. Examine under the phase microscopy for protoplast fonnation.
Note: EDTA must be added at the same time as the lysozyme, that excess lysozyme or
divalent ions can inhibit lysis, and that the cells must be freshly harvested after good
aeration.
Alternate method
Escherichia coli and a variety of its mutant sub-strains are used in most of the
experiments. The use of penicillin and sucrose was suggested by the possible analogy
between penicillin and lysozyme as lytic agents and by the finding that hypertonic
sucrose would interrupt bacteriolysis of Bacillus megalerium by lysozyme. ' In addition,
spherical bodies had been casually noted in other applications of penicillin, and many
authors have emphasized its use in the production of L-fonns. The following procedure
was adopted after empirical trials and can doubtless be further improved.
Protocol
I. Grow the bacteria overnight in tubes with 10 ml of broth (Difco penassay medium) at
37°C, on a rotator.
2, Samples of 3 ml of the grown culture, add directly to 10 ml of broth supplemented
with penicillin (1000 ulml), sucrose (20%), and magnesium sulfate (0.25%).
3. In 2-3 h, the cells are quantitatively converted into spheres.
4. During this interval the optical density (measured at 650 nm) is increased about 50%,
but the total count (spheres or rods) remained constant. .5. The spheres promptly are lysed upon dilution of the suspension in distilled water, and
they are therefore regarded as "protoplasts."
Note
I. The indicated supplements are in substantial excess, and nearly optimal yields of
protoplasts can be obtained with 5% sucrose, 100 unit penicillin and 0.1 % MgS0 4 .
The high magnesium requirement may depend partly on binding with the sodium
citrate used in the compounding of the penicillin preparation. The Mg++ can be
replaced by Ca++ but was preferred, to minimize precipitation of phosphate. In the
absence ofMg++, protoplasts are fanned but are less well preserved, and a third or less
of the initial rods is recovered. In the absence of sucrose or comparable stabilizer, the
cells are almost totally lysed, leaving only debris. In the absence of penicillin, the rods
grow at nearly the nonnal rate, i.e., are not appreciably inhibited by 20% sucrose.
2. Protoplasts are also fanned in minimal medium, but the transfonnation has been
irregular and incomplete in the trials to date. Cells exposed to penicillin under
conditions not supporting growth, e.g., in sucrose buffer, were not transfom1ed into
spheres.
3. The transfonnation of rods into spheres was observed in small droplets each
containing ten to twenty cells, immersed in an oil chamber. Each rod gave rise to a
62
single sphere; even cells about to divide were inhibited in further growth and division.
The progression of stages as observed with dark phase contrast at IOOOx.
Observations
Results
Inferences
63
Exercise 14 - Preparation of bacterial spheroplasts
Introduction
Spheroplasts are spheriod bodies fanned after partial loss of the bacterial cell wall; they
are osmotically fragile, and undergo lysis in media of low osmotic pressure. The cells of a
number of unrelated species of Gram-negative bacteria could be lysed by lysozyme, in the
presence of ethylenediaminetetraacetic acid (EDTA) with tris buffer (tris-(
hydroxymethyl) aminomethane) at pH 8. It is the purpose of this communication to
demonstrate that treatment of Gram negative bacteria with the Iysozyme-EDTA-tris
buffer system may yield osmotically fragile celis, without so complete a destruction of the
rigid cell wall as to cause the production of true spheroplasts.
Spheroplasts - have their cell wall only partially removed. Several groups have developed
various ways to create potentially viable spheroplasts by using penicillin, lysozyme and
ampicillin methods. These methods allow the spheroplasts to revert back to their original
growth competent rod-shaped morphology once the stress is relieved. The inner and outer
membranes of gram-negative bacteria are usually obtained from spherop lasls. The
lysozyme treatments normally used for converting cells to spheroplasts were originally
developed with exponential phase cells and have proven to be ineffective with cells
grown under other conditions. A procedure has therefore been developed which renders
variously grown cells completely susceptible to lysozyme. This procedure has been tested
on various strains of Escherichia coli at all stages of growth in minimal medium. from the
early exponential to the late stationary phase. It has been tested on stationary phase
cultures which ceased to grow because of limiting aeration, limiting carbon source,
limiting amino acids, and limiting nicotinic acid. Very efficient conversion of cells to
spheroplasts was observed in all cases. The resulting spheropJasts are an excellent source
for subsequent membrane separations.
Materials
• E. coli active culture
• Sucrose
• MgSO,.7H20,
• Ampicillin! Penicillin
• Phosphate buffer
• LB medium
Protocol
Method I
I. Grow culture overnight at 37°C in shaking water bath.
2. Dilute the culture in phosphate buffer supplemented with the following reagents (I :5:
Reagents used at the following concentration to induce spheroplast formation in E.
coli: OAM sucrose, 8 mM MgSO,.7H20, and penicillin at 50 microgram/ml.
3. Dilute this culture 5 fold in LB, with each treatment containing the following reagents
as follows:
Reagent
Treatment
64
Control
Sucrose
MgSO•.7H,O
Penicilin
Sucrose
Sucrose and Penicilin
PeniciUn
+
+
+
+
+
4. Cultures were then incubated at 37°C at 250 rpm until spheroplasts were fonned
(approximately 2 h).
5. The transition from rod to spherical morphology was observed by using a phase
contrast microscope at lOOx magnification.
Method II
1. Harvest cells from I mt of stationary phase E. coli culture by centrifugation and wash
in Iml O.OIM phosphate buffer, pH 7.0.
2. Pellet the cells by centrifugation at 10,000 rpm and re-suspend in 500
sucrose solution made in PB to induce plasmolysis.
~l
of a 0.5 M
3. Add lysozyme to the cell suspension (50 mg! ml).
4. After incubation in a 37°C water bath for 1- 2 h, dilute the suspension I: I with PB and
add EDTA to a final concentration of 10 mM.
5. Incubate at 37°C in water bath for 10 min.
6. Detennine the transition of the rod-shaped bacteria into spheres by light microscopy/
phase contrast microscopy to measure progression of the spheroplasts fonnation.
7. When ~80% of the cells are converted to spheroplast, stop the reaction by petleling
the cells at 500xg for 15 min.
8. Wash the spheroplasts in 0.25 M sucrose in PB, pellet at 5000 rpm for 15 min and resuspend in same solution.
Observations
Results
Inferences
65
Exercise 15 - Study of the morphology of yeasts
Introduction
Yeasts are single celled fungi; spherical to oval in shape and reproduce by process of
budding. Yeasts are a heterogenous group of fungi that superficially appear to be
homogeneous. Yeasts grow in a conspicuous unicellular fann that reproduces by fission,
budding, or a combination of both. True yeasts reproduce sexually, developing ascospores
or basidiospores under favourable conditions. The identification of these fungi is based
upon a combination of morphological and biochemical criteria. Morphology is primarily
used to establish the genera.
Yeast cells exhibit great diversity with respect to cell size, shape and colour. Even
individual cells from a particular yeast strain of a single species can display
morphological and colour heterogeneity. This is mainly due to alterations of physical and
chemical conditions in the environment. S. cerevisiae cells are generally ellipsoidal in
shape ranging from 5 to I 0 ~m at the large diameter and I to 7 ~m at the small diameter.
Mean cell volumes are 29 or 55 ~3 for a haploid or a diploid cell, respectively; cell size
increases with age. Unstained yeast cells can hardly be visualized by light microscopy. At
1000 fold magnification, it may be possible to see the yeast vacuole and cytosolic
inclusion bodies. By phase-contrast microscopy, together with appropriate staining
techniques, several cellular structures can be distinguished. Fluorochromic dyes can be
used with fluoresecence microscopy to highlight features within the cells as well as on the
cell surface.
Structure-specific dyes for yeast cells
Dye
Structures visualized
Comments
Methylene Blue
Whole cells
Non-viable cells stain blue
Aminoacridine
Cell walls
Indicator of surface
potential
Calcofluor white
Bud scars
Chitin in scar fluoresces
Neutral red
Vacuoles
Vacuoles stain red-purple
Iodine
Glycogen deposits
Glycogen stained red-brown
Materials
•
Yeast culture on solid agar or
Kluyveromyces spp.)
•
Glass slide
•
Burner
•
70% alcohol
•
Microscope
III
broth culture (Saccharomyces spp. and
Protocol
a) Positive staining
6. Prepare two smears on clean and grease free microscopic slides and heat fix over the
gas flame.
7. Flood the smear with the methylene blue or crystal violet solution for 1-2 min.
8. Gently wash the slide with water to remove the excess dye.
9. Blot dry followed by drying over the gas flame of the stained smear on the slide and
examine under the microscope (100x. oil immersion objective).
10. Draw and identify the shape and arrangement of the bacterial cells in the circles
below.
b) Negative staining
6. Take two clean and grease free microscopic slides.
7. Transfer two loopful of broth culture and one loopful of nigrosin solution on the slide
and mix thoroughly with the help of inoculating loop.
8. Spread unifonnly the mixture of bacterial culture and nigrosin dye to get a thin and
transparent smear on the slide.
9 . Allow to air dry ofthe smear and examine under the microscope (lOOx, oil immersion
objectives
10. Draw and identify the shape and arrangement of the bacterial cells in the circles
below.
Observations
67
Results
Inferences
68
Exercise 16 - Study of tbe morpbology of moulds
Introduction
The classification of fungi includes both yeasts and molds. Both of these groups have
useful microorganisms and pathogenic microorganisms within them. Molds have the
following characteristics filamentrous fungi, hyphae, a single filament grows into
multiple filaments called mycelium, hyphae develop from fungal spores that can produce
either sexually or asexually, fungi are decomposers their hyphae release enzymes that
break down nutrients in their environment which then are absorbed, likes a slightly moist
environment, can grow in temperatures ranging from _6°C to 50°C. In this exercise three
fungi Aspergillus, Rhizopus stolonifer, and Penicillium notatum, will be studied.
Aspergillus is commonly found 0 in soil and hay, but when the spores are inhaled they
can cause a respiratory disease called Aspergillosis. Also this fungus can produce an
Aflatoxin (when it infects peanuts) that is a potent carcinogen. Rhizopus stolonifer is also
known as " Black Bread Mold." Penicillium nota tum is a blue gray mold that is used to
produce the chemotherapeutic agent Penicillin.
Direct Mounts - Direct mounts are made in order to study yeast morphology
microscopically and to detennine purity of the isolates. Lactophenoi Mount is prepared
on a clean glass microscope slide. Lactophenol cotton blue is a stain commonly used for
making semipermanent microscopic preparation sof fungi. It stains the cytoplasm of fungi
and provides a light blue background against which walls of hyphae can be seen. It
consists of 4 components phenol (fungicide), lactate (cleaning agent), cotton blue (stain
fungal cytoplasm) and glycerine. Lactophenol blue (phenol+lactic acid+cotton blue =
lactophenoi cotton blue staining) has 3 important functions when observing filamentous
fungi. Phenol destroys contaminants compounds that have the fungi, the acid lactic
conserves the fungi structures by provoking osmotic gradient change with relation of the
interior of the fungi forming a protective layer. The cotton blue is the one that gives the
color and pennit to visualize the fungi under optic microscope. It adheres to the chitin of
the fungi walls of hiphas and conidios. Use of the Lactophenol Cotton Blue Stain
Droppers in preparing a wet mount of a mold or yeast will stain the specimen light blue
and allow it to be easily visualized by microscopy. Lactophenol Cotton Blue Stain
solution consisting of glycerol 40%, phenol 20%, lactic acid 20%, cotton blue dye
(Poirrier's blue) 0.05%, and water.
Materials
•
Fungal culture (Mould culture on solid agar surface or in broth culture)
•
Lactophenol cotton blue stain (Lacto phenol cotton blue mount: Phenol - 20 gm;
Lactic acid - 20 ml; Glycerine - 40 ml; Cotton blue - 0.05gm; Distilled water - 20 ml.
Add in order to prepare the dye.)
•
Needle
•
Glass slide
•
Coverslip
•
Burner
69
•
70% alcohol
•
Microscope
Protocol
I. Place a drop of lactophenol cotton blue on a clean and dry glass slide.
2. Transfer a small portion of fungal material into the drop using a inoculating needle
and mix the content.
3. Gently place a cover slip over the dye mix to avoid trapping of air bubble inside.
4. Examine ueder the lOx objective first and then under 40x objective of microscope.
Observations
Results
Inferences
70
Exercise 17 - Detection and enumeration of bacteriophages in cheese whey
Introduction
Phages cause lysis of lactic acid bacteria used in cheese and fermented milks production.
The virulent bacteriophages specific for lactic acid bacteria can be isolated and purified
from cheese whey. They show distinct plaque sizes, morphology by electron microscope
examination, the dimensions etc. The phage heads are elongated and hexagonal in shape,
and the flexible tails appeared periodically cross-striated. On infection, phage is adsorbed
on the bacterial surface by the free end of the tail. The bacteriophage in cheese whey is
enumerated on soft agar overlay method after making suitable dilution using suitable host
Lactococcus spp., where phage plaques (clear zone of lysis of host cells) are fonned on
bacterial lawn. Further, the bacteriophage can further be enriched using host bacteria in
liquid broth, where phages are multiply in host cells and the number of specific
bacteriophage is increased. Size of bacteriophages is in nanometer range and hence, the
morphology of phages is camed out under electron microscopy.
Materials
•
Bacterial cultures - Lactococcus lactis subsp. lactis, subsp. cremor;s and diacetyiactis,
Lactobacillus case;, Escherichia coli.
•
Tryptone soy medium with 0.3% yeast extract (TSY) for culturing L. lactis subsp.
lactis and the other bacterial strains.
•
M broth (gil): tryptone, 20; glucose 5; NaCI, 4; sodium acetate, \.5 ; CaCh H20, 0.15;
MgSO.7H20, 0.2; and MnSO., 0.05; adjusted to pH 7.0 before being autoclaved at
121 °C for 15 min
•
Skim milk (II % TS) and heated at 100°C for 20-30 min.
•
Cheese whey samples - centrifuged at 5000 rpm and filtered through a Millipore
membrane (0.45 11m pore size).
Protocol
1. Inoculate Lactococcus spp. into the heated skim milk, with and without the cheese
whey filtrates.
2. Propagate bacteriophage on the sensitive strain Lactococcus spp. growing in M broth.
3. Use agar plates of this medium for plaque isolation.
4. Purify the phages by replaquing on sensitive lactocci strain.
5. Test the phage against Lactobacillus case; and E. coli (do not show any evidence of
bacterial lysis).
6. Grow Lactococcus spp. for 18 h in skim milk at 30°C for inoculation to M broth.
7. Inoculate by a second transfer (2.5% inoculum) into fresh M medium.
8. After 2 h, add phage at a mUltiplicity of infection of about I, and incubate continued
until lysis occurred.
9. filter the clear Iysates (10' to 10' PfUlml) through the Millipore membranes to
eliminate debris and store at 4°C.
10. Detennine the phage titers on M medium by the double-layer agar method.
71
Morphological features ofthe bacteriophages
I. Concentrate the bacteriophage and re-suspend in 0.1 M ammonium acetate.
2. Perfonn the negative staining with a saturated solution of uranyl acetate in water.
3. Touched a small drop of phage suspension by the surface of carbon-coated Formvar
film covering the electron microscope grid.
4. Remove excess material by absorption with filter paper, and floated the grid briefly on
uranyl acetate for 20 S, remove, and touch to a filter paper surface.
5. Examine the grid in a electron microscope at original magnifications of x50,OOO and
x75,OOO.
6. Negatively stain some specimens with 2% phosphotungstic acid in aqueous solution
brought to pH 7.0 with NaOH.
Observations
Results
Inferences
72
Exercise 18 - Application of computer software in bacterial identification
Introduction
Classification is orderly arrangement of individuals into units (taxa) on the basis of
similarity. Each unit (taxon) is homogenous and different from all others. Identification is
the matching of an unknown against known in a classification, using minimum number of
diagnostic characters. Identification of bacteria is based on the phenotypic and genotypic
characters. Phenotypic characters include morphological features, growth requirements
and physiological and biochemical activities. Genotypic methods used including DNA
base ratio, DNA-DNA hybridization, rRNA homology and DNA based typing methods.
Computers are emp loyed in the collection and analysis of this data. Polyphasic
identification is the integration of these various techniques for identification of unknown
bacteria. Few computer based polyphasic identification systems have been developed.
Most computer based identification systems use only a subset of taxonomic infonnation
available for a particular group of bacteria. A wide and increasing range of computerized
systems for the identification of bacteria is now reported in the scientific literature and
allied to commercial kits. Recently, genomic analyses have proven successful in defining
taxa within different microbial groups. The use of 16S rRNA has been proposed as the
most reliable tool for allocating bacterial strains to families and genera. Moreover, other
molecular methods, such as multi locus sequencing typing, have also been needed for the
assignation of strains to species. However, the use of these molecular methods for routine
analyses and rapid diagnoses, which are usuaUy required for clinical and environmental
analyses involving high numbers of samples or strains, remains impractical. Identification
procedures for routine practices need to be simple, low cost, and rapid in order to be
effective and successful. In this way, conventional biochemical identification provides the
optimal approach for microbial identi fication in lieu of complete genomic
characterization. Evidently, genomic analyses should be carried out for studies of
systematic bacteriology or biodiversity when molecular taxonomical criteria are needed.
The main approach to the identification of an unknown bacterium involves detennination
of its relevant characters and the matching of these with an appropriate database known
taxa. This database may be known as a probability matrix, or identification matrix. The
ideal objective is to assign a name to the unknown that is not only correct but also
predictive of some or all its natural characters. Traditional identification and classification
of microorganisms are usually based on numerical taxonomy that was introduced at the
end of the 1950s. Numerical taxonomic methods were applied extensively for
classification and identification in subsequent years, and they were strengthened by the
extensive use of computers in various research fields. Later, computt:r·assislt:d
identification programs were developed for the identification of bacterial groups, based
mostly on phenotypic data obtained by traditional methods or raw data from commercial
kits. Computerized identification can be achieved in several ways, numerical codes and
probabilistic identification schemes provide more flexible system than those sequential
73
systems (e.g., dichotomous keys). Computerized identification can be achieved in several
ways; numerical codes and probabilistic identification are the most popular approaches.
Bacterial Identification, first described by Bryant (1 986), is a program that provides a
novel way of enhancing the teaching of systematic bacteriology and numerical
identification procedures. A student is assigned an unknown isolate fro l11 a list of bacteria.
The student's objective is to identify the unknown isolate using the least number of tests
from the set of tests available. Simultaneously the computer tries to identify the unknown
using optimized and random selection of tests. A student can compare their progress
against that of the computer. The program has been improved and the data matrix on
which it was based has been revised by the addition of 14 more species, an additional test
and by updating the original probability matrix . The program, Bacterial Identification. is
available as 'freeware' and has been placed with organizations that distribute such
software. A computer program was developed to identify anaerobic bacteria by using
simultaneous pattern recognition via a Bayesian probabilistic model. The system is
intended for use as a rapid. precise, and reproducible aid in the identification of unknown
isolates. The program operates on a data base of 28 genera compri sing 238 species of
anaerobic bacteria that can be separated by the program. Input to the program consists of
biochemical and gas chromatographic test results in binary format . The system is fl exible
and yields outputs of: (i) most probable species, (ii) significant test results conflicting
with established data, and (iii) differential tests of significance for missing test results.
Numerical codes are usually based on positive (+)1 negative (-) reaction. They are applied
to a relatively small set of characters that have been selected for their good diagnostic
value and are applied to clearly defined taxa. Numerical codes require determination of a
series of characters states and the conversion of the binary results into a code number that
is then accessed against the identification database. Such identification systems are
particularly appropriate for the analysis of test results obtained when commercial
identification kits are used.
API 20E kit generates a unique 7 digit number form a battery of 21 tests. The tests are
divided into groups of three and the results are coded 1,2,4 for positive results for tests
in each group. These values are then used to produce a score that reflects the test results,
which can be accessed against the identification system. Organisms that generate profile
number that are not in the identification system can be tested against appropriate
computer assisted probabilistic identification systems. Numerical codes have proved to be
convenient and effective. particularly for well-studied groups.
Probablistic Identification
Probabilistic schemes are designed to assess the likelihood of an unknown strain's
indentifying to a known taxon. Theoretically. the taxa are treated as hyperpheres in an
attribute space (a·space), in which the dimensions are the characters. The center of the
hypersphere (taxon) is defined by the centroid (the most typical representative), and the
critical redius encompasses all the members of each taxon. Ideally. each taxon will be
distinct from any others if the identification matrix has been well constructed. To obtain
an identification, the diagnostic characters for an unknown strain are determined and its
74
position in the a-space calculated. If it falls within the hyperspace (taxon) of a know
taxon, it is identified. Thus in essence, probabilistic identification systems allow for an
acceptable number of "deviant" characters in both the known tax and the unknown strain.
Such task is typically based on the calculation of probabilities via Bayes' theorem. This
approach essentially generates a quantitative value, known as Willcox's identification
scores, which measures the similarity of an unknown isolate with those in the data matrix.
The most computer assisted identification systems are based on Willcox 's implementation
of Bays theorem,
P(t;IR) ~ {P(Rlt;)} / {LP(Rlt;)}
Where, P(tiIR) is the probability that an unknown isolate, giving a pattern of test resul st R,
is a member of taxon (group of bacteria) ti and P(Rlt;) is the probability that the unknown
has a pattern R, given that it is a member of taxon ti. Bays theorem incorporates prior
probabilities; these are expected prevalence of strains included in the identification
matrix. For bacterial, most authors give all taxa an equal chance of being isolated and,
therefore, the prior probabilities for all taxa are set to 1.0 and omitted from the equation.
The above equation therefore can be re-expressed as
Lj* = LJ LLi
Where, the probabilities are now referred to a Identification Scores or Willcox scores.
The identification scores for each taxon are nonnalized value and Lj * for all taxa sums to
one. Identification of an unknown isolate is achieved when Li* for one taxon exceed a
specified threshold value.
The span of characteristics that can be used for bacterial identification includes the
following properties: cultural, morphological, physiological, biochemical, nutritional,
chemotaxonomic, serological, inhibitory tests, genotypic properties, chromatographic
properties and electrophoretic properties. Among these sets, biochemical tests have
remained the typical first step for bacterial identification. Biochemical test measures the
ability of an unknown microorganism in metabolizing substrates (e.g. sugar, amino acids,
etc.). Such catalysis gives rise to detectable color change that is the basis for bacterial
identification. Bacterial identification through the use of biochemical tests can be
performed by either manual or automated approaches. The fanner requires analytical
skills of technologists in reading and interpreting the biochemical results while the latter
makes use of computers for bacterial identification. Several probabilistic identification
as
Bacterial
Identificatifier
programs
have
been
published
such
(http://staff.medschool.soton.ac.ukltnb/pib.htm), BBACTID, BACTIC, CIBAC, Gideon,
IDENTIFY, MICRO-IS, MATIDEN, Identmpm, no-name, The Identifier etc.
Identax is a computer-assisted identification of microorganisms by using only result s
obtained from conventional biochemical tests. Idenlax improves current microbial
identification software and provides a multiplatfonn and user-friendly program. It can be
executed from any operating system and can be downloaded without any cost from the
Identax website (www.identax.org). Identax has two main features. The first is the fast
identification of unknown bacterial strains from phenotypic data , represented as the
75
dichotomous results (positive or negative) ofa set of biochemical tests. The second main
feature is the generation of dichotomous trees that will allow the isolation of one taxon
from the others with the lowest possible number of tests. The first feature is the most
adequate from the point of view of decision support systems, as the software
recommends. in real time, the test with the most discriminative potential. It can also
detect and handle false positives and show the present candidates. The second feature
consists of the generation of a dichotomous tree. Each node represents a test, and its two
branches correspond to a negative or to a positive result from the test. This tree offers an
overview of the search space and allows rapid identification without the need for a
computer.
Identification of Lactobacillus spp. isolates by API method
Materials
• Bacterial culture: Lactobacillus spp. cultures and isolated cultures
• MRS agar plates
• API 50 CHL kit (API CH50, Biomerieux, France).
• Carbohydrate profile of API 50 CHL kit - 0: control, 1: GLYcerol, 2: ERYthritol, 3:
D-ARAbinose, 4: L-ARAbinose, 5: D-RiBose, 6: D-XYL ose, 7: L-XYLose, 8: DADOnitol, 9: Methyl-pD- Xylopyranoside, 10: D-GALactose, II: D-GLU cose, 12 : DFRUctose, 13: D-MaNnosE, 14: L-SorBosE, 15: L-RHAmnose, 16: DULcitol, 17:
INO sitol, 18: D-MANnitol, 19: D-SORbitol, 20: Methyl-nD-Mannopyranoside, 21:
Methyl-aD-Gluco pyranoside, 22: N-Acetylglucosamine, 23: AMYgdalin, 24:
ARButin, 25: ESCulin ferric citrate, 26: SALicin, 27: D-CELIobiose, 28: DMAltose, 29: D-LACtose, 30: D-MELibiose, 31: D-SACcharose, 32: D-TREhalose,
33: INUlin, 34: D-Melezitse, 35: D-RAFfinose, 36: Amidon, 37: GLYcogen, 38:
XYLitol, 39: GEN tiobiose, 40: D-TURanose, 41: D-LYXose, 42: D-TAGatose, 43:
D-FUCose, 44: L-FUCose,45: D-ARabitol, 46: L-ARabitol, 47: potassium glucinate,
48: potassium 2-ketogluconate, 49: potassium 5-keto glucinate.
Protocol
1. Grow the culture on MRS agar plate by streaking overnight at 37' C.
2. Transfer two loopful colonial cultures in 10 ml CHL broth.
3. Mix the content homogenously.
4. Transfer 100 "I of the mix in the tubule of each well with different sugars in API kit.
5. Fill the capalue pfthe each well with sterile mineral.
6. Incubate at 37'C for 24 h in moist API kit chamber.
7. Note the sugar utilization pattern by observing the change of colour of medium.
8. Analysis the sugar utilization data using online API software/API website for
identification of the cultUre.
76
Observations
Results
a) Isolate I:
VI flY. GOOD
11)( NllflCAllON
.
..
.
Slrlp
APl50CH.. VS.1
--
.....
..
.. . .
.
- ~- ---~
-~
- ••• -+ •• •• + + ........ - ++ . + ... +++ + .. + ... _.+.+. - - - .. + - + ••• • +. -
... 1
"'II
T
0.1
T
,
...
I
Meat....
ltctott.cilus pwacMMtl ..., pwtcHei 1
"'II
'
I.It
T_._
J
1~-'1'$:c .,,,,1
b) Isolate II:
..
Vl...HY GOOD ....
IOf ..
N.llFICA
rl{lN
. . . ..
-
.
-
.. .
--
_.... __.. .. ---.---- -
...
--~
API SO CH. VS.1
Strip
_ •• --+ _. --+ + ... ~. - .- .-. --- _ •• ++ + +_. -+ _ •• • _ • • - - _. + --
lIoIa
Loet....._f.
1
1 ____ -
mentum 1
1::.::..-Inferences
"'II
T
9U
1.0
%11
T
U
8.15
T08t..........
1
IL=~
25% 1MLZ '1% 1
~
..
- ----- --_._-_.
Exercise 19 - Examination of ultra-structure of microbial cell using Scanning
Electron Microscopy (SEM)
Introduction
Electron Microscopy (EM) is an important viewing technique for the study of ultrastructures of microorganisms. EM uses a beam of electrons to fonn an image of a
specimen. In contrast to light microscopy which uses visible light as a source of
illumination and optical (glass) lenses to magnify specimens in the range between
approximately IOta 1,000 times their original size, EM is operated in the vacuum and
focuses the electron beam and magnifies images with the help of electromagnetic lenses.
The electron microscope takes advantage of the much shorter wave length of the electron
(e.g. , A = 0.005 nm at an accelerating voltage of 50 kV) when compared to the
wave lengths of visible light (A ::= 400 om to 700 11m). When the acce lerating voltage is
increased in EM, the wavelength decreases and resolution decreases. In other words,
increasing the velocity of electrons results in a shorter wavelength and increased
resolving power.
Scanning Electron Microscopy (SEM) is a powerful method for the investigation of
surface structures of microorganisms. This technique provides a large depth of field,
which means, the area of the sample that can be viewed in focus at the same time is
actually quite large. Furthennore. three-dimensional appearing images may be more
appealing to the human eye than the two-dimensional images. SEM specimen preparation
harbours various risk factors that can easily distort the integrity and ultra-structure. The
basic steps involved in SEM samp le preparation include surface cleaning, stabilizing the
samp le with a fixative, rinsing, dehydrating, drying, mounting the specimen on a metal
holder, and coating the sample with a layer of a material that is electrically conductive.
C leaning of specimen
The proper cleaning of the surface of the samp le is important because the surface can
contain a variety of unwanted deposits, such as dust, si lt, and detritus, media components,
or other contaminants, depending on the source of the biological material and the
experiment that may have been conducted prior to SEM specimen preparation. If these
deposits are not removed prior to fixation, this material may get pennanently fixed to the
specimen surface and it will be almost impossible to remove later. The specimen should
be quickly rinsed in a suitable butTered solution of the appropriate pH, temperature, and
osmotic strength close to the milieu from which the specimen has been removed. Perhaps
the best way to clean the surface of bacteria is to carefully rinse them three times for 10
min in 0.1 M cacodylic acid butTer (PH 7.2±O.2) at room temperature.
Stabilization of specimen
Stabilization is typically done with fixatives. Fixation can be achieved by perfusion and
microinjection, immersions, or with vapours using various fixatives including aldehydes,
osmium tetroxide, tannic acid, or thiocarbohydrazide. Simple chemical fixation by
immersing the specimen in 1.5% glutaraldehyde solu.tion prepared in 0. 1 M cacodylic
78
acid buffer (PH 7.2±0.2) and incubated at 4°C overnight. The use of a post-fixative (e.g.
osmium) has been described to improve hulk conductivity of the specimen.
Rinsing of specimen
After fixation, sample is rinsed to remove excess fixative . Best protocol is to rinse in 0.1
M cacodylic acid buffer (PH 7.2±0.2), starting with one time for 10 min, and then three
times for 20 min at 4°C. Samples can be stored in this EM buffer for several months
because the buffer contains arsenic which inhibits the growth of unwanted
microorganisms. However, it is strongly suggested for changing the cacodylic acid buffer
at least monthly if the samples are to be stored in thi s buffer for longer period.
Dehydrating the specimen
It is performed with either a graded series of acetone or ethanol. Dehydrating includes the
immersion of the specimens in 50% acetone for 5 min , 70% acetone for 10 min, 80%
acetone for 10 min, 90% acetone for 15 min, and 100% acetone (CaCh dried) twice for 20
min at 4°C. It allows water in the samples to be slowly exchanged through liquids with
lower surface tensions.
Drying of specimen
Scanning electron microscope operates with a vacuum. Thus, the specimens must be dry
or the sample will be destroyed in the electron microscope chamber. Critical Point Drying
using carbon dioxide (liquid carbon dioxide as the transitional fluid) which is th e go ld
standard for SEM specimen drying. Carbon dioxide is removed after its transition from
the liquid to the gas phase at the critical point, and the specimen is dried without
structural damage. Alternately, simple air-drying or freeze drying after fixation, rinsing,
and dehydration can be carried out.
Mounting of specimen
After drying, specimen is mounted on a holder that can be inserted into the scanning
electron microscope. Samples are typically mounted on metallic (aluminium) stubs using
a double-sticky tape. A reorientation proves difficult and can result in significant damage
to the sample.
Coating of specimen
Coating increases the conductivity in scanning electron microscope and prevents the
build-up of high voltage charges on the specimen by conducting the charge to ground.
Specimen is coated with a thin layer of approximately 20 nm to 30 Dm of-a conductive
metal (e.g., gold, gold-palladium, or platinum) using sputtercoater apparatus.
Materials
•
Bacterial culture: Lactobacillus spp.
•
MRS broth
•
Refrigerated centrifuge
•
0.1 M cacodylate buffer (pH7.2±0.2)/ phosphate buffered saline (PBS; pH 7.2)
•
2.5% glutaraldehyde in cacodylate buffer
•
Ethanol (30, 50, 70, 90 and 100%)
79
Exercise 20 - Examination of ultra-structure of microbial cell using
Transmission Electron Microscopy (TEM)
Introduction
Transmission Electron Microscopy (TEM) has the advantage over SEM that cellular
structures of the specimen can be viewed at very high magnifications. However, TEM
sample preparation for microorganisms is longer and more difficult than that for SEM and
includes additional steps such as post-fixation, the embedding of sample in a resin, the
sectioning of samples, and the staining of semi-thin and ultrathin sections. Specimen
preparation for TEM includes eight major steps: cleaning, primary fixation, rinsing,
secondary fixation, dehydration, infiltration with a transitional solvent, infiltration with
resin and embedding, and sectioning with stai ning. The first two steps are essentially the
same as those described for SEM specimen preparation.
Cleaning the surface of the specimen
The sample is rinsed carefully three times for 10 min in 0.1 M cacodylic acid buffer (pH
7.2±0.2) at room temperature to clean the surface from contaminants.
Primary fixation of the specimen
Bacteria can be chemically prefixed by immersing the specimens in 1.5% glutaraldehyde
solution in 0.1 M cacodylic acid buffer (PH 7.2±0.2) and incubated at 4°C overnight.
Rinsing of the specimen
Specimens is washed in 0.1 M cacodylic acid buffer (PH 7.2±0.2), starting with one time
for 10 min, and then three times for 20 min at 4°C to remove excess glutaraldehyde from
the samples.
Secondary fixation of the specimen
Fixation can help preserving the structure of the specimen with no alterations from the
living state. Fixation is important for protecting the specimen during steps such as
embedding, sectioning and exposure to the TEM electron beam which operates at higher
accelerating voltages also. Minimizing the risks of artefact induction includes the
selection of the most appropriate fixation protocol for a particular specimen. Bacteria can
be successfully stabilized by postfixation with I % osmium tetroxide in 0.1 M cacodylic
acid buffer (PH 7.2±0.2) for 1.5 h at room temperature (immersion fixation).
Dehydrating the specimen
Bacteria can be dehydrated in a graded series of ethanol by following protocol:
dehydration in 50% ethanol for 5 min, 70% ethanol for 10 min, 80% ethanol for 10 min,
90% ethanol for 15 min, and 100% ethanol twice for 20 min at room temperature. The
process allows the water in the samples to be slowly exchanged through liquids with
lower surface tensions.
Infiltration of specimen with transitional solvent
Replacement of the dehydration solution by another intermediary solvent (i.e., propylene
oxide) is necessary. This process is essentially an alcohol substitution. The immersion in
81
Protocol
1. Grow the Lactobacillus spp. in MRS broth upto late log phase.
2. Harvest the cells by centrifugation (10,000 rpm for 15 min at 4°C).
3. Wash the cells for 2 times using 0.1 M cacodylate butTer (pH7.2±0.2).
4. Fix the cells in 2.5% butTered glutaraldehyde at 4°C for 2 h.
5. Remove the excess glutaraldehyde solution.
6. Fix with 2% osmic acid solution for 2 times at 4-6°C for 2 h.
7. Dehydrate bacteria using series of ethanol (30, 50, 70 and 90%) for one time,
respectively for 5-10 min at 4°C and finally with absolute ethanol/acetone at 4°C for
30-45 min or at room temperature for 1 h.
8. Air-dry the sample - mount the sample on aluminium stub coated with copper strip
and keep in dessicator over night.
9. Coat the dried sample with goldlgold-platinim using ion coater (100-200A) using 6
rnA ion current at fine vacuum of 0.05-0.07 toor for 2-4 min.
10. Load the stub with specimen on specimen holder and insert into SEM and observe
through SEM under vacuum at 15 kV.
Observations
Results
Inferences
80
propylene oxide twice for 20 min at room temperature is sufficient before attempting to
embed the specimens in a resin.
Inflltration witb resin and embedding of specimen
Bacteria can be embedded in a variety of different media depending on the use (e.g.,
conventional TEM or immuno TEM). For conventional TEM, the epoxy resin Durcupan
ACM is quite suitable. The following protocol can be used: Immersion in propyleneoxideIDurcupan-ACM (1:1, v/v) at room temperature overnight (use gloves and a fume
hood, and leave the specimen container open for the propylene oxide to evaporate). The
next day, the specimen is immersed in a freshly prepared Durcupan ACM mixture (pure)
and left for 2 h at room temperature. A second Durcupan ACM mixture (pure) is then
prepared and used as the embedding medium (free of air bubbles). Polymerization of the
epoxy mixture can be achieved by placing the specimens in a drying cabinet for 2 days at
40°C and for an additional 2 days at 60°C. Leaving the samples after heat polymerization
for an additional 1-2 weeks at room temperature can improve the subsequent cutting
experience as the resin blocks continue to harden during the time.
Sectioning and staining of the specimen
The procedure for cuning specimens into semi-thin and ultrathin slices (sections) is
known as microtomy and ultramicrotomy, respectively. Semithin sections (0.5 ~m to 2
~m) were typically stained with toluidine blue for I min on a hot plate (70-90°C),
examined by light microscopy, and used for identifying the specimen within the resin
block before proceeding with ultra-microtomy. Ultrathin sections (about 70 - 90 om) were
typically stained with uranyl acetate followed by lead citrate and observed through TEM .
A) Rapid Method TEM
Materials
•
Culture: Staphylococcus aureus
•
Culture concentration: 10
solution at I: 14-15
•
2% glutaraldehyde in 0.1 M phosphate buffer saline (PBS)
•
2% uranyl acetate
•
Osmium tetraoxide
•
Acetone (50%, 70%, 90% and 100%)
•
Epoxy resin
7
-
10 8 bacteria per test in a ratio of bacteria and fixative
• Reynold's stain
Protocol
I. Harvest the bacterial cells by centrifugation and wash twice with phosphate buffer
saline (PBS).
2. Fix the bacteria for 20 minutes in 2% glutaraldehyde (GA) in 0.1 M PBS and wash 6
times with distilled water.
3. Stain with 2% uranyl acetate (UA) for 5 minutes and wash the bacteria 3 times with
distilled water.
82
4, Expose the stained bacteria to osmium tetraoxide (OT) for 5 minutes and discard the
excess OT.
S. Dehydrate the stained bacteria using the series of acetone (50%, 70%, 90% and
100%) for one time, respectively for 5 minutes.
6, Polymerize with pure epoxy resin in embedding oven at 75°C for 2 h and 90°C for 2
h, after the bacteria infiltrated by mixture of acetone and epoxy resin (1: I) for IS
mm.
7, Trim the blocks and cut to 80-90 run ultra thin sections and mount on 200 mesh thin
bar copper grids,
8. Stain the specimens with Reynold's stain for 1-2 min.
9. Examine the specimen at 39000x magnification by using rEM at an accelerating
voltage of90 KY,
B) General Method TEM
Materials
•
0, I M Cacodylate buffer
•
2% GA in 0, 1M Cacodylate buffer
•
2% OT in 0, I M Cacodylate buffer
•
Ethanol (50, 70, 90 and 100%)
•
Epoxy resin
•
Oven (60°C)
•
Reynold's stain
•
•
2% uranyl acetate
Ultra-microtome
Protocol
I, Harvest the bacterial cells by centrifugation and wash twice with cacodylate buffer,
2, Fix the bacteria for overnight in 2% GA solution in 0, I M Cacodylate buffer,
3, Wash the fixed bacteria with O,IM Cacodylate buffer for 3 times each of20 min,
4, Expose the fixed bacteria to 2% OT solution in 0, I M Cacodylate buffer for I hand
wash the bacteria with 0, I M Cacodylate buffer for 3 times each of20 min,
5, Dehydrate the OT exposed bacteria using the series of ethanol (50,70, 90 and 100%)
for one time, each 20 ·min respectively.
6, Polymerize with pure epoxy resin in the embedding oven ·at 60°C for 18 h after the
bacteria infiltrated by mixture of ethanol and epoxy resin (I : I) for I h,
7, Trim the blocks and cut to 80-90 nm ultra thin sections and mounted on 200 mesh thin
bar copper grids,
8. Stain the specimens with UA and Reynold' s stain at 10 minutes, respectively.
9, Examine the specimen at 50,OOOx magnification in TEM at an accelerating voltage of
90KY,
83
Observations
1
I
!
Results
Inferences
84
Viva voice
I)
2)
3)
4)
S)
6)
7)
S)
9)
10)
II)
12)
13)
14)
15)
16)
17)
18)
19)
20)
21)
22)
23)
24)
25)
<For spore staining weakly positive dye is used' - why?
What are "Acid-fast organisms"?
Define endospore; vegetative form; germination of spores.
Define Microscopic Factor.
Describe the characteristics of bacterial spores.
Differential staining technique - differentiating bacteria types by observing the
amount of stain they absorb.
Draw the diagram of Ocular and Stage micrometer, the scaling details and
positions/ locations in a compound microscope.
Free spores resist ordinary dyes such as methylene blue, crystal violet and
carbolfuchsin why?
Gram stain is used to differentiate types of bacteria depending on their abilities to
retain a particular stain-explain.
Gram-negative organisms will be decolorized by the alcohol and are subsequently
stained by the safranin and appear red or pink - why?
Gram·positive organisms will not be easily decolorized and thu s retain the purple
stain of crystal violet - why?
In the negative stain the unstained halo· like material surrounding the cells would
represent the capsule surrounded by a dark background - why?
Name two gram positive and two gram negative bacteria present in milk.
Some species of bacteria do not stain readily by simple stain or Gram stain
procedures (such as the Mycobacterium) - why?
What genera of bacteria fonn spores?
What is a differential stain?
Why do we stain microorganisms before viewing them with a microscope?
Why is immersion oil used to view microscopic organisms?
Why do we stain microorganisms before viewing them with a microscope?
Name some basic and acidic stains.
Define simple staining.
What makes a bacterium acid fast?
Why is heat necessary for staining of acid fast bacteria and endospores?
What is the purpose ofheat fixing before staining?
What are the advantages of negative staining?
85
References
1. Yadav 1.S., Grover, S., Batish V.K. 1993. A Comprehensive Dairy Microbiology.
METROPOLITAN, New Delhi - 110002, India.
2. HiMedia 2003. The HiMedia Manual for Microbiology and Cell Culture Laboratory
Practice, HiMedia Laboratories Pvt. Ltd., Mumbai.
3. Standard Methods for Examination of Water and Wastewater, 15the Edition,
American Public Health Association, New York, 1995.
4. Mandai, S. and Puniya, A.K. 2009. Practical Manual on Environmental Microbiology.
National Dairy Research Institute, Kamal, Haryana, India.
5. Puniya, A.K., Mandai, S. and Tomar, S.K. 2009. Practical Manual on Fundamental of
Microbiology. National Dairy Research Institute, Kamal, Haryana, India.
6. Manual of Microbiological Methods by The Society of American Bacteriologists,
McGRAW-HILL BOOK COMPANY, INC, 1957.
7. http://student.ccbcmd.edulcourseslbio 141 Ilecguide/unit I Ishape/u 1coccus.html
8. www.bmb.leeds.ac.uklmbiology/uglugteachlnewde ...
86
>
,
•
~ G5460
I
1111 11111 1111111111 1111 1111
Call No.576.8(02) 02