1. The document discusses different types of culture media used to grow microorganisms in the laboratory, including solid media, semisolid media, and liquid broths. 2. Solid media like agar plates are used for isolating pure cultures, observing colony morphology, and storing cultures long-term. Semisolid media allow studying bacterial motility. 3. Broths are used for growing cultures for assays and biochemical tests since bacterial growth is visible through turbidity. 4. Agar is commonly used to solidify media due to its properties of being bacteriologically inert and allowing surface growth of colonies.

1. Chapter 4
TYPES OF MICROORGANISMS MEDIA
Dr. Yousef Elshrek

2. • The study of microorganisms requires techniques for
isolating cells from natural sources and growing
them in the laboratory on synthetic media.
• Thus, developments of synthetic culture media and
culture techniques have played important roles in the
advancement of this field.
• Microbiologists use bacterial culture media for many
purposes and applications.
• Media are used to
1. Isolate and identify bacteria
2. reveal their metabolic properties
3. Allow long-term storage of pure cultures.

3. • Taxonomic descriptions of bacteria commonly
include
1. Information about their cultural requirements.
2. Species that are poorly characterized are
frequently those most difficult to culture under
laboratory conditions.
• Indeed, Koch’s second postulate requires
culturing of a suspected pathogen in pure form.
• In the slide you will learn about composition
and types of culture media and how different
types of media can be used to study the
properties of bacteria.

4. • MEDIA
• General and specialized media are required for
bacterial growth and for characterization.
• The media are, in fact, research tools.
• The basic procedures can be applied to almost
any type of assay or culture requirement for
propagation of obligate aerobes or faculatative
anaerobes.
• Obligate anaerobes are poisoned by oxygen,
and specialized procedures are needed for their
maintenance.

5. • BACTERIA NUTRITIONAL REQUIREMENTS
• The ability to study different types of bacteria ultimately
relies upon knowing their nutritional requirements.
• The bacteria with which they are most familiar are
generalists (which are able to use a wide range of nutrients)
and/or nutrients that are commonly available.
• Some bacteria can synthesize all of their growth
requirements from common mineral nutrients and simple
carbohydrates.
• However, some bacteria are classified as auxotrophs (a
mutant strain of microorganism having nutritional
requirements additional to those of the normal organism)
because, even given a carbohydrate carbon source, they
cannot synthesis one or more organic molecules required
for their growth – these molecules must be also provided in
growth media.

6. • However, if a sample swabbed from mouth were
inoculated on a plate of common culture medium, only a
small percentage of the hundreds of different bacteria will
grow and form colonies.
• This is because most bacteria are fastidious, meaning that
they have very specific and/or complex nutritional
requirements.
• These species do not grow because they cannot use one or
more nutrients in the form provided in the medium (e.g.,
they might require H2S rather than SO4 as a sulfur source),
have requirements for very specific types of nutrients
(such as certain complex organic molecules), and/or
require unusual growth conditions (such as growth in
living cells or at high temperature or pressure).

7. • Presently know very little about many of these
bacteria because nobody knows how to grow
them under artificial laboratory conditions.
• One factor that greatly influences bacterial
growth is their oxygen requirements.
• Clearly, the techniques used to culture and
study an obligate an aerobe must be different
from those used when culturing an aerobe.

8. • TYPES OF BACTERIAL GROWTH MEDIA
1. Nutrient Agar(NA)
2. Nutrient Broth(NB)
3. Trypic Soy Broth(TSB)
4. Tryptic Soy Agar(TSA)
• MEDIA REQUIREMENTS
• Bacteria display a wide range of nutritional and
physical requirements for growth including
1. Water
2. A source of energy
3. Sources of carbon, nitrogen, sulfur, phosphorus
4. Minerals, e.g., Ca2+, Mg2+, Na+.
5. Vitamins and growth factors.

9. • Microorganisms may be grown in liquid, solid or
semisolid media.
• Liquid media are utilized for growth of large numbers of
organisms or for physiological or biochemical studies and
assays.
• Some species, such as Streptococcus or Staphylococcus,
often demonstrate typical morphologies only when grown
in liquid media.
• Solid media are useful for observations of characteristic
colonies, for isolation of pure cultures and for short-term
maintenance of cultures.
• Usually, the preparation of a solid medium for growth
simply includes the addition of 1 to 2% agar to a solution
of appropriate nutrients.
• Agar is a complex carbohydrate extracted from marine
algae that solidifies below temperatures of 45C. It is not a
nutritional component.

10. • Usually, bacteria are grown in complex media, because
we simply do not know enough about the organism or
organisms to define all of their requirements for growth
and maintenance.
• Neither the chemical composition nor the
concentrations of substrates are defined.
• Media frequently contain nutrients in the form of
extracts or enzymatic digests of meat, milk, plants or
yeast.
• For fastidious organisms we must often use delicious-
sounding concoctions such as tomato juice agar or
chocolate agar, or something less appetizing (but
nutrient-rich) such as brain-heart infusion broth or
blood agar.

11. • There is no single medium or set of physical
conditions that permits the cultivation of all
bacteria, and many species are quite fastidious,
requiring specific ranges of
1. pH
2. osmotic strength
3. temperature
4. presence or absence of oxygen.
• The requirements for growth of bacteria under
laboratory conditions are determined by trial
and error.

12. • Using a rich, complex of culture bacteria medium, such
as tryptic soy agar or broth, so that a wide variety of
possible unknowns can be mixed into the same culture
and grown on the same plates.
• Agar plates will be used for isolation and some assays,
and for short term maintenance of cultures.
• Agar slant tubes will be used for long term maintenance
of isolates.
• Broths (liquid media) will be used to grow isolates for
some assays or for the assays themselves.
• Therefore , their types of media as following:
1. Solid.
2. Semisolid
3. Broth

13. • SOLID MEDIA
• Solid media are more versatile (adaptable) in their usage.
1. Promote surface growth.
2. Used to isolate pure cultures.
3. Ideal for culture storage.
4. Helpful in the observation of biochemical reactions.
5. Used to make slants, deeps, and plates (named by medium).
6. Bacteria may be identified by studying the colony character.
7. Mixed bacteria can be separated.
8. Solid media is used for the isolation of bacteria as pure
culture.
9. Agar is most commonly used to prepare solid media.
• This type of media is prepared by adding a solidifying agent
(agar 1.5 -3%).
• Prepared mainly in Petri dishes, but also in tubes and slopes.
• After growth the bacterial colonies are visible. e.g. blood agar,
chocolate agar, MacConkey agar.:

14. • Agar is polysaccharide extract obtained from seaweed.
• Agar is an ideal solidifying agent as it is :
1. Bacteriologically inert, i.e. no influence on bacterial growth..
2. It is transparent
3. Somewhat like gelatin.
4. It melts at 970C and solidifies at 370C.
5. Comes as sold powder and then adding water to it.
6. Colony morphology, pigmentation, hemolysis can be
appreciated.

15. • SEMISOLID AGAR
• Contains small amounts
of agar (0.5-0.7%).
• Used to check for motility
and also used as a
transport media for fragile
organisms.
• Can have semisolid agar
in Petri dishes or in tubes.
In tubes it is usually
slanted to increase surface
area, e.g. SIM

16. • LIQUID (BROTH) NO AGAR
• Mostly used for biochemical tests (blood
culture, Broth culture).
• Growth of bacteria is shown by turbidity
in medium. e.g. Nutrient broth, Selenite F
broth(A medium for the selective
enrichment of Salmonella spp from both
clinical and food samples. It is a buffered
Lactose Peptone Broth to which Sodium
Biselenite is added as the selective
agent. Subcultures should be made from
the top 1/3 of the broth after not more
than 24 hours incubation as after this
time there is a loss of selectivity),
alkaline peptone water.
• Used for inoculum preparation , blood
culture, for the isolation of pathogens
from mixture

17. • PROPERTIES OF AGAR
• Simple (basal, ordinary): Culture Media: are
media that contain the basic nutrients (growth
factors) that support the growth of bacteria without
special nutrients, and they are used as basis of
enriched media. e.g. Nutrient broth, nutrient agar,
peptone water. They are for the growth of non-
fastidious organisms like E. coli.
• Enriched Culture Media: are media that are
enriched with: Whole blood e.g. blood agar. Lysed
blood (heated to 80C) e.g. Chocolate agar
• Selective Media: it is a media, which contains
substances that prevent or slow the growth of
microorganisms other than the bacteria for which
the media is prepared for.

18. • Differential Media (indicators): Contains indicators, dyes,
etc, to differentiate microorganisms. e.g. MacConkey agar,
which contains neutral red (pH indicator) and is used to
differentiate lactose fermenter and non-lactose fermenter. (e.g.
E. coli and Salmonella).
• Chocolate Agar:
• (Non selective media) blood agar prepared by heating blood
to 95C until medium becomes brown or chocolate in color
heating the blood releases broth X and V growth factors and
also destroys the inhibitors of V factor (Haemophilus
influenzae requires two accessory growth factors: factor X
(hemin) and factor V (NAD, nicotinamide adenine
dinucleotide). The X and V factor requirement is usually
demonstrated by the absence of growth on porphyrin and
NAD deficient but otherwise nutritionally adequate media
except near paper disc impregnated with X and V factors. ).
• These factors are required for the growth of most species of
Haemophilus and also Neisseria gonorrhoear.

19. • Chocolate agar with the addition of bacitracin
becomes selective, most critically, for the
genus Haemophilus. Another variant of
chocolate agar called Thayer-Martin agar
contains an assortment of antibiotics which
select for Neisseria species.

20. • Mueller Hinton Agar:
• rich medium that
support the growth of
most microorganisms.
• It is commonly used for
antibiotic susceptibility
testing: disk diffusion
antibiotic susceptibility;
antibiotic serum level
measurements; MBC
determination.
Mueller Hinton Agar

21. • Salmonella Shigella (SS) Agar: isolation and
differential medium for pathogenic Gram
negative bacilli in particular, Salmonella and
Shigella. Inhibitor for Coliforms.
Salmonella Shigella (SS) Agar

22. • Triple Sugar Iron Agar (TSI):
this a key medium for use in beginning the
identification of a Gram- negative bacilli of
the enteric group. It contains
1. Glucose (0.1% )
2. Lactose (1%)
3. Sucrose (1%).
4. Peptone (2%) as nutritional sources.
5. Sodium thiosulfate serves as the electron
receptor for reduction of sulfur and
production of h2s.
6. Detects fermentation of sucrose, lactose,
glucose, as well as production of hydrogen
sulfide and /or gas.
7. Phenol red is the pH indicator; ferric
ammonium citrate is H2S indicator.
Triple Sugar Iron Agar (TSI)

23. • TYPES OF CULTURE MEDIA
• MacConkey agar (Selective and differential
media)
• MacConkey Agar: an inhibitory and differential
medium used to distinguish lactose fermenting
Gram- negative organism from non fermentation.
• Crystal violet, bile salts and neutral red are
inhibitor agent.
• Neutral red is the PH indicator.

25. • Streak a plate of MacConkey's agar with
the desired pure culture or mixed culture.
• If using a mixed culture use a streak plate
or spread plate to achieve colony isolation.
• Good colony separation will ensure the
best differentiation of lactose fermenting
and non-fermenting colonies of bacteria.
• Streak plate of Growth of Escherichia coli
and Serratia marcescens on MacConkey
agar.

26. • Growth of E. coli, which
ferments lactose, appears
red pink on the agar.
Growth of S. marcescsens,
which does not ferment
lactose, appears colorless
and translucent.
• Both microorganisms grow
on this selective media
because they are gram-
negative non-fastidious
rods

27. • MACCONKEYAS SELECTIVE
•MACCONKEYAS DIFFERENTIATE
•Lactose positive
•Lactose negative

28. • MANNITOL SALT AGAR (MSA)
• Mannitol salt agar (MSA) is both a selective and
differential medium used in the isolation of
staphylococci.
• It contains 7.5% sodium chloride and thus selects
for those bacteria which can tolerate high salt
concentrations.
• MSA also distinguishes bacteria based on the
ability to ferment the sugar mannitol, the only
carbohydrate in the medium.
• Staphylococci can withstand the osmotic pressure
created by 7.5% NaCl, while this concentration
will inhibit the growth of most other gram-
positive and gram-negative bacteria.

29. • Additionally, MSA contains mannitol
and uses phenol red as a pH indicator
(pK = 7.8) in the medium.
• At pH levels as following:
1. Below 6.9, the medium is a yellow
color.
2. In the neutral pH ranges (6.9 to 8.4)
the color is red.
3. Above pH 8.4, the color of phenol
red is pink.
A) Staphlococcus aureus: large yellow halo around growth indicates
fermentation of mannitol.
B) Staphlococcus epidermidis: Growth but not color change to the
media indicating no fermentation of mannitol.
C) Staphlococcus saprophyticus: small yellow halo around growth
indicates fermentation of mannitol. (10% of S. saprophyticus
ferment mannitol
D) E. coli: no growth. Inhibited by the 7.5% NaCl.

30. 4. When mannitol is fermented by a bacterium,
acid is produced, which lowers the pH and
results in the formation of a yellow area
surrounding an isolated colony on MSA.
5. A nonfermenting bacterium that withstands the
high salt concentration would display a red to
pink area due to peptone breakdown .
6. In clinical samples, mannitol positive isolates are
suggestive of Staphylococcus aureus and should
be tested further.

31. • PROTOCOL
• Streak a plate of mannitol salt agar with appropriate
culture using the quadrant streak plate method to
obtain isolated colonies.
• Well-isolated colonies will provide the best results in
the biochemical differentiation bacteria using MSA.
• (A)Staphylococcus aureus
•(B) Staphylococcus epidermidis, and
•(C) Escherichia coli streaked on a mannitol
salt agar plate.
•The mannitol fermenting colony (yellow) is
S. aureus.
•The mannitol nonfermenting colony (pink) is S. epidermidis.
•The growth of E. coli was inhibited by the high salt concentration.

32. • MANNITOL SALT AGAR (MSA)
SELECTIVE MEDIA
• Selective for staphylococci

33. MANNITOL SALT AGAR (MSA) DIFFRENTIATE MEDIA
Differentiate between pathogenic and non pathogenic
staphylococci

34. • SOME NOTES
• Media color change demonstrates mannitol
fermentation, not colony color.
• This is particularly important as many
micrococci are pigmented.
• Inoculated plates that are kept refrigerated may
exhibit color loss over time.
• Some instructors have found that reincubation
may bring back some color.
• Others have indicated that pouring plates
thicker lessens the color loss.

35. • PHENYL ETHYL ALCOHOL AGAR
• SELECTIVE MEDIA
• Brewer and Lilley developed a selective medium
containing phenyl ethyl alcohol (PEA)
1. Allowed growth of gram-positive organisms,
particularly cocci, while inhibiting most gram-
negative bacteria and fungi.
2. PEA(benzylcarbinol,C6H5CH2CH2OH,CH2CH2OH)
Found in volatile oils of many flowers such as
roses, is colorless and soluble in water.

36. 3. With a boiling point between 219 and 222oC, it
can be synthesized by the reduction of ethyl
phenyl acetate with sodium in absolute alcohol.
4. Brewer observed a selective phenomenon when
5 ml of 1:30 PEA in acetone was placed on a pad
on a petri dish lid covering an agar inoculated
with various microorganisms.
5. He found that under this situation, gram-negative
organisms (Pseudomonas aeruginosa and
Proteus) did not overgrow while the gram-
positive organism (Staphylococcus aureus) grew
abundantly.
6. Lilley and Brewer researched the optimum
concentration of PEA for differential inhibition
and found it to be 0.25%

37. 7. In this study, PEA was also found to show
fungistatic activity.
8. When 16 molds and yeasts were tested, only
Candida albicans grew on Sabouraud’s
medium containing 0.25% PEA.
9. Isolation of anaerobic gram-positive cocci
from marine animals could be done by using
a modified glucose blood liver agar with 0.3%
PEA .
10. Organisms grown on basic media containing
PEA do not change genetically.

38. 11. Organisms grown on media with PEA show normal
growth characteristics when subcultured on a medium
without PEA.
12. PEA agar with 5% sheep blood is used in microbiology
laboratories to inhibit gram-negative bacteria,
specifically Proteus species, in specimens containing a
mixed bacterial flora.
13. Five percent sheep blood is added to the base medium
to enhance the growth of anaerobic bacteria.
14. Most gram-positive and gram-negative anaerobes grow
on PEA agar medium, especially in mixed culture, and
morphology of colonies is similar to that on blood agar
plates, however, a longer incubation time is necessary
to detect the more slowly growing and pigmented
anaerobes.

39. • PURPOSE
1. PEA agar is a selective medium that is used for
the isolation of gram-positive Staphylococcus
species and Streptococcus species from clinical
specimens or specimens that contain mixtures
of bacterial flora.
2. Typically PEA agar is used to inhibit the
common contaminants such as Escherichia coli
and Proteus species.
3. PEA agar may be prepared with and without
5% sheep blood supplement.

40. 4. PEA agar with 5% sheep blood is used to
isolate most gram-positive and gram-
negative anaerobes from enteric samples. It
is used to inhibit facultative gram-negative
rods, preventing Enterobacteriaceae from
overgrowing the anaerobes and inhibiting
swarming of Proteus and Clostridium
septicum
5. PEA agar is used for purulent specimens and
when mixed infections are suspected.

41. Gram
reaction
Organism
Growth
response
Swarming
inhibition
Gram negative Escherichia coli Inhibited N/A
Gram negative Proteus mirabilis Markedly inhibited
Yes,no
spreading
Gram negative Pseudomonas aeruginosa Partially inhibited N/A
Gram negative Salmonella enteritidis Inhibited N/A
Gram negative Enterobacter aerogenes Inhibited N/A
Gram positive Staphylococcus aureus Good N/A
Gram positive Streptococcus pyogene Good N/A
Gram positive Streptococcus pnuemoniae Good N/A
Gram positive Clostridium perfringens Partially inhibited N/A
Gram positive Enterococcus faecalis Good N/A
Gram positive Bacillus sp. Good N/A
Gram positive Micrococcus luteus Good N/A
Table 1. Examples of growth of some gram-negative
and gram-positive bacteria on PEA agar

42. • THEORY
• PEA agar is a selective medium that permits the growth of
gram-positive cocci while inhibiting most gram-negative
organisms.
• PEA acts on gram-negative bacteria by altering their
membrane permeability, allowing influx of otherwise blocked
molecules, and allowing leakage of large amounts of cellular
potassium that ultimately results in disruption or inhibition of
DNA synthesis.
Pancreatic digest of casein 15 g
Papaic digest of soybean meal 5.0 g
Sodium chloride 5.0 g
β-Phenylethyl alcohol 2.5 g
Agar 15 g
Distilled water 1.0 liter
Table 2 PEA agar typical composition (g/liter)

43. • MEDIA PREPARATIONS
1. Suspend the first five ingredients in 1 liter of
distilled water.
2. Mix thoroughly.
3. Heat with frequent agitation and boil for 1
minute to dissolve completely.
4. Autoclave the medium at 121oC for 15
minutes at 15 psi.
5. Final pH of the medium should be 7.3 + 0.2
at 25oC.

44. 6. After sterilization, pour the melted medium
into sterilized petri plates (approximately 20
to 30 ml per plate) and let it solidify before
use.
7. Prepared medium is clear to slightly hazy
and pale yellow.
8. Prepared plates can be stored in the
refrigerator for up to 4 weeks before use.
9. Allow the medium to come to room
temperature before inoculation.

45. • PEAAGAR WITH 5% SHEEP BLOOD
TYPICAL COMPOSITION (G/LITER)
Pancreatic digest of casein 15 g
Papaic digest of soybean meal 5 g
Sodium chloride 5 g
β-Phenylethyl alcohol 2.5 g
Sterile defibrinated sheep blood 50 ml
Agar 15 g
Distilled water 1.0 liter
Table 3 PEA agar typical composition
(g/liter) with 5% sheep blood

46. 1. Suspend all ingredients except sheep blood in 1
liter of distilled water and mix thoroughly.
2. Heat with frequent agitation and boil for 1 minute
to dissolve completely.
3. Autoclave the medium at 121oC for 15 minutes at
15 psi. Final pH of the medium should be 7.3 ± 0.2
at 25oC.
4. Cool to 45oC and add 5% sterile defibrinated blood
and mix well.
5. Quickly pour the melted medium into sterilized
petri plates (approximately 20 to 30 ml per plate)
and let it solidify before use.
6. Prepared medium appears firm, opaque, and red in
color.

47. 1. Prepared plates could be stored in the
refrigerator up to 1 week before use.
2. Allow the medium to come to room
temperature before inoculation.
• PEA agar medium is also commercially
available as premixed powder from biological
supply companies.
• The manufacturer’s instructions should be
followed to prepare the plates.
• This media can also be purchased as premade
agar plates from biological supply companies.

48. • PROTOCOL
1. Inoculation. Aseptically transfer potentially
mixed cultures onto the surface of the agar using
a four-way streaking technique.
2. Depending on the objectives of the study, either a
confluent growth or a four-way streaking
technique can be used.
3. Incubation. Incubate plates for 24 to 48 hours at
35oC ± 2oC in an appropriate atmosphere.

49. 1. In some cases, a longer incubation, up to 1 week,
may be needed.
2. PEA blood agar plates can be incubated under
aerobic, anaerobic, and 5% CO2 atmosphere based
on the type of microorganisms being studied.
3. Incubation in high CO2 atmosphere allows the
detection of bacteria which require an increased CO2
concentration and also results in better growth of
almost all of the other pathogens.
• INTERPRETATION OF RESULTS.
• After proper incubation, growth of isolated colonies or a
group of colonies may be observed.
• Gram-positive bacteria demonstrate good growth (Fig. C1 and
C2) while most gram-negative bacteria do not grow or are
partially inhibited (Fig. B1 and B2).

50. PEA agar plates with 5% sheep blood:
1. an uninoculated PEA agar plate with 5% sheep blood
2. a PEA agar plate with 5% sheep blood inoculated with Escherichia coli, a gram-negative
bacteria, incubated under 5% CO2 for 48 hr at 35oC ± 2oC (growth inhibited),
3. a PEA agar plate with 5% sheep blood inoculated with Staphylococcus aureus, a gram-
positive bacteria, incubated under 5% CO2 for 48 hr at 35oC ± 2oC (growth exhibite
PEA agar plates:
1. (A2) an uninoculated PEA agar plate
2. (B2) a PEA agar plate inoculated with Escherichia coli, a gram-negative bacteria, incubated
under aerobic conditions for 48 hr at 35oC ± 2oC (growth inhibited)
3. (C2) a PEA agar plate inoculated with Enterococcus faecalis, a gram-positive bacterium,
incubated under aerobic conditions for 48 hr at 35oC ± 2oC (growth exhibited).

51. • NOTICE
1. PEA agar with 5% sheep blood should not be
used for determination of hemolytic reactions as
irregular patterns may be observed. Organisms
should be subcultured onto tryptic soy agar with
5% sheep blood to examine hemolysis.
2. Some gram-positive cocci may be slightly
inhibited by PEA and many require incubation
up to 48 hours for sufficient growth to be visible
(2).
3. Due to nutritional variation, some strains may
be encountered that grow poorly or fail to grow
on PEA agar medium.

52. 1. Pesudomonas aeruginosa (a gram-negative
bacteria) is not inhibited on this medium.
2. In order to control for the viability of the
organisms used, a control nutrient agar or
other nonselective medium should be used in
parallel.
3. It is important to remember that this medium
inhibits the growth of gram-negative
bacteria. Tiny observable colonies on PEA
agar may be gram-negative microorganisms
and are often confined to first quadrant on a
streak plate.

53. • EOSIN-METHYLENE BLUE (EMB) AGAR
• Eosin-methylene blue (EMB) agar was first developed by
Holt-Harris and Teague in 1916.
• They used EMB agar to clearly differentiate between the
colonies of lactose fermenting and nonfermenting
microbes.
• In the same medium, sucrose was also included to
differentiate between coliforms that were able to ferment
sucrose more rapidly than lactose and those that were
unable to ferment sucrose.
• Lactose fermenter colonies were either black or
possessed dark centers with transparent and colorless
outer margins, while lactose or sucrose nonfermenters
were colorless.

54. • EMB agar was shown to be more sensitive
and stable and differentiated between sugar
fermenters and nonfermenters faster when
compared to other agars.
• In 1918, Levine described an EMB agar that
differentiated between fecal and nonfecal
types of the coli aerogenes group.
• It also differentiated between salmonellae
and other nonlactose fermenters from the
coliforms.

55. • Present day Bacto EMB agar is a combination of
the EMB agar described by Holt-Harris and
Teague and Levine.
• It contains lactose and sucrose (as described by
Holt-Harris and Teague) and also contains Bacto
peptone and phosphate (as described by Levine).
• The two indicator dyes, eosin and methylene blue,
are used in a ratio to impart minimum toxicity but
provide best differentiation.

56. PURPOSE
• Eosin-methylene blue agar is selective for gram-
negative bacteria against gram-positive bacteria.
• In addition, EMB agar is useful in isolation and
differentiation of the various gram-negative
bacilli and enteric bacilli, generally known as
coliforms and fecal coliforms respectively.
• The bacteria which ferment lactose in the
medium form colored colonies, while those that
do not ferment lactose appear as colorless
colonies.

57. • EMB agar is used in water quality tests to
distinguish coliforms and fecal coliforms that
signal possible pathogenic microorganism
contamination in water samples.
• EMB agar is also used to differentiate the
organisms in the colon-typhoid-dysentery
group: Escherichia coli colonies grow with a
metallic sheen with a dark center, Aerobacter
aerogenes colonies have a brown center, and
nonlactose-fermenting gram-negative bacteria
appear pink.

58. • Theory EMB agar contains peptone, lactose,
sucrose, and the dyes eosin Y and
methylene blue; it is commonly used as
both a selective and a differential medium.
• EMB agar is selective for gram-negative
bacteria.
• The dye methylene blue in the medium
inhibits the growth of gram-positive
bacteria; small amounts of this dye
effectively inhibit the growth of most gram-
positive bacteria.

59. • Eosin is a dye that responds to changes in pH,
going from colorless to black under acidic
conditions.
• EMB agar medium contains lactose and
sucrose, but not glucose, as energy sources.
• The sugars found in the medium are
fermentable substrates which encourage
growth of some gram-negative bacteria,
especially fecal and nonfecal coliforms.

60. • Differentiation of enteric bacteria is possible due to
the presence of the sugars lactose and sucrose in the
EMB agar and the ability of certain bacteria to
ferment lactose in the medium.
• Lactose-fermenting gram-negative bacteria (generally
enteric) acidify the medium, and under acidic
conditions the dyes produce a dark purple complex
which is usually associated with a green metallic
sheen.
• This metallic green sheen is an indicator of vigorous
lactose and/or sucrose fermentation ability typical of
fecal coliforms.
• A smaller amount of acid production, which is a result
of slow fermentation (by slow lactose-fermenting
organisms), gives a brown-pink coloration of
growth.

61. • Colonies of nonlactose fermenters appear as
translucent or pink.
RECIPE as described in the Difco manual
dyes eosin Y
Substances Quantity in grams
peptone 10
lactose 05
sucrose 05
dipotassium phosphate 02
agar 13.5
eosin Y 0.4
methylene blue 0.065
Distilled water t o bring final volume to 1 liter

62. • Adjust pH to 7.2.
• Boil to completely dissolve agar.
• Sterilize in an autoclave for 15 minutes at 15 psi
(121C).
• Cool to 60C.
• If any precipitate is apparent in the medium,
disperse by gently swirling before pouring into
sterile Petri dishes (1).
• EMB agar is commercially available in premixed
form from biological supply companies.
PROTOCOL
• Obtain an EMB agar plate and streak it with the
appropriate bacterial culture using the quadrant
streak plate method.
• This will result in the isolation of individual

64. • COMMENTS
• The concentration of agar may be increased to 5%
(by using an additional 3.65 g of agar per 1 liter of
medium, refer to the recipe section) to inhibit the
spreading of Proteus .
• If the sucrose-containing EMB medium is
used, Proteus colonies will also show the
characteristic metallic sheen if they are inhibited
from spreading by the higher concentration of agar .
• Besides being used as a fermentation indicator
medium to differentiate gram-negative enteric
bacteria, EMB can also be used for testing strains of
bacteria for sensitivity to phage.

65. • In this case, 5 g of NaCl per liter is to be added into
the medium, and the medium is to be made without
added sugars to a final concentration of 1% as in
the typical EMB.
• This medium is then designated as EMBO agar .
• SUMMARY
• EMB can be used as selective media for gram
negative bacilli / rods and as differential to
differentiate between groups of enteric bacteria.
• Differentiate between groups of enteric bacteria
• The term coliforms is used in the USA to indicate
the presence of enteric bacteria in water , foods and
other samples.

67. 5. The addition of sucrose permitted the
earlier detection of coliform bacteria that
ferment sucrose more rapidly than lactose.
6. Adding sucrose also aided the identification
of certain gram-negative bacteria that could
ferment sucrose but not lactose.
7. In 1940, Difco Laboratories, Sulkin and
Willet, and Hajna described a similar triple
sugar ferrous sulfate medium for the
identification of enteric bacilli.

68. • The current formulation of triple sugar iron
medium is essentially the same as Sulkin and
Willet except that phenol red is used as the pH
indicator instead of brom thymol blue,
Tryptone has been replaced by a combination
of Bacto Peptone and Proteose Peptone, and
yeast extract has been added.

69. • PURPOSE
• Triple sugar iron (TSI) agar is a tubed differential
medium used in determining carbohydrate
fermentation and H2S production.
• Gas from carbohydrate metabolism can also be
detected.
• Bacteria can metabolize carbohydrates aerobically
(with oxygen) or fementatively (without oxygen).
• TSI differentiates bacteria based on their
fermentation of lactose, glucose and sucrose and on
the production of hydrogen sulfide.
• TSI is most frequently used in the identification of
the Enterobacteriaceae, although it is useful for
other gram-negative bacteria.

70. • THEORY
1. TSI contains three carbohydrates: glucose
(0.1%), sucrose (1%), and lactose (1%).
2. TSI is similar to Kligler's iron agar, except that
Kligler's iron agar contains only two
carbohydrates: glucose (0.1%) and lactose (1%).
3. Besides the carbohydrates mentioned, the
medium also contains beef extract, yeast
extract, and peptones which are the sources of
nitrogen, vitamins and minerals.
4. Phenol red is the pH indicator, and agar is used
to solidify the medium.
5. During preparation, tubes containing molten
agar are angled.

71. 6. The slant of the medium is aerobic, while the deep
(or butt) is anaerobic.
7. When any of the carbohydrates are fermented, the
drop in pH will cause the medium to change from
reddish-orange (the original color) to yellow.
8. A deep red color indicates alkalization of the
peptones.
9. Sodium thiosulfate in the medium is reduced by
some bacteria to hydrogen sulfide (H2S), a colorless
gas.
10.The hydrogen sulfide will react with ferric ions in
the medium to produce iron sulfide, a black
insoluble precipitate.
11.Based on carbohydrate utilization and hydrogen
sulfide production, a TSI slant can be interpreted in
several ways:

72. • GLUCOSE FERMENTER.
1. The tube reaction is alkaline over acid (K/A)
signifying that only glucose is metabolized.
2. The bacteria quickly metabolized the glucose,
initially producing an acid slant and an acid butt
(acid over acid; A/A) in a few hours.
3. The Emben-Meyerhof-Parnas pathway was used
both aerobically (on the slant) and anerobically (in
the butt) to produce ATP and pyruvate.
4. On the slant, the pyruvate was further metabolized
to CO2, H2O, and energy.
5. After further incubation (18 hours) the glucose was
consumed, and because the bacteria could not use
lactose or sucrose, the peptones (amino acids) were
utilized as an energy source aerobically, on the
slant.

73. 6. Utilization of peptones causes the release of
ammonia (NH3) increasing the pH resulting in the
pH indicator, phenol red, turning from yellow to
red.
7. In the anerobic butt, the bacteria use the Embden-
Meyerhof-Parnas pathway to metabolize the
glucose producing ATP and pyruvate, which is
converted into stable acid endproducts, thus the
butt remains acidic.
8. The results would be recorded as alkaline over
acid (K/A).
9. Bacteria producing a K/A reaction with or without
gas include: Citrobacter freundii* , Citrobacter
koseri*, and Morganella morganii*.* = variable
reactions

74. • GLUCOSE, LACTOSE AND/OR SUCROSE
FERMENTER
1. The tube reaction is acid over acid (A/A) indicating
that glucose, lactose and/or sucrose have been
metabolized.
2. The bacteria quickly metabolized the glucose,
producing an acid slant and an acid butt in a few hours.
3. The Emben-Meyerhof-Parnas pathway is used both
aerobically (on the slant) and anerobically (in the butt)
to produce ATP and pyruvate.
4. On the slant, the pyruvate is further metabolized to
CO2, H2O, and energy.
5. After further incubation (18 hours) the glucose was
consumed, and then the bacteria utilized lactose and/or
sucrose, maintaining an acid slant.

75. 1. The results are recorded as acid over acid (A/A).
2. If the medium were incubated longer, over 48
hours, the lactose and sucrose would be depleted,
and the slant would revert to an alkaline pH due to
metabolism of the peptones.
3. In the anerobic butt, the bacteria convert pyruvate
into stable acid endproducts, thus the butt remains
acidic.
4. The bacteria commonly producing an A/A reaction
with or without gas include: Enterobacter
aerogenes, E. cloacae, Escherichia coli, Klebsiella
oxytoca, and K. pneumoniae.

76. • GLUCOSE, LACTOSE AND SUCROSE
NONFERMENTERS.
1. The tube reaction is either alkaline over
alkaline (K/K) or alkaline over no change
(K/NC) indicating that all three sugars have
not been metabolized.
2. The difference between K/K and K/NC) is
subtle.
3. Some nonenteric bacteria, such as the
pseudomonads, are unable to ferment
glucose, lactose, or sucrose.
4. These bacteria derive energy from peptones
either aerobically or anaerobically

77. 6. Utilization of peptones causes the release of ammonia
(NH3) resulting in the pH indicator, phenol red,
turning from pink to red.
7. Nonglucose fermenters can produce two possible
reactions.
8. If the bacteria can metabolize peptones both
aerobically and anaerobically, the slant and butt will
be red (alkaline over alkaline; K/K).
9. If peptones can only be metabolized aerobically, the
slant will be red and the butt will exhibit no change
(K/NC).
10. Bacteria producing K/K or K/NC include:
Acinetobacter spp. and Pseudomonas spp.

78. • GAS PRODUCTION.
1. Gas production (CO2 and O2) is detected by
splitting of the agar.
2. In some cases, so much gas is produced that
the agar is pushed to the top of the tube.
3. Bacteria commonly producing an A/A
reaction with gas include: Enterobacter
aerogenes, E. cloacae, Escherichia coli,
Klebsiella oxytoca, and K. pneumoniae.
However, some strains do not produce gas.

79. • GLUCOSE FERMENTER AND HYDROGEN
SULFIDE PRODUCTION.
1. The tube reaction is alkaline over acid (K/A) with
black precipitate.
2. The bacteria quickly metabolized the glucose, initially
producing an acid slant and an acid butt (acid over
acid; A/A) in a few hours.
3. The Emben-Meyerhof-Parnas pathway is used both
aerobically (on the slant) and anerobically (in the butt)
to produce ATP and pyruvate.
4. On the slant, the pyruvate is further metabolized to
CO2, H2O, and energy.
5. After further incubation (18 hours) the glucose was
consumed, and because the bacteria could not use
lactose or sucrose, the peptones (amino acids) were
utilized as an energy source aerobically, on the slant.

80. 6. Utilization of peptones causes the release of ammonia (NH3)
resulting in the pH indicator, phenol red, turning from yellow
to red. In the anerobic butt, the bacteria use the Embden-
Meyerhof-Parnas pathway to metabolized the glucose
producing ATP and pyruvate, which is converted into stable
acid endproducts, thus the butt remains acidic.
7. The black precipitate indicates that the bacteria were able to
produce hydrogen sulfide (H2S) from sodium thiosulfate.
8. Because H2S is colorless, ferric ammonium citrate is used as
an indicator resulting in the formation of insoluble ferrous
sulfide.
9. Formation of H2S requires an acidic environment; even
though a yellow butt cannot be seen because of the black
precipitate, the butt is acidic.

81. 10.The results would be recorded as alkaline over acid (K/A),
H2S positive. Bacteria producing a K/A with H2S include:
Citrobacter freundii*, Edwardsiella tarda, Proteus mirabilis*,
and Salmonella spp*. Bacteria commonly producing an A/A
with H2S include: Citrobacter freundii*, Proteus mirabilis*,
and P. vulgaris*.* = variable reactions.
• GLUCOSE NONFERMENTER HYDROGEN SULFIDE PRODUCER.
1. The tube appears as alkaline over no change (K/NC) with a
black precipitate (H2S)
2. The reduction of thiosulfate in KIA and TSIA requires H+.
3. Nonfermenters cannot produce an acid environment from
the fermenation of the carbohydrates.
4. Cysteine and perhaps other organic sulfate molecules are
metabolized to pyruvic acid, ammonia, and H2S.
5. Nonfermentative H2S positive reaction is strongly suggestive
of members of the genus Shewenella.

83. Pancreatic digest of casein USP (see Note) 10.0 g
Peptic digest of animal tissue USP (see Note) 10.0 g
Glucose 1.0 g
Lactose 10.0 g
Sucrose 10.0 g
Ferrous sulfate or ferrous ammonium sulfate 0.2 g
NaCl 5.0 g
Sodium thiosulfate 0.3 g
Phenol red 0.024 g
Agar 13.0 g
Distilled water 1,000 ml
RECIPE
Table ( ) Media composition of TSI

84. • Note: The following combination of ingredients
can substitute for the first two components listed:
beef extract, 3.0 g; yeast extract, 3.0 g; and
peptone, 20.0 g.
• Combine ingredients, and adjust the pH to 7.3.
• Boil to dissolve the agar, and dispense into tubes.
• Sterilize by autoclaving at 121°C for 15 min.
Cool in a slanted position to give a 2.5-cm butt
and a 3.8-cm slant.TSI agar is also available
commercially.

85. • PROTOCOL
1. Use a straight inoculating needle
to pickup an isolated colony.
2. Inoculate the TSI slant by first
stabbing the butt down to the
bottom, withdraw the needle, and
then streak the surface of the
slant. Use a loosely fitting closure
to permit access of ai
3. Read results after incubation at
37°C for 18 to 24 h. Three kinds of
data may be obtained from the
reactions.

86. • SUGAR FERMENTATIONS
• Acid butt, alkaline slant (yellow butt, red slant):
glucose has been fermented but not sucrose or
lactose.
• Acid butt, acid slant (yellow butt, yellow slant):
lactose and/or sucrose has been fermented.
• Alkaline butt, alkaline slant (red butt, red slant):
neither glucose, lactose, nor sucrose has been
fermented.
• GAS PRODUCTION
• Indicated by bubbles in the butt.
• With large amounts of gas, the agar may be broken
or pushed upward.

87. • HYDROGEN SULFIDE PRODUCTION
• Hydrogen sulfide production from thiosulfate is indicated
by a blackening of the butt as a result of the reaction of
H2S with the ferrous ammonium sulfate to form black
ferrous sulfide.
• The black precipitate indicates that the bacteria were able
to produce hydrogen sulfide (H2S) from sodium
thiosulfate.
• Because H2S is colorless, ferric ammonium citrate is used
as an indicator resulting in the formation of insoluble
ferrous sulfide.
• Formation of H2S requires an acidic environment; even
though a yellow butt cannot be seen because of the black
precipitate, the butt is acidic.
• The results would be recorded as acid over acid (A/A),
H2S positive

88. • KLIGLER IRON AGAR (KIA)
• Note the relative amounts of sugars in KIA according to the
table seen above. By the degree of acid produced from
fermentation, differentiation can be made between non-
fermenters, glucose-fermenters (which produce a relatively
small amount of acid) and those which ferment both glucose
and lactose (producing a relatively large amount of acid which
diffuses throughout the medium and easily overneutralizes the
aerobic deamination reaction in the slant).
• Organisms which produce hydrogen sulfide from the reduction
of thiosulfate are easily detected; the H2S reacts with the iron in
the medium to produce ferrous sulfide, a black precipitate.
• The medium is inoculated with the needle, first stabbing down
the center to the bottom of the tube and then streaking up the
slant.
• Incubation is for one day at 37°C. The various combinations of
reactions are explained and illustrated below.
• (Tube "C" is the uninoculated control tube which shows an
orange (neutral) reaction throughout.)

89. corresponding
tube no. above 1 2 3 4* 5**
deamination of
amino acids
(aerobic alkalin
e rx.)
+ + + + +
glucose
fermentation
(minor acid rx.)
– + + + +
lactose
fermentation
(major acid rx.)
– – – + +
H2S production
(black color) – – + – +**
typical
examples
Pseud
omon
as
(a
non-
enteri
c)
Morga
nella,
Provid
encia,
Shigel
la
Citrobact
er,
Salmonel
la,
Proteus,
Edwardsi
ella
E. coli,
Enteroba
cter,
Klebsiella
coliform
strains of
Citrobact
er that
are H2S+,
H2S+ E.
coli,
lactose+
Salmonel
la

90. • Tube 4: Much gas is often seen for this tube, evidenced
by cracks in the medium. Also, lactose fermenters
which are methyl red-negative may show a "reversion"
toward an alkaline reaction as neutral products are
formed from some of the acid.
• This appears as shown in tube 4A where a slight
reddening of the slant occurs as the alkaline
deamination reaction becomes no longer over-
neutralized by acid from fermentation. How might
such a tube appear after two or more days of
incubation? (Recall the methyl red test.)
• Tube 5: Enough acid can be produced to cause the
black iron sulfide precipitate to break down and not be
seen. In this case, the tube will look like no. 4.

91. TSI
INGREDIENTS FUNCTION
RESULT/INTERPRETATION
Phenol Red a pH indicator:
below 6.8 it is
yellow
above 82., it is red
Phenol red turns yellow in an acid environment. It thus
indicates whether the acids of fermentation have been
produced. Failure to turn the butt yellow indicates that no
fermentation has occured, and that the bacterium is an
obligate aerobe.
0.1 % glucose if only glucose is
fermented, only a
small amount of
acid is produced
If only glucose is fermented, only enough acid is produced to
turn the butt yellow. The slant will remain red.
1.0 % lactose
1.0% sucrose
if the culture can
ferment either
lactose (lac+)
and/or sucrose
(suc+), a large
amount of acid is
produced
a large amount of acid turns both butt and slant yellow, thus
indicating the ability of the culture to ferment either lactose or
sucrose
FeSO4
(ferrous sulfate)
A source of iron
and sulfur
A few bacteria are capable of reducing the SO4= to H2S
(hydrogen sulfide).
The iron combines with the H2S to form FeS (ferrous sulfide)
a black compound. This will turn the butt black. Thus, a
black butt indicates H2S production.
Table ( ) FUNCTION and RESULT/INTERPRETATION for TSI

92. • BISMUTH SULFITE AGAR
1. Bismuth Sulfite Agar is used for the selective isolation of Salmonella
spp. Salmonellosis continues to be an important public health problem
worldwide. Infection with non-typhi Salmonella often causes mild,
self-limiting illness. Salmonellosis can result from consumption of
raw, undercooked, or improperly processed foods contaminated with
Salmonella. U. S. federal guidelines require various poultry products
to be routinely monitored before distribution for human consumption.
2. Bismuth Sulfite Agar is a modification of Wilson and Blair formula.
The typhoid organism grows abundantly on the medium, forming
characteristic black colonies.
3. Gram-positive bacteria and coliforms are inhibited on Bismuth Sulfite
Agar. The inhibitory action of Bismuth Sulfite Agar permits the use of
a large inoculum, increasing the possibility of recovering pathogens
that may be present in small numbers.
4. Bismuth Sulfite Agar is generally accepted for routine detection of
most Salmonella spp. Bismuth Sulfite Agar is used for the isolation of
S. typhi and other Salmonella spp. from food, feces, urine, sewage,
and other infectious materials.
5. Bismuth Sulfite Agar is a standard methods medium for industrial
applications and the clinical environment.

93. • PRINCIPLES
• Enzymatic Digest of Casein, Enzymatic Digest of Animal
Tissue, and Beef Extract provide sources of nitrogen, carbon,
and vitamins required for organism growth.
• Dextrose is the carbohydrate present in Bismuth Sulfite
Agar.
• Disodium Phosphate is the buffering agent.
• Bismuth Sulfite Indicator and Brilliant Green are
complementary, inhibiting Gram-positive bacteria and
coliforms, allowing Salmonella spp. to grow.
• Ferrous Sulfate is used for H2S production. When H2S is
present, the iron in the formula is precipitated, and positive
cultures produce the characteristic brown to black color with
metallic sheen.
• Agar is the solidifying agent.

95. Enzymatic Digest of Casein 5 g
Enzymatic Digest of Animal Tissue 5 g
Beef Extract 5 g
Dextrose 5 g
Disodium Phosphate 4 g
Ferrous Sulfate 0.3 g
Bismuth Sulfite Indicator 8 g
Brilliant Green 0.025 g
Agar 20 g
Final pH: 7.5 ± 0.2 at 25 C
Table ( ) Media of Bismuth Sulfite Agar composition
Formula may be adjusted and/or supplemented as required to meet performance
specifications.

96. • MEDIA PREPARATION
1. Suspend 52.0gm of the dehydrated culture media in
1 liter of distilled or deionized water.
2. Stir to mix thoroughly.
3. Heat to boiling to dissolve completely,
approximately 1 minute.
4. Do not overheat.
5. Do not autoclave.
6. Cool to 45-50 degrees C.
7. Mix thoroughly before pouring into petri plates.
8. Use poured plates the same day.

97. • SLANT AGAR AND BROTH
MEDIA
• Growth of bacterial cultures on agar
slants and in broths can provide us
with useful information concerning
motility, pigmentation and oxygen
requirements. While there is variation
even among individual strains of the
same species, some characteristics are
distinctive, thus can aid in the
beginning steps of identification.
• All samples were grown on trypticase
soy agar (TSA) for 48 hours at 37o
C. Click on each image to see a larger
view.
Figure ( ) Inoculation
method of slant agar

98. This is a slant of Staphylococcus aureus.
Note the even pattern of growth which
follows the line of inoculation. The wider
portion at the bottom is due to the presence
of a small amount of condensation.
This is a slant of Bacillus subtilis.
Note the spreading pattern of
growth.

99. • MEDIUM PREPARATION
1. The medium is prepared differently for slants than Petri
dishes.
2. Sterilization is done with the agar in the tubes; Petri
dishes are pre-sterilized before sterilized agar is poured
into them.
3. Measure the amount of water needed and put it in a pot.
4. Heat it on a stove until it is almost boiling. Add dry
ingredients and stir the mixture slowly until they
dissolve. Before adding agar, mix it with a small amount
of cold water to prevent lumping. Use caution when
adding agar to the hot liquid since it can foam and
overflow the pot. Add small amounts of agar at a time
and stir to evenly distribute the agar. Turn off the heat
after bringing the agar to boil.

100. • STERILIZING TUBES
1. Place test tubes without the caps on a test tube rack. Fill the test tubes
by transferring about 5 milliliters -- about .17 ounce or 1 teaspoon --
of the molten agar from the pot using a pipette, a small funnel or a
syringe. Place all the caps loosely on the test tubes -- the agar won't be
sterilized if they are sealed tight -- and sterilize all the tubes for about
25 minutes at 250 degrees Fahrenheit.
2. Slanting
3. When the agar is still hot, tilt the rack holding the test tubes on a solid
surface or a thick book, making sure the medium inside the tubes is at
a slanted position. Allow the medium to cool and solidify at this angle,
which increases the surface area of the agar.
4. Storage
5. Tighten the caps of the test tubes after the agar has cooled. The slants
are ready for use once the agar has solidified. They can be stored at
room temperature or in the refrigerator for future use.
6. Inoculation
7. Inoculate the slant by transferring cells with an inoculating loop from
a single-colony microorganism on a plate to the slant's surface. Move
the loop across the surface of the slant and cap the tubes. Incubate the
slant until there is evidence of growth, then put the tube in a
refrigerator.

101. • TERMS USED FOR GROWTH ON NUTRIENT
SLANTS
• Abundance of growth - the amount of growth is
designated as none, slight, moderate, or large
• Pigmentation – chromogenic bacteria may produce
intracellular pigments that are responsible for the color of
the colonies on the agar surface. Other bacteria produce
extracellular soluble pigments that are excreted into the
medium and that also produce a color. Most
microorganisms are nonchromogenic and will appear
cream, white, or gray.
• Optical characteristics - these characteristics are based on
the amount of light transmitted through the growth:
opaque (no light transmitted), translucent (partial
transmission), or transparent (full transmission).

102. • THE APPEARANCE OF THE SINGLE LINE
STREAK OF GROWTH ON THE AGAR SLANT.
• Filiform – continuous, threadlike growth with smooth
edges
• Echinulate – continuous threadlike growth with
irregular edges
• Beaded – nonconfluent to semi-confluent colonies
• Effuse – thin, spreading growth
• Arborescent – treelike growth
• Rhizoid – rootlike growth
Figure ( ) Different pattern bacterial growth on slant agar

103. • BROTHS
1. When bacteria are grown in broths such as trypticase soy broth
(TSB), they may exhibit patterns of growth ranging from a
sediment at the bottom of the tube, turbid growth throughout the
tube, or a pellicle (thick growth at the top of the tube).
2. Pellicle formation is sometimes due to a affinity for oxygen, but
may also be the result of hydrophobic compounds present in the
cell wall or the general formation of dry, light colonies.
3. Also, if an organism produces and releases soluble pigments,
these will spread into the broth and change its color.
4. Here are two examples of growth patterns in broth after 48 hours
incubation at 37o C:
•This broth contains the acid-fast species
Mycobacterium smegmatis. Note the pellicle on
the surface of the broth which forms due to the
high concentration of hydrophobic mycolic acids
embedded in the cell wall of this species.

104. • This broth contains Serratia marcescens, a gram-
negative rod. Observe the turbid appearence of
the broth and the red color present in both the
sediment and pellicle, which is the result of the
nonsoluble pigment prodigiosin produced by this
bacterium.
TERMS USED FOR GROWTH IN NUTRIENT BROTH
•Uniform fine turbidity – finely dispersed growth throughout (cloudy)
•Flocculent – flaxy aggregates dispersed throughout
•Pellicle – thick, padlike growth on the surface
• Sediment – concentration of growth at the bottom of the broth culture may be
granular, flaxy, or flocculent
Ring formation – a ring of growth on the surface

105. • CLOSTRIDIUM DIFFICILE AGAR
• Clostridium difficile causes gastrointestinal infections
in humans that range in severity from asymptomatic
colonization to severe diarrhea, antibiotic-associated
diarrhea, and pseudomembranous colitis (PMC).
• Nosocomail infection, both symptomatic and
asymptomatic, occurs through transient cross-infection
of C. difficile on the hands of healthcare workers as
well as through contact with contaminated
environmental surfaces.
• In 1979, George et al. isolated C. difficile using CCFA
Medium, a modification of McClung Toabe agar.
Levett described Clostridium difficile Agar which is a
modification of CCFA Medium with an egg yolk agar
base and reduced concentrations of cycloserine and
cefoxitin.

106. • PRINCIPLE
1. Proteose peptone supplies amino acids and other
nitrogenous compounds necessary for the growth of
anaerobic bacteria, including C. difficile.
2. Sodium chloride is a source of essential electrolytes and
maintains osmotic equilibrium.
3. Fructose is an energy source.
4. Monopotassium and disodium phosphates are buffering
agents which maintain the pH of the medium.
5. Clostridium difficile Agar is both selective and
differential.
6. The growth of C. difficile raises the pH of the medium
causing the neutral red indicator to turn a yellow color;
this can be observed in the colonies and the surrounding
medium.

107. 7. C. difficile also produces a characteristic yellow
fluorescence which can be observed under long wave
ultraviolet light.
8. Egg yolk reduces the toxic effect of organic peroxides
which may accumulate in the medium and serves as a
substrate for detection of lecithinase and lipase activity.
9. Some species of Clostridium produce lecithinase and/or
lipase; C. difficile does not.
10. Cycloserine and cefoxitin are selective agents.
11. Cycloserine is active against Escherichia coli ,other gram-
negative bacilli, and streptococci.
12. Cefoxitin is a broad-spectrum antibiotic which is active
against a variety of gram-positive and gram-negative
bacteria, with the exception of Enterococcus faecalis and
C. difficile.
13. Agar is a solidifying agent.

108. REAGENTS (CLASSICAL FORMULA)*
Proteose
Peptone
40.0 g Magnesium
Sulfate
0.1 gm
Fructose 6.0 gm Neutral Red 0.03 gm
Disodium
Phosphate
5.0 gm Cefoxitin 0.016 gm
Sodium
Chloride
2.0gm Egg Yolk
Suspension
100.0 ml
Monopotassiu
m Phosphate
1.0 gm Agar 20.0 gm
Cycloserine 0.25 gm Demineralize
d W ater
900.0 ml
pH 7.6 ±0.2 , 25°C
*Adjusted as required to meet performance standards.

109. • PROCEDURE
• Prior to use, reduce the plates for a minimum of 24 hours by
placing them in an anaerobic jar at room temperature.
• Inoculate specimens for anaerobic culture on both selective and
nonselective media.
• Incubate anaerobically at 33-37°C for 48-72 hours.
• Following incubation, examine the plate for flat, circular colonies
with filamentous edges that demonstrate a yellow zone extending
2-3 mm from the edge of the colony.
• Inspect suspicious colonies under long wave ultraviolet light for
yellow fluorescence.
• Confirm anaerobic growth by subculture of colonies
representative of C. difficile to a blood agar plate incubated at
33-37°C in ambient air.
• Consult appropriate references for additional tests to confirm the
presence of C. difficile

110. • POTATO DEXTROSE AGAR
• Potato Dextrose Agar (PDA) is used for the
cultivation of fungi.
• Potato Dextrose Agar (PDA) is a general purpose
medium for yeasts and molds that can be
supplemented with acid or antibiotics to inhibit
bacterial growth.
• It is recommended for plate count methods for
foods, dairy products and testing cosmetics.
• PDA can be used for growing clinically
significant yeast and molds.
• The nutritionally rich base (potato infusion)
encourages mold sporulation and pigment
production in some dermatophytes

111. • PRINCIPLE OF PDA
• Potato Dextrose Agar is composed of dehydrated
Potato Infusion and Dextrose that encourage
luxuriant fungal growth.
• Agar is added as the solidifying agent.
• Many standard procedures use a specified amount
of sterile tartaric acid (10%) to lower the pH of
this medium to 3.5 +/- 0.1, inhibiting bacterial
growth.
• Chloramphenicol acts as a selective agent to
inhibit bacterial overgrowth of competing
microorganisms from mixed specimens, while
permitting the selective isolation of fungi.
• Note: Do not reheat the acidified medium, heating
in the acid state will hydrolyze the agar.

112. • USE OF PDA
• Potato Dextrose Agar is used for the detection of
yeasts and molds in dairy products and prepared
foods.
• It may also be used for the cultivation of yeasts
and molds from clinical specimens.
• Potato Dextrose Agar with TA (Tartaric Acid) is
recommended for the microbial examination of
food and dairy products.
• Potato Dextrose Agar with Chlortetracycline is
recommended for the microbial enumeration of
yeast and mold from cosmetics.
• Potato Dextrose Agar with Chloramphenicol is
recommended for the selective cultivation of
fungi from mixed samples.

113. • COMPOSITION OF PDA
• In lab preparations
1. 200 gm potato infusion( is equivalent to 4.0 gm of potato extract).
2. 20 gm Dextrose.
3. 20 gm agar.
4. 1000ml distilled water.
• To prepare potato infusion
1. Boil 200 g sliced, unpeeled potatoes in 1 liter distilled water for
30 min.
2. Filter through cheesecloth, saving effluent, which is potato
infusion (or use commercial dEhydrated form).
3. Mix with Dextrose, Agar and Water and boil to dissolve.
4. Autoclave 15 min at 121°C.
5. Dispense 20-25 ml portions into sterile 15 × 100 mm petri dishes.
6. Final pH, 5.6 ± 0.2.

114. • PREPARATION PDA FROM COMMERCIAL
1. Add 39 gm of Commercial PDA Powder (20 gm dextrose,
15 gm agar, and 4 gm potato starch) to 1000ml distilled
water.
2. Boil while mixing to dissolve.
3. Autoclave 15 min at 121°C.
4. In addition, Potato Dextrose Agar with Chlortetracycline
contains: 40.0 mg Chlortetracycline
5. In addition, Potato Dextrose Agar with Chloramphenicol
contains: 25.0 mg Chloramphenicol
6. Final pH of 5.6 +/- 0.2 at 25 degrees C.
7. In addition, Potato Dextrose Agar with TA contains: 1.4
gm Tartaric Acid
8. Final pH of 3.5 +/- 0.3 at 25 degrees C.

115. • COLONY CHARACTERISTICS ON PDA
• After sufficient incubation, isolated colonies
should be visible in the streaked areas and
confluent growth in areas of heavy inoculation
Aspergillus flavus: Powdery masses of yellow-green spores on the
upper surface and reddish-gold on the lower surface.

116. • STANDARD PLATE COUNT (VIABLE
COUNTS)
1. A viable cell is defined as a cell which is able to
divide and form a population (or colony).
2. A viable cell count is usually done by diluting
the original sample, plating aliquots of the
dilutions onto an appropriate culture medium,
then incubating the plates under proper
conditions so that colonies are formed.
3. After incubation, the colonies are counted and,
from a knowledge of the dilution used, the
original number of viable cells can be
calculated.

117. 4. For accurate determination of the total number of
viable cells, it is critical that each colony comes from
only one cell, so chains and clumps of cells must be
broken apart.
5. However, since one is never sure that all such
groups have been broken apart, the total number of
viable cells is usually reported as colony-forming
units (CFUs) rather than cell numbers.
6. This method of enumeration is relatively easy to
perform and is much more sensitive than
turbidimetric measurement.
7. A major disadvantage, however, is the time
necessary for dilutions, platings and incubations, as
well as the time needed for media preparation.

118. • TERMS USED FOR GROWTH ON NUTRIENT
AGAR PLATES
1. Size – pinpoint, small, moderate, large
2. Pigmentation – color of colony
3. Optical properties
• a. opaque
• b. translucent (clear)
• c. shiny
• d. dull

119. • SABOURAUD AGAR FOR FUNGAL GROWTH
PROTOCOLS
• Sabouraud (pronounced sah-bū-rō′) agar medium was
developed by the French dermatologist Raymond J. A.
Sabouraud in the late 1800’s to support the growth of
fungi that cause infection of the skin, hair, or nails,
collectively referred to as dermatophytes.
• Sabouraud’s medical investigations focused on bacteria
and fungi that cause skin lesions, and he developed many
agars and techniques to culture pathogenic moulds and
yeasts, such as dermatophytes and Malassezia.
• He particularly desired that all mycologists detail their
exact media formulations, temperatures and times of
incubation of specimens, in order to standardize the
field’s observations and thus reduce differences in
appearance as a possible source of error in identification .

120. • PURPOSE
• Historically, Sabouraud agar was developed to
support the studies of dermatophytes, which require
long incubation periods (weeks).
• There were two driving forces behind Sabouraud’s
development of this medium:
• The need to avoid bacterial contamination while
culturing dermatophytes and other fungi
• The need to provide a medium that would yield
reliable results for fungal identification across
laboratories.
• Sabouraud agar is a selective medium that is
formulated to allow growth of fungi and inhibit the
growth of bacteria.

121. • The available means of inhibiting bacterial growth
in Sabouraud’s pre-antibiotic era was an acidic
medium (pH 5.6).
• However, the addition of antibiotics to the acidic
medium to inhibit bacteria (and sometimes
saprophytic fungi, depending on the particular
antibiotics used) is common in currently used
versions.
• Glucose is present at the high level of 4% in
Sabouraud’s formulation to assist in vigorous
fermentation and subsequent acid production by any
bacteria present.
• High acid concentrations eventually serve to inhibit
all bacterial growth.

122. • THEORY
• The medium is complex but contains few
ingredients.
• Peptones, as soluble protein digests, are sources
of nitrogenous growth factors that can vary
significantly according to protein source.
• Sabouraud’s original formulation contained a
peptone termed “Granulée de Chassaing,” which
is no longer available (This may be why the
standard name for this medium is “Sabouraud
agar, modified.”) Variations in

123. pigmentation and sporulation can be
consistently observed if one uses Sabouraud
medium prepared with consistent ingredients,
because morphology can vary slightly based
on the peptones used.
• Both Difco and BBL Sabouraud agars use
pancreatic digests of casein as their peptone
source.
• Although Sabouraud originally used the sugar
maltose as an energy source, glucose (or
dextrose, as it used to be called), is currently
used, and agar serves to solidify the medium.

124. • RECIPES AND PROTOCOLS
• Sabouraud agar can be purchased from a variety of commercial
sources, either as the original recipe (Sabouraud agar, modified),
or in a slightly altered version termed “Sabouraud agar,
Emmons.” The neutral pH of the Emmons modification seems
to enhance the growth of some pathogenic fungi, such as
dermatophytes.
• Per liter of medium:
1. 10gm Peptone.
2. 40 gm Glucose.
3. 15 gm Agar
4. Combine all ingredients in ~900 ml of deionized water.
5. Adjust to pH 5.6 with hydrochloric acid and adjust final
volume to 1 liter
6. Autoclave 20 minutes at 121°C, 15 lb/in2.
7. Cool to ~45 to 50°C and pour into petri dishes or tubes for
slants.

125. • EMMONS MODIFICATION OF SABOURAUD
AGAR
• Per liter of medium: Neo-peptone, 10 g
• Glucose, 20 g Agar, 20 g
• Follow steps 1 through 4, above, except adjust the
pH to the range of 6.8 to 7.0 with hydrochloric acid
before autoclaving, cooling, and pouring.
• Either Sabouraud agar or its Emmons version can be
made more selective by adding antibiotics.
• Commonly used are gentamicin, which inhibits
gram-negative bacteria, and/or chloramphenicol,
which inhibits a wide range of gram-positives and
gram-negatives, and cycloheximide, which inhibits
primarily saprophytic fungi but not dermatophytes or
yeasts.

126. • Chloramphenicol and gentamicin are used at 50
mg/liter (dissolved in 10 ml of 95% ethanol before
adding to molten media) and cycloheximide at 0.5
g/liter (dissolved in 2 ml of acetone first).
• Antibiotics should only be added after media has
been autoclaved and then cooled to ~45 to
50°C. Keep all plates at 4°C until they are used,
regardless of whether they contain antibiotics.
• Sabouraud agar plates can be inoculated by
streaking (see the Streak Plate Protocol for an
explanation of this method), as with standard
bacteriological media, or by exposing the medium
to ambient air.

127. • Typically, molds are incubated at room
temperature (22 to 25°C) and yeasts are
incubated at 28 to 30°C or 37°C if suspected of
being dimorphic fungi.
• Incubation times will vary, from approximately 2
days for the growth of yeast colonies such as
Malasezzia, to 2 to 4 weeks for growth of
dermatophytes or dimorphic fungi such as
Histoplasma capsulatum. Indeed, the incubation
time required to acquire fungal growth is one
diagnostic indicator used to identify or confirm
fungal species.

128. • SABOURAUD DEXTROSE AGAR
• Hardy Diagnostics Sabouraud Dextrose Agar,
Sabouraud Dextrose Broth, and Sabouraud Dextrose
Agar, Emmons are recommended for the isolation,
cultivation, and maintenance of non-pathogenic and
pathogenic species of fungi and yeasts.
• Sabouraud Dextrose Agar with Chloramphenicol,
Sabouraud Dextrose Agar with Chloramphenicol and
Gentamicin, and Sabouraud Dextrose Agar with
Chloramphenicol and Tetracycline are recommended
for the selective isolation of fungi and yeasts from
clinical and nonclinical specimens.

129. • Sabouraud Dextrose Agar was formulated by Sabouraud in
1892 for culturing dermatophytes.
• The pH is adjusted to approximately 5.6 in order to enhance the
growth of fungi, especially dermatophytes, and to slightly
inhibit bacterial growth in clinical specimens.
• This medium is recommended for mold and yeast counts by the
Association of Official Analytical Chemists and the
Compendium of Methods for the Microbiological Examination
of Foods.
• Sabouraud Dextrose Broth is a modification of the original
formulation made without agar.
• Sabouraud Dextrose Agar, Emmons is a modification of the
original formulation. Emmons originally formulated this
modification, which reduces the amount of dextrose, and
neutralizes the medium to a pH of approximately 7.0.
• Chloramphenicol, gentamicin, and tetracycline are selective
agents added to inhibit bacterial overgrowth of competing
microorganisms while permitting the successful isolation of
fungi and yeasts.

130. • Sabouraud Dextrose Medium contains digests of
animal tissues (peptones) which provide a nutritious
source of amino acids and nitrogenous compounds
for the growth of fungi and yeasts.
• Dextrose is added as the energy and carbon source.
• Chloramphenicol and/or tetracycline may be added
as broad spectrum antimicrobials to inhibit the
growth of a wide range of gram-positive and gram-
negative bacteria.
• Gentamicin is added to further inhibit the growth of
gram-negative bacteria.
• Sabouraud Dextrose Medium is not recommended
for the cultivation of dermatophytes, dematiaceous
fungi, and mucormycetes (formally zygomycetes).
Also, it is a poor promoter of conidiation (see
"Limitations" section below).

131. • FORMULA
• Ingredients per liter of deionized water:
Sabouraud Dextrose Agar:
Dextrose 40.0gm
Pancreatic Digest of Casein 5.0gm
Peptic Digest of Animal Tissue 5.0gm
Agar 15.0gm
Final pH 5.6 +/- 0.2 at 25ºC
Final pH 5.6 +/- 0.2 at 25ºC

132. • In addition,
1. Sabouraud Dextrose Broth is the same
formulation as above, without agar added.
Final pH 5.6 +/- 0.2 at 25ºC.
2. Sabouraud Dextrose Agar with Chloramphenicol
contains 50.0mg of chloramphenicol.
Final pH 5.6 +/- 0.3 at 25ºC.
3. Sabouraud Dextrose Agar with Chloramphenicol
and Gentamicin contains 50.0mg of
chloramphenicol and 5.0mg gentamicin
4. Final pH of 5.6 +/- 0.3 at 25ºC.

133. 5. Sabouraud Dextrose Agar with
Chloramphenicol and Tetracycline contains
50.0 mg of chloramphenicol and 10.0mg of
tetracycline.
6. Final pH of 5.6 +/- 0.3 at 25ºC.
7. Sabouraud Dextrose Agar, Emmons has only
20.0gm of dextrose.
8. Final pH of 6.9 +/- 0.2 at 25ºC.
• Adjusted and/or supplemented as required to
meet performance criteria

134. • SABOURAUD DEXTROSE BROTH
• Sabouraud Dextrose Broth is used for the
cultivation of fungi from sterile and non-sterile
products.
• Sabouraud Dextrose Broth is a modification of
Dextrose Agar described by Sabouraud.
• Sabouraud Dextrose Media are used for cultivating
pathogenic & commensally fungi and yeasts.
• The high dextrose concentration and acidic pH of
the formulas permit selectivity of fungi.
• Sabouraud Dextrose Broth is used for the
determination of fungistatic activity in sterile and
non-sterile pharmaceutical, food & beverage, and
cosmetic products.

135. • PRINCIPLES OF THE PROCEDURE
• Enzymatic Digest of Casein and Enzymatic Digest of
Animal Tissue provide the nitrogen and vitamin source
required for organism growth in Sabouraud Dextrose Broth.
• The high concentration of Dextrose is included as an energy
source.
• FORMULA / LITER
• Enzymatic Digest of Casein ................................................ 5
g
• Enzymatic Digest of Animal Tissue..................................... 5
g
• Dextrose ............................................................................. 20
g
• Final pH: 5.6 ± 0.2 at 25 C
• Formula may be adjusted and/or supplemented as required to
meet performance specifications

136. • MARGIN – THE APPEARANCE OF THE OUTER EDGE
OF THE COLONY
• a. entire – sharply defined, even, smooth
• b. lobate – marked indentation (lobed)
• c. undulate – wavy indentation
• d. serrate or erose – tooth-like appearance
• e. curled
• f. rhizoid – root-like
• g. filamentous – threadlike, spreading edge
ELEVATION – THE DEGREE TO WHICH THE COLONY
GROWTH IS RAISED
a. flat – elevation not discernible
b. raised – slightly elevated
c. convex – dome-shaped
d. umbonate – raised, with elevated convex center region
e. pulvinate – very convex

137. • FORM OF COLONIES
• Circular.
• Irregular.
• Filamentous.
• Rhiziod.

138. • ELEVATION
• Raised
• Convex.
• Flat.
• Unbonate.
• Crateriform.

139. • MARGIN
• Entire.
• Undulate.
• Filiform.
• Curled
• Lobate

140. • TURBIDIMETRIC MEASUREMENT
• A quick and efficient method of estimating the number of
bacteria in a liquid medium is to measure the turbidity or
cloudiness of a culture and translate this measurement into
cell numbers.
• This method of enumeration is fast and is usually preferred
when a large number of cultures are to be counted.
• Although measuring turbidity is much faster than the
standard plate count, the measurements must be correlated
initially with cell number.
• This is achieved by determining the turbidity of different
concentrations of a given species of microorganism in a
particular medium and then utilizing the standard plate
count to determine the number of viable organisms per
milliliter of sample.

141. • A standard curve can then be drawn (e.g., this lab
protocol section), in which a specific turbidity or
optical density reading is matched to a specific
number of viable organisms.
• Subsequently, only turbidity needs to be measured.
• The number of viable organisms may be read directly
from the standard curve, without necessitating time-
consuming standard counts.
• Turbidity can be measured by an instrument such as
a colorimeter or spectrophotometer. These
instruments contain a light source and a light detector
(photocell) separated by the sample compartment.
• Turbid solutions such as cell cultures interfere with
light passage through the sample, so that less light
hits the photocell than would if the cells were not
there.

142. • Turbidimetric methods can be used as long as each
individual cell blocks or intercepts light; as soon as the
mass of cells becomes so large that some cells effectively
shield other cells from the light, the measurement is no
longer accurate
• Before turbidimetric measurements can be made, the
spectrophotometer must be adjusted to 100% transmittance
(0% absorbance).
• This is done using a sample of uninoculated medium.
Percent transmittance of various dilutions of the bacterial
culture is then measured and the values converted to
optical density, based on the formula: Absorbance
(O.D.) = 2 - log % Transmittance.
• A wavelength of 420 nm is used when the solution is clear,
540 nm when the solution is light yellow, and 600-625 nm
is used for yellow to brown solutions.

143. • DIRECT MICROSCOPIC COUNT
• Petroff-Hausser counting chambers can
be used as a direct method to determine
the number of bacterial cells in a culture
or liquid medium.
• In this procedure, the number of cells in
a given volume of culture liquid is
counted directly in 10-20 microscope
fields.
• The average number of cells per field is
calculated and the number of bacterial
cells ml-1 of original sample can then be
computed.
• A major advantage of direct counts is the
speed at which results are obtained.
• However, since it is often not possible to
distinguish living from dead cells, the
direct microscopic count method is not
very useful for determining the number of
viable cells in a culture.
Petroff-Hausser counting
chambers

144. • MATERIAL
1. Seven 9-ml dilution tubes of nutrient broth
2. Six nutrient agar plates
3. 1.0 and 10 ml pipets
4. Glass spreader
5. 95% ethyl alcohol in glass beaker (WARNING:
Keep alcohol away from flame!!)
6. Overnight broth culture of Serratia marcescens

145. • PROCEDURE: (WORK IN PAIRS)
• A. SPREAD PLATE TECHNIQUE
• Prepare serial dilutions of the broth culture as shown in the
figure from a previous lab exercise (Isolation of Pure
Cultures). Be sure to mix the nutrient broth tubes before
each serial transfer. Transfer 0.1 ml of the final three
dilutions (10-5, 10-6, 10-7) to duplicate nutrient agar plates,
and label the plates
• Spread the 0.1 ml inoculum evenly over the entire surface
of one of the nutrient agar plates until the medium no
longer appears moist. Return the spreader to the alcohol.
• Repeat the flaming and spreading for each of the remaining
five plates.
• Invert the six plates and incubate at room temperature until
the next lab period (or ~ 48 hours, whichever is the
shortest). Remember that only plates with 25 – 250
colonies are statistically valid.

146. • B. TURBIDIMETRIC METHOD
• Using the spectrophotometer, determine the optical
density (O.D.) of the assigned broth culture at 600
nm. Note, you may have to use one of your serial
dilutions of the broth culture to get a good reading.
• Record results.
• C. DIRECT MICROSCOPIC COUNTS
• MATERIAL:
• Petroff-Hausser counting chamber
• Cover slips
• Sterile diluents (nutrient broth or sterile saline)
• Pasteur pipets

147. • PROCEDURE: (WORK IN PAIRS)
• BE EXTREMELY CAREFUL HANDLING PETROFF-
HAUSSER COUNTING CHAMBERS!
1. Clean P-H counting chamber with 70% alcohol an let air dry.
2. Mix culture well and apply a single drop to counting chamber
with Pasteur pipet. Examine the counting chamber using high
power, oil immersion objective.
3. Make a preliminary estimation of the concentration of cells
from the overnight culture of Serratia marcescens using the
following formula: Therefore, if you counted an average of 15
cells per small square, then you would have a final
concentration of 3.0 x 108 cells/ml.

148. • You may have to adjust downward using one of
your initial serial dilutions so that the counts per
small square are in the 5 to 15 cell range.
• Once this is done, make sure to allow time for
cells to settle and move focus through the
suspension (i.e., up and down) so as to count all
cells within the small square “box”. Most cells will
have attached to the bottom and/or top glass
interface. You can also check the depth, which is
20 μm. The small square should also be 50 by 50
μm.
• Count the number of bacterial cells in at least 10
small squares. Variability should be less than +/-
10%.

149. • SECOND PERIOD
• MATERIAL:
• 1. Colony counter
• PROCEDURE:
• Remember to pull plates and refrigerate after 48
hours max. Either then or next lab period, count the
number of colonies on each plate, calculate an
average and record results.
• Compare results from the standard plate counts with
P-H direct microscopic counts.
• Compare results from the standard plate counts and
direct microscopic counts with that of optical
density while considering the graph provided.
Which data are the most robust and why? Which
data yields the highest counts and why?

150. Plate # 10-6 10-7 10-8
1
2
3
Average
RESULTS:
Dilutions
Number of colony-forming units per ml ___________

151. • THE PREPARATION OF SPREAD AND STREAK
PLATES
• Bacteria are found just about everywhere, and most of them
are nonpathogenic.
• Others are just plain harmful, pathogenic forms.
• Still others are harmless as long as they maintain their
personal space, but become a threat when they get into areas
other than their natural habitat.
• E. coli , for example, are natural residents of large intestines.
• There they cause no harm and actually help by assisting with
waste processing, vitamin K production, and food absorption.
• When E. coli or some of the other types of microorganisms
leave their normal habitats and enter areas where they are not
normally found, they can cause disease.

152. • Contamination of foods by E. coli or other
microorganisms is a serious threat to health.
• How can we test for organisms such as E. coli that might
cause microbial contamination? What if we find that the
organisms are present in some substances - how can we
determine the degree of contamination of the material?
• The rate of microbial spoilage depends upon the chemical
composition of the affected substance(s) and the types of
microorganisms causing the infection.
• Freezing, boiling and secure packaging help prevent
contamination.
• Improper handling, such as employees returning to
processing areas from the bathroom without washing their
hands, can cause serious contamination.

153. • Improper slaughter and packaging procedures
can also cause contamination.
• Careless beef processing has apparently caused
recent outbreaks of a lethal form of E. coli .
Animal feces containing E. coli were included
in beef processing along with the beef body
tissues.

154. • EXERCISE )1)
1. Each member of a two-person team needs to
obtain a clean, closed Petri dish that contains
nutrient agar.
2. Each team needs to select one culture solution of
an unknown organism.
3. Make a note of the identification code on the
unknown container. Keep the solution closed
until it is time to use it.
4. Working with microbial cultures requires the use
of aseptic technique to prevent the contamination
of both the laboratory as well as its personnel.
5. All materials and media used for the growth of
microbes must be sterilized prior to use.

155. 6. While working with the cultures, the spreaders,
inoculation loops and other materials must be
kept sterile by flaming them both before and
after their use.
7. Culture tubes must be flamed when opened and
also prior to closing.
8. Observe the location of the Bunsen burner on
your lab table. You will use the burner flame to
sterilize the opening of your unknown culture
tube, the glass spreader and the wire inoculation
loop.
9. Mark the outer bottom cover of the Petri dish
(use tape or grease pencil) with your name.

156. • Petri dishes must be stored upside down (agar
hanging from the small lid) in the incubator, to
prevent moisture from washing away the
organisms growing on the surface of the
nutrient agar.
Figure ( ) Petri dishes must be stored upside down (agar hanging from the
small lid) in the incubator, to prevent moisture from washing away the
organisms growing on the surface of the nutrient agar

157. • Use this procedure for preparing a growth plate of
the unknown solution by means of the spreader
method:
1. Turn the Petri dish right side up,
2. Open the unknown culture tube, and flame its
opening,
3. Open the lid of the Petri dish only part way: just
enough so that you can pour the unknown on the
agar surface; make a puddle a little smaller than the
size of a dime,
4. Close the Petri dish,
5. Flame the opening of the unknown culture tube and
close it, then
6. Take the glass elbow (called a spreader) from its
container of alcohol, tapping as much alcohol as
possible off its surface against the inside wall of its
container,

158. 6. Carefully flame the elbow and hold it until it cools
slightly,
7. Open the Petri dish just enough to admit the glass
elbow,
8. Use the sterile spreader to spread the food solution
evenly over the surface of the Petri dish,
9. Close the Petri dish,
10. Reflame the glass elbow, let it cool, and return it to
the alcohol solution,
11. Secure the Petri dish with several pieces of tape and
Place upside-down (agar hanging) Petri dish in
incubator.
• Next session you will look at the growth of colonies on
the surface of the plate to see if your sample was
contaminated.

159. • EXERCISE )2(
1. Observe the sample plates of Escherichia coli, Serratia
marcescens and Micrococcus luteus.
2. Compare the size, shape, height, color, and other
features of their colonies and record your observations.
3. Remember! Each colony is a group of many hundreds to
thousands of individual organisms.
• EXERCISE (3(
1. Each member of a two-person team needs to obtain
another clean, closed Petri dish that contains nutrient
agar.
2. Each team needs to use the same unknown culture that
they used for procedure #1.
3. Keep the solution closed until it is time to use it.

160. 4. Again observe the location of the Bunsen burner
on your lab table.
5. Once again use the burner flame to sterilize the
opening of your unknown culture tube when
you are ready to open it, and before you close it
again.
6. You must also remember to flame the
inoculation loop before and after its use.
7. Be certain to remember to mark the outer
bottom cover of the Petri dish (use tape or
grease pencil) with your name.

161. 4. Recall that Petri dishes must be stored upside
down (agar hanging) in the incubator, to prevent
moisture from washing away the organisms
growing on the surface of the nutrient agar.
5. You will now use an alternative method for
preparing a bacterial growth plate: the streak
method.
• Turn the Petri dish right side up,
• Open the unknown culture, and flame the
opening,
• Flame the inoculation loop and let it cool until the
red color disappears,

162. • Place the loop end of the inoculation wire into the
unknown culture,
• Withdraw the inoculation loop,
• Flame the opening of the tube,
• Close the tube,
• Carefully streak the inoculation loop across the
agar using the pattern shown below (NOTE: DO
NOT break the surface of the agar),
• Close the Petri dish,
• Flame the loop,
• Tape the Petri dish shut and
• Place the Petri dish upside down in the incubator.