Chapter 4 TYPES OF MICROORGANISMS MEDIA

Dr. Yousef Elshrek

• 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.

• 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.

• 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.

• 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.

• 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).

• 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.

• 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.

• 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.

• 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.

• 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.

• 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

• 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.:

• 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.

• 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

• 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

• 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.

• 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.

• 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.

• 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

• 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

• 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)

• 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.

• 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.

• 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

• MACCONKEY AS SELECTIVE

•MACCONKEY AS DIFFERENTIATE

•Lactose positive

•Lactose negative

• 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.

• 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.

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.

• 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.

• MANNITOL SALT AGAR (MSA)

SELECTIVE MEDIA

• Selective for staphylococci

MANNITOL SALT AGAR (MSA) DIFFRENTIATE MEDIA

Differentiate between pathogenic and non pathogenic

staphylococci

• 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.

• 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.

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%

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.

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.

• 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.

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.

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

• 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)

• 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.

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.

• PEA AGAR 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

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.

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.

• 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.

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).

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).

• 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.

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.

• 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.

• 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.

• 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.

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.

• 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.

• 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.

• 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.

• 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.

• 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

• 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

• 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.

• 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.

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.

• 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.

• 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.

• 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.

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:

• 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.

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

• 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.

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.

• 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

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.

• 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.

• 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.

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.

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.

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

• 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.

• 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.

• 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.

• 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

• 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.)

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

• 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.

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

• 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.

• 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.

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.

• 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.

• 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

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.

• 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.

• 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.

• 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).

• 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

• 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.

• 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

• 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.

• 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.

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.

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.

• 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

• 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

• 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.

• 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.

• 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.

• 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.

• 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.

• 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.

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.

• 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

• 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 .

• 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.

• 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.

• 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

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.

• 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.

• 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.

• 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.

• 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.

• 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.

• 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.

• 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).

• 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

• 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.

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

• 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.

• 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

• 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

• FORM OF COLONIES

• Circular.

• Irregular.

• Filamentous.

• Rhiziod.

• ELEVATION

• Raised

• Convex.

• Flat.

• Unbonate.

• Crateriform.

• MARGIN

• Entire.

• Undulate.

• Filiform.

• Curled

• Lobate

• 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.

• 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.

• 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.

• 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

• 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

• 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.

• 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

• 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.

• 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%.

• 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?

Plate # 10-6 10-7 10-8

1

2

3

Average

RESULTS:

Dilutions

Number of colony-forming units per ml ___________

• 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.

• 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.

• 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.

• 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.

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.

• 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

• 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,

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.

• 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.

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.

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,

• 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.