FOS 4222 FOOD MICROBIOLOGY

LABORATORY MANNUAL

SPRING SEMESTER, 2008

M-W 1:00-3:50

Dr. ANITA WRIGHT Room 214 Aquatic Food Products Building

Phone: 392-1991, Ext. 311 Lab Ext. 312

Email: acw@ufl.edu

Website: http://fshn.ifas.ufl.edu/faculty/ACWright/FOS4222.html

Teaching Assistants: Dr. Venata Vedum-Mai and Melissa Jones

FFOOOODD

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TABLE OF CONTENTS

LAB EXERCISES PAGE

Section 1:

LAB EXERCISE 1: INTRODUCTION 3

LAB EXERCISE 2: MEDIA 6

LAB EXERCISE 3: MICROSCOPY 11

LAB EXERCISE 4: ENUMERATION 17

LAB EXERCISE 5: MPN 21

LAB EXERCISE 6: E. COLI 24

LAB EXERCISE 7: FUNGI 27

LAB EXERCISE 8: SALMONELLA 29

Section 2:

LAB EXERCISE 9: IMMUNOASSAYS 34

LAB EXERCISE 10: DNA PROBE 38

LAB EXERCISE 11: PCR 41

LAB EXERCISE 12: MOLECULAR TYPING 46

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LAB EXERCISE 1: INTRODUCTION TO THE LABORATORY

OVERVIEW: These lab exercises in this mannual are designed to supplement your course work

and will chronologically follow (for the most part) the sections you are studying in class. They

should provide practical “hands on experience” with standard microbiology, as well as recently

developed rapid methods and molecular techniques. Basic principles of experimental controls,

data presentation, and interpretation will be explored.

Two very important issues in microbial science are safety and data handling. We will be working

with appropriate precautions and equipment for pathogens, but remember ALL bacteria are

potential pathogens for some people or at some doses. If you are not sure about a procedure:

ALWAYS ASK!

Lab Safety Rules:

1. Absolutely no food or drink

2. No mouth pipetting

3. Always wear gloves and lab coat

4. Report any spills/contamination

5. Disinfect area before and after

6. Dispose of gloves and all contaminated materials in biohazard

7. Breakable or sharp or sharp objects are disposed to sharps container

8. WASH HANDS BEFORE LEAVING!

Failure to respect safety rules will result in expulsion from lab.

Lab Safety Assignment:

1. Fill out Safety form

2. Locate fire extinguisher

Grading:

Lab notebook with answers to study questions provided for each lab (20%)

Lab report (20%)

Quizzes on current lab (10%) Note: You are required to read lab assignments before class.

Two Exams (25% each = 50% total)

Be Prepared! Lab quiz will be given prior to each lab based on the material that will be

covered that day.

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Lab notebook: Laboratory data are the backbone of science, and your records should be

accurate and precise. Scientific papers are generally composed of a number of experiments.

Your notes should be sufficient such that you can later examine them and comprehend what,

why and how you did something. Data should be recorded such that you (or anyone else) will

be able to repeat the experiment from the description in your notebook. Some laboratory

work, especially where data may be patentable or subject to government regulation, will

require bound notebooks to be dated, signed and witnessed daily. All pages must be written

in ink on sequentially numbered pages, and copies made on a regular basis. In practice, raw

data is generally recorded outside of the notebook and then entered later (but not too much

later). Pictures, tables, and other tabulations are permanently attached and are not removed

from the notebook. Multiple notebooks are used when working on independent projects. The

following elements should be present the recording of all experiments:

1. Date and Title of experiment.

2. Statement of purpose: Brief description of rational for experiment

3. Methods: Detailed step by step outline of experiment to include all variables and

controls. Include anything that might be relevant: source of reagents, strains,

temperatures, phase of the moon, etc. The object of this section is to provide a history of

what you have done so that you or anybody else should be able to read and follow the

protocol and reproduce the data. This section should be written before beginning any

experiment. If you are following the protocol from the lab manual, you can cut and paste

this from the lab manual. If there are deviations to the protocol that occur, such as a tube

that was omitted or the wrong incubation temp., etc., record those in your results. Once a

standard protocol has been established it can be referred to by a reference name or

number

4. Results: Record everything! Including negative data, screw ups, changes in protocol,

contamination. You do not know what might become important later. For example if you

are doing plate counts, include the actual number of colonies at each dilution. Making

tables ahead of time in your notebook is useful so you do not forget to record anything.

5. Data analysis: This section should be well-labeled so that it is easy to comprehend. For

example, if you have been using a code for your strains (#1-10), your table or chart

should be converted back to the strain names so it does not have to be deciphered every

time you go back to it. Publishable data requires statistics, minimally mean, Standard

deviation, p values.

6. Discuss you interpretation of results. This section should record any thoughts you have

about the meaning of the data. Discuss how the data compares to expected results based

on prior data or your own or in the literature. Are comparisons of strains statistically

valid. Include implications for future experiments.

7. Provide answers to study question in your notebook.

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Lab report: Lab report will cover the last two laboratory sections on PCR and molecular tying.

A 4-6 page (double space) lab report will discuss the methods and results of these experiments.

Send in word format to Maria (chatz@ufl.edu).

1. Statement of Purpose (1-2 pages): brief description of the purpose of the experiment.

Should include some background on the topic and a rationale for the particular

variables you are investigating. State hypothesis if possible. Direct quotes use

quotation marks and any outside supporting material must be referenced (use ASM

format).

2. Methods Description (1 page): Describe protocol in your own words so that someone

else could do the experiment. Divide in to sections such as the following: Bacteria,

culture conditions, biofilm assay. Include reagents and manufacturer if known.

3. Results (1-2 pages): Describe results in writing without interpreting but do not draw

any conclusions. Prepare tables or graphs to incorporate raw data and analysis as

needed.

4. Discussion. Conclusions (1 page): Discuss what the results and draw conclusions. Did

you prove your hypothesis? Why or why not? How do your results compare to other

data in the literature? What future experiments would you recommend?

5. References: Two required

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LAB EXERCISE 2: MEDIA

Isolation and cultivation of specific microorganisms is essential to the study of microbiology. In

the environmental food, bacteria exist in complex communities that are extremely diverse and

are rapidly altered with changing conditions. Robert Koch is credited with the concept of a “pure

culture”, which allowed microbiologists to characterize a particular organism independently of

related flora. This development was essential to the study of infectious disease and to

establishing “Koch’s Postulates. His first observation of bacterial colonies on solid medium was

on a potato. Later agar was incorporated into broth medium to provide solid support and the

addition of a variety of components into the medium. The method for inoculation of bacteria

varies with the target organism and the matrix from which the organism is isolated. Several

methods are listed below:

Agar Inoculation:

1. Spread plating: Liquid sample (usually 100 µl) on agar plate is spread evenly across

the surface using a sterile hockey stick.

2. Pour plating: Liquid sample (usually 1 ml) is pipetted into an empty sterile petri dish,

and liquid agar (at 50°C) is pored into dish and swirled to distribute the sample evenly

throughout the agar.

3. Streak plating: Method for isolation of individual colonies. Using sterile loop (flamed)

streaking sample across one edge of the plate two or three times. Re-flame and make

another two or three parallel streaks by drawing the loop across the initial streaks once

at a right angle. Flame the loop and repeat as before, this time pulling the loop through

the second set of streaks.

4. Stabbing: Inoculate by stabbing strait down the center of the media in slant tube.

Ensure that the stab is as straight as possible to help prevent misinterpretation of

bacteria spreading

Selective/differential media were developed to provide the ability to differentiate among

different genera or even species of bacteria. These agars are generally used for “presumptive”

identification and require further confirmatory assays for positive identification. The use of these

media requires application of appropriate microbial controls to ensure proper media reaction.

Both a positive control and a negative are required. Most bacterial culture media resemble the

natural substrate on which the microorganisms usually grow. The content of a particular medium

will depend upon then nutrient requirements of the microorganisms of interest. There are several

different forms of media, which include liquid or broth medium and solid or semisolid agars.

Broth medium contains nutrients dissolved in water and is used often as enrichment for more

robust growth of bacteria. Solid medium contains nutrients in water and a solidifying agent. The

most common solidifying agent is agar, which is a complex polysaccharide composed of

galactose and galacturonic acid. It is found in many marine plants and is extracted commercially

from certain marine algae. Agar is resistant to enzymatic hydrolysis by most microorganisms;

however, a few species, mainly in the marine environment, do hydrolyze agar. Silica gel (from

inorganic chemical silicic acid) can be used in place of agar as a solidifying agent and is also

used when microorganisms need to be cultivated on media that are inorganic matter free.

Bacteria generally grow in colonies on solid media, but slant tubes can be inoculated by stabbing

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bacteria into the agar in order to observe chemical reactions, such as for the production of acid or

H2S. Semisolid media have a lower concentration of solidifying agent, which results in a more

jellylike consistency, and is used for motility or gelatin-hydrolysis testing of bacteria. The

nutritional requirements of bacteria vary widely. For example, autotrophs may require only

simple inorganic substrates while more fastidious bacteria (including most human pathogens)

require complex vitamins and growth factors just to survive and multiply. Necessary nutrients to

support bacterial growth always include the following: a) Water, b) Nitrogen (inorganic salts,

peptides, proteins, amino acids); c) Energy or carbon source (carbohydrates, peptides, proteins,

amino acids) ; d) Accessory growth factors (i.e., blood for pathogens, milk for milk

microorganisms).

Food microbiologist use media to detect and differentiate microorganisms from the food we

consume. The types of media are as varied as the organisms that they target. General categories

of media include the following:

1. Non-Selective Media. Multipurpose media that incorporate a wide variety of nutrients and

growth factors for general cultivation of most bacteria.

2. Enrichment Media. Microorganisms to be analyzed are often outnumbered or out-

competed by other microbes (especially in food, water, and soil). Enrichment media

increase the number of target microorganisms while suppressing the growth of competing

microflora.

3. Elective Media. Satisfy the minimum nutritional requirements and are particularly useful

if the microorganism has unusual nutritional requirements. “Wild” yeast cells can be

isolated by using lysine agar on which other microorganisms can not grow unless they

utilize lysine as their sole source of nutrition.

4. Selective Media. Generally support the growth of several species, but also include

inhibitory agents which restrict the growth of undesired microorganisms.

5. Differential Media. Contains reagents (usually dyes) that react with pH changes or other

metabolic consequences of growth to produce reactions that are indicative of particular

species. Colonies will have species-specific characteristics which are easily recognized. For

example, haemolytic and non-haemolytic microorganisms can be easily distinguished by

examining for zones of lysis formed on blood agar plates. Unfortunately these reactions are

not always 100% reliable and generally further tests are required for identification.

Note: Media for food microbiology are commonly both selective and differential.

Some media can be purchased already prepared, but proper preparation of most media is an

essential skill in the food laboratory. The following criteria apply to all reagents and media used

in microbial analysis:

Preparation criteria:

1. Reagents and chemicals must be of suitable grade and composition.

2. Purchase smaller 500g quantities to ensure minimal exposure to atmosphere.

3. Store media in a clean, dry, cool place away from sun (no longer than two years).

4. List dates of receipt and first opened on each container.

5. Solutions should be prepared using laboratory pure (distilled or deionized) water.

6. Use borosilicate glassware or other non-reactive vessels.

7. Weigh media to the nearest 0.1 gram.

8. Stir continuously with heat to prevent scorching and provide an even mixture.

9. Check pH before and after sterilization.

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10. Clearly label all media preparations accordingly before sterilization.

Sterilization criteria:

1. Media with sugars should be filtered or autoclaved for < 10 minutes.

2. Total heat exposure should not exceed 45 minutes for any medium.

3. Adequately space items for proper sterilization to take place.

4. Indicator strips are run with each autoclave operation to ensure proper sterilization.

5. Media should be removed immediately.

6. Sterilize rehydrated media within 2 hours (30 minutes optimal) of preparation.

7. Media is never re-autoclaved.

8. Maintained sterilization records (batch number, date, etc).

9. Cool in water bath between 46-50oC for not more than 4 hours.

10. Discard Durham fermentation tubes with any air bubbles.

Storage of Reagents and Media:

1. Store prepared media in clean, dry areas without light exposure.

2. Store media containing sugars at room temperature for not more than 7 days.

3. Store prepared media under refrigeration (generally not to exceed 4 weeks).

4. Refrigerated media brought to room temperature 4 hours prior to use.

5. Tubes with water loss should be discarded.

Media should be checked for contamination prior to use.

Cultures can fail to grow or produce unexpected or invalid results when reagents are prepared

incorrectly. Common errors include following:

1. Incorrect weighing of reagents or measuring of water volume.

2. Deterioration of media because of exposure to heat, moisture, or oxidation.

3. Water impurities such as heavy metals, pesticides or chlorine.

4. Contamination of glassware with detergents or chemicals.

5. Incomplete mixing.

6. Overheating of some media can produce caramelization of sugars or breakdown of

indicator dyes.

7. Improper pH determination leading to non-ideal growth conditions.

Media are often sterilized by autoclaving. An autoclave is a chamber in which steam sterilization

is used to add heat, humidity, and pressure to the chamber. By increasing the pressure within the

chamber the boiling point of liquids is increased and sterilization time is decreased. Autoclave

sterilization requires 121ºC and 15 psi for 15 to 60 min. Media are autoclaved in a container that

will accommodate at least twice the volume to be prepared in order to prevent boiling over.

Procedure:

Day 1: Media Pouring Contest and Autoclave Field Trip.

Media preparation:

1. Measure 5 grams ± 0.1 grams tryptone, 2.5 grams ± 0.1 grams yeast extract, and 5 grams

± 0.1 grams NaCl, using the weigh boats and scales provided in the back room. Carefully

poor it into 1 liter media bottle or flask.

2. Add 500 ml of distilled water.

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3. Tilt the bottle at a 45º angle and drop a stir bar into it.

4. Place the bottle onto the center of the hotplate and allow it to stir sat a slow speed (do not

use the heat) until reagents are in solution.

5. Add 7.5 grams of agar.

6. Briefly stir the mixture and bring to autoclave. Agar will go into solution during

autoclaving. (Some media require heating until agar is dissolved before autoclaving.)

Plate pouring:

1. After autoclaving media is generally cooled to 45°-50°C to prevent condensation of

plates. We will use pre-autoclaved and pre-cooled media that will be provided in the

waterbath.

2. Separate plates into stacks of four (save the plastic sleeves for storage).

3. Pore the warm media (about 50°C) into the plate until the bottom is completely covered.

Pouring media that is too hot will produce excess condensation and water on the plates.

4. Stack up to 20 plates on top of each other and allow to solidify. This helps to prevent

condensation from building up on the lids.

5. Once solid, invert plates so that the agar is on the top and allow to sit overnight. Sitting

overnight allows some of the excess water to evaporate.

6. The next morning place the plates back into the plastic selves, tape shut and label with

the date and name of the media.

7. Incubate 10 uninoculated plates at room temperature to check for sterility at 24 and 48

hours.

8. Refrigerate to store.

Days 1-3: Isolation and selective media.

Materials:

A. L. agar-

This is a non-selective agar that will support growth of most aerobic heterotrophic

bacteria and will serve as a positive control for the viability of the organisms assayed.

B. TCBS agar –

Thiosulfate-citrate-bile salts-sucrose agar is a combination selective and differential

media. Bile salts provide the selective reagent by preventing the growth of most gram

positive strains of bacteria. The fermentation of sucrose by Vibrio cholerae produces

yellow colonies due to a change in pH, while most other Vibrios produce green

colonies since they do not ferment sucrose.

C. EMB agar –

Eosin-methylene blue agar (Levine): Lactose utilization and eosin and methylene

blue dyes permit differentiation among enteric lactose fermentors and nonfermentors

as well as the identification of Escherichia coli. E. coli colonies are blue-black with a

metallic green sheen caused by the large quantity of acid that is produced and the

precipitates of dyes onto the growth’s surface.

D. TSI/LIA agar slants –

Triple sugar-iron (TSI) is designed to differentiate among the different groups of

Enterbacteriaceae, which are capable of fermenting glucose with the production of

acid. TSI slants contain lactose, sucrose, glucose and the acid base indicator phenol

red. Phenol red indicates the production of acid by a change in color of the medium

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from orange-red to yellow. Typical salmonella will have a red top with a yellow butt.

In most cases H2S is produced which will appear as a black precipitate.

Lysine iron agar- Salmonellae will produce an alkaline reaction from the

decarboxylation of lysine to cadaverine. LIA contains brom thymol blue, which turns

yellow with the production of acid. Typical salmonella will have a purple top with a

yellow butt. In most cases, there is also production of H2S, which will appear as a

black precipitate.

E. Motility Agar. Motility can be observed with the microscope, but observations can be

difficult because there is very little contrast between the bacterial cell and its

environment in live, unstained cells. The MOT Agar Motility Test determines

motility when culture growth radiates out from the line of inoculation in semi-solid

medium. Non-motile organisms will grow only on the inoculation line. It is

important to insert and remove the needle in a single, straight line.

1. Streak each of the control strains and unknown bacteria to the different media listed

above and incubate at the 35°C. Label all plates on the bottom and include date, your

initials, organism, and type of media. Record the observed reactions for each species at

24 and 48h. 2. Using the same organisms from above, transfer the bacteria via sterile flamed needle by

inoculating the MOT agar with a single straight line into the tube and incubate at room

temperature. Observe the tubes at 24 and 48 hours. Check the tube for growth (motility)

that radiates out from the inoculation line.

STUDY QUESTIONS:

1. Why is it necessary to avoid condensation on solid medium?

2. What are the advantages of spread plating? Stabbing?

3. What are sources of contamination of microbial media?

4. Why are bile salts used in selective media?

5. What would be the expected results for these strains for media incubated at

room temp? at 4º C?

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LAB EXERCISE 3: THE MICROSCOPE AND STAINING TECHNIQUES

Compound microscope: The compound microscope (Figure 1) consists of two separate lens

systems and is the most widely used in the field of microbiology. The first lens system in the

compound microscope, called the objective, is nearest to the specimen being viewed. The

objective magnifies the

specimen and produces a

real image within the

body tube. The objective

lens system is comprised

of both convex and

concave lenses, which

correct various chromatic

and spherical aberrations

inherent to a simple

convex lens. Typically

there are at least three

objectives: the low-

power objective (16mm),

the high-dry objective

(4mm), and the oil-

immersion objective

(1.8mm). Focal length

of the objective is

determined by the distance (mm) used to produce the real image. The shorter the focal length of

the objective, the shorter will be the working distance between the objective and specimen. The

ocular lens consists of the eyepiece that you look through. The ocular magnifies the real image

in the body tube to produce a virtual image that is seen by the eye while viewing the specimen.

The total magnification can be calculated by multiplying the magnification of the objective by

the magnification of the ocular. The objective magnification is usually found engraved on the

side of the objective mount and the ocular magnification is usually found on the eyepiece.

Resolving power is the ability to show two closely adjacent points as both distinct and separate.

Resolving power is a function of both the wavelength of the light used to view the specimen and

the numerical aperture of the lens system. Therefore, the shorter the wavelength of light, the

smaller will be the structure that is clearly visible. However, decreasing wavelength to increase

resolution is limited due to the narrow spectrum of visible light (400 to 750nm). The greatest

increase in resolution can be achieved by increasing the numerical aperture, which is defined

as the function of the effective diameter of the objective in relation to its focal length and the

refractive index of the space between specimen and the objective. To decrease the refraction

between the slide and objective, immersion oil is used because it has roughly the same refractive

index as the glass slide. This decrease in refraction increases the light rays entering the objective

and results in a greater resolution and clearer image.

Fluorescence microscope: Fluorescence microscopy requires a special type of powerful

illumination source, usually a mercury lamp. The light from the lamp passes through special

colored filters, which only allow light with distinct wavelengths to pass. This narrow band of

light hits the bacterial specimen. Certain compounds in the specimen (either natural compounds

Figure 1. A. Compound Microscope

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or fluorescent stains) capture

the light and reflect it back up

as light with a lower energy. This

reflected light is detected either

with the viewer's eye, or with

sensitive detectors. Because this

type of microscopy uses reflected

light on a dark background, very

small amounts of light (and of your

sample) can be seen. Fluorescent

compounds include natural

compounds such as chlorophyll, as

well as certain DNA-binding dyes

such as ethidium bromide and

DAPI. Sometimes fluorescent stains

are attached to the constant region

of antibodies, to generate very

specific, very sensitive bacterial

tags.

STAINING TECHNIQUES

Bacteria are almost colorless and require staining to increase the contrast in their color with their

surroundings. Stains generally react with the cell wall, not the background. Three advantages to

staining include: 1) Providing contrast between the bacteria and the background to determine cell

morphology; 2) Permitting the study of internal structures, such as the cell wall, vacuoles, or

nuclear bodies; and 3) Allowing higher magnification as a result of the increased contrast.

Simple stain. The Simple Stain is the easiest method, but lacks the power of discrimination seen

with Gram and other stains. Simple staining is routinely used to determine size, shape, and

arrangement of bacterial cells, as well as for direct microscopic cell counts. Typical dyes are

crystal violet, safranin, carbol fuchsin, and methylene blue. These dyes are generally salts that

include a colored ion or chromophore. For example, methylene blue is actually methylene blue

chloride, and the color is derived from the positively charged (cationic) methylene blue ion. The

interior of a bacterial cell has a slight negative charge in medium at neutral pH combines with

the positively charged methylene blue ion. Basic dyes have a positively charged color-bearing

ion, while an acidic dye will be negatively charged.

Gram stain: Hans Christian Joachim Gram, a Danish physician invented the Gram Stain method

in 1884. The Gram Stain is a differential stain that permits the division of bacterial species into

two distinct groups: gram positive and gram negative. The two cell groups differ in permeability

of the cell surface layers, and in cell wall composition. Gram positive cell walls have much more

extensive peptidoglycan layers than do gram negative organisms, while gram positives lack the

double membrane seen with gram negatives. Both gram-positive and gram-negative bacteria take

up crystal violet and iodine, but the crystal violet-iodine complex is trapped inside the gram-

positive cell by the dehydration and reduced porosity of the thick peptidoglycan layer. Cell walls

Figure 2. Fluorescence Microscope

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of gram-negative bacteria contain more lipids than gram-positive cell walls, and are more soluble

in alcohol and acetone. It is theorized that these lipids are removed from the cell walls during the

decolorizing step or that due to their higher lipid content, the gram-negative cells may contain

fewer acidic reactive sites for binding with the basic dye.

Four different solutions are used in a Gram stain:

1. The basic dye (crystal violet) stains

bacterial cells with a purple blue color.

2. The mordant (Iodine) increases the affinity

between the bacterial cell and the dye.

Acids, bases, and metallic salts are examples

of a mordant. A mordant complex is

required to prevent certain cells from losing

the stain during the decolorization step.

3. The decolorizing agent (95% ethanol)

removes the dye from some of the stained

cell. Cells with thick peptidoglycan layers

will resist destaining and remain blue.

4. The counterstain (safranin) is another basic

dye but of a different color from the initial

stain and is used to stain the decolorized

cells. The counterstained cells will appear

red or pink.

Stained cells that retain the basic dye following decolorization are termed gram-positive, while

those that are decolorized are termed gram-negative.

Spore stain: Species of the genera Bacillus and Clostridium produce a highly resistant body

called an endospore (a cell with a tough spore coat). The endospore provides the bacteria with

the capability to survive long periods of time in high temperatures or toxic chemicals.

Endospores are also resistant to standard staining techniques, and heat must be applied along

with a suitable stain in order to stain the endospore. Malachite green dye used in conjunction

with steam is a common spore stain and is not removed from the endospore by washing. Safranin

counterstain stain is then applied to stain the cell interior light red and contrast with spore coat

green stain. The spore stain can also be used to visualize fungal endospores from molds and

yeast, which are a diverse group of heterotrophic organisms comprising the higher fungi. Many

are saprophytes and digest dead organic matter and waste, while others are parasitic and obtain

their nutrients from the tissues of the other organisms. Most fungi, such as the molds, are

multicellular, but yeasts are unicellular.

Fluorescent staining: Chromofluors are chemicals that adsorb light at one wavelength to

produce excited electrons, that emit light at another wavelength = fluorescence. To visualize

these stains a fluorescent microscope is required, which will illuminate with ultraviolet light to

produce the excitation wavelength required for the chromofluor emission. Filters are used to

isolate emitted light at a particular wavelength and separate it from other wavelengths traveling

to the ocular lens. A dark field condenser is used to create dark background. Chromofluors can

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be used to directly stain molecules within the cells, such as DNA or RNA, or they may be

coupled to other probe molecules (usually antibodies) that bind specific components of the cell.

DAY 1: GRAM AND SPORE STAIN. In this laboratory we will examine three bacterial

species using differential staining procedures which will include the Gram and spore stains. A

demonstration of fluorescent microscopy will also be presented.

IMPORTANT: Notes on the use of the microscope

� Never dust a lens by blowing on it as harmful saliva will be deposited on lenses.

� Use only lens paper to clean after use - Never use facial tissues to clean lenses.

� Avoid touching lenses. Light fingerprints can seriously degrade image quality.

� Use proper immersion oil on immersion objectives as specified by manufacturer. Avoid

getting immersion liquid on non-immersion objectives.

� Keep microscopes covered when not in use.

Procedures:

A. Preparation and Fixation of Bacteria.

1. With a wire loop, place a small drop of each of the three bacterial suspensions on a

clean slide. (see Figure below)

2. For multiple samples use grease pen to prepare individual circles for each sample.

3. Spread the drop on the slide to form a thin film.

4. Allow film layer on slide to dry by holding high above (film side up) a Bunsen flame

(Don’t over heat!).

5. When film is dry, pass the slide three times through Bunsen flame. This is called heat

fixing. The purpose of heat fixing is to kill the microorganism, coagulate the

protoplasm of the cell, and cause it to adhere to the slide.

B. Gram Stain Procedure

1. Cover the entire heat fixed smear with crystal violet dye and let stand for 30 seconds.

2. Tilt slide so as to drain off excess crystal violet to run off into sink and rinse very

gently for 1-3 seconds with deionized water, drain.

3. Cover the smear with iodine solution (mordant) and let stand for 30 seconds.

4. Tilt slide to drain off excess iodine solution and rinse gently with deionized water.

5. Dry slide between blotting paper.

6. Flood slide with ethanol (decolorizing solution) for 10-20 seconds.

7. Rinse ethanol from slide with deionized water to stop the decolorizing process.

8. Cover the smear for 20 seconds with safranin counterstain.

9. Wash slide with running water, gently blot dry, and allow to air dry.

10. Examine the stained preparation using oil immersion objective.

11. Examine and make drawings the stained microorganisms for the following:

• Size and shape

• Arrangement (singly, pairs, chains, clusters, etc.)

• Pleomorphisms (the same organism having more than one form)

• Gram-positive vs. gram-negative coloration

Important note: The Gram stain should not be taken as an absolute indicator of gram-positive

or gram-negative because, in certain instances, the Gram stain will not give the characteristic or

expected reaction. The Gram stain is based on how quickly cells lose the crystal violet-iodine

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complex during the decolorization step. It is possible for Gram-positives to appear Gram-

negative reaction (or vise versa) under certain conditions. Factors that could possibly affect

Gram stain reaction include the following:

1. Overheating the slide during the fixation step causing cells to burst.

2. Too many cells: The higher the number of cells the longer the decolorization takes or it

may be incomplete altogether. This would result in the gram-negative cells appearing to

be gram-positive.

3. The extent to which the smear is washed. Excess dye remaining from too little washing or

over-decolorization if over washing occurs.

4. The age of the culture suspension from which the smear is made.

C. Spore stain procedure:

1. Heat fix bacterial and fungal suspension as described above in Part I.

2. Cover smear with malachite green and place over boiling water for five minutes.

(Keep smear saturated by adding additional malachite green if stain boils off.)

3. After 5 min, remove slide and cool. Gently wash with deionized water for 20 sec.

4. Counterstain with safranin dye for 30 seconds.

5. Gently wash with deionized water and blot dry.

6. Examine the stained endospore preparation using oil immersion objective.

7. Make drawings of the stained microorganism for the following:

• Size and shape

• Arrangement

• Polymorphism

• Presence of spores

16

STUDY QUESTIONS:

1. Diagram the difference in the Gram negative and Gram positive cell walls and explain how

these difference contribute to the results of the gram stain.

2. Why would you see differences in morphology of cells within a pure culture?

3. Based on your previous knowledge of microbiology, which strain would you identify as

Escherichia coli, Bacillus subtilus, or Staphylococcus aureus?

4. What other test would you suggest for confirmed identification of species?

17

LAB EXERCISE 4: ENUMERATION OF MICROORGANISMS

In order to determine food product safety the total number of bacteria within a sample is

frequently determined. This laboratory will examine advantages and disadvantages of two

methods for enumerating microorganisms in foods: Direct Microscopic Count (DMC) and

Standard Plate Counts (SPC).

Direct microscopic count is also referred to as the Petroff-Hausser method and uses slides

embedded with grids of etched squares that contain a specified volume of microorganisms to be

counted. The etched area is covered with a glass slip at a fixed distance from the etched surface,

and bacteria are counted using the high dry (40X) objective of the light microscope. This method

is only accurate for food samples containing a large number of microorganisms (4 x 106 to 2 x

107 bacteria/ml). The major sources of error are inaccuracies in diluting samples and in filling the

etched chambers. Also food particles mask the microorganisms and lead to an underestimation.

The DMC has been used most extensively with milk. A measured volume of milk is spread over

the surface of the area over the etched grid, dried, stained, and viewed under oil-immersion. To

determine the number of bacteria per field, clusters, clumps, or chains are counted as one

bacterial cell because each cluster would give rise to a single colony if plated on solid medium.

Limitations of this method include the inability to determine low concentrations of cells, to count

motile cells, and to distinguish viable and non-viable microorganisms (unless the method is

combined with the use of vital fluorescent dyes described below for viability determination).

Advantages are that results are obtained rapidly and inexpensively with relatively simple

equipment.

Standard plate counts. Bacteria may also be enumerated by either spread or pour plating to solid

media for standard plate counts. Plating to agar media allows determination of viability, as

numbers of bacteria are calculated by the number of colony forming units (CFUs). Samples are

serially diluted and dilutions are applied to agar plates. For spread plates samples are spread over

the agar surface with a glass rod, while the pour plate method uses molten agar mixed with

sample and poured onto the agar surface. With the SPC procedure we assume that a single cell

will give rise to a separate colony; however, this method may not always measure the actual

total number of organisms. Cells in pairs or clusters can still produce a single colony and thus

underestimate actual number of bacteria. Also, not all strains of microorganisms will always

grow on all media under all conditions. Bacteria that are stressed or injured may become

noncultureable and require special recovery media or specific environmental conditions to grow.

It is important to remember that determining colony counts from a food product will depend on

the growth characteristics of organism itself. Some species may spread or swarm over the plate

and present difficulties in determining a single CFU. Swarming is the coordinated migration of

multicellular colonies that results in the production of non-separate colonies. Therefore, the plate

count technique is usually an estimation of the actual number of living bacteria in a sample. The

counts obtained by these methods should not be reported as total viable cell counts but rather as

CFU per unit of sample. Although there are inherent inaccuracies in this method, the plate

count is still the most accurate and widely used method for enumerating bacteria.

Rules for Counting Colonies on Plates and Recording Data: To calculate the Aerobic Plate

Count, multiply the total number of colonies counted by the reciprocal of the dilution factor. The

18

dilution factor is simply the amount of sample transferred over the total volume of the sample

from which it was taken.

1. How many to count. To obtain the Aerobic Plate Count, count duplicate plates from dilutions

that produce 25 to 300 colonies and average the two counts. Use a Quebec colony counter

equipped with magnification and a guide plate ruled in cm2.

2.Consecutive dilutions. If plates from two consecutive serial dilutions yield 25 to 250 colonies,

compute the count per milliliter for each dilution by multiplying the number of colonies per plate

by the dilution used. Report the arithmetic average as the Aerobic Plate Count per milliliter,

unless the higher computed count is more than twice the lower one. If this is the case report the

lower computed count as the Aerobic Plate Count per milliliter or per gram, as applicable.

3. No plate with 25 to 250 colonies. When number of CFU per plate exceeds 250, for all

dilutions, record the counts as too numerous to count (TNTC) for all but the plate closest to 250,

and count CFU in those portions of the plate that are representative of colony distribution. Mark

calculated APC with EAPC to denote that it was estimated aerobic plate count from counts

outside of 25-250 per plate range.

4. All plate with fewer than 25 colonies. If plates from all dilutions yield fewer than 25

colonies each, record the actual number of colonies on the lowest dilution and report the count as

the Estimated Aerobic Plate Count per milliliter or per gram.

5. Plates with no colonies. If plates from all dilutions of any sample have no colonies and

inhibitory substances have not been detected, report the count as less than (<) one times the

corresponding lowest dilution. For example, if no colonies appear on the 1:100 dilution, report

the count as “less than 100 (<100) Estimated Aerobic Plate Count” per milliliter or per gram.

6. Spreaders. If spreaders occur on the plate(s) selected, count colonies on the representative

portions of the plate where colonies are well distributed in spreader free areas. If all the plates

prepared from the original samples have excessive spreader growth or are known to be

contaminated or are otherwise unsatisfactory, report as “Spreaders” (Spr) or “Laboratory

Accident” (LA). Inhibitory substances in a sample may be responsible for the lack of colony

formation.

7. Computing and recording counts. To compute the Aerobic Plate Count, multiply the total

number of colonies or the average number per plate by the reciprocal of the dilution used.

Record the dilutions used, and the number of colonies counted or estimated on each plate. When

colonies on duplicate plates and/or consecutive dilutions are counted and the results are averaged

prior to recording, round off counts to two significant figures (one decimal place) only at the

time of conversion to the Aerobic Plate Count.

Sample preparation is essential to enumeration of bacteria from solid foods. Liquid foods like

milk can be examined directly, but solid food samples are generally homogenized in culture

media or buffer prior to enumeration. The volume of food will depend upon the matrix and

expected level of bacterial contamination. A weighed amount of solid food is blended

mechanically with diluent such as phosphate buffered saline (PBS). Generally 1 to 25 grams of

food are blended with equal volume of liquid in a blender or stomacher. Liquid foods are added

directly to the diluent in measured amounts. Sterile water is not used as diluent because it can

cause cell destruction due to osmotic lysis. Adequate sample mixing and changing pipettes to

avoid carry over between dilutions is essential to prevent miscalculations.

19

Day 1. STANDARD PLATE COUNTS:

A. Serial Dilutions: Before samples are enumerated, they usually need to be diluted or there will

be too many colonies on a plate to count. Decimal or serial 10 fold dilutions are employed

because they are easier to manipulate mathematically (See Figure 1 below). Samples (1ml) are

mixed or vortexed and transferred to dilutions tubes (9 ml) using a fresh sterile pipette with each

transfer. Dilution protocol is shown below. Samples will be plated in duplicate to agar plates in

order to determine CFU/ml.

Figure 1. Serial dilutions

Materials:

Sample culture (log or stationary?)

Phosphate Buffered Saline

Pipettes

Test tubes

Glass spreaders

Alcohol

Quebec colony counter

L agar (LA)

Procedure:

1. For sample, prepare 6 dilution tubes with 9 ml sterile PBS, using aseptic technique. Label

a series of dilution tubes ranging from 10-1

to 10-6

for each sample, with corresponding

plates that are labeled for 10-2

to 10-7

.

2. Use a well-mixed sample so that the test portion represents the entire lot.

3. Using aseptic technique, pipette 1.0 ml of sample into 9-mL PBS dilution tube. Recap

tube and mix the test tube thoroughly for 10 seconds. Using new pipette, apply 0.1-mL of

this dilution to a previously marked Petri dish. Flame spreader and gently spread liquid

around plate. Plate each dilution in duplicate.

4. Using new pipette, continue dilutions from first diluted sample and repeat as above until

the end of the dilution series.

5. Allow plates to dry and then invert and incubate at 35°C for 24 hours.

6. Calculate the number of bacteria from plate counts and convert the CFU/ml to

LogCFU/ml. Average the numbers obtained from all groups and calculate the standard

deviation.

DAY 2: Direct Microscopic Count

In this lab, DMC will be used to directly enumerate the number of bacteria in a samples

derived either from liquid food products. Numbers obtained for all lab groups will be compiled

and used to determine standard deviation. One of the most common counting chambers for DMC

20

is the hemocytometer, also used for counting blood cells. The counting grid is divided into 25

small squares, further divided into 16 smaller ones (Fig. 1). The depth of the hemocytometer

chamber, 0.1 mm differs from Petroff-Hausser counter which is 0.02 mm, and thus calculations

will also differ between methods.

Figure 1. Hemocytometer grid

Materials:

Gloves

Biohazard waste container

Hemacytometer slide and glass cover slip

Microscope

Disposable pipettes and Pipettors

Methyl violet stain

Bacterial culture

Disinfectant

Water bottle

PBS dilution tubes (9 ml)

Procedure

1. Add 1 drop of methyl violet stain to empty test tube

2. Add 1 ml of sample or 1:10 diluted sample to test tubes with dye.

3. Place 10µl of stained cells onto the notch of the grid-etched slide. Cover drop with a

cover glass and read the number of bacteria using the 40X objective.

4. The cells to be counted will appear green with a purple border. If there are more than

100 cells per 1mm large square, count the next dilution of sample.

5. Count the cells in the large 1mm center square containing 25 small squares (See

Figure 1 below). If there is more than one cell per small square, count all the cells in

five small squares (four corners and the center). Regard a clump of cells as one cell.

Multiply number obtained by 5 to estimate the total for 1 mm2.

6. If there is less than one cell per square, count all 25 large squares.

7. Bacteria/ml = the number of bacteria in the 25 squares x 104 x dilution factor.

8. Count both chambers and use the mean of the two counts.

9. Rinse slides with disinfectant and then water to biohazard waste .

STUDY QUESTIONS:

1. How do the two methods of enumeration compare in terms of sensitivity of detection?

2. What does the standard deviation indicate?

3. Do the numbers of cells obtained by DMC indicate they were all viable?

4. Can the food matrix influence the results?

Large central square = 1mm2

with 25 small squares further

subdivided into 16 squares

21

LAB EXERCISE 5: MOST PROBABLE NUMBER AND INDICATOR ORGANISMS

Most probable number (MPN) is a procedure to estimate the population density of viable

microorganisms in a test sample. It applies the theory of probability to positive growth responses

in a standard dilution series or end point titration. Growth of bacteria is obtained in enrichment

broth, which is generally a non-selective medium that encourages the growth of injured or

stressed cells. A positive growth response is indicated by turbidity or gas production in

fermentation tubes. The number of sample dilutions to be prepared is based on the expected

population within the sample. Generally tenfold dilution is used with replicates of 3, 5 or 10 test

tubes for each dilution in the MPN series. When a higher number of tubes are inoculated in the

series, the confidence limits of the MPN are narrowed. Most reliable results occur when all

tubes at the lower dilutions are positive and all tubes at the higher dilutions are negative. For

large microbial populations, the MPN value is generally not as precise as population numbers

derived from direct plating methods, and it should be emphasized that MPN values are only

estimates. MPN values are, however, particularly useful when low concentrations of organisms

(<100/g) are encountered in such materials as milk, food, water and soil or where the matrix may

interfere with obtaining accurate colony counts.

Coliforms. MPN is commonly used to estimate the numbers of coliform bacteria in food. These

organisms are defined as “gram-negative, non-sporeforming, facultative rods that ferment lactose

with acid and gas formation within 48 hrs at 35°C”. These organisms are natural flora of the

intestines of warm blooded animals, including humans. Collectively, coliforms are referred to as

indicator organisms because they indicate the presence of animal or human fecal

contamination. The historical definition of this group has been based on the methods used for

their detection, primarily lactose fermentation. Although E. coli is nearly always found in fresh

fecal pollution from warm-blooded animals, other coliform organisms may be found in the

absence of E. coli. The genera Escherichia, Enterobacter, Klebsiella, and Citrobacter usually

represent the majority of coliform isolates, with Enterobacter the most frequently isolated. It is

important to note that not all coliforms originate from sewage, and E. coli may be more readily

affected by conventional water treatment than other coliforms. Differentiation of coliform types

is valuable in determining the source of increased coliform densities. Large numbers of coliforms

of the same type in water source suggests that multiplication has occurred. Industrial waste

containing high concentrations of bacterial nutrients are capable of promoting growth of

coliforms in effluents and receiving waters. Elevated temperatures (45°C) are needed to

discriminate organisms of fecal origin from others in the coliform group. Thus, incubation at

45°C is used to determine fecal coliform numbers, while total coliforms are determined by

incubation at 35°C.

E. coli detection is considered to be a more accurate measure of fecal contamination and requires

confirmatory assays or the use of media, such as EMB, which is selective and differential for the

species. E. coli LST-MUG broth combines lactose medium (LST) with a substrate of

methylumbelliferone glucose (MUG) to release fluorescent compound 4-methylumbelliferone in

the presence of the enzyme glucuronidase, which is produced by the majority of E. coli (94%)

but not by other coliforms.

22

Day 1: Total and Fecal coliform MPN. In this exercise you will perform coliform MPNs from

a water sample or from samples brought to class. Fecal coliform detection is a simple 24-48 h

test using A-1 medium with a Durham tube. A-1 is a differential medium that uses the

production of gas, from the fermentation of lactose, to indicate the presence of coliforms.

Durham tubes are small tubes that are placed upside down inside larger tube to catch some of the

gas that is produced during lactose fermentation. Samples of different amounts are inoculated

directly into replicate tubes, alternatively samples can be diluted (usually 10 fold) and volumes

of diluted samples can be used to inoculate tubes. In this case we will use 3 replicate tubes at

each of 4 dilutions for a 3-tube MPN with 4 dilutions.

Materials:

A-1 MPN tubes

Whirl-Pak bags

10-ml pipettes

Pipettes and tips

Portable UV light

UV protective goggles

EM agar plates

Wooden applicator sticks

Procedure:

1. Two racks with A-1 Medium with Durham tubes will be provided.

2. Set up and label a three-tube MPN rack with a row for each of four dilutions for the Total

coliform MPN. The first row of three tubes should contain 10 ml of 2X or 1X A-1 media for

liquid samples, depending on the total volume of sample. The remaining three rows of three

tubes should contain 10mL of 1X A-1 media. (Solid samples will all use 10ml of 1X

medium).

3. Shake sample bag (Whirl-Pak) or tube 25 times within 7 seconds in a one-foot arc to mix

sample. Use a well-mixed sample so that the test portion represents the entire lot, dilute

sample 10-1

, 10-2

, 10-3

.

4. Inoculate each of the three tubes in the first row containing either 2X or 1X A-1 media with

either 10 ml or 1 ml of undiluted sample, respectively.

5. Inoculate each of the three tubes in the next rows (containing 1X A-1 media) with 1.0 diluted

water sample.

6. Incubate all tubes at 35 o

C (±0.5oC) for 48 hours for total coliform determination.

7. Repeat the above MPN method for the Fecal Coliform MPN with a second set of A-1 MPN

tubes, and incubate tubes at 35oC (±0.5

oC) for 1 hour.

8. After 1-hour incubation, transfer these tubes to 44.5oC (±0.2

oC).

9. At 24 and 48 hour total incubation time, record gas production as positive. All tubes

exhibiting a positive result from Fecal coliform MPN should be streaked for isolation onto

EMB Medium for a confirmation of E.coli. Incubate plates at 35°C for 24h and record

results.

23

DAYs 2 and 3.

Computation of MPN Results:

1. Write down the number of positive tubes for all dilutions at 24 and 48 hours. To obtain 3

digit number for MPN table, begin with the first dilution that is completely negative or the

last dilution with positives and the next two dilutions with positives. If all have positives,

start with most diluted sample. (as an example: 3-2-0).

2. Most Probable Number (MPN) values are determined from established tables. Look up the

corresponding three-digit number on the MPN table. This number should be reported as an

MPN of Fecal Coliforms per g or ml of sample.

LAB WRITE-UP: 1. How did the numbers compare for the different assays?

2. How did the numbers compare at 24 and 48h?

3. Which numbers would you expect to be higher: total, fecal, or E.coli MPN?

4. How can you adjust the sample size for MPN determinations?

5. What are some limitations of fecal coliform MPN?

24

LAB EXERCISE 6: Escherichia coli

Eschericia coli is a gram negative bacillus in the family Enterobacteriaceae, which can be

classified as diarrheogenic or nondiarrheogenic according to the effects on the human host. The

normal flora of the human intestine harbors nondiarrheogenic E. coli that are considered

relatively harmless to the host. These generally harmless bacteria are frequently used as

indicators of fecal contamination. Most strains will grow under conditions for fecal coliform

analysis, but are differentiated by the following characteristics:

1. Methyl red positive

2. Voges-Proskauer negative

3. Does not use citrate as sole carbon source

4. Indole positive are E. coli type 1 and associated with mammalian intestines

Methods of detection: Typical E. coli assay include the following:

1. IMViC tests: determine indole, methyl red, V-P, citrate characteristics but are

time-consuming

2. MacConkey’s agar: selective/differential agar used to detect coliforms such as E.

coli in milk and water. It contains peptone, bile salts, NaCl, and lactose with an

indicator dye to indicate fermentation. The inhibitory action of bile salts on the

growth of gram-positive organisms allows for the isolation of gram-negative

bacteria. Incorporation of the carbohydrate lactose, and the pH indicator neutral

red permits differentiation of enteric bacteria on the basis of their ability to

ferment lactose. Colonies that ferment lactose will appear pink or red, while non-

lactose fermentors will be colorless. Typical appearances are as follows: E. coli –

red, non-mucoid; Klebsiella- pink, mucoid; Salmonella- colorless.

3. EMB (described in Lab 3)

4. Violet Red Bile Agar: modification of MacConkey’s and includes crystal violet to

inhibit gram positives and produce more red E. coli colonies.

5. MacConkey’s Sorbitol Agar (MSA): uses sorbitol instead of lactose fermentation

as an indicator.

6. MUG ASSAY: uses the substrate 4-methlumbelliferyl PD-glucuronidide (MUG),

which will fluoresce upon enzymatic degradation. Most (93%) E. coli produce the

enzyme B- glucuronidase, which is detected in this fluorogenic assay.

7. E.coli Petrifilm (3M Company): Contain ready-made media enriched with

standard nutrients, a gelling agent, and indicator dyes. Includes Violet Red Bile

(VRB) agar, which detects glucuronidase activity.

Pathogens: Positive results for E. coli may indicate not only fecal contamination but also the

presence of human pathogens. Six groups of E. coli are known as diarrheogenic including

enteropathogenic (EPEC) strains that attach to the brush border of the intestinal epithelial cells

and cause a specific type of cell damage called effacing lesions. Effacing lesions represent

destruction of brush border microvilli adjacent to adhering bacteria. Enterotoxigenic (ETEC)

strains produce two distinct enterotoxins, which are responsible for diarrhea and distinguished by

their heat stability: heat stable enterotoxin (ST) and heat-labile enterotoxin (LT). The

enteroinvasive (EIEC) strains cause diarrhea by penetrating and multiplying within the intestinal

epithelial cells. The enteroaggregative (EaggEC) strains show unique localized regions with a

25

“stacked brick” appearance. The diffuse adhering (DAEC) strains adhere over the entire surface

of the epithelial cells and usually cause disease in immunocompromised or malnourished

children. The enterohemorragic E. coli (EHEC) include E. coli 0157:H7. The first E. coli

0157:H7 outbreak occurred 1982 at fast-food restaurants in Oregon and Michigan. The first case

documented was from a young girl who died from hemolytic uremic syndrome (HUS) as a result

of eating an undercooked cheeseburger. During an investigation, the CDC team traced the

bacteria back through the meat slaughtering and distribution system to the farm, pinpointing its

reservoir in cattle. Entrance into the human food supply is through the contamination of meat and

produce with fecal material. EHEC is considered a major health concern in the US and it is

responsible for an estimated 10,000- 20,000 infections and 250 deaths per year. This dangerous

serotype of ubiquitous and normally harmless genus has the ability to attach to human intestinal

cells and produces potent toxins that produce bloody diarrhea. It may also enter the blood stream

and the renal system, producing high mortality particularly in children. There is no cure for an

infection with this toxin. Although bacteria are killed by antibiotics, toxin is already present by

the time symptoms appear. Thus, antibiotics are not recommended, as they can set the stage for

complications by destroying those harmless bacteria that compete the pathogen. The incubation

period of hemorrhagic colitis usually lasts for 3 to 4 days with a range of approximately 2 to 8

days.

It should be noted that EHEC is not detected in many of the standard tests for E. coli. Nearly

all E. coli (93%) ferment sorbitol with the exception of EHEC, which exhibits clear colonies on

MacConkey sorbitol agar. It also does not produce B- glucuronidase and therefore is negative for

the MUG assay. EHEC is also unique from other E. coli strains in terms of growth

characteristics: Although EHEC grows rapidly between 30-40ºC, it shows slow growth at 44–

44.5ºC. Therefore, it is not be detected in standard screening procedures for fecal coliforms

involving these temperatures ranges. However, EHEC resembles other pathogens, such as

Salmonella, in that it can be acclimated to grow at very low pH (3.7- 4.0), such as that found in

apple juice.

Day 1. The purpose of this lab is to compare alternate methods for E. coli detection. Obtain

sample and bacterial controls from your instructor. Alternatively, water sample or swabbed

surface expected to be positive for E. coli may be used. Determine E. coli counts by 1) SPC by

spread plating to petrrifilm vs. MacConkey’s Sorbitol Agar and by 2) MPN using MUG broth

and streaking to MacConkey’s.

Materials: Petrifilm, MSA, MUG-MPN tubes (10 ml-1X), pipettes, glass spreader, inoculation

loops, burners, alcohol, water bath, and incubators.

Prodedure:

1. Use a well-mixed sample so that the test portion represents the entire lot, dilute sample 10-1

,

10-2

, 10-3

.

2. Distribute samples to MPN-MUG tubes as described in Lab 4 and incubate tubes at 35°C

(±0.5°C) for 1 hour and then transfer these tubes to 44.5°C (±0.2°C) water bath.

3. Spread Plate samples to MacConkey’s in duplicate as described in Lab 4.

4. Place the 3M Petrifilm (Note: Petrifilm should be stored below 8°C) on a flat surface and

dispense 1mL of your sample onto the center of the Petrifilm.

5. Distribute the sample on the Petrifilm in duplicate by applying downward pressure using a

spreader to press the top film over the sample.

26

6. Incubate the Petrifilm plates at 35°C in stacks placed in a horizontal position with the clear

side of the plate up. The stacks should not exceed 20 plates.

Day 2 and 3. Record results at 24 and 48h

1. SVC on MSA: determine logCFU/ml as described in Lab 4.

2. Petrifilm: All of the colored dots indicate a colony and should be counted. The

circular growth area is 20 cm. You may count the number of colonies in a square

area and multiply by 20 to get the total number of colonies.

3. MPN-MUG: Examine the (MUG) test tubes for fluorescence by darkening the

room and placing an ultraviolet light over the tube. (POSSIBLE EYE

DAMAGE!!! WEAR SAFETY GLASSES!). For safety reasons please do not

directly look into the ultraviolet light. Examine the turbidity of the (MUG) test

tubes from the incubator at 44.5°C. Streak all turbid samples after 24h incubation to

MSA and incubate at 35°C.

Record results for SPC and MPN as logCFU/ml and post on board.

Sample

Group

E. coli Numbers

Plate Counts (CFU/ml) MPN (MPN/ml)

Petrifilm SPC- MSA MUG 24h MUG 48h MSA

1

2

3

4

5

6

7

8

9

Reference:

Harrigan, W. F. 1998. Laboratory Methods in Food Microbiology. Academic Press, London

Lab Write-up:

1. Compare results from the different assays and time points.

2. Samples 1-5 and 6-10 are from identical sources respectively and represent duplicate

samples. Calculate MPN using 3 vs. 15 tube replicates. How do these results differ

statistically?

3. What are the advantages of using MPN-MUG assay?

4. How did the initial numbers of E. coli in sample alter the results?

5. Would any of the tests detect EHEC?

27

LAB EXERCISE 7: FUNGI - MOLDS AND YEAST

Molds and Yeast are a diverse group of heterotrophic organisms comprising the higher fungi.

Many are saprophytes, which digest dead organic matter and waste, while others are parasitic

and obtain their nutrients from the tissues of the other organisms. Most fungi, such as the molds,

are multicellular. Yeast are unicellular fungi closely related to the molds. They are ellipsoidal,

spherical, or cylindrical cells, and are several times larger than the average bacterial cell.

Because of their large size, numerous structures and inclusions may be readily seen using a light

microscope.

In the food industry, the presence of molds or yeast may be either beneficial or detrimental.

Molds are used in the manufacture of cheeses such as Blue, Camembert, and Stilton as a ripening

substrate. They are also employed to produce enzymes and acids as in the case of various

Oriental foods such as soy sauce and miso. Yeast are used in the manufacture of a number of

foods through fermentation, such as beer, wine, vinegar, and some cheeses, which are surface

ripened. Yeast, like the molds, are grown for their enzymatic action in foods, but may also serve

a source of single cell protein. A special strain of Saccharomyces cerevisiae, which produces

large amounts of carbon dioxide, is added to bread dough to make it rise.

Mold spoilage is frequently found in nuts and oilseeds and in refrigerated foods such as cheese

and cured meats. Rhizopus nigricans is the most common bread mold, but several other species,

which thrive in bread, are Penicillium, Aspergillus, and Monilla. The contamination of bread by

Monilla, a pink bread mold, is extremely troublesome because it is difficult to eliminate from a

bakery once it has become established. Bacillus species are the most likely to contaminate Rye

bread, which hydrolyze proteins and starch and give the bread a stringy texture. Yeast can cause

spoilage in certain foods such as sauerkraut, pickles, meats, and other foods. Fresh fruit juices,

because of their high sugar and acid content, provide an excellent growth environment for molds

and yeast. Penicillium expansum, which grows on apples, produces a toxin, patulin, that can

contaminate cider. Other molds in foods are capable of producing other toxic compounds called

mycotoxins, of which aflatoxins are an example.

The body of a fungus is called a thallus. The thallus consists of a mass of intertwining branching

threads called mycelium. A single strand of these thread-like structures is called a hypha. Much

of the mycelium of each mold grows within or on the surface of the growth medium and

functions to extract nutrients for its survival and growth. This is termed vegetative mycelium.

Growing above this vegetative mycelium are specialized fruiting structures that produce asexual

or sexual spores.

Molds and yeast are grouped into four classes, based on their mode of sexual reproduction:

1. Phycomyces resembles algae but differ in lacking chlorophyll. This class includes both

aquatic and terrestrial molds in which sexual spores, when produced, are borne exposed.

The growth of molds in this class tend to spread over the entire surface of the culture

plates, appearing not a single colony but as a fibrous coarse mass, whose borders are

limited only by the walls of the petri plate. The vegetative mycelium of the Phycomyces

is coenocytic, or nonseptate (without crosswalls) and consists of a continuous mass of

multinucleate protoplasm within the tube-like cell walls of the mold. The fruiting, stalk-

like structures of the Phycomyces produce sporangiospores, an asexual spore enclosed in

sporangia, or spore case. Less frequently, molds produce sexually by the fusion of two

28

hyphae, which give rise to gametes. From these gametes, sexual spores are produced. In

certain Phycomyces genera, the sexual spores are formed singly and without a covering

and are termed zygospores.

2. The class Ascomycetes are characterized by the formation of sexual spores called

ascospores. These ascospores are produced in a sac –like ascus with several spores being

formed. Some genera of Ascomycetes do, however, produce asexual spores. These spores

are termed condiospores and they are unenclosed. Molds of the class Ascomycetes tend

to form discrete colonies on culture plates.

3. The class Basidiomycetes are the fleshy fungi and bear sexual spores, termed

basidiospores, in highly developed structures called basidia.

4. The class Deuteromycetes (or Fungi Imperfecti) represent a taxonomic “dumping

ground” for those molds and yeast in which no sexual reproduction has been observed.

Within this class are pathogenic fungi such as the genus Trichophyton, which is the cause

of athlete’s foot, and the species Candida albicans, the cause of a throat infection known

as thrush.

Procedure:

Yeast morphology and growth

1. Prepare a suspension of baker’s yeast as described on the back of package. Allow

to grow for about 45 minutes at room temperature.

2. Make a methylene blue stain of the yeast culture and observe by using a

microscope.

3. Make drawings of all observations.

4. Inoculate three tubes of yeast extract sucrose broth with baker’s yeast and

incubate at 50, 20

0 (room temp.) and 43

0C.

Molds morphology Growth

1. Observe all mold cultures for characteristics such as color, texture, reverse color,

2. Make a slide mount of each mold culture. Take a small amount of mold growth

and suspend it in water on a clean slide and cover with coverslip. Starting with

low power magnification, use all three objectives, if necessary, to observe the

following:

• Mycelium

• Spores if present

• Fruiting heads

• Special structures

3. Make drawings of all observations.

4. Select one of the mold cultures and inoculate some spores onto a Potato Dextrose

agar plate and a Czapek’s agar plate (general purpose media for the cultivation of

both molds and yeast). Incubate plates at room temperature in a dark area. Do not

invert mold plates during incubation. Check growth in 5 and 7 days.

5. Divide one Potato Dextrose agar (PDA) plate into three sections. Using one mold

culture, the baker’s yeast and a bacterial culture provided, inoculate each of the

three sections on each plate. Incubate the plates and record results in 2 days and

again at day 5.

29

LAB EXERCISE 8: DETECTION OF SALMONELLA

Salmonella are facultatively anaerobic, Gram-negative, rod-shaped bacteria. Salmonella are

motile by peritrichous flagella, grow on citrate as a sole carbon source, and generally produce

hydrogen sulfide (H2S), although H2S-negative serovars do exist. Well-established culture

methods for the detection of Salmonella in various foods are available from the following:

1. FDA, Bacteriological Analytical Manual (BAM): http://vm.cfsan.fda.gov/~ebam/bam-

toc.html

2. USDA, Microbiological Laboratory Handbook:

http://www.fsis.usda.gov/Science/Microbiological_Lab_Guidebook/index.asp

3. American Public Health Association. 2001. Compendium of Methods for the

Microbiological Examination of Foods. 4th Edition.

4. Culture and rapid methods validated by the AOAC

Currently, there are over 2,500 recognized serovars of Salmonella, however 10 serovars are

responsible for the majority of illness in humans, known as salmonellosis. Salmonella is widely

distributed throughout nature with the primary reservoir being the intestinal track of mammals,

birds and reptiles. Salmonellosis is a communicable disease and is readily transmitted from

animals to humans, but most often is acquired through the ingestion of contaminated food

products. The actual number of cases may be between 400,000 and 4 million/yr. Clinical

symptoms include stomach pain, diarrhea, headache, and fever accompanied by chills. After

ingestion, bacteria reproduce in the small intestine, and onset of illness usually occurs 8 to 48

hours after ingestion. Symptoms can persist for 3 to 5 days. Salmonellosis can be fatal for the

very young, infirm, and those with compromised immune systems. Salmonellosis is usually

contracted from food products that have been abused during handling. Abuse can occur through

exposure to warm temperatures and/or cross-contamination

Media and test kits: The following media are often used for the enrichment/isolation of

Salmonella from food.

Primary enrichment media:

a. Universal pre-enrichment broth (UPB): Used for the recovery of sublethally injured

Salmonella and Listeria from food products. Sodium and potassium phosphate buffers

provide sufficient buffering capacity to prevent the rapid decrease in pH, allowing for the

repair of injured cells which may be sensitive to low pH. UPB also contains peptones,

essential ions, dextrose as an energy source, and sodium pyruvate to aid in the recovery

of stressed cells.

Secondary (selective) enrichment media:

a. Tetrathionate (TT) broth: Tetrathionate and excess thiosulfate suppress coliform

microorganisms and other accompanying bacteria, whereas all tetrathionate-reducing

bacteria (e.g. salmonellae and Proteus) can multiply more or less normally in this

medium. Acidic tetrathionate decomposition products are formed, which are neutralized

by calcium carbonate. TT broth contains bile salts, which largely inhibit all

microorganisms that do not normally live in the intestine. The addition of brilliant green

suppresses, above all, the Gram-positive microbial flora. An iodine solution is also added

to provide an inhibitory effect.

b. Rappaport-Vassiliadis (RV) medium: This medium is a modification of the Salmonella

Enrichment Broth with lower concentrations of malachite green and magnesium chloride

30

to improve the growth of Salmonella at 43 °C. Peptone from soymeal is also used for the

same reason. The pH of RV medium is 5.2 for increased selectivity.

Isolation/plating media:

a. Bismuth sulfite (BS) agar: Brilliant green and bismuth largely inhibit the accompanying

bacterial flora. Colonies of H2S-positive salmonellae exhibit blackening due to the

formation of iron sulfide. Reduction of bismuth ions to metallic bismuth produces a

metallic lustre around the colonies.

b. Xylose lysine desoxycholate (XLD) agar: Degradation of xylose, lactose and sucrose to

acid causes phenol red to change its color to yellow. Production of hydrogen sulfide is

indicated by thiosulfate and iron (III) salt, which react to form a precipitate of black iron

sulfide in the colonies. Bacteria which decarboxylate lysine to cadaverine can be

recognized by the appearance of a purple coloration around the colonies due to an

increase in pH. These reactions can proceed simultaneously or successively, this may

cause the pH indicator to exhibit various shades of color or it may change its color from

yellow to red on prolonged incubation. The culture medium is weakly inhibitory.

c. Hektoen enteric (HE) agar: When compared with other selective culture media (e.g. SS

Agar, BPL Agar and Bismuth Sulfite Agar), HE agar has the advantage that it only

slightly inhibits the growth of Salmonella and Shigella, thus giving high yields of these

microorganisms, but at the same time ensures adequate inhibition of accompanying

microorganisms. Lactose-positive colonies have a clearly different color from lactose-

negative colonies due to the presence of the two indicators bromothymol blue and acidic

fuchsin. This color difference is also observed for colonies, which can only slowly

ferment lactose due to the presence of sucrose and salicin. These reactive compounds can

be fermented more easily - false-positive pathogenic results are thus avoided. The

combination of thiosulfate as a reactive compound with an iron salt as an indicator causes

H2S-positive colonies to become black in color. The mixture of bile salts suppresses the

growth of most of the accompanying microorganisms.

Differential media:

a. Triple sugar-iron (TSI): Designed to differentiate among the different groups of

Enterobacteriaceae, which are capable of fermenting glucose with the production of acid.

TSI slants contain lactose, sucrose, glucose and the acid base indicator phenol red.

Degradation of sugar and accompanying acid production are detected by the pH indicator

phenol red, which changes its color from red-orange to yellow, on alkalinization it turns

deep red. Thiosulfate is reduced to hydrogen sulfide by several species of bacteria, the

hydrogen sulfide reacts with an iron salt to give black iron sulfide.

b. Lysine iron agar (LIA). Lysine is decarboxylated by LDC-positive microorganisms to

give the amine cadaverine, which causes the pH indicator bromocresol purple to change

its color to violet. As decarboxylation only occurs in an acidic medium (below pH 6.0),

the culture medium must first be acidified by glucose fermentation. This medium can

therefore only be used for the differentiation of glucose-fermenting microoganisms.

LDC-negative, glucose-fermenting microoganisms cause the entire culture medium to

turn yellow. On prolonged incubation alkalinization of the culture medium surface may

occur, resulting in a color change to violet. H2S production causes a blackening of the

culture medium due to the formation of iron sulfide.

31

Enterotube II:

The BBL™ Enterotube™ II is a prepared multimedia tube for the rapid identification of

Enterobacteriaceae. Each compartment of the Enterotube™ II tests for a specific

biochemical trait of the microorganism. After incubation, each compartment is observed

for color change, which indicates a positive result. The results are recorded and a score

for each result is tabulated. Using the provided score card, the scores are combined to

form either a 5 or 6 digit code. The resulting code can be used to identify the unknown

microorganism. In some cases, further biochemical testing is needed to differentiate

among several possible identifications.

Reveal for Salmonella:

The Reveal for Salmonella test system is a presumptive qualitative test that detects the

presence of Salmonella organisms in foods within 21 h total testing time. Foods are

enriched with a proprietary resuscitation medium called Revive and then selectively

enriched with either Selenite Cystine or Rappaport-Vassiliadis selective media. The

enriched culture is used to inoculate the detection device, which initiates a lateral flow

through a reagent zone containing anti-Salmonella antibodies conjugated to colloidal gold

particles that capture antigens present in the culture. The antigen-antibody complex

migrates farther and is captured by an additional anti-Salmonella antibody, causing the

colloidal gold to precipitate and form a visual line, indicating a positive result. A

procedural control line also will form regardless of the presence of Salmonella organisms

to indicate the test is working properly.

PROCEDURE: The following procedure is based on the U.S. Food and Drug Administration’s

Bacteriological Analytical Manual (BAM) Online, updated September 2005. A flow chart has

been provided as an overview to the analytical procedure for the isolation of Salmonella.

DAY 1

a) Each group will be given a samples that contain Salmonella positive control, a non-

Salmonella negative control and two unknowns that have been incubated in Universal

pre-enrichment broth (UPB) for 24 h at 35 ± 2.0°C.

b) For each culture: Transfer 1.0 ml of the UPB enrichment to a 9.0 ml tube of tetrathionate

(TT) broth. Transfer 0.1 ml of the UPB enrichment to a 9.9 ml tube of Rappaport-

Vassiliadis (RV) medium.

c) Incubate RV medium 24 ± 2 h at 42 ± 0.2°C (circulating, thermostatically controlled,

water bath). Incubate TT broth 24 ± 2 h at 35 ± 2.0°C.

DAY 2

a) Vortex tubes and streak broths for isolation on bismuth sulfite (BS) agar, xylose lysine

desoxycholate (XLD) agar, and Hektoen enteric (HE) agar. In addition, streak a

MacConkey plate (to be used for the Enterotube II). Incubate plates 24 ± 2 h at 35°C.

b) Save the RV medium for use on DAY 4 at 4°C.

DAY 3

a) Examine BS, XLD, and HE plates for presence of colonies that resemble Salmonella.

32

TYPICAL Salmonella COLONY MORPHOLOGY:

1. Hektoen enteric (HE) agar. Blue-green to blue colonies with or without black

centers. Many cultures of Salmonella may produce colonies with large, glossy

black centers or may appear as almost completely black colonies.

2. Xylose lysine desoxycholate (XLD) agar. Pink colonies with or without black

centers. Many cultures of Salmonella may produce colonies with large, glossy

black centers or may appear as almost completely black colonies.

3. Bismuth sulfite (BS) agar. Brown, gray, or black colonies; sometimes they have

a metallic sheen. Surrounding medium is usually brown at first, but may turn

black in time with increased incubation, producing the so-called halo effect.

ATYPICAL Salmonella COLONY MORPHOLOGY: In the absence of typical or suspicious

Salmonella colonies, search for atypical Salmonella colonies as follows:

1. HE and XLD agars. Atypically a few Salmonella cultures produce yellow

colonies with or without black centers on HE and XLD agars. In the absence of

typical Salmonella colonies on HE or XLD agars after 24 ± 2 h incubation, then

pick 2 or more atypical Salmonella colonies.

2. BS agar. Atypically some strains produce green colonies with little or no

darkening of the surrounding medium. If typical or suspicious colonies are not

present on BS agar after 24 ± 2 h, then do not pick any colonies but reincubate an

additional 24 ± 2 h. If typical or suspicious colonies are not present after 48 ± 2 h

incubation, then pick 2 or more atypical colonies.

b) Pick 2 or more colonies of Salmonella from each selective agar. Lightly touch the very

center of the colony to be picked with sterile inoculating needle and inoculate TSI slant

by streaking slant and stabbing butt. Without flaming, inoculate LIA slant by stabbing

butt twice and then streaking slant. Since lysine decarboxylation reaction is strictly

anaerobic, the LIA slants must have deep butt (4 cm). Store picked selective agar plates

at 5-8°C. Again without flaming, inoculate a tube of tryptic soy broth (TSB).

c) Incubate TSI and LIA slants and TSB at 35°C for 24 ± 2 h. Cap tubes loosely to maintain

aerobic conditions while incubating slants to prevent excessive H2S production.

d) Select an isolated colony from the MAC plate and use to inoculate an Enterotube II as

demonstrated by the instructor. Incubate at 35°C for 24 ± 2 h.

e) Transfer the RV medium (saved from DAY 3) to a fresh RV tube by

DAY 4 a) Examine/interpret the TSI and LIA slants as follows:

a. Salmonella in culture typically produces alkaline (red) slant and acid (yellow) butt,

with or without production of H2S (blackening of agar) in TSI.

b. In LIA, Salmonella typically produces alkaline (purple) reaction in butt of tube.

Consider only distinct yellow in butt of tube as acidic (negative) reaction. Do not

33

eliminate cultures that produce discoloration in butt of tube solely on this basis.

Most Salmonella cultures produce H2S in LIA. Some non- Salmonella cultures

produce a brick-red reaction in LIA slants.

b) Examine and score the Enterotube II. Use the codebook to determine the identity of your

unknown microorganism.

c) Describe your results, including any color changes seen in broth, agar plates and slants.

What do these color changes tell you about the isolated microorganism(s)?

d) What was the identity of your unknown?

e) Were there any differences in the results obtained from enrichment in TT broth versus

enrichment in RV broth? If so, describe those differences and why they might have

occurred

STUDY QUESTIONS:

1. What is the lowest initial Salmonella concentration detected by this FDA protocol?

2. The USDA’s method for evaluating meat products for Salmonella uses a different

medium for primary enrichment. What is the name of this broth and why does the use of

it seem odd? (HINT: What are the biochemical traits of Salmonella?) Explain the

rationale behind using this broth for the enrichment of Salmonella.

3. Are there any other bacteria in the Enterobacteriaceae that can produce typical colonies

on the plating media used in this lab? If so, what are they and how could you determine

if they are Salmonella or not using conventional culture techniques?

34

LAB EXERCISE 9: Immunoassays

Agglutination assay: Antibodies or immunoglobulins (Ig) are proteins produced by the immune

system in response to infectious disease. They are generally specific for a particular organism

and can bind with high affinity to a particular antigen on the cell surface. Common types of

antigens include LPS (O antigen), capsular polysaccharide (K antigen), flagella (H antigen).

These structures contain epitopes (the antibody-binding portion of the antigen) that are specific

to a particular species or strain of bacteria and bind the unique Fab segment of the antibody.

Some antigens also bind less specifically to the tail of Fc portion of the antibody and can detect

all strains of a species. The antigen-antibody binding is commonly used for detection of

foodborne pathgens. One of the simplest tests is the agglutionation assay, which is detected

visulally due to the clumping of bacteria and antibody. An example of an agglutination assay is

the one used for Staphylococcus aureus. These Gram positive, halophilic cocci can be highly

pathogenic and produces a variety of toxins, which can be lethal to humans. It is responsible for

20-50% of all foodborne illness outbreaks in the United States. The bacterium is sensitive to heat

and nearly all sanitizing agents. Therefore, detection of S. aureus in processed foods is an

indication of poor sanitation, and post-processing contamination is usually the result of human

contamination or contamination from unsanitary surfaces. Foods that are suspected to be the

source of “staph” food poisoning are examined for the presence of the organism and/or its

enterotoxins. Microbiological tests for S. arueus include the use of a differential/selective Baird-

Parker medium. Colonies should appear as black, shiny, convex colonies with a narrow white

margin, surrounded by a clear zone.

The BBL Staphyloside Latex Test is an agglutination assay that can be used to detect

bacterial surface structures that are unique to pathogenic S. aureus. One such component is

coagulase enzyme, which the catylses the fomation of fibrin clots from the fibrinogen in plasma.

It is produced as both a soluble and surface-associated form and is also referred to as clumping

factor. Another surface

factor is Protein A,

which is able to bind the

non-specific or Fc end

of IgG antibody

molecules and thus

render them inactive.

The Staphyloside Latex

test consists of blue

latex particles coated

with human fibrinogen

and IgG. Colonies of

staphylococci which

have clumping factor or

Protein A present, will cross linking the latex and produce visible agglutination of latex particles.

The agglutination will occur notably with S. aureus. If neither clumping factor nor Protein A is

present, no agglutination will occur, and the result will be regarded as negative. The reaction is

illustrated here.

35

1.Bind Ab

2.Add Ag

3.Wash Unbound Ag

4.Add labeled Ab

5.Wash unbound Ab

6. Appearance of green color as

increasing concentrations of

enzyme convert substrate,

indicating increased

concentrations of antigen

ELISA assay: Another very powerful immunoassay is the Enzyme-Linked ImmunoSorbent assay

(ELISA). This method is also referred to as a “sandwich” ELISA, and the basic principles are

outlined in this figure.

Basically in Step 1 a high

affinity “capture” antibody

(Ab) that is specific for the

target organisms is absorbed

to the surface of a microtiter

plate. If cells or antigen (Ag)

are present that correspond to

this antibody, they will be

captured by the antibody in

Step 2. In Step 3 bound Ag

will remain and unbound will

be removed during washing.

In order to detect the bound

antigen, a labeled secondary

antibody conjugated to an

enzyme label and also specific

for the antigen completes the sandwich in Step 4. This antibody will also bind the cells or

antigen and all unbound labeled antibody is removed in the final washing in step5. The enzyme

label can then be used for detection of a positive sample. In this assay the enzyme converts the

substrate to a green color. Thus, presence of any potential pathogens will be noted by the

appearance of a green color. The antibodies in this assay may detect all strains within a species

or may be specific for a particular serotype.

Antibody Lateral Flow devices: In these assays, reagents are loaded into one end of the device

and the sample moves through the paper by lateral diffusion (wicking) until it reached the spot

where antibody markers are bound on the

paper. Anitbodies are gold conjugated

and the complex of antibody and antigen

continues to move and bind second

antibody which captures complex. The

captured complex then aggregates and

forms visual indication markers for

positive sample.

Antibody Lateral Flow Devices

• Reagents are load in one end

of the device

• Sample moves through the

paper where antibody

markers are bound.

• Visual indicator marks

presences.

Positive ResultPositive

Control

36

Day 1: Salmonella detection assays

Microbiological detection of Salmonella spp may require days and may not always produce

species-specific results. An alternative method is the enzyme-linked immunosorbent assay

(ELISA) such as the TECRA® Salmonella Visual Immunoassay. Although, kits like TECRA®

are rapid, other kits have been developed that require fewer steps and therefore give results more

rapidly. An example of this is the Visual Immunoprecipitate Assay (VIP)® Salmonella kit.

While these kits are faster, they utilize different ELISA concepts. This kit uses a proprietary

reagent system to form an antigen-antibody-chromogen complex. When the Salmonella antigen

comes in contact with the antibody-chromogen complex, a precipitate forms and causes a color

change, which is indicated by the colored line that appears in the detection window.

Immunological protocols:

Each group will be given two unknown cultures, and results for positive and negative

controls will be provided by instructor. The purpose of this lab will be to identify the

Salmonella spp. IMPORTANT:

Change your pipette tip between reagent bottles. DO NOT use the same tip.

Reveal Lateral Flow Device

1.Using the dropper, place 5 free-flowing drops of each unknown into the sample port on

the Reveal for Salmonella device.

2.Read the results after 15 minutes at room temperature. Make sure that the device is

level and is not disturbed during incubation.

TECRA® Salmonella Visual Immunoassay ELISA

1. Heat the broth cultures for 15min in a boiling water bath. After 15 min vortex the tubes

before use.

2. The four Removawells that are already coated with the capture antibody. One for each

sample, one for positive control and one for the negative control (label them accordingly

on the top of the plastic cling wrap).

3. Using a new pipette tip for each sample add 200µL of the controls and samples into

individual wells. Cover the wells with plastic cling wrap film and incubate for 30 min at

35-37°C.

4. Quickly invert the holder, emptying its contents into a contaminated waste container.

Remove residual liquid by striking the wells firmly several times face down on a piece

of absorbent paper towel. This is important for effective removal of sample residue.

5. Add 0.5 mL of wash solution in each well, taking care not to trap air bubbles in the

bottom of the wells. Wash and completely empty the wells a total of three (3) times as

outlined above.

6. Ensure the Removawells are empty before proceeding.

7. Add 200µL of Conjugate [4] to each well. Cover the holder with plastic cling wrap and

incubate for 30 min at 35-37°C.

8. Empty the wells and wash them thoroughly a total of four (4) times as described in steps

6 and 7.

37

9. Ensure the Removawells are empty before proceeding. 10. Add 200µL of Substrate [6] to each well. Incubate at 20-25°C (room temperature).

11. At 10min the positive control should be equivalent to either Panel 4 on the Color Card.

Results can be read visually using the Color Card.

Day 2 Agglutination assays

Polyvalent Salmonella flagellar (H) test:

1. Add 2.5 ml formalinized physiological saline solution to 5 ml TSB enrichment.

2. Place 0.5 ml of appropriately diluted Salmonella polyvalent flagellar (H) antiserum in a

13 x 100 mm serological test tube.

3. Add 0.5 ml of the TSB enrichment.

4. Prepare saline control by mixing 0.5 ml formalinized physiological saline solution with

0.5 ml formalinized antigen.

5. Incubate the test mixture and the control mixture in a 48-50°C water bath. Observe at 15

min intervals and read final results in 1 h..

Staphylococcus aureus agglutionation assay

Materials:

Baird-Parker medium, plates

BBL Staphyloside Latex Test

Inoculating loops

Procedure:

1. You will be given a positive control (S. aureus), a negative control (E. coli) and two

unknowns grown on L agar.

2. Streak these strains for isolation to the Baird-Parker plates and incubate overnight at

35°C. Observe and record the colony appearance the next day.

3. BBL Staphyloside Latex Test: (Warning: The reagents contain material from human

origin. All reagent materials have been tested for HIV or Hepatitis C, but extreme

care must still be taken when handling all materials. The reagents also contain

azide, which is very toxic by inhalation, in contact with skin, and if swallowed.

Contact with acids liberates very toxic gas. After contact with skin, wash

immediately with plenty of water.)

4. Assay positive and negative control strains and unknown strains using the latex test.

(NOTE: Better results are obtained from non selective media.)

5. Mix the latex reagent by shaking; expel any latex from the dropper.

6. Dispense 1 drop each of Test Latex onto four circles on the reaction card, and add 1 drop

of Control Latex onto another four circles.

7. Using a microbiological loop pick up and smear 5 suspect colonies from each of the

strains onto a Test Latex circle and mix bacteria into the Test Latex reagent.

8. Spread to cover the circle.

9. Repeat step 8-9 with the Control Latex for each of the strains.

10. Pick up and gently rock the card for 20 seconds and observe and record agglutination

reactions under normal lighting conditions.

11. Dispose of the reaction cards in the orange biohazard bucket.

38

Interpretation of test results:

• Positive: A positive result is obtained if agglutination of the blue latex particles is

seen in 20 seconds in the test circle, with no agglutination in the control circle.

• Negative: A negative result is obtained if no agglutination occurs and a smooth

suspension remains at 20 seconds in the test circle.

Lab Write-up QUESTIONS: 1. What are the cellular components that are detected by the S. aureus latex test?

2. What are the corresponding host components?

3. What is the difference between Fab and Fc regions of the antibody?

4. If both the test latex and negative control latex agglutinated, is that a false positive? False

Negative?

5. Why the TECRA® ELISA procedure termed a sandwich ELISA.

6. What are possible advantages of the TECRA® ELISA procedure over the VIP® kit?

7. What antigens might be the targets for antibodies is this kit?

8. Can immunoassays discriminate virulent vs. avirulent strains?

39

LAB EXERCISE 10: DNA PROBE DETECTION OF VIBRIOS

DNA probes are used for Colony Blot Hybridization to directly enumerate bacteria in

food samples. In this procedure, colonies from standard plate counts are lifted from the L-agar

plates to a sterile filter paper and then hybridized with a DNA oligonucleotide probe that is

specific for a particular bacterial species. DNA probes are enzymatically labeled with an alkaline

phosphatase enzyme. The alkaline phosphatase label to detect probe-positive colonies is

visualized by color change of a substrate (NBT/BCIP solution) that reacts with this enzyme.

Postive colonies produce brown/bluish colonies, when substrate is cleaved by the enzyme, while

negative colonies appear yellow. This protocol is used for enumeration of bacterial colonies on

spread plates to detect probe positive colonies in a background of other bacteria. It can be used in

conjunction with growth on different media, although it will not work well on differential agar

containing dyes.

Vibrio species are gram-negative, halophiles that are common to estuarine environments. V.

vulnificus remains one of the leading causes of seafood related deaths in humans with a mortality

rate of approximately 50%. Disease can result from the consumption of raw or undercooked

shellfish, such as oysters and clams, which harbor the bacteria and lead to fatal septicemia

(cellular destruction due to accumulation of bacteria in the blood). Exposure of open cuts to the

bacterium in seawater can also lead to wound infections. Fortunately, most people are not

susceptible to this disease, and fatalities are primarily in individuals with underlying disorders

that compromise the immune system, including HIV/AIDS, cancer, diabetes, liver disease or

hemochromatosis. Vibrio species are quite variable biochemically and it is difficult to

discriminate species based on traditional differential media, such as TCBS (see lab 2). Also

growth on selective media generally requires pre-enrichment in broth used for end-point titration

of MPN enumeration (see lab 3). Unfortunately, this process requires days and lacks species

specificity. This probe is derived from the cytolysin gene (vvh), and has been shown to be very

species specific.

PROCEDURE: Positive and negative controls should be prepared with each batch of samples

and processed simultaneously with unknown samples. Incubation should be done in a water bath

or hybridization oven that is accurate within ± 1º C.

DAY 1:

1. Scrub and shuck 4-6 oysters and weigh oyster meat in sterile Petri dish (need at least

50g). Place oysters in sterile blender jar. Add an equal weight or volume (1g: 1ml) of

sterile phosphate buffered saline (PBS) to blender jar, using sterile grad cylinder.

2. Homogenize oyster sample for 2 min on hi-speed and add 1 ml of oyster homogenate to 9

ml of APW and continue to serially dilute using the 9 ml APW dilution tubes to the 10-3

dilution.

3. Plate 100 µL from each dilution tube to duplicate plates each of LA and TCBS, yielding 2

LA and 2 TCBS plates per dilution (Remember: plating 100 µL increases the dilution on

the plate, i.e plating 100 µL from a 10-6

dilution tube will result in 10-7

plate.)

4. Incubate plates (media side up) at 37ºC for approximately 24 hours.

DAY 2:

1. Count colonies on L agar and TCBS plates and place plates in cabinet at room

temperature.

40

2. Record numbers and determine CFU/g for comparison to colony blot hybridization

results.

DAY 3:

Preparation of Colony Lifts:

1. Label 85 mm Whatman #541 filters with pencil to record your group number and the

dilution of the plate you are blotting. Handle filters aseptically.

2. Overlay colonies on spread plate with filter (pencil side up) so that there are no bubbles

under filter by first lining up one edge of the filter with one edge of the plate. (DO NOT

pick up filter and replace on agar.). Use a “hockey stick” to lightly press the filter directly

against the agar surface and remove bubbles.

3. Allow filter to sit on plate for 15 min to ensure transfer of smaller colonies.

4. Place 100 mm Whatman #3 filter onto microwaveable tray and saturate with lysis

solution (about 3ml) using squeeze bottle

5. Aseptically lift the #541 filter from the agar plate using tweezers and overlay the #541

filter onto the wet #3 filter with colony side up ensuring contact and no bubbles between

filters.

6. Microwave filters on medium to high heat until the filters are dry but not brown (about 3-

5 min.).

7. Place #541 filters into a clean container containing ammonium acetate neutralization

buffer (about 20 ml per filter) colony side down and incubate at room temperature, with

shaking (125 rpm) for 5 min.

8. Drain the neutralization buffer and rinse in 1x SSC buffer (20 ml per filter). Swirl briefly,

drain and repeat.

9. Place filters, colony side up, onto absorbent paper and dry completely at room

temperature over night. Dried filters may be stored at room temp. in a sealed plastic bag.

Alternatively, filters are to be processed immediately and drying is not necessary.

Proteinase K treatment:

1. The proteinase K working solution (40 µg/mL) will be prepared for you by your TA.

Solution is pre-warmed to 42ºC.

2. Transfer filters to a container with about 5 ml per filter of proteinase solution.

3. Incubate with shaking at 42° C for 30 min.

4. Empty the container and rinse with shaking in1x SSC buffer (20 ml/ filter) at room temp

for 10 min. Repeat 3 times.

5. Filters can be dried and stored indefinitely.

41

Hybridization Demonstration:

1. Include control filters with each bag. Place 5-10 filters into a bag (Whirl Pack 4.5” X 9”)

or tupperware container and add 100 ml of pre-warmed hybridization buffer (56° for

VVAP). Exclude air from bag and seal or roll up.

2. Incubate with shaking (125 rpm) at temperature for 30 min.

3. In new bag add probe (5 picomoles/filter, usually 3 to 20 µl of probe) to pre-warmed

hybridization buffer (25-100 ml).

4. Transfer filters with forceps to fresh buffer with probe. Exclude air from bag and seal or

roll up. Incubate for 1 h with shaking (125 rpm) at 56°C.

5. Transfer filters to container with pre-warmed (56°C) 1x SSC/1%SDS using 10 ml/filter

with shaking (125 rpm) at appropriate temp for 10 min.

6. In the same container, rinse five times in 1x SSC buffer (20ml/filter) at room temperature

for 5 min. each rinse. Filters can be combined at this point with appropriate increase in

the volume of buffer.

Probe Development Demonstration:

1. In a new petri-dish, develop each filter in 2 ml of NBT/BCIP solution.

2. Incubate at room temperature (or at 35°C for faster results). Cover to omit light. Check

development of the positive control every 1/2 hr.

3. To stop further development, place filters in new container and rinse 3 times at room

temp with DI water for 10 min. each.

NOTE: be sure to dispose of the NBT/BCIP correctly. It is a hazardous waste.

4. Place the filters on an absorbent paper in the dark and allow to dry. Positive colonies will

have a bluish or brown color. Negative colonies should appear colorless or yellow. The

filters should be stored in the dark to prevent color change

STUDY QUESTIONS:

1. Create a table comparing your TCBS counts and your colony blot hybridization counts

for each dilution. Do these counts differ? In what ways? Give an explanation of your

results.

2. What accounts for the specificity of colony blot hybridization?

3. Is colony blot hybridization more specific than traditional plating? than the MPN

method? Why or why not?

4. We saw in the ELISA lab that by increasing the speed of an assay we decreased the

sensitivity. Is the same true when comparing traditional plating to colony blot

hybridization? Why or why not? (hint: base your answer on your results)

42

LAB EXERCISE 11: POLYMERASE CHAIN REACTION (PCR) LAB

PCR is a method for rapid amplification of short pieces of DNA using DNA polymerase, and

was first described by Kleppe, Ohtsuka, and Kleppe in 1971. Saiki and colleagues went on to

develop a primer-directed enzymatic amplification of DNA using a thermo-stable polymerase.

PCR is one of the most widely applied procedures in biology today. Applications include the

following:

• Identification of species using species- or type-specific primers

• Molecular characterization: DNA fingerprinting

• DNA Sequencing: single primer is used to amplify one strand to be sequenced

• Cloning of targeted genes

• Gene expression studies using reverse transcriptase PCR (RT-PCR) to detect RNA

transcripts of specific genes

• Quantification of bacteria or expressed genes

PCR amplification is one of the most powerful diagnostic tools available for

identification of food borne pathogens, providing extremely rapid and specific detection. PCR

primers are currently available for every major pathogen, and some of the assays require less

than an hour. PCR will often detect as little as one microorganism in a sample. Some sample

media can be very complex with a large background of other organisms, such as fecal samples.

Thus, this method is exceptionally useful for finding the “needle in the haystack”.

This procedure is based on 1) the specificity of DNA sequences, called primers, which

serve as species-specific markers, and 2) the thermal stability of the Taq polymerase enzyme.

In PCR reactions, a gene target is selected that is specific for the organism or gene of interest.

For example, PCR primers for Salmonella are based on the virulence gene invA. Thus, detection

is not only specific for the species but may also determine if strains have virulence potential.

Through a series of enzymatic reactions, a small amount of DNA is amplified exponentially to

produce thousands of DNA fragments that can then be visualized on an agarose gel. Electrical

charge is used to separate DNA fragments by size on the gel. There is a positive charge at one

end of the gel box and a negative charge at the other end. DNA is negatively charged and will

migrate toward the positive electrode. Smaller fragments of DNA move through the gel more

quickly than larger fragments, which allows for separation by size. After migration, DNA-

specific dyes are then used to visualize the PCR product.

The reaction mixture is prepared in small volumes of 10 to 100µL and contains the

following components:

a) Template or target DNA (10 to 1000 ng) from the microorganism of interest that you are

tying to detect in your sample

b) Primers or short pieces of DNA (14-45 bases) that correspond to your target DNA and

match up or hybridize in different directions at 2 separate locations that are usually 500

to 3000 bp apart.

c) Taq polymerase is a heat stable enzyme that functions optimally around 72ºC, but is

stable at higher temperatures, which are needed for denaturation of DNA. Thus the

enzyme is added only once but is repeatedly “reused” for the multiple cycles of the

amplification reactions

d) dNTPs are nucleotide bases (ATP,CTP,GTP,TTP) or building blocks of DNA.

e) Buffers and salts at concentrations that are optimized for the reaction. Generally the

MgCl2 concentration is adjusted for optimization.

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The basic reaction is performed in a programmable incubator called thermocycler, and

the procedure is outlined in the following steps and shown in Figure 1:

a) Denaturation: The DNA is heated to 94-96°C to denature or separate the double

stranded template DNA.

b) Annealing: The mixture is rapidly cooled to 30-60°C, allowing the primers to match up

and bind to specific sites on the target DNA.

c) Elongation: The Taq polymerase extends the DNA strand begining where the primer

binds to produce a new strand of DNA. 72°C is the optimum temperature for enzymatic

activity.

d) Cycling: The steps above are repeated in the same sequence for 25-35 cycles to produce

multiple new strand of DNA.

e) Final elongation: The elongation temperature is held for about 10 min. to maximize

reactions.

Figure 1. Polymerase Chain Reaction

Potential Problems include the following:

a) Interfering compounds: Heavy metals and other compounds can interfere with the

enzymatic activity of Taq polymerase. Thus, sample preparation includes a step

where DNA is extracted and isolated from the rest of the cellular material.

44

b) Sample size: Some organisms may be present in very low numbers in foods.

Although PCR can detect 1 organism in a sample, generally very small samples are

analyzed. These problems can be addressed by testing multiple samples and

enrichment of samples.

c) Viability question: DNA can be extremely stable under certain conditions and PCR

may detect DNA that is present from organisms that are not alive

d) Quantification: The amount of product produced roughly reflects that amount of

template. Therefore more heavily contaminated samples will produce more fragments

and exhibit a stronger band. However, exact quantification requires more complex

procedures or special equipment.

PROTOCOL:

Ground rules for molecular biology:

1. Always were gloves (DNases on hands destroy DNA, some of the reagents used are

carcinogens).

2. Be aware of cross-contamination (change tips frequently and NEVER use the same tip

for different reagents).

3. Always run controls (positive and negative).

4. Due to the extreme sensitivity of the PCR assay, false positive are possible if samples are

cross contaminated. False negatives are possible due to experimental error usually

arising from pipetting problems. Therefore, positive and negative controls are extremely

important, and data is only acceptable when controls show the expected results.

Colonies from the previous lab have been selected for DNA extraction and PCR analysis.

The PCR primers are based on sequences from the invA gene and should produce a PCR product

about 457 bp in length. You will perform PCR amplification with your Salmonella strain. Each

group will perform PCR on the following samples:

1. Positive control: DNA from a known Salmonella strain that is positive for invA

2. Negative control for specificity: DNA from V. vulnificus

3. Negative control for contamination: PCR reaction with everything except the target

DNA. Water will be added to compensate for the volume.

DAY 1:

Material needed:

Salmonella culture

Microcentrifuge tubes

Sterile PBS

DNA templates from target strain (which you will be extracting)

PCR tubes

PCR master mix reaction: water, DNA, Taq, dNTPs, buffer, and primers.

PCR racks

10-20 µl pipettors

200 µl pipettors

Filtered pipette tips

Centrifuge

Heat Block

Thermocycler

45

DNA extraction:

1. Pipette 1 ml from each of the two (Vibrio spp. and Salmonella spp.) L Broth cultures into

two microcentrifuge tubes. Label them accordingly.

2. Centrifuge your tubes at 10,000 rpm for 10 min.

3. Pour off the supernatant, taking care not to dislodge the pellet.

4. Add 400 µl of sterile PBS to each microcentrifuge tube.

5. Vortex the tubes until the pellet is completely resuspended.

6. Boil the tubes at 100°C for 5 min in the heat block.

7. Centrifuge tubes at 13,000 rpm for 3 min.

8. Transfer supernatants to sterile microcentrifuge tubes taking care not to disrupt the pellet.

The supernatants contain your DNA, DO NOT DISCARD!!

PCR amplification procedure:

1. Prepare a master mix in a separate tube on ice. Reaction mixture given below is for 1

sample. You will need to prepare a mixture for 4 samples (multiply volumes by 4). Write

in the final volume before starting and check off each item as it is added.

Volume Final Volume

Primer forward 1.0 µL

Primer reverse 1.0 µL

dNTP 2.5 µL

10x buffer 2.5 µL

Water 16.75 µL

Add last: Taq 0.25 µL

**Do not add Taq until you are ready to add the DNA to your PCR tube and you

are ready to go into the thermocycler.

Keep all your reagents on ice.

2. Label PCR tubes to indicate sample and lab group.

3. Add 24µL of master mix to each tube.

4. To your PCR tube, add 1µL of your template (either DNA or water).

5. Thermocycler should be started and preheated. Once temperature is up, place tubes in

thermocycler, and start program. This gives you a “hot start”.

DAY 2:

Material needed:

Gel boxes with combs and trays

Power units

UV source

Camera/film

1% Agarose

1X TAE

Loading dye

Ethidium Bromide (EtBr)

PCR rack

10-20 µl pipettes

Filtered tips

Gel electrophoresis:

46

1. Add 5µL EtBr (CARCINOGEN!! WEAR GLOVES!) to a flask containing previously

prepared 1% agarose gel that is kept warm in the water bath.

2. Set gel tray into gel box with rubber ends forming a seal with the side of the gel box.

3. Insert the comb with the 12 pronged-end facing downward.

4. Pour the gel into the gel tray. Avoid making bubbles in the gel. If bubbles do form use a

sterile pipette tip to pop them immediately, before the gel sets.

5. When gel is completely solidified (approximately 15 min), remove comb and rotate gel

tray so the wells are on the same side of the gel box as the negative electrode.

6. Carefully load 12µL of the samples to any lane but the first on the gel.

7. Carefully load 5µL of marker to the first lane.

8. Run gel at 100mV for ½ h.

9. View gel on UV light (WEAR SAFETY GLASSES!).

Molecular weight markers are run simultaneously to estimate the size of the fragments.

Ethidium bromide is added to the gel to bind DNA and make it visible under UV light.

Lab Questions

1. On what basis do you select primers used to identify target DNA, and how do they

determine the specificity of PCR?

2. Define positive/negative controls and explain why they are important.

3. Why do you need a thermostable enzyme?

4. From the PCR products we obtained above, what other steps could be done to further

characterize the DNA and the strains from which they were derived?

5. Define template, primers, and elongation.

Helpful website:

http://dlab.reed.edu/projects/vgm/vgm/VGMProjectFolder/VGM/RED/RED.ISG/

47

LAB EXERCISE 13: MOLECULAR TYPING

A variety of methods are available for the molecular typing of bacterial species. These

methods generally provide DNA fragments of varying sizes that can be used like a barcode to

discriminate either different species or strains of the same species. This genotypic

characterization can be used to profile isolates from different geographic locations or from

different sampling sources to establish geographic distribution among the strains or trace

outbreak strains. The obtained profiles are usually analyzed using special imaging and data

analysis software. One of the most widely accepted methods is Pulsed Field Gel

Electrophoresis (PFGE). This method is the basis of PULSE NET, a CDC database for

identification and tracking of bacterial pathogens. PFGE examines the entire bacterial

chromosome and uses restriction enzymes cut up or digest chromosomal DNA into large

fragments that can be visualized by agarose gel electrophoresis. The number of fragments is

determined by the number of sites for restriction enzyme digestion in the chromosome. These

large fragments will not enter an agarose gel using standard electrophoresis, but can be separated

on a gel by alternating, pulsed electrical fields that help them move through the gel. The method

requires precise calibration and technique in order to compare results from different sources.

Many strains are “untypeable” by this method because they are not sensitive to the enzymes used

for digestion under these conditions. Therefore, alternative strategies have been explored for

more rapid and reliable typing systems.

One of the alternatives to PFGE is repetitive extragenic palindromic DNA PCR (rep-

PCR). These repetitive DNA element are distributed at various sites outside of specific genes

throughout the chromosome and can serve as targets for PCR amplification. These extragenic

sequences are panlidromic (same sequence in both directions, i.e., ATGCCCT followed by

TCCCGTA) and therefore PCR primers derived from these sequences can be used to amplify

adjacent DNA in both directions. Different types of repetitive elements, such as ERIC and BOX,

have been identified in gram negative bacteria and are used for molecular typing at the genomic

level. PCR products provide DNA “fingerprints” that discriminate interstrain variation based on

amplicon size and intensity (Figure 1).

Figure 1. Repetitive-DNA-sequence-based PCR. Primers hybridize with genomic repetitive

elements and amplify intervening sequence to produce multiple PCR products.

48

In this lab, we will compare the genetic profiles of V. vulnificus strains isolated from oysters and

clinical case of septicemia. Rep-PCR genomic typing will be performed using DiversiLab

Microbial Typing System (Bacterial Barcodes, Inc., Houston, TX). Extracted DNA will be

amplified by PCR using the DiversiLab Salmonella kit (DL-SE01).

PROTOCOL You will do PCR simultaneously on an unknown strain of V. vulnificus, a positive control, a

negative control for a total of 3 PCR reactions/group.

1. Positive controls: DNA from 8 different V. vulnificus strains (one per group; 4 from

oyster and 4 from environmental).

2. Negative control: Provided in the DiversiLab Kit.

3. Positive control : Provided in the DiversiLab Kit.

Material needed:

DNA templates

Gel electrophoresis

Equipment: PCR racks, forceps, 10-20 µl pipettors (6), 100-200 µl pipettors (6), Filtered tips

(6 boxes each size), PCR tubes (6 containers with 10 in each), 1.5 ml tubes (6 containers with

10 in each), 6 gel boxes with combs, tray, 6 power units, UV source, Camera/film.

DAY 1:

PCR amplification procedure:

1. Prepare a master mix in a separate tube on ice. Reaction mixture given below is for 1 sample.

You will need to prepare a mixture for 4 samples. In other words multiply volumes by 4 and

add to one tube. Write in the final volume before starting and check off each item as it is

added. Keep all your reagents on ice.

Volume Final Volume

Rep-PCR MM1 18.0 µL

GeneAmp® 10x PCR Buf. 2.5 µL

Primer Mix P 2.0 µL

AmpliTaq DNA Polymerase 0.5 µL

2. Label PCR tubes to indicate sample and group. Add DNA, positive, and negative control to

each tube.

3. Add 23.0 µL of master mix to each tube.

4. To each PCR tube, add 2µL DNA, 2µL of the positive, and 2µL of the negative controls.

5. Vortex and return tubes in ice.

6. Program and preheat thermocycler. Once temperature is up, pause and place tubes in

thermocycler. Start program. This gives you a “hot start.”

DAY 2:

Gel electrophoresis:

The PCR reaction produces multiple DNA fragments for each strain and will be electrophoresed

on an agarose gel. DNA is prepared in a “loading buffer” which contains tracking dye and

glycerol to prevent the DNA from diffusing out of the well. The DNA is applied to the gel,

which is placed in an electrical field, and DNA migrates to the positive electrode due to the net

negative charge of DNA. The agarose acts as a molecular sieve with the shorter fragments of

49

DNA migrating faster. Molecular weight markers are run simultaneously to estimate the size of

the fragments. Ethidium bromide (Carcinogen!! Dispose properly and use gloves!) is added to

the gel to bind DNA and make it visible under UV light.

1. Prepare 1x TAE buffer from 20x stock.

2. Prepare agarose gel (2% in TAE buffer or 2 g in 100 ml). Microwave to melt, cool to 50C,

add 10 µl EtBr (CARCINOGEN!! WEAR GLOVES!), and pour gel.

3. Add loading buffer to PCR product at a ratio of 1:6. Vortex and centrifuge.

4. Fill gel box with buffer, place gel in box.

5. Carefully load 12 µl of sample or markers to each lane on gel.

6. Run gel at 100mV for ½ h.

7. View gel on UV light (POSSIBLE EYE DAMAGE!!! WEAR SAFETY GLASSES!)

Rep-PCR Demonstration in Building 866

The PCR products of the 8 V. vulnificus strains tested will be loaded on the Bacterial Barcode

chip, to be analyzed by the Bioanalyzer (As shown in the flow chart below).

4. Dendrogram are based on similarity of banding patterns and indicate the relatedness of

the DNA targets among different strains.

1. PCR amplicons are loaded on the chip 2. Chip is loaded in the Bioanalyzer

3. Band size and intensity is calculated as amplicons are scanned during microcapilary electrophoresis

50

References 1. Beyer, W., F. Mukendi, M. Kimmig, and P. Böhm, 1998. Suitability of repetitive-DNA-

sequence-based PCR fingerprinting for characterizing epidemic isolates of Salmonella

enterica serovar Saintpaul. J. Clin. Microbiol. 36:1549-1554.

2. Jensen, M. A., J. A. Webster, and N. Straus, 1993. Rapid Identification of Bacteria on the

Basis of Polymerase Chain Reaction-Amplified Ribosomal DNA Spacer Polymorphism.

Appl and Environ Microbiol. 59(4):945-942.

3. Versalovic, J., F. de Bruijn, and J. Lupski, 1998. Repetitive sequence-based PCR (rep-

PCR) DNA fingerprinting of bacterial genomes. In: Bacterial Genomes: Physical

Structure and Analysis. Chapman & Hall: New York.

Lab Questions

1. How do the banding patterns on the gels relate to the origin of the strains?.

2. What are some sources of variability found in genetic typing assays?

3. Why where we able to use rep-PCR Salmonella typing kit for V. vulnificus typing?

4. Which assay would you expect to be more specific: regular PCR, rep-PCR or ELISA?

Which assay would be more sensitive?