LABORATORY EXERCISES

to accompany

MICROBIOLOGY LABORATORY

BIO 209

Professor Susan C. Kavanaugh

Bluegrass Community & Technical College

Spring 2016

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FUNDAMENTAL REQUIREMENTS FOR LABORATORY SAFETY

 Learn about the hazardous properties of all the materials used in your work and observe the applicable safe handling, storage, disposal, and emergency procedures.  treat all substances are hazardous.  confine drinking, eating, chewing gum or tobacco, smoking and the storage and disposal of food, beverages, and tobacco products to uncontaminated or designated areas outside of the laboratory. Absolutely no food or drink may be present in the laboratory.  wear the appropriate personal protective equipment for the activities being conducted. As a minimum protection, wear a lab coat or gown, gloves, and safety glasses whenever chemicals, radiochemicals, or biohazards are being used in the laboratory.  wear close-toed shoes providing full coverage, no open shoes such as sandals.  keep exits, passageways, and access to safety equipment like emergency eyewash stations and showers or fire extinguishers free from obstruction.  be familiar with the locations and operating procedures for safety and emergency equipment such as fire extinguishers, first aid kits, emergency eyewash stations and showers*, fire alarm pull stations, emergency telephones##, and emergency exits.  wash hands before leaving the laboratory and remove lab coat or gown before leaving the laboratory and before entering other areas, particularly eating facilities.  tie back or otherwise restrain long hair or loose articles of clothing or jewelry when working with chemicals, biohazards, radioactive material, flames, or moving machinery.  use mechanical pipetting devices-never pipette by mouth.  perform procedures involving the liberation of volatile materials or aerosols of a toxic or flammable nature in a fume hood.  know the location of the laboratory safety manual and the Material Safety Data Sheets (MSDS) notebook.  report all accidents, dangerous incidents, and suspected occupational diseases and seek preventative measures to avoid recurrences.**

*(recommended duration = 15 minutes with lukewarm water) ##(the nearest phone is in Room 244A) **(report forms are on the instructor’s desk)

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Microbiology Laboratory Specific Guidelines

Your1. Your lab bench lab be areanch isa rtoea be is cleanedto be clea ned with Lysol disinfectant before lab begins, after each procedure, withand Lysol befor disinfectante you leav ebefore. lab begins, after each procedure, and before you leave. For2. liquidFor liqui cultured c ultspills,ure immediatelyspills, immedia tely saturate the area with Lysol and cover with paper towels. Allow saturatecontact thewit areah the with disi Lysolnfect aandnt forcover at least 10 minutes. Wipe away all liquid with clean paper towels. with paper towels. Allow contact with theDispose disinfectant of all for pa peat leastr tow 10els in the trash. minutes. Wipe away all liquid with clean paper towels. Dispose of all paper towelsFor3. surfacesFor in sur thef acontaminatedtrash.ces c ontamin byated inoculating by inocula ting loops, pipettes, swabs, test tube lids, Petri dishes or other loops,such itpipettes,ems, imm swabs,ediat testely tubespra ylids, the surface with Lysol, and cover with a paper towel. Allow contact with Petri dishes or other such items, immediatelythe disinfe cspraytant f theor a surfacet least 10with mi nutes. After this time, the area should be wiped with paper towels which Lysol,are then and disposed cover with into a paper the tra towel.sh. Allow contact with the disinfectant for at least 10 minutes. After this time, theAll4. broken areaAll brokeshould glasswaren beglasswa wiped shouldr withe sho be paper ulddisposed be disposed of into the sharps containers or cans with the biohazard towelsoflabe intols. thewhich Non sharps are-contaminat then containers disposeded or bro cansintoke nthe g lass must be swept up by the instructor with a broom and dustpan. trash.with the biohazard labels. Non- contaminatedContaminate brokend broke glassn gla mustss m ustbe sweptbe disi nfected first using the Lysol disinfectant and covered with paper uptowe by lsthe for instructor at least with 10 mia broomnutes beandfor e being swept up by the instructor as previously described. Glass dustpan.microsc Contaminatedope slides, sta brokenined sm glassears, must and cover slips should also be disposed of into the sharps containers. be disinfected first using the Lysol disinfectant and covered with paper Swabs,towels5. Swa for tonguebs, at tleastong depressors,ue 10 d minutesepres sors,and before disposable and disposa ble pipettes should be disposed of in the plastic trays containing pipettesbeingdisinfe swept cshouldtant up pr bebyovided thedisposed instructor at e aofc hin asst theude nt bench. plasticpreviously trays described. containing Glass disinfectant microscope providedslides, stained at each smears, student and bench. cover slips Usedshould6. Use gloves alsod g lovesbe should disposed should be disposed of be into disposed the of into of into the designated biohazard trash bag. Used gloves should not be sharps containers. thethrow designatedn into th biohazarde regular trashtrash. bag. Used gloves should not be thrown into the regular trash. Nothing7. Nothing should shoul be thrownd be thr intoown the into small the small sinks at each student bench. sinks at each student bench. Any8. Anclothingy clot thathing becomes that bec contaminatedomes contaminate d must be removed immediately, placed in a biohazard bag, mustand abeutocla removedved. immediately, Lab coats m placeday not in be a worn outside of the Microbiology Lab. biohazard bag, and autoclaved. Lab coats may not be worn outside of the UsedMicrobiology9. Use Petrid P dishe triLab. culturesdi sh cult mustures bemus t be discarded into designated buckets containing biohazard bags, to be discardedautoclave intod prio designatedr to dispos bucketsal. Glasswa re cannot be discarded into these containers. containing biohazard bags, to be autoclaved prior to disposal. Glassware Usedcannot10. Usetest be d tubesdiscarded test torube other sinto or specimens theseother spec otherimens other than Petri dishes should be placed in the designated area for thancontainers.used Petri suppl dishes ies in should the corn be eplacedr at the in end of the front counter of the lab. the designated area for used supplies in the corner at the end of the front counter of the lab.

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INTRODUCTION TO CULTURING, MEDIA, AND ASEPTIC TECHNIQUES

You may be unaware of the number and variety of microorganisms (microbes) found everywhere in our environment, including the human body. In this laboratory you will learn new techniques and make observations which relate to the concepts of microbiology. Most of the microorganisms that you will use in these laboratories are normal inhabitants of our environment and our bodies. These microbes are called normal microbiota for the environment in which they normally reside. Health professionals need this knowledge in order to be able to distinguish normal flora from a possible infectious agent when interpreting microbiological reports. They also need to understand how normal microbiota can occasionally cause an infection when they invade a different area of the body, or when the patient's immune responses have been compromised.

Microorganisms are found almost everywhere. In these first laboratory exercises you will be introduced to aseptic techniques, the procedures followed by microbiologists and healthcare workers to prevent contamination from outside sources and to prevent introduction of potentially disease-causing microbes (pathogens) into the human body. The methods for handling previously sterilized materials, for taking samples, for handling cultures, and for disposal of contaminated materials are all designed to prevent the spread of microbes from one area to another. Pay close attention to the details in the written procedures and to the instructor's demonstrations to prevent contamination of your cultures, yourself, your environment, and the other people in your laboratory as well as prevention of infecting people outside of the laboratory, such as your friends and family. These techniques can be applied not only here in the microbiology laboratory, but also throughout your career, and in your daily life.

Most of the laboratory exercises performed in this course will involve a two-step process. During the lab session you will set up the cultures and then after these cultures have incubated for the appropriate length of time (usually 24 to 48 hours) you will need to observe the growth and record your observations and results.

Wait for the instructor to demonstrate the procedures described and to make the specific assignments.

In the rest of the exercises in the course, you are dealing with living bacteria, so it is very important to follow the procedures exactly to avoid contamination or infection. The following precautions are especially important:

1. Always wash your hands with the antiseptic soap provided before you begin and after you have finished each procedure.

2. Always wipe off your work area with the disinfectant provided before you begin and after you have finished each procedure.

3. Always wear gloves when handling cultures or specimens.

4. Discard all used materials in the appropriate designated place after you are done. Put all used materials and cultures into the special containers for contaminated material. Never put any used materials back into the supply area.

5. Do not lay liquid broth cultures, test tubes, swabs, or pipettes down on the tabletop or touch anyone with them.

6. Hold the lid of the culture (Petri plate over the surface while you are inoculating the surface and then immediately replace.

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7. The cultures you will observe after the 24-48 hour incubation period will have a high concentration of bacteria on them. Even though they are "normal inhabitants" of the environment or human body, they can cause an infection if they get into an open cut or sore or transmitted to the mouth, hair, or eyes from your hands because of the large number of bacteria present. Thus, it is extremely critical that the Petri dishes be examined when the covers are in place. Never hand one to someone else with the lid removed.

8. Always carry test tubes in the test tube racks provided, not in your hands. Do not pick up test tubes by their caps.

9. A disposable, fluid-resistant, full-length, long-sleeved lab coat must be worn at all times in the lab. The coat must be removed before leaving the room for any reason. If the lab coat becomes contaminated, it must be removed, put into a biohazard bag, and autoclaved before disposal into the trash.

10. If a spill occurs, notify the instructor immediately and decontaminate the area right away.

11. Long hair must be pulled back.

12. Closed-toed shoes must be worn in the lab at all times. No sandals are permitted.

13. If you have any doubts or questions about what you are doing; ASK THE INSTRUCTOR FOR HELP!

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BACTERIAL MEDIA

Objectives: After completion of this laboratory experiment, the student will be able to:

1. Perform a commonly used method of isolating bacteria in pure culture - the streak plate method. 2. Perform essential aseptic techniques. 3. Use selective media to isolate an organism from a mixture of organisms. 4. Transfer microorganisms from liquid nutrient broth to an agar plate using a pipette or an inoculating loop.

In this exercise you will use different types of culture media to grow various species of bacteria from a mixed culture. To study microorganisms properly, we have to be able to grow them. To accomplish this, it is necessary to transfer the specimens to an environment that will simulate the same conditions under which they occur in nature. Nutritional requirements vary widely from one species of bacteria to another and in many cases are not clearly known. Much has been accomplished concerning the duplication of conditions necessary for the cultivation of microorganisms, and most microbes can now be cultivated on or in artificial media. Ingredients in media are intended to supply the nutritional and growth requirements of microorganisms so that the cultures studied will present characteristics comparable to those that exist in nature.

1. Primary or Isolation Media: Media used for primary inoculation of specimen; usually prepared in Petri dishes so they can be streaked to obtain isolated colonies of any organisms present. Media used routinely in most laboratories are: Trypticase soy agar (TSA) and Nutrient agar

2. Enrichment Media: Media that has been enriched by the addition of extra ingredients to enhance the growth of fastidious microbes.

Examples: blood agar chocolate agar

3. Selective Media: Media used to grow one particular type of bacteria from a mixed culture by inhibiting the growth of the other bacterial species.

Examples: Phenylethyl alcohol (PEA) agar-selects for gram-positive bacteria Mannitol salt agar-selects for staphylococci MacConkey agar-selects for gram-negative bacteria Eosin methylene blue agar-selects for gram-negative bacteria

4. Differential Media: Media used to distinguish between species of bacteria which may look exactly alike or very similar by other methods, such as the Gram stain, or on TSA.

Examples: MacConkey agar – distinguishes between lactose fermenters and non-lactose fermenters Mannitol salt agar- distinquishes between Staphylococcus aureus and other Staphylococcus species Eosin methylene blue – distinguishes between E.coli and other enteric bacilli

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The media that you will be using in this experiment are:

TSA = trypticase soy agar; nutrient primary isolation media; will grow many types of bacteria (both gram-positive and gram-negative bacteria)

PEA = phenylethyl alcohol agar; selective media; grows only gram-positive bacteria. The phenylethylalcohol is inhibitory to gram-negative bacteria.

MAC = MacConkey agar; selective media; grows only gram-negative bacteria; gram-positive bacteria are inhibited by the crystal violet dye in the agar. MacConkey agar is also used as differential media to distinquish between lactose-fermenting and non-lactose fermenting bacteria. Incorporation of lactose, bile salts, and phenol red indicator causes lactose-fermenters to appear red, whereas non- lactose fermenters will appear colorless or transparent.

MSA = Mannitol salt agar; selective media; grows only Staphylococcus bacteria. 7.5% salt is inhibitory to most other bacteria. Mannitol salt is also differential media used to distinguish between Staphylococcus aureus and other Staphylococcus species. Mannitol fermentation with subsequent acid production by S. aureus is indicated by a change in the color of the phenol red indicator to yellow.

EMB = Eosin methylene blue; selective media; grows only gram-negative bacilli. Eosin methylene blue is also differential media used to distinguish E.coli from other gram-negative enteric bacilli. E.coli ferments the lactose in the agar, causing acid production, which precipitates the eosin and methylene blue dyes. This results in a metallic blue-black color with a greenish sheen. Other gram-negative enteric bacilli will appear pink or transparent.

BAP = Blood agar plate; enrichment media used to grow a variety of fastidious microorganisms such as Streptococcus. Blood agar is also used to demonstrate different types of hemolysis: beta hemolysis = complete lysis of the red blood cells by streptolysin 0 and streptolysin S enzymes alpha hemolysis = incomplete lysis of red blood cells resulting in the breakdown of hemoglobin, which produces a greenish halo around the bacterial colonies gamma hemolysis = no lysis of the red blood cells; no significant change in in the color of the agar surrounding the colonies

Specimens submitted to the laboratory for microbiological examination often contain a mixture of microorganisms. In order to study the characteristics of a microorganism, it is first necessary to separate it from other microorganisms present in the mixture; we must isolate the suspected organism in pure culture. A pure culture is one in which all of the cells present in the culture originated from a single cell type. The streak plate method is the method classically used for isolating a pure culture from a mixed culture.

With this method you will attempt to purify a mixed broth culture containing several different species of bacteria. Once isolated, the bacterial colonies can be differentiated from each other.

An essential component for isolating a pure culture is aseptic technique, which involves the transfer of microorganisms from one environment to another in such a way that neither you nor the environment around you is contaminated with the specimen that you are transferring and that the pure culture you are making is not contaminated with other organisms from the environment. In the aseptic preparation of pure cultures, the transfers are usually made with sterile inoculating loops or needles or with sterile pipettes. Your instructor will first demonstrate the aseptic techniques to be used.

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SPECIMEN HANDLING

Objectives: After completing this exercise the student should be able to:

1. obtain a throat swab specimen 2. explain the effect of drying on swab specimens prior to their inoculation onto bacteriological media 3. describe correct collection and handling procedure for the following specimens: throat swabs, wound swabs, CSF, peritonal/pleural/synovial fluids, blood cultures, sputum, sputum for AFB, cultures for gonorrhea, stools, urines.

**Assignment: Read the article entitled "Know your Specimen Collection Techniques to avoid Errors" by Mahesh C. Goel, D.V.M., Ph.D. You will be held responsible for the material in this article. The article is on reserve in the LCC Library and is also available on-line through the LCC Library's homepage. Here is how to access this item: *start from the library's homepage at http://www.bluegrass.kctcs.edu/lrc/ereserves *click on BSL 214 ( instructors name) *Username: Will be announced at the first lab meeting (type exactly as shown; case sensitive) *Password: Will be announced at the first lab meeting (type exactly as shown; case sensitive) *click on the article you want: “Know Your Specimen Collection Techniques”

The proper handling of specimens for microbiological analysis requires:

(1) aseptic collection techniques (2) the use of appropriate containers (3) suitable means for preservation (4) suitable means of transporting specimens to the laboratory.

All specimens should be handled aseptically and treated as potentially infectious. In cases of spillage or contamination of the outside of a container, some form of disinfection should be carried out immediately.

SPECIMEN HANDLING: Throat swabs

Materials:

1. Two blood agar plates (BAP). 2. Sterile cotton swabs. 3. Tongue depressors to hold the tongue down during specimen-taking. 4. Sterile test tube with a previously inoculated throat swab that has been left to dry out. 5. candle (CO2) jar for incubation

Procedure:

1. Obtain a throat specimen from your assigned partner's throat with a sterile swab. Place the sterile swab against the back wall of the throat gently and move it up and down. 2. Inoculate a blood agar plate with the throat specimen. Streak it out using the streak plate method. 0 3. Incubate in a candle jar for increased CO2 at 37 C for 24-48 hours. 4. Take the previously inoculated, dried out throat swab and inoculate the second BAP. Streak for isolation and incubate in the candle jar at 370C for 24-48 hours. 5. Record the amount of growth on each plate in the Results and Observations.

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THROAT CULTURE RESULTS and OBSERVATIONS

Estimated amount of growth* Fresh culture

Dried culture

*0 = no growth 1+ = a few colonies 2+ = a moderate # of colonies 3+ = heavy growth (almost solid – no distinct colonies)

Study Questions:

1. What difference did you notice between the culture grown from the fresh throat swab and the one grown from the dried-up throat swab?

2. What explains the difference between the amount of growth on the two cultures?

3. Give two methods that would be used to prevent the loss of microbes after collection of the specimen.

4. What type of hemolysis did you observe?

SPECIMEN HANDLING: Urine Samples

Materials:

1. Urine sample containing Staphylococcus epidermidis, a gram-positive coccus in clusters and Escherichia coli, a gram-negative bacillus.

2. One plate of trypticase soy agar (TSA) (primary isolation media).

3. One phenylethylalcohol agar (PEA) plate (selective media for the growth of gram positive bacteria).

4. One MacConkey (MAC) agar plate (selective/differential media for the growth of gram negative bacilli).

5. One eosin methylene blue (EMB) agar plate (selective/differential media for the growth of gram negative bacilli; growth of Escherichia coli has a green metallic sheen)

6. One mannitol salt agar (MSA) plate (selective/differential media for the growth of staphylococcus species)

7. (1) inoculating loop

8. (1) sterile transfer pipette

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Procedure:

1. Disinfect your bench top with the disinfectant provided.

2. Using a marker, label the bottom (contains the agar) of each Petri dish with (a) your name, (b) date, (c) class and section number and (d) description of the specimen.

3. Obtain a sample of urine. Be sure the urine is well mixed beforehand. This can be done by gently swirling the cup.

4. Remove a drop of urine from the cup using a pipette or an inoculating loop using proper aseptic technique.

5. Lift the lid of the Petri dish just enough to get the pipette tip or loop inside. Place a drop of urine in the top half section.

6. Using your inoculating loop, streak back and forth in the pattern demonstrated by your instructor, using proper aseptic techniques. Do this for each of the 5 agar plates.

7. Invert the agar plates and incubate the streak plates at 37 Centigrade (body temperature) for 24 - 48 hours.

URINE CULTURE RESULTS and OBSERVATIONS

Record your observations on each type of culture media:

Trypticase soy agar:

Phenylethylalcohol agar:

MacConkey agar:

Eosin methylene blue agar:

Mannitol salt agar:

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STUDY QUESTIONS

1. Explain the difference between normal microbiota and pathogenic microbes. Is Staphylococcus epidermidis normal microbiota or a pathogen? E.coli?

______

______

______

2. Under what circumstances can normal microbiota become pathogenic?

______

______

______

3. Explain the importance of the aseptic techniques used in microbiology as they relate to your career as a health care practitioner.

______

______

______

______

4. Describe five aseptic techniques that you used during this laboratory exercise.

a. ______

______

b. ______

______

c. ______

______

d. ______

______

e. ______

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5. What is the purpose of trypticase soy agar? What type of bacteria will grow on TSA?______

______Phenylethylalcohol agar? ______

______MacConkey agar? ______

______Eosin methylene blue agar? ______

______Mannitol salt agar?______

______

6. What is the purpose of the streak plate technique?

______

______

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PREPARATION OF A BACTERIAL SMEAR

As you use this procedure throughout this course, remember these precautions for achieving the best results:

1. Use fresh cultures between 24-48 hours old, whenever possible.

2. When making smears, use a medium-sized drop of water and a small amount of bacteria. Mix the bacteria in the drop quite well with an inoculating needle, and spread it out thinly. A smear that is too thick will not only be difficult to stain properly but it will also be very difficult to observe individual bacterial cells under the microscope.

Materials: glass slide Bunsen burner tube of sterile water slide warmer gloves pencil inoculating needle sterile transfer pipette (“transpette”) inoculating loop culture of Staphylococcus epidermidis and Escherichia coli

a. Take your streak plates from the last lab period and examine them for the two different colony types. The TSA plate should have well-isolated Staphylococcus epidermidis (Gram-positive) and Escherichia coli (Gram-negative) colonies. The PEA and MSA should only have one colony type (S.epidermidis), and the MAC and EMB should only have one colony type (E.coli).

b. Assemble the materials necessary for making the smears.

c. With a pencil, label two glass slides on the frosted end with the names of the respective test organisms: Staphylococcus epidermidis and Escherichia coli.

d. Using the aseptic techniques demonstrated by the instructor put a medium-sized drop of water on the slide in the center, using a sterile pipette or an inoculating loop. Transfer a small amount from a single, well-isolated colony from the Petri plate to the drop of sterile water on the slide. When transferring an isolated colony from the streak plate, an inoculating needle rather than a loop is used.

e. Touch the inoculating needle to the center of a well-isolated colony. You may use any one of your plates. However, if you use a selective agar, remember that the bacterial type that did not appear to grow is only inhibited. Therefore, you should touch the needle to the very top or edge of the colony without going too deep. DO NOT TOUCH THE AGAR SURFACE! Transfer the colony aseptically to the appropriately labeled glass slide and thoroughly mix the bacteria with the drop of sterile water on the slide.

f. Repeat the procedure for the other colony type.

g. Let each smear air-dry thoroughly and then heat-fix gently using either the flame of a Bunsen burner or a slide warmer. Heat-fix the bacteria onto the slide by passing the slide, smear side up, quickly through the flame of the bunsen burner 4-5 times. Avoid getting the slide too hot; this will cause distortion of the morphology of the cells. This step will keep your smear from washing off of the slide during the staining procedure.

These smears will be used to perform the Gram Stain procedure.

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THE GRAM STAIN

Objectives: After completion of this laboratory exercise, the student will be able to:

1. Explain the technique and theory of the Gram Stain. 2. Describe bacterial cell morphology. 3. Explain the importance of the Gram stain as an important step in the identification of a bacterial species. 4. Properly perform a Gram stain.

Individual bacterial cells exhibit morphology typical of their species: size, shape, and arrangement of cells. These can be demonstrated by making a smear on a glass slide, then staining the smear with a suitable dye. The use of a stained smear permits microscopic examination of the smear with the oil immersion lens, which gives the greatest magnification, revealing the size, shape, and arrangement. The study of individual bacterial cells is thus frequently one of the first steps in the identification of bacteria.

In this exercise you will use the Gram stain. This is called a differential stain, because it not only shows bacterial morphology but allows differentiation of different bacterial types since different species react differently to the stain. The differential Gram stain gives information about the bacterial cell wall, which may be gram-positive or gram-negative. Gram-positive bacteria will appear purple, the color of the primary stain, crystal violet. Gram- negative bacteria will appear pink-red, the color of the counterstain, safranin. The Gram stain is especially useful as one of the first steps in the identification of a bacterial species, since it reveals both the morphology and the Gram reaction of the bacteria.

The bacteria may show the following shapes: coccus/cocci(spherical), bacillus/bacilli(rod-shaped), or spirillum/spirilli(curved or spiral). The cells may assume a characteristic arrangement: some occur singly, others appear in pairs (diplo-), chains (strepto-), or clusters (staphylo-).

Materials:

1. slides of Staphylococcus epidermidis (Gram +)and Escherichia coli (Gram -) 2. wash bottle (with tap water) 3. rinse bucket 4. clothespins (slide holders) 5. absorbent mat 6. glass marker 7. reagents used in the Gram stain: crystal violet Gram's iodine 95% ethyl alcohol safranin

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The Gram Stain Procedure

1. Add crystal violet stain until the slide is completely covered. Stain for one minute.

2. Do not drain the stain off of the slide before rinsing, because the crystal violet will form dye crystals on the slide. Dilute the crystal violet stain on the slide with a gentle stream of water from a wash bottle. Then tip the slide and drain off the stain, and continue rinsing until all the purple color has washed off of the slide. Drain off excess rinse water. If viewed under the microscope at this point, all bacterial cells will appear purple.

3. Cover the slide with Gram's iodine solution and let it stand for one minute. This step will not change the color of the cells; the iodine forms a complex with the crystal violet in the cell wall. Rinse with water, using the wash bottle.

4. Decolorize the smear by letting 95% ethyl alcohol run down over the slide, which should be held at an angle with the clothespin until the purple stain no longer is being visibly removed from the slide. This step should only take a few seconds. (NOTE: a thick smear will take longer to decolorize than a thin one.)

5. Quickly rinse the slide with water. At this stage, if viewed under the microscope, gram-positive bacteria will still appear purple and gram-negative bacteria will appear colorless.

6. Add safranin, the counterstain, to cover the slide. Stain for two minutes. At this stage, if viewed under the microscope, gram-positive bacteria will still appear purple, and gram-negative bacteria will appear the color of the counterstain, pink-red.

7. Rinse with water, and let the slide air-dry or blot gently (DO NOT RUB) with bibulous paper. The slide must be completely dry before adding oil for observation under the oil-immersion lens.

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USE OF THE MICROSCOPE

Objectives: After completion of this laboratory, the student will be:

1. acquainted with the basic principles of compound light microscopy. 2. able to properly use the low power, high power, and oil immersion objectives. 3. able to exercise the steps necessary for proper care of a microscope.

In microbiology, the small size of the microorganisms requires that you become a microscopist. Development of this skill requires practice and experience. The purpose of this exercise is to allow you to become familiar with the use of the microscope. At first you are all thumbs, but with patience and practice, you will become better as time progresses.

First, familiarize yourself with the parts of the microscope and their functions. Refer to your textbook for complete descriptions. Starting at the base of the microscope and following the path of light upward: Illuminator = lamp or light source Substage condenser = a lens system located below the microscope stage that directs (“condenses”) the light rays through the specimen Iris diaphragm = controls the amount of light that can pass through the condenser; integrated into the condenser itself and is usually controlled by a rotating ring or a lever Mechanical stage = platform with clips that hold the specimen (microscope slide) in place; the slide can be moved up/down and side to side using stage knobs Objective lenses = primary lenses that magnify the specimen Body tube = transmits the image from the objective lens to the ocular lens Ocular lens (eyepiece) = remagnifies the image received from the objective lens Coarse adjustment/focusing knob = used initially to bring the desired image into view Fine adjustment/focusing knob = used to make final focus adjustments to the image

There are two sets of lenses that make up the magnification system in a compound light microscope. The objective lenses provide the initial magnification of the specimen. This "real image" is then projected up through the body tube to the ocular lens, which magnifies the real image 10X. This is the image that is seen by your eyes.

Microscopes for bacteriological use are usually equipped with at least three objectives: (1) low power (10X magnification) (2) high power (40-45X) (3) oil immersion (100X) The desired objective is rotated into place by a revolving nosepiece.

To calculate the total magnification, the power of the ocular lens (10X) is multiplied by the power of the objective being used (10X, 40X, or 100X).

Proper illumination is a major part of compound light microscopy. The amount of light entering the objective lens is regulated in three ways: (1) raising or lowering the amount of light coming from the lamp or light source, (2) opening or closing the iris diaphragm (3) focusing the light up through the objective is controlled by raising or lowering the condenser.

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With increasing magnification, the objective lens requires more light. For example, when the oil immersion objective is used, the maximum amount of light possible is necessary. To achieve this, the lamp must be turned up all the way, the condenser is raised up to stage level, and the iris diaphragm is opened completely. The lamp, condenser, stage, objective, and ocular lenses must be kept clean to achieve optimal results. The lenses are highly susceptible to scratching, so they must be cleaned carefully. This can be done by moistening a piece of lens paper with special lens cleaner, wiping off the lens, and then drying it off with a piece of dry lens paper. To clean oil from the lenses on stage, use the same procedure until no oil is seen on the lens paper.

Precautions: 1. Do not touch the lenses with your fingers. Always use special lens cleaning paper. 2. Do not force the adjustments. If you have problems making adjustments, consult the instructor before proceeding. 3. Always clean off the lenses and stage with special cleaner and lens paper before putting your microscope away. 4. After each use, the following steps should be followed:

a. clean off all lenses and the stage d. rotate the 4X or 10X objective into place b. make sure the light is turned off e. wrap the cord around the base c. lower the condenser and the stage f. cover the microscope with a plastic cover

Procedure:

1. Place the microscope on your desk and identify the different parts of the microscope and their function. Refer to your textbook for a diagram and description of each microscope part, and the path of light through the microscope.

2. Obtain a stained bacterial smear from your instructor, or use one of the smears that you have prepared yourself. Make sure that the smear side is up before placing it on the microscope stage.

3. Place the slide on the stage with the smear centered over the opening.

4. Rotate the low-power (10X) objective into position. For initial coarse focusing, first use the large coarse adjustment knob. The fine adjustment knob, the smaller knob, can then be used to complete your focusing.

5. After examining the smear under low power, rotate the nosepiece until the high-dry objective (40X) snaps into place. You should only have to refocus slightly, using the fine adjustment knob.

6. Note the increased size of the bacterial cells and the decreased number of cells present per microscopic field.

7. For practice focusing with the oil-immersion objective (100X), place a drop of immersion oil on the slide, over the area of the smear. Lower the oil-immersion objective slowly until it just touches the oil.

8. Next bring the specimen into a fuzzy focus very slowly with the coarse adjustment knob, and then into sharp focus with the fine adjustment knob. The field will come in and out of view quickly.

9. If the microscope is parfocal, an alternate method is to find the smear with the low power objective (10X) or high power objective (40X) and then carefully switch over to the oil immersion lens.

10. Sketch and describe the appearance of the cells on the Results Sheet.

11. Remove the slide when finished and put it into one of the cans labeled "for glassware only." 16

RESULTS

1. After performing the Gram Stain procedure on your bacterial smear, use the oil-immersion objective to examine the bacteria. You should see a mixture of two different species of bacteria: one gram-positive, and one gram-negative. Sketch the appearance of each type of cell. Describe the morphology and give the gram reaction. (Be sure to use the correct terminology in describing the morphology.*)

(*morphology means size, shape, and arrangement)

Morphology: ______Gram reaction: ______

Morphology:______Gram reaction:______

17

2. Examine the unknown pre-prepared Gram stains provided by the instructor. Sketch a few cells of each, describe the morphology, and give the gram reaction.

A. Morphology: ______Gram reaction: ______

B. Morphology: ______Gram reaction: ______

C. Morphology: ______Gram reaction: ______

D. Morphology: ______Gram reaction: ______

18

STUDY QUESTIONS

1. What conclusion can you make about the relationship between the size of the microscopic field (average number of organisms per field) and the magnification used?

______

______

______

2. How do you determine the actual total magnification of the specimen you are looking at? (Show your calculations for each of the three objectives.)

______

______

a. low power objective:

b. high-dry objective:

c. oil-immersion objective:

3. Why do you have to use the oil-immersion objective to view bacteria?

______

______

4. Describe the type of information the Gram stain can give:

the microbiologist

the physician.

19

5. Fill in the following table:

Appearance of bacterial cell after each step (color) Steps Gram positive cell Gram negative cell

crystal violet

Gram's iodine

95% alcohol

safranin

6. What is the function of the Gram's iodine (the mordant) in the Gram stain?

7. What is the function of the safranin counterstain?

8. What is the function of the 95% alcohol decolorizer?

9. Explain the chemistry behind how the Gram stain distinguishes between gram-positive and gram-negative bacteria.

20

Special Staining Techniques: Acid-fast Bacilli Stain, Capsule Stain, Endospore Stain, and Flagella Stain

Objectives: After completion of this laboratory exercise, the student will be able to:

1. Identify special bacterial structures: capsules, flagella, and endospores. 2. Explain the significance of these bacterial structures in diagnosis and identification of disease. 3. Perform or describe the techniques that identify these special structures. 4. Identify acid-fast bacilli on a stained preparation. 5. Explain the significance of acid-fast bacilli in a specimen.

Some bacteria possess cell walls and other structures that are best demonstrated by methods other than the Gram Stain. This exercise deals with a differential stain for the special type of waxy cell walls possessed by Mycobacterium and with methods used to demonstrate endospores, capsules, and flagella. In addition to their value in identification of certain bacteria, demonstration of these structures is important for your understanding of the basic structure and function of bacterial cells in disease processes.

Bacterial Capsules

The capsule is a gelatinous, slimy material surrounding the bacterial cell. In many cases the capsule helps protect the cell against phagocytosis. Thus potential pathogens are protected from the body's natural defenses and are more likely to cause disease than non-capsulated strains. The capsule also allows bacteria to adhere to surfaces, such as mucous membranes and teeth. Other functions of a capsule include protection from dehydration and loss of nutrients. In this exercise, capsules are demonstrated by the negative stain, in which the capsule shows up as a clear area or halo surrounding the cell against the dark background of nigrosin stain.

Bacterial Flagella

Flagella are structures that enable bacteria to be motile. They may occur singly at one end, in tufts at one or both ends, or arranged all around the cell. monotrichous = a single flagellum amphitrichous = a single flagellum at both ends of the cell lophotrichous = two or more flagella at one or both ends of the cell peritrichous = flagella distributed over the entire cell

The number and arrangement of flagella can be used to help identify bacteria.

Flagella are demonstrated by special stains using mordants that increase the width of the flagella and are then stained with carbol-fuchsin so that they may be seen with the microscope. NOTE: The pink color of the microbes is due to the color of the primary carbol-fuchsin stain, and is NOT an indication of a gram reaction, as in the Gram stain procedure.

Bacterial Endospores

Endospores are very resistant structures that are formed by certain bacteria under adverse conditions. Two genera of gram-positive bacilli (rods) are endospore-formers: Bacillus and Clostridium. Endospores enable the organism to survive drying and lack of nutrients, so they can exist in dust and soil for many years. Endospores are the most resistant form of life known. Their presence in dust accounts for much of the laboratory contaminants. The very thick spore wall does not stain easily, so the endospores will appear in Gram stains as unstained areas inside the cell. To stain the spores themselves, carbol-fuchsin stain is heated so that it will be absorbed by the wall of the endospore so that they appear red. The vegetative part of the cell will decolorize upon rinsing with 95% ethanol and can then be counterstained with methylene blue or brilliant green for contrast.

21

Acid-Fast Bacilli

The cell walls of the genus Mycobacterium, which includes the pathogens of tuberculosis and leprosy, are different from most other types of bacterial cell walls because they are waxy and stain poorly, if at all. However, they will take up the acid-fast stain. This stain uses carbol-fuchsin to which phenol has been added. The cell wall then resists decolorization with acid-alcohol. (alcohol plus hydrochloric acid; thus the name "acid-fast") The end result is an organism that retains the carbol-fuchsin color. Other organisms will decolorize with the acid-alcohol and will take up the counterstain brilliant green or methylene blue. Mycobacterium species are therefore often called "acid-fast bacilli" (AFB).

Materials: Prepared demonstration slides of capsules, flagella, endospores, and acid-fast bacilli

Procedures:

I. Capsule stain by the negative method Examine the demonstration slides with oil immersion for the presence of capsules. They should appear as tiny, unstained, “halos” around the bacteria cells. The bacteria may be seen inside the capsule as tiny blue bacilli.

II. Flagella stain: Examine the demonstration slides under oil-immersion for the presence of flagella. They should appear as thin, whip-like "tails". Remember, this in not a gram stain, and the color does not designate a gram reaction.

III. Endospores stain Examine the demonstration slides under oil-immersion for bacterial endospores. They will appear as small pink or colorless circles or ovals inside the streptobacilli.

IV. Acid-fast stain Examine the demonstration slides with oil-immersion for the presence of the acid-fast organisms (“AFB” = acid-fast bacilli). They should appear as clumps (“cords”) of tiny, fuschia-colored bacilli. Other, non- acid fast bacteria will appear blue.

Acid-fast stain: (Kinyoun method) 1. Prepare a smear of Mycobacterium. These cells are waxy, so your smear preparation will not mix with the water very easily. Air-dry and heat fix. 2. Cover the slide with carbol-fuchsin with phenol (Kinyoun) and leave it in the stain for 5 minutes. 3. Rinse with water. 4. Decolorize with acid-alcohol for a few seconds; rinse immediately. (Be sure to use acid-alcohol, not 95% alcohol) 5. Counterstain with methylene blue for 3 minutes. 6. Rinse with water and blot dry. 7. Examine with oil-immersion for the presence of the acid-fast organisms. They should appear as tiny, fuschia-colored bacilli in clumps called “cording”. Other microbes will appear blue.

22

RESULTS SHEET SPECIAL STAINS

Examine the special stains provided by the instuctor. Draw the appearance of the structures. Describe the appearance of the structure and the bacterial cell. Label your diagrams.

I. Capsules

II. Endospores

III. Acid-fast bacilli

IV. monotrichous flagellum

amphitrichous flagella

lophotrichous flagella

peritrichous flagella

23

STUDY QUESTIONS SPECIAL STAINS

I. What is the importance of performing these special stains? What information do they give you?

______

______

______

______

II. a. Is a bacterium that possesses a capsule always considered a pathogen?

______

______

b. What are the functions of a capsule?

______

______

III. a. Why are endospores important to a bacterial cell? Under what conditions are they formed?

______

______

b. What genera of bacteria can produce endospores?

______

c. Give an example of the genus and species of four(4) pathogenic bacteria that produce bacterial endospores.

______

______

______

IV. a. What are the genera that the acid-fast stain is used to identify?

______

b. Name two diseases that can be diagnosed with the aid of the acid-fast stain.

______

______

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V. What is the function of flagella?

______

______

VI. Write a brief explanation of why each one of the following bacterial structures requires a "special" staining technique in order to be observed. (Explain why they cannot be demonstrated using a Gram Stain.)

a. capsule

b. endospore

c. acid-fast bacilli

d. flagella

rev 5/97 tt 25

Soil Serial Dilution-Plate Count

Objectives: After completion of this laboratory, the student should be able to: 1. Perform serial dilutions of soils 2. Determine the number of viable bacterial cells in a soil sample using both the pour plate and spread plate methods 3. Characterize colony morphology of soil bacteria

Materials: 1. 450C water bath 2. Analytical Balance 3. Aluminum weigh dish 4. Plastic weigh dish 5. Oven for drying soil 6. 25g of student soil 7. 4 test tubes containing 9 ml of sterile water 8. 15 sterile 1 ml serological pipettes 9. 6 empty Petri plates 10. 3 Actinomycete agar plates 11. 6 screw cap test tubes containing melted nutrient agar (in 450C water bath) 12. 1 milk dilution bottle containing 95 ml of sterile water 13. Plastic spoons 14. funnels

Pour Plate Procedure for Bacteria: 1. Label six empty Petri dishes as 1A, 1B, 2A, 2B, 3A, and 3B.

2. Label the test tubes #1-4

3. Using an analytical balance, weigh out 10g of soil into an aluminum weigh dish

4. The instructor will place the aluminum weigh dish into an oven for 24 hours to remove the moisture from the soil. The following lab you will weigh your dry soil.

5. Weigh out another 10g of soil into a plastic weigh dish

6. Place the 10g of soil into the milk dilution bottl e containing 95 ml of sterile water. Tighten the cap and shake vigorously 30 times. Your soil has been diluted 10 times (10-1 w/v).

7. Using a sterile pipette, aseptically transfer 1ml of the bacterial culture from the milk dilution bottle into test tube #1. This soil has been diluted 10 times (10-2). Discard the pipette in the disinfectant.

8. Take a fresh pipette and mix tube #1 by pipetting up and down several at least 5 times . Using the same pipette transfer 1 ml from tube #1 to tube #2. This soil has been diluted 100 times (10-2). Discard the pipette in the disinfectant.

9. Take a fresh pipette and mix tube #2 by pipetting up and down at least 5 times . Using the same pipette transfer 1 ml from tube #2 to tube #3. This soil has been diluted 1000 times (10-3). Discard the pipette in the disinfectant.

10. Take a fresh pipette and mix tube #3 by pipetting up and down at least 5 times . Using th e same pipette transfer 1 ml from tube #3 to tube #4 . This soil has been diluted 10,000 times (10 -4). Discard the pipette in the disinfectant. 26

11. Using a fresh pipette transfer 0.1 ml o f liquid suspension from tube #1 to plate #1A. Discard this pipette into the disinfectant.

12. Using a fresh pipette transfer 1 ml of liquid suspension from tube #2 to plate #1B and 0.1 ml of liquid suspension to plate 2A. Discard this pipette into the disinfectant.

13. Using a fresh pipette transfer 1 ml of liquid suspension from tube #3 to plate 2B and 0.1 ml of liquid suspension to plate 3A. Discard this pipette into the disinfectant.

14. Using a fresh pipette transfer 1 ml of liquid suspension from tube #4 to plate #3B. Discard this pipette into the disinfectant.

15. Remove a screw cap tube with agar from the 45 0C water bath and pour the agar immediately into plate 1A aseptically. Gently rotate the plate to evenly distribute the bacterial cells in the medium.

16. Repeat step 15 for the remaining Petri plates.

17. Once the media is solidified, invert the plates and incubate them at room temperature for one week in your student drawer.

Pour Plate Follow-up Lab 1. Count all colonies (surface and subsurface colonies) on the plates using a colony counter. Count only the plates that have between 30-300 colonies (to be statistically significant). Plates that have more than 300 colonies cannot be counted and are referred as too numerous to count (TNTC) . Plates that have fewer than 30 colonies cannot be counted and are referred as too few to count (TFTC).

2. If the plate you have chosen to count has closer to 30 colonies, you may count the entire plate. However, if the plate you chose to count has closer to 300 colonies, you may want to use the following method to estimate the total number of colonies: a. choose ten square centimeters at random and count all of the colonies in each square b. total the ten squares and divide by 10 to get the average # of colonies/cm2 c. multiply this number by the area of the Petri dish = πr2 (π = 3.14, r = 5 cm) This will give an estimate of the total number of colonies on the entire plate.

3. Calculate the number of bacteria per gram of original sample by using the following formula:

a) Number of cells/g= number of colonies on entire plate X dilution factor (reciprocal of dilution) OR b) Number of cells/g= number of colonies from 10 squares X πr2 X dilution factor 10 For example, if your final count is 150 CFU on a plate with 10-5 dilution your answer would be: 150 = 15 X 106 CFU/10g moist soil 10-5

4. Next, calculate how many microorganisms are in one gram of dry soil. To do this, divide your answer by your soil dry weight. For example, if 10g of soil weighed 7g after drying, your answer would be:

15 X 106 CFU/10g moist soil = 2.14 X 106 CFU/g dry soil 7g dry soil

27

Spread Plate Procedure for Actinomycetes: 1. Label the three actinomycete agar plates 1, 2, and 3.

2. Take a fresh pipette and mix tube #1 by gently pipetting up and down. Use this pipette to transfer 0.1 ml of liquid suspension from tube #2 to plate #1. Discard this pipette into the disinfectant.

3. Sterilize a glass spreader by dipping it into 70% isopropanol and passing it through the Bunsen burner flame (Do not keep the glass spreader in flame for more than a few seconds; it will melt!)

4. Allow the glass spreader to cool and gently spread the 0.1 ml of bacterial suspension uniformly across the surface of the agar plate.

5. Take a fresh pipette and mix tube #2 by gently pipetting up and down. Use this pipette to transfer 0.1 ml of liquid suspension from tube #2 to plate #2. Discard this pipette into the disinfectant.

6. Repeat steps 3-4 above

7. Take a fresh pipette and mix tube #3 by gently pipetting up and down. Use this pipette to transfer 0.1 ml of liquid suspension from tube #3 to plate 3. Discard this pipette into the disinfectant.

8. Repeat steps 3-4 above

9. Invert the plates and incubate them at room temperature for two weeks in your student drawer.

Spread Plate Follow-up Lab 1. Follow procedures for the pour plate follow -up lab above. Record your actinomycete counts/g of soil sample in the table below

28

Record your bacterial counts/g of sample in the table. Plate # Dilution Amount Dilution # of Soil dry Bacteria (CFU/g) tube plated (ml) factor colonies weight 1A 1B 2A 2B 3A 3B

Record your actinomycete counts/g of sample in the table. Plate # Dilution Amount Dilution # of Soil dry Actinomycete tube plated (ml) Factor colonies weight (CFU/g) 1

2 3

1 ml 1 ml -2 1 ml -3 1 ml -4 1 ml -5 10-1 10 10 10 10

Soil Sample 10g #1 #2 #3 #4

H20 H20 H20 H20 H20 95 ml 9 ml 9 ml 9 ml 9 ml

Volume of sample 1.0 ml to be added onto 0.1 ml 0.1 ml 1.0 ml 0.1 1.0 ml ml plates

Bacteria Plate 1A 1B 2A 2B 3A 3B Actinomycete Plate #1 #2 #3 Dilution 10-2 10-3 10-3 10-4 10-4 10-5

Dilution factor 103 103 104 104 105 105

rev 10/2012 EW

29

IDENTIFICATION OF GRAM-POSITIVE COCCI

Objectives: After completion of these laboratory exercises, the student will be able to:

1. Name the medically significant Gram-positive cocci. 2. List the media and biochemical tests that are commonly used to identify Gram-positive staphylococci and Gram-positive streptococci. 3. Explain the theory behind the following tests for the identification of Gram-positive staphylococci: mannitol salt agar, catalase, coagulase. 4. Describe the actions of the enzymes catalase and coagulase as they relate to microbial metabolism and pathogenicity. 5. Define hemolysis, hemolysin. 6. List the three types of hemolysis produced by Gram-positive streptocci on blood agar media and describe the appearance of each type. 7. List the medically significant streptococci that produce each of the three types of hemolysis. 8. Explain how the production of hemolysis relates to pathogenicity. 9. Identify the type of hemolysis produced by various species of streptococci on blood agar.

BIOCHEMICAL TESTING FOR THE IDENTIFICATION OF GRAM-POSITIVE COCCI

In a clinical microbiology, specimens from infected patients are cultured, and then pathogens must be distinguished from normal and transient microbiota. Normally, the first step in this identification process is to perform a microscopic examination of the morphology and staining characteristics of the suspected pathogen by performing stains such as the Gram stain. However, the problem is that through a microscope, there is often too much similarity between organisms to rely on microscopic descriptions alone. For example, there are numerous bacterial species that are gram-positive cocci.

Therefore, further testing must be done to identify bacteria. These include the use of selective and differential media, and biochemical tests.

IDENTIFICATION OF STAPHYLOCOCCI

Staphylococci are gram-positive cocci in clusters. After a Gram stain has determined that the organism to be identified is a gram-positive coccus in clusters, the tests for identification of staphylococci can be performed. (Note: other species of bacteria can also have biochemical activity similar to that of the staphylococci, such as production of the enzymes catalase and coagulase; therefore, a test is meaningless with first performing the Gram stain.) It is important to be able to distinguish Staphylococcus. aureus from other staphylococcus species. Staphylococcus aureus can be part of the normal flora of the skin and upper respiratory tract, but it is also a potential pathogen. S. aureus is one of the most common causes of nosocomial (hospital-acquired) infections. Other species of staphylococci, such as S. epidermidis and S.saprophyticus, are also part of the normal flora, but are not normally pathogenic.

30

Biochemical tests used to identify Staphylococci:

Three biochemical tests that are commonly used to isolate, differentiate, and identify Staphylococci are: 1. mannitol salt 2. catalase 3. coagulase

1. Mannitol salt agar (MSA) is a type of selective and differential medium that can be used to isolate staphylococcus species from a specimen. MSA is selective for staphylococci because of the high salt content; only staphylococci will grow on mannitol salt agar. All other organisms are inhibited. MSA is also differential for staphylococci: S. aureus will cause the agar to turn yellow because of the fermentation of the carbohydrate mannitol in the agar; other species of staphylococci (such as S. epidermidis) will not change the color of the agar because they do not ferment mannitol, and it will remain red.

2. Staphylococci are capable of producing the enzyme catalase. This enzyme can be tested for by mixing the bacteria in question with a drop of hydrogen peroxide. If catalase is being produced, the following chemical reaction will occur: catalase 2H2O2 + bacterium -----> 2H2O + O2

The oxygen that is liberated will produce a bubbling effect.

3. As a potentially pathogenic organism, S. aureus produces an invasive enzyme, coagulase. This enzyme is capable of coagulating plasma. This clot may protect the bacteria from phagocytosis and isolate them from the body's defenses. Coagulase production can be tested for by mixing the bacteria in question with sterile plasma. This mixture is allowed to incubate at body temperature (37°) for several hours. If the mixture coagulates, the test is positive for coagulase.

** In summary, S. aureus is catalase positive and coagulase positive, with yellow growth on mannitol salt agar. Other species of staphylococci, such as S. epidermidis, are catalase positive and coagulase negative, with red growth on mannitol salt agar.

31

MANNITOL SALT AGAR for the selection and differentiation of Staphylococcus species

1. Obtain a mannitol salt agar plate that has been divided into three sections. Label the bottom of the plate with your name, date, course, and section number.

2. Label one section "A", the second section "B", and the third section "C".

3. Aseptically streak out the unknown organism "A" on that third of the plate. Repeat the procedure for unknown organism "B" and "C".

4. InvertInver tthe the plate plate and and incubate incubate for for 24-48 24-48 hours at 37C. hours at 37 degrees Celsius 5. After the incubation period, observe each section of the agar for bacterial growth. Staphylococci can tolerate high concentrations of salt and will grow on MSA; other organisms will not grow well, if at all..

6. Also observe each section of the plate for a change in the color of the agar. The presence of a distinct yellow color indicates fermentation of the mannitol sugar by S. aureus. Other staphylococci species will not change the color of the agar.

7. Record your results on the Results Sheet.

8. Discard the used culture plates into the buckets marked "For Plastic Petri Dishes Only".

32

SLIDE CATALASE TEST for the detection of Staphylococcus species

1. Obtain three clean, glass microscopic slides.

2. Label the first slide “A”, the second “B”, and the third “C”.

3. Aseptically place a drop of hydrogen peroxide onto each slide.

4. Using a sterile inoculating needle, aseptically transfer a visible amount of unknown organism "A" to the hydrogen peroxide on slide "A" and mix. Observe for the immediate production of vigorous oxygen bubbling, which indicates a positive catalase test. Little or no bubbling is a negative catalase test. Record your observation on the Results Sheet.

5. Sterilize your transfer needle and repeat Step #4 for organism "B" and "C" and record your results.

6. Discard the slides in disinfectant.

7. The presence of vigorous oxygen bubbling indicates that the hydrogen peroxide has been broken down by the enzyme catalase. Little or no oxygen bubbling is a negative for catalase activity. All Staphylococci produce strong catalase activity.

TUBE COAGULASE TEST for the detection of pathogenic Staphylococcus aureus

1. Obtain three (3) small test tubes containing sterile rabbit plasma.

2. Label each tube with a piece of tape with your name, date, course, and section number. Label one tube "A", the second tube "B", and the third tube "C".

3. Using a sterile inoculating loop, transfer a loop-full of unknown organism "A" into tube A.

4. Repeat Step #3 with unknown organisms "B" and "C".

5. Incubate the inoculated plasmas at 370C for 6-24 hours.

6. Observe each tube for coagulation of the plasma by tilting the tube slightly. If the plasma is still liquid, the test is negative for coagulase activity. If the plasma has coagulated, it will be semi-solid, and the test is considered positive for coagulase activity.

7. Record your results on the Results Sheet.

8. Place the culture tubes into a rack in the corner for "Items to be Autoclaved".

33

RESULTS SHEET

UNKNOWN ORGANISM: A B C

growth on mannitol salt agar (yes or no)

color of mannitol salt agar (yellow or red)

Slide catalase test: bubbles(+) or little/no bubbling (-)

Tube coagulase test:

plasma coagulated(+) or liquid (-)

IDENTIFICATION OF UNKNOWN ORGANISM:

8/93 ge rev 5/97 tt

34

IDENTIFICATION OF STREPTOCOCCI

If a Gram stain performed on a patient's specimen or from a culture shows the presence of gram-positive cocci in pairs or chains, this morphology is typical of streptococci. Streptococci are responsible for more infectious disease processes than any other type of bacteria. Therefore, differentiation and identification of streptococci is an important step in diagnosis.

There are many different species of streptococci, which makes them more difficult to identify. One method for the differentiation of streptococci is to divide them into groups based on their action on blood agar. This action is called “hemolysis”, which means “breakdown of red blood cells”. Streptococci produce enzymes called "hemolysins" that cause this breakdown. The type of hemolysis on blood agar is the most important test in the identification of the different groups of streptococci. The three groups of streptococci are:

1. beta-hemolytic streptococci 2. alpha-hemolytic streptococci 3. non-hemolytic (gamma) streptococci

Beta-hemolytic streptococci produce colonies on blood agar that are surrounded by a relatively clear zone of hemolysis in which the red blood cells in the agar are completely lysed. Many serious infections such as pharyngitis, scarlet fever, impetigo, rheumatic fever, and glomerulonephritis are caused by the beta-hemolytic species Streptococcus pyogenes. Another beta-hemolytic streptococcus species, Streptococcus agalactiae, is often the cause of bacterial meningitis in newborns, and can also cause childbirth sepsis. (This is due to the fact that S. agalactiae is present in the vaginal normal flora of up to 25% of all women.)

Alpha-hemolytic streptococci produce colonies on blood agar that are surrounded by a greenish zone of hemolysis, due to the incomplete breakdown of the hemoglobin in the red blood cells. Streptococcus pneumoniae is an example of a pathogenic alpha-hemolytic streptococcus. S. pneumoniae causes pneumonia, ear infections (otitis media), and meningitis. Other alpha-hemolytic streptococci are primarily normal flora, such as Streptococcus salivarius and Streptococcus mutans, found in the mouth. Collectively, these non-pathogenic streptococci are called "viridans” strep.

Gamma or non-hemolytic streptococci do not produce any hemolysis on blood agar. Enterococcus faecalis is an example of a non-hemolytic streptococcus that is normally found in the intestinal tract, and is therefore included in a group of streptococci called the "enterococci". These enterococci can migrate to other areas of the body to cause conditions such as urinary tract infections or peritonitis.

35

After determination of the type of hemolysis produced by a streptococcus colony on blood agar, further biochemical tests should be performed to identify the species of streptococcus. For example, the tests used to identify the various species of beta-hemolytic streptococci are different from those used to identify the alpha- hemolytic streptococci. The following is a summary of some of the biochemical tests commonly used to identify streptococcus species:

Beta-hemolytic Strep Alpha-hemolytic Strep Gamma-hemolytic Strep

bacitracin optochin bile esculin sensitivity sensitivity hydrolysis

hippurate growth in 6.5% salt hydrolysis

LANCEFIELD ANTIGENIC GROUP (SEROLOGICAL) TYPING

Beta-hemolytic streptococci and enterococci possess chemicals called CH (carbohydrate) antigens. The presence and type of CH antigen can be demonstrated by extraction of the antigen from the cell, and reacting it with antibodies specific to each antigen. Lancefield found thirteen different antigenic groups, A-O. Of these, Groups A, B, and D are most commonly implicated in human infections. Groups C, F, and G are also occasionally cultured from patients.

Group Major Species

A S. pyogenes

B S. agalactiae

D E. faecalis* S. faecium* S. durans* S. avium*

(*enterococci)

36

IDENTIFICATION OF BETA-HEMOLYTIC STREPTOCOCCI

The two most common beta-hemolytic streptococcal pathogens are Streptococcus pyogenes and Streptococcus agalactiae. It is important to differentiate these two beta-hemolytic strep species from other beta-hemolytic strep and from each other for a correct diagnosis.

TESTING FOR BACITRACIN SENSITIVITY

S. pyogenes is sensitive to the antibiotic bacitracin, whereas other beta-hemolytic strep are not. When a paper disk impregnated with bacitracin is placed on a blood agar plate upon which S. pyogenes is growing, there will be a zone of inhibition around the bacitracin disk where the S. pyogenes cannot grow. This is a positive test for S. pyogenes.

Observe the demonstration blood agar plates of:

1) beta-hemolytic S. pyogenes (also known as “Group A” strep by Lancefield typing), sensitive to bacitracin 2) beta-hemolytic strep species that is resistant to bacitracin. (further I.D. required)

In summary: S. pyogenes = beta-hemolytic, sensitive to bacitracin resistant to bacitracin = other species of beta-hemolytic streptococci; (*further ID required)

If the organism is a beta-hemolytic streptococcus that is resistant to bacitracin, the next step in the identification process is to perform further testing to determine whether it is a Group B strep such as S. agalactiae or some other beta-hemolytic strep such as Groups C, F, or G.

TESTING FOR HIPPURATE HYDROLYSIS

The hippurate test is used in the identification of beta-hemolytic Group B streptococci (S. agalactiae) by detecting the ability of the organism to hydrolyze (break down) hippurate.

Procedure: 1. To a hippurate test tube, add 3-4 drops of distilled water. 2. Using a heavy inoculum (a full loop) from an 18-24 hour culture, make a heavy suspension of the organism in the Hippurate Reagent with a standard inoculating loop. 3. Incubate the tube for 2 hours at 37 degrees C. 4. After the 2 hour incubation period, add 2 drops of the Ninhydrin Indicator solution to the Hippurate Reagent/organism mixture. Ninydrin acts as an indicator to detect glycine, a byproduct of hippurate Hydrolysis. 5. Reincubate at 37 degrees C for 30 minutes. Observe the tubes at 10 minute intervals for the appearance of a deep blue/violet color, which is a positive test. The color change will usually appear in 10-15 minutes after the Ninhydrin Indicator solution has been added. A negative reaction is indicated by a faint blue color or no color change.

Observe the demonstration of the hippurate hydrolysis test:

1) hippurate (+) S. agalactiae (Group B strep) 2) hippurate (-) (*further I.D. req.)

In summary:

S. agalactiae = beta-hemolytic, bacitracin (R), hippurate hydrolysis (+) beta-hemolytic, bacitracin (R), hippurate hydrolysis (-) = other beta-hemolytic streptococcus species (*further I.D. required*.) 37

IDENTIFICATION OF ALPHA-HEMOLYTIC STREPTOCOCCI

The most common human pathogen in the alpha-hemolytic streptococci group is Streptococcus pneumoniae (also called the pneumococcus). Most other species of alpha-hemolytic strep are usually normal flora of the oral cavity or upper respiratory tract. As a group, these streptococci are called "viridans" strep. This group consists of at least ten different known species, including S. mutans, the oral bacteria implicated in the formation of dental caries. To differentiate S. pneumoniae from the viridans streptococci, one of the biochemical tests often used is the optochin sensitivity test.

TESTING FOR OPTOCHIN SENSITIVITY

The optochin sensitivity test is similar to the bacitracin sensitivity test, except that the disk used is impregnated with the chemical optochin. The presence of a zone of inhibition around the optochin disk is a presumptive identification of S. pneumoniae.

In summary: optochin sensitive = S. pneumoniae optochin resistant = possible viridans streptococci (*further I.D. required)

Observe the demonstration of the optochin sensitivity tests:

1. alpha-hemolytic, optochin sensitive S. pneumoniae 2. alpha-hemolytic, optochin resistant strep (*further I.D. required.)

38

IDENTIFICATION OF NON-HEMOLYTIC STREPTOCOCCI

The major pathogens in the non-hemolytic (gamma) streptococcus group are the Group D enterococci, such as E. faecalis, S. faecium, S. durans, and S. avium. The most accurate tests for identification of enterococci are the bile esculin (BE) hydrolysis test and growth in 6.5% salt.

TESTING FOR BILE ESCULIN HYDROLYSIS

BE media can be made into agar plates or slants. The surface is then inoculated with the suspected organism and incubated for 24-48 hours. If blackening of the media occurs, the test is positive for bile esculin hydrolysis, and the organism can be identified as part of the group of streptococci called the enterococci.

However, some streptococci that are BE + are not enterococci species. Therefore, another test must be done to differentiate these strep species from the true enterococci. The test used for this purpose is the 6.5% NaC1 tolerance test.

TESTING FOR GROWTH IN 6.5% SALT

The salt can be incorporated into an agar plate or a tube of broth. The media is then inoculated with the strep, incubated for 24-48 hours, and checked for growth. If growth occurs, the organism is an enterococcus.

In summary: bile esculin hydrolysis positive, growth in 6.5% salt = enterococcus group Group D bile esculin hydrolysis positive, no growth in 6.5% salt = non-enterococcus group

Observe the demonstration of the bile esculin hydrolysis and growth in 6.5% salt tests : 1. non-hemolytic, BE (+), salt (+) Group D Enterococcus 2. non-hemolytic, BE (+), salt (-) non-enterococcus

39

Unknown Streptococci RESULTS SHEET

Unknown Bacitracin Hippurate Optochin Bile Growth in # and Sensitivity Hydrolysis Sensitivity Esculin 6.5% salt type of Hydrolysis hemolysis

1

2

3

4

5

IDENTIFICATION:

#1 ______

#2 ______

#3 ______

#4 ______

#5 ______

40

IDENTIFICATION OF GRAM-NEGATIVE COCCI and COCCOBACILLI

If a gram stain performed from a specimen or culture shows the presence of gram-negative cocci or coccobacilli, this morphology is typical of several genera, including Neisseriae, Hemophilus, and Moraxella (formerly Branhamella) Most of these bacteria are normal microbiota of the respiratory, digestive, and genitourinary tracts of humans. However, several species are pathogenic, including Neisseria gonorrhoeae (the “gonococcus”) and Neisseria meningitidis (the “meningococcus”), Hemophilus influenzae, and Moraxella catarrhalis. These microbes often appear as small, kidney-bean shaped diplococci, often seen inside phagocytes on a smear from a clinical specimen, or as small bacteria that have a typical “in between” morphology called “coccobacilli”.

These bacteria grow best on enrichment media such as chocolate agar in an increased CO2 atmosphere. Some are also extremely sensitive to cold; clinical specimens sent to the lab for possible isolation of Neisseriae must not be refrigerated. Specimens typically collected for detection of Neisseriae include cerebrospinal fluid (CSF), cervical or urethral swabs. Hemophilus and Moraxella specimens are most often respiratory or eye samples.

Preliminary identification can be done after 24-48 hours of incubation by gram-staining and testing suspicious colonies for oxidase activity.

THE OXIDASE TEST Procedure

1. Grow a culture of the suspected bacteria on chocolate agar.

2. Put a drop of oxidase reagent directly onto an area of the plate where there are isolated colonies.

3. Wait up to 60 seconds and observe for a color change to dark purple-black.

Results:

Study Questions: 1. If the Gram stain shows gram negative cocci, and the oxidase test is positive, what genus does this bacterium belong to?

______

2. If the Gram stain shows gram negative cocci or coccobacilli, and the oxidase test is negative, what genera might this bacterium belong to?

______

41

IDENTIFICATION OF NEISSERIA SPECIES USING THE API NH system

The API NH system for identification of Neisseria, Hemophilus, and Moraxella species consists of microcupules containing dehydrated test medium. The media are rehydrated by filling them with a heavy saline suspension of bacteria. The strip is then incubated and observed for color changes, which indicates the metabolism of the medium.

Procedure:

1. Set up an incubation tray and lid. Dispense tap water into the bottom of the tray using a squeeze bottle, to provide a humid atmosphere. Record the specimen number on the end flap.

2. Open a pouch and remove an API strip. Place the strip into the incubation tray. The strip should be at room temperature before using.

3. Open an ampule of NaCl 0.85% medium. Using a sterile swab, inoculate the sterile saline with bacteria taken from a culture of the suspected bacteria. This inoculum should be taken from a fresh (18-24 hr) culture on recommended media. Transfer enough inoculum into the saline so that a heavy suspension is achieved. The turbidity should be equivalent to or greater than a No. 4 McFarland standard. Suspensions should be used immediately after preparation.

4. Use a sterile pipette to fill the first seven cupules about 2/3 full with the bacterial suspension. For the last three cups with a box around them, fill the cup all the way up.

5. Cover the first seven cups (those that are underlined) with mineral oil.

6. Place a plastic lid on the tray.

o 7. Incubate the test strip at 37 C for 2 hours in aerobic conditions in a non-CO2 incubator.

Reading the Strip:

1. First, on the result sheet provided, record all reactions as positive (+) or negative (-) before the addition of reagents. (Refer to the Reading Table provided)

2. Add one drop of ZYM B reagent to microcupules 8 and 9 (LIP/ProA and PAL/GGT).

3. Add one drop of JAMES reagent to microcupule 10 (BGAL/IND).

4. Wait three minutes, and then read these reactions according to the Reading Table, and record on the results sheet. Note: If the LIP reaction is blue (+), interpret the ProA reaction as negative, whether the ZYM B reagent has been added or not.

42

READING TABLE

Test Color for positive test result

PEN Penicillinase yellow, yellow-green, yellow-blue

GLU Glucose yellow or orange FRU Fructose yellow or orange “ MAL Maltose yellow or orange“ SAC Saccharose/Sucrose yellow or orange“

ODC Ornithine decarboxylase blue

URE Urease pink-violet

LIP Lipase blue

PAL alkaline phosphatase yellow

BGAL beta galactosidase yellow

ProA proline arylamidase orange

GGT gamma glutamyl transferase dark orange

IND indole pink

HINT: If the test reads any color other than that clearly defined as “positive”, call it negative.

43

Interpretation of Test Results:

Identification is obtained with a numerical profile. To determine the numerical profile, the test results are divided into groups of three on the results sheet. A value of 1, 2 or 3 is assigned to each of the three tests in the group. By adding the three values together for each group, a 4-digit number is obtained. Note: do not code the first test (penicillinase) Example: the first group consists of the tests GLU – FRU – MAL.

Looking up this 4-digit number in the profile list provided or on https://apiweb.biomerieux.com (user name and password required) will give the identification of the organism.

Results:

1. The number of the unknown organism you were assigned:

2. The API NH numerical profile obtained for your organism:

Study Questions:

1. WhatWha tis is the the identification identification of o yourf your organism according to the API NH profile index? organism according to the API NH profile index?______blank______space ______

2. IsI sthis thi sorganism organism normal normal microbiota microbiot aor or a a pathogen? If a pathogen, what type of infectious diseases does it pathogen?cause? If a pathogen, what type of infectious diseases does it cause? blank space ______

______

RESULTS SHEET

Unknown # Identification 1 2 3 4

44

LIST OF API NH NUMERICAL PROFILES

0001 Neisseria cinerea/gonorrhoeae 5424 Haemophilus influenzae 0002 Neisseria meningitidis 5520 Haemophilus parainfluenzae 0010 Branhamella catarrhalis 5560 Haemophilus parainfluenzae 1001 Neisseria gonorrhoeae 5620 Haemophilus influenzae/parainfluenzae 1002 Neisseria meningitidis 5624 Haemophilus influenzae 1003 Neisseria meningitidis 5720 Haemophilus parainfluenzae 1020 Haemophilus influenzae 5724 Haemophilus parainfluenzae 1024 Haemophilus influenzae 5760 Haemophilus parainfluenzae 1103 Neisseria spp. 7000 Neisseria spp. 1224 Haemophilus influenzae 7001 Neisseria spp. 1420 Haemophilus influenzae 7003 Neisseria spp. 1424 Haemophilus influenzae 7020 Haemophilus spp. 1426 Haemophilus influenzae 7022 Haemophilus spp. 1620 Haemophilus influenzae 7024 Haemophilus influenzae/parainfluenzae. 1624 Haemophilus influenzae 7060 H. aprophilus/paraphrophilus/parainfluenzae 1626 Haemophilus influenzae 7062 Haemophilus aphrophilus/paraphrophilus 1720 Haemophilus parainfluenzae/influenzae 7100 Neisseria spp/Haemophilus parainfluenzae 3001 Neisseria spp. 7101 Neisseria spp. 3003 Neisseria spp. 7103 Neisseria spp. 3020 Haemophilus influenzae 7120 H.aphrophilus/paraprophilus/parainfluenzae 3024 Haemophilus influenzae 7122 H.aphrophilus/paraprophilus/parainfluenzae 3026 Haemophilus influenzae 7124 Haemophilus parainfluenzae 3100 Neisseria spp/Haemophilus parainfluenzae 7160 H.aphrophilus/paraprophilus/parainfluenzae 3101 Neisseria spp. 7162 Haemophilus aphrophilus/paraphrophilus 3103 Neisseria spp. 7164 Haemophilus parainfluenzae 3120 Haemophilus parainfluenzae 7220 Haemophilus parainfleunzae/influenzae 3200 Haemophilus somnus 7224 Haemophilus influenzae/parainfluenzae 3204 Haemophilus somnus 7260 Haemophilus parainfluenzae 3220 Haemophilus influenzae 7300 Haemophilus parainfluenzae 3224 Haemophilus influenzae 7320 Haemophilus parainfluenzae 3320 Haemophilus parainfluenzae 7322 Haemophilus parainfluenzae 3324 Haemophilus parainfluenzae/influenzae 7324 Haemophilus parainfluenzae 3360 Haemophilus parainfluenzae 7326 Haemophilus parainfluenzae 3420 Haemophilus influenzae 7340 Haemophilus parainfluenzae 3422 Haemophilus influenzae 7360 Haemophilus parainfluenzae 3424 Haemophilus influenzae 7362 Haemophilus parainfluenzae 3426 Haemophilus influenzae 7364 Haemophilus parainfluenzae 3520 Haemophilus parainfluenzae/influenzae 7420 Haemophilus influenzae/parainfluenzae 3524 Haemophilus influenzae/parainfluenzae 7424 Haemophilus influenzae 3560 Haemophilus parainfluenzae 7426 Haemophilus influenzae 3620 Haemophilus influenzae 7460 Haemophilus parainfluenzae 3622 Haemophilus influenzae 7500 Haemophilus parainfluenzae 3624 Haemophilus influenzae 7520 Haemophilus parainfluenzae 3626 Haemophilus influenzae 7522 Haemophilus parainfluenzae 3720 Haemophilus parainfluenzae/influenzae 7524 Haemophilus parainfluenzae/influenzae 3724 Haemophilus influenzae/parainfluenzae 7540 Haemophilus parainfluenzae 3760 Haemophilus parainfluenzae 7560 Haemophilus parainfluenzae 4002 Neisseria meningitidis 7562 Haemophilus parainfluenzae 4003 Neisseria meningitidis 7564 Haemophilus parainfluenzae 5001 Neisseria polysaccharea/spp 7620 Haemophilus influenzae/parainfluenzae 5002 Neisseria meningitidis 7624 Haemophilus influenzae/parainfluenzae 5003 Neisseria meningitidis 7626 Haemophilus influenzae/parainfluenzae 5041 Neisseria lactamica 7660 Haemophilus parainfluenzae 5060 Haemophilus aphrophilus/paraphrophilus 7700 Haemophilus parainfluenzae 5103 Neisseria spp. 7720 Haemophilus parainfluenzae 5120 H.parainfluenzae/aphrophilus/paraphrophilus 7722 Haemophilus parainfluenzae 5122 H.aphrophilus/paraprophilus/parainfluenzae 7724 Haemophilus parainfluenzae 5160 H.aphrophilus/paraprophilus/parainfluenzae 7726 Haemophilus parainfluenzae 5162 Haemophilus aphrophilus/paraphrophilus 7740 Haemophilus parainfluenzae 5320 Haemophilus parainfluenzae 7760 Haemophilus parainfluenzae 5324 Haemophilus parainfluenzae 7762 Haemophilus parainfluenzae 5360 Haemophilus parainfluenzae 7764 Haemophilus parainfluenzae 5420 Haemophilus influenzae/parainfluenzae

45

IDENTIFICATION OF ENTEROBACTERIACEAE

Enteric bacteria are gram-negative bacilli (the Enterobacteriaceae). They are microbes whose normal habitat is the intestinal tract of humans and other animals, birds, and reptiles. Examples of some of the more common enteric bacilli are Escherichia coli, Enterobacter, Salmonella, and Shigella. Whereas E. coli and Enterobacter are usually normal flora, Salmonella and Shigella are enteric pathogens. These various genera of enteric bacilli can be differentiated and identified by using selective and differential media and biochemical tests.

Identification of Enteric Bacteria

The API 20E system is a miniaturized version of the conventional test tube procedures for identifying enteric bacteria. The system contains 20 or more different biochemical tests. Each microcupule consists of dehydrated media that is reconstituted by adding several drops of a bacterial suspension. The strip is then incubated at 37ø C for 18-24 hours and read.

Procedure:

PREPARATION OF STRIPS

1. Using aseptic technique, inoculate a tube of sterile water with a loopfull of the organism provided by the instructor.

2. Set up an incubation tray and lid. Dispense tap water into the bottom of the tray using a squeeze bottle, to provide a humid atmosphere.

3. Remove one API strip from the sealed packet and place the strip into the incubation tray. Label the end of the strip.

4. Using a sterile pipette, fill each microtube with the bacterial suspension prepared in step #1.

5. Fill both the microtube and the cupule of the |CIT|, |VP|, and |GEL|.

6. Upon completion of all the inoculations, completely cover the cupule of the ADH, LDC, ODC, H2S, and URE with mineral oil.

7. Place the plastic lid on the tray and incubate the strip in aerobic conditions at 370C for 18-24 hours.

READING THE STRIPS

1. Record all reactions not requiring the addition of reagents. This will be all tubules except TDA, VP, and IND. Interpretation of reactions are given in the reading table provided.

2. After recording the above reactions, add one drop of 10% ferric chloride to the TDA tubule. The reaction should be immediate.

3. Next, add one drop of solution A (40% potassium hydroxide) to the VP tubule. Then add one drop of solution B (6% alpha-naphthol). This reaction may take up to 10 minutes.

4. Last, add one drop of Kovac's reagent to the IND tubule. This reaction should occur within two minutes.

46

INTERPRETATION OF RESULTS (IDENTIFICATION)

1. Using a marker, mark the strip off in groups of three tubules.

2. Within each group of three tubules, assign the following numbers:

tubule #1 = 1 tubule #2 = 2 tubule #3 = 4

3. To obtain the identification number for your organism, add up the numbers within each separate group of tubules that corresponds to a positive reaction. For example: API 20E identification number = 5146572

4. Once the identification number has been obtained, you can look it up in the API 20E Analytical Profile Index or on https://apiweb.biomerieux.com. (user name and password required)

STUDY QUESTIONS:

1. What is the identification of each organism according to the API 20E Analytical Profile Index? (Fill in the chart below)

2. Are these organisms normal enteric microbiota or enteric pathogens? Under what circumstances can they become pathogenic?

______

______

RESULTS SHEET:

Unknown # API # Identification (Genus and species)

47

API 20E SYSTEM READING TABLE (Interpretation of reactions)

TUBE POSITIVE NEGATIVE

ONPG any yellow color clear or colorless

ADH red or orange-red yellow/yellow-orange

LDC red or orange-red yellow/yellow-orange

ODC red or orange-red yellow/yellow-orange

CIT turquoise or dark blue light green or yellow

H2S blackening of the media no blackening present

URE pink or coral (red-orange) yellow/no pink or coral

TDA dark reddish-brown light red-brown or yellow

IND red yellow/no red color

VP bright pink pale pink or no pink color

GEL diffusion of the black granules no diffusion/black granules remain throughout the cupule clumped together at the bottom yellow or yellow green (any yellow color blue or blue green GLU is +) MAN yellow or yellow green (any yellow color blue or blue green INO yellowis +) or yellow green (any yellow color blue or blue green SOR yellowis +) or yellow green (any yellow color blue or blue green RHA yellowis +) oryel yellowlow or green yellow (any-gr eeyellown color blue or blue green SAC yellowis +) or (any yellow yell owgreen co l(anyor is yellow+) color blue or blue green MEL yellowis +) or yellow green (any yellow color blue or blue green AMY yellowis +) or yellow green (any yellow color blue or blue green ARA yellowis +) or yellow green (any yellow color blue or blue green is +) *NOTE: The other tests listed after ARA with a dotted line around the cupule (OX, NO2, N2, MOB, McC, OF-O, OF-F) are optional and can be used for further differentiations. You will leave these blank.

7/89 cf rev 5/97 tt

48

MICROBIAL GROWTH CURVE AND DIRECT AND INDIRECT MEASURES OF MICROBIAL GROWTH

Objectives: After completion of this laboratory, the student should be able to: 4. Compare and contrast direct and indirect methods for bacterial enumeration using plate counts and optical density, respectively. 5. Determine the generation time of bacteria 6. Define the 4 major phases of the growth curve

Materials: 15. Spectrophotometers 16. (3) TSA plates for spread plating per pair of students 17. 9ml tubes of sterile water for diluent 18. tubes of 5 ml sterile TSB broth 19. empty sterile test tubes 20. glass spreader rods per pair 21. container of alcohol per pair 22. Kim wipes 23. subsamples of E.coli from 0-24 hrs of incubation 24. sterile pipettes and pipetters per pair 25. test tube racks

Prelab procedures performed by instructor/staff

1. E.coli is added to 50ml of TSB and incubated at room temperature overnight. 2. 100ul of E.coli from the 50ml culture is transferred to a 250ml flask of TSB. 3. A 5ml aliquot from the flask is removed before incubation (T=0) 4. The remaining 245 ml is incubated for 8 hours at 37C in a shaking incubator (performed by instructor) 5. 5ml aliquots are removed at various times (T0 – T24 hours) (performed by instructor) 6. All aliquots are placed immediately in the refrigerator 7. Each pair of students is then given one E.coli sample labeled T0—T24 that contains varying concentrations of E.coli Indirect Measure of Microbial Growth Using a Spectrophotometer

2. Each student group will be provided with an E.coli sample labeled with the number of hours that it has incubated (Ex: T2 or 2 hours of incubation). 3. The original undiluted E.coli sample will be used in this procedure. 4. The wavelength of the spectrophotometer is set at 600nm (this the optimal wavelength absorbed by E.coli) 4. Place Place a ablank blank sample sampl e(this (thi ssample sampl e contains sterile TSB broth, but no E.coli), provided by the instructor into contains sterile TSB broth, but no theE.coli), spectr providedophotom etbyer the an dinstructor put on the into cov er. Blanking is used to ensure that only the bacteria are measured and notspectrophotometer the matrix (broth) and. Clea putn tonhe theout sicover.de of the test tube with a kim wipe to remove fingerprints, etc. before zerBlankingoing the isb lusedank. to ensure that only the bacteria are measured and the matrix (broth). Clean the outside of the test 5. tube M withix y oura kim E.c wipeoli sa tom removeple with a serological pipette by gently pulling it up and down. fingerprints, etc. before the blank. 6. With a Kim wipe clean the outside of the E.coli sample tube.

49

7. Place it into the spectrophotometer chamber and put on the cover.

8. Record your absorbance value and record it into the table below.

9. IfI fyour your absorbance absorbance readingreading isis overover 0.80.8 you will need to dilute it: ( to ensure that you have an accurate readiyoung) will need to dilute it: ( to ensure that you have an accurate reading) To dilute your E.coli sample take out 2ml of your original E.coli sample using a sterile pipette and place it into an empty test tube. a. Add 2 ml of sterile TSB broth to the 2ml of E.coli. You have just diluted the E.coli sample by ½. b. Take a reading of your diluted E.coli. c. You must multiply this value by 2 to get the absorbance reading for the original undiluted sample. Record this value in the table. 10. If Iyourf your absorbance absorbanc ereading reading is i stills st ioverll ov er 0.8, you will need to dilute it again, using 2 ml of the diluted 0.8, you will need to dilute it again, samusingple fr 2om ml st ofep the #9 dilutedand an otsampleher 2 mfroml of sterile TSB. Multiply this absorbance value by 4 and record in the tablstepe. #9 and another 2 ml of sterile TSB. Multiply this absorbance value by 4 and Direct Mrecordeasur ine theof M table.icrob ial Growth Using Serial Dilutions and Spread Plating

1. (See table below) Perform a serial dilution as follows: a. Mix the E.coli broth given to you by your instructor by gently pipetting the contents up and down using a sterile serological pipette. b. Using that same pipette, remove 1ml of the bacteria sample and place it into tube #1. Gently mix tube contents by pipetting up and down. c. Using another pipette, remove 1 ml of sample from tube #1 and place it into tube #2. Gently mix contents by pipetting up and down. d. Continue these steps through tube #8.

3. After performing your serial dilution, for groups with samples T0-T2, remove 0.1 ml of the solution from tube #2, 3, and 4 and place onto TSA plates labeled 103, 104, and 105.

5. If your group was given samples T3 or above, remove 0.1 ml of the solution from tubes #6, 7, and 8 and place onto TSA plates labeled 107, 108, and 109.

6. Dip a glass spreader rod into alcohol and quickly pass it through a Bunsen burner flame; allow to cool for 10 seconds.

7. Use this glass rod to gently spread the inoculum over the entire surface of the plate. Repeat this process for each of your plates.

50

1 ml 1 ml -6 -7 1 ml -8 1 ml -2 1 ml -3 1 ml -4 -5 1 ml 10 1 ml 10-1 10 10 10 10 10 10

#8 # 1 #2 #3 #4 #5 #6 #7

H20 H20 2 2 H20 H20 H20 H 0 H20 H 0 E.coli 9 ml 9ml 9 ml 9ml 9 ml 9 ml 9 ml 9 ml sample

0.1 ml 0.1 ml 0.1 ml 0.1 ml 0.1 ml 0.1 ml

Petri Dish 1 Petri dish 2 Petri dish 3 Petri dish 4 Petri dish 5 Petri dish 6

Dilution 10-2 10-3 10-4 10-6 10-7 10-8

Dilution Factor 103 104 105 107 108 109

Spread Plate Follow-up Lab 5. Count all colonies on the plates using a colony counter. Count only the plates that have between 30-300 colonies (to be statistically significant). Plates that have more than 300 colonies cannot be counted and are referred as too numerous to count (TNTC) . Plates that have fewer than 30 colonies cannot be counted and are referred as too few to count (TFTC).

6. If the plate you have chosen to count has closer to 30 colonies, you may count the entire plate. However, if the plate you chose to count has closer to 300 colonies, you may want to use the following method to estimate the total number of colonies:

b. choose ten square centimeters at random and count all of the colonies in each square

c. total the ten squares and divide by 10 to get the average # of colonies/cm2

d. multiply this number by the area of the Petri dish = πr2 (π = 3.14, r = 5 cm)

This will give an estimate of the total number of colonies on the entire plate.

7. Calculate the number of bacteria per ml of original sample by using the following formula:

c) Number of cells/ml= number of colonies on entire plate X dilution factor (reciprocal of dilution)

8. Once all groups have their counts and absorbance values, place them in the table below.

51

Incubation time Absorbance Dilution factor # of colonies CFU/ml T0 T1 T2 T3 T4 T5 T6 T7 T8 T9 T24

9. Next you will determine the generation time and graph the results following the instructions given.

How to Calculate Generation (Doubling) Time

To calculate the number of generations a culture has undergone, cell numbers must be converted to logarithms. Standard logarithms are based on 10. The log of 2, which is 0.301, is always part of the equation used because one cell divides into two when they undergo binary fission. (Refer to Appendix B in the Tortora Microbiology textbook.)

loglog number number of cells at the end ( minuslog num ) ber of log of numbercells of cells( m atinus the beginning) ce lls at the (over) at the 0.301end beginning number of generations = 0.301

For example: Given the initial concentration of cells (C0) at t=0 is 1,000 cells and the concentration of cells after 7 hours (C) is 30,000 cells:

STEP 1. Determine the log10 of 30,000 by using Excel, log tables, or the log key on a scientific calculator. log10 of 30,000 = 4.477 STEP 2. Determine the log10 of 1,000 = 3 STEP 3. Plugging these values into the above equation, 4.477 – 3/0.301 = 4.9 generations per 7 hours

Next, determine the number of minutes it takes for a generation to be created. To calculate the generation time: 60min/hr60min/ xhr #of x #hrsof h(over)rs number of minutes/generation = generationsnumber of g enerations

For example: 60min/hr60min/hr x x 7 7 hrs hr s(over). 4.9 generations 4.9 generation s = 86 minutes/generation

In other words, every 86 minutes, the number of cells would double.

52

How to Graph CFU’s and Absorbance Results Using Excel

Set up a table in excel with your class results, following the example below. This is an example only!! Time E.coli Absorbance E.coli Plate Counts

T1 0.001 5.60 T2 0.05 5.13 T3 0.5 6.03 T4 0.98 6.62 T5 2 9.88 T6 3.2 9.89 T7 3.2 8.81 T8 3.2 8.50

1. Choose the Insert tab at the top of the Excel menu

2. Click on Charts

3. Choose line graph (ex: “line with markers graph”)

4. Right click on the chart (above)

5. Choose Select Data

6. Now highlight the E.coli Plate Counts and E.coli Absorbance column only

7. Create a title for the chart (ex: E.coli Growth Curves by Absorbance and CFUs) and label the horizontal axis (ex: Incubation time in hours) using text boxes (choose Insert tab, Text box)

8. To add a secondary vertical axis, click on the graph line in the chart and choose “format data series” from the drop-down menu

a. Click on add 2nd axis

9. Place a text box on the chart labeled “Generation time”.

10. Place a text box on the chart indicating which line shows plate counts and which line color shows absorbance.

11. Label the growth phases (Lag, Log, Stationary, and Death Phase) for the plate counts using text boxes.

12. Your chart format should look something like the example below

G e n Stationary e r a t i o n 53 t i m E.coli Growth Curve Absorbance and CFU's [Type a quote from the document or the summary of an 12.00 3.5 interesting point. You can position[Type a quotethe text from box the anywhere document inor the summarydocument. of Usean the Drawinginteresting Tools point. tab to You change can the 3 formattingposition the of textthe pullbox anywherequote 10.00 textin the box.] document. Use the Death Drawing Tools tab to change the formatting of the pull quote text box.] 2.5 Log 8.00

2

l CFU/m

Log 10 10 Log 6.00

ty si n e D al c i Lag Op 1.5 E.coli Plate Counts

4.00 E.coli Absorbance 1

2.00 0.5

0.00 0 1 2 3 4 5 6 7 8 E.coli Incubation Time Over 8 Hours

54

Study Questions 1. When we speak of microbial “growth” how is it different than referring to the growth of other organisms like animals? ______

______

2. Briefly describe each major phase of microbial growth Lag Phase: ______

______

Log Phase: ______

______

Stationary Phase: ______

______

Death Phase: ______

______

3. Compare and contrast direct plate counting with indirect counts using a spectrophotometer. For example, which gives a more accurate reading? Which is easier to perform? Can you describe any other similarities or differences?

______

______

______

______

______

4. Why is it useful to plot bacterial growth on a logarithmic graph versus a linear graph?

______

______

55

VIRAL PLAQUE ASSAY

Objectives: After completion of this part of the laboratory exercise, you should be able to:

1. Describe the effect of bacteriophages on bacteria 2. Be able to perform a viral plaque count to determine the number of bacteriophages in a sample.

Materials: 1. Sample culture containing competent E. coli host 2. Bacteriophage 3. 9 test tubes containing 9 ml tryptone broth 8 4. 5 test tubes containing 2 ml tryptone soft agar 5. 5 petri plates containing tryptone hard agar 6. 1 ml sterile serological pipettes H20 9 ml Procedure: 1. Label all tryptone broth tubes 10-1 through 10-9 2. Label all tryptone soft agar tubes 10-4 through 10-8 3. Label all tryptone hard agar plates 10-4 through 10-8 4. Using a sterile, 1 ml serological pipette, aseptically transfer 1 ml of bacteriophage culture into tube 10-1. Continue performing a ten-fold serial dilution. 5. Aseptically add two drops of the E. coli culture and 1 ml of 10-4 tryptone broth phage dilution into the 10-4 tryptone soft agar. 6. Mix the tube quickly by gently flicking it or rolling it between your palms. 7. Pour the contents into the 10-4 hard tryptone agar plate. 8. Swirl the plate and let it stand at room temperature till the media solidifies 9. Repeat steps 5-8 for the 10-5 to 10-8 tryptone broth phage dilutions. 10. Invert and incubate the plates at 370C for 24 hours. 11. Count all plaques on plates using a colony counter. Count the plates that have 30-300 plaques (to be statistically significant). Plates that have more than 300 plaques cannot be counted and are referred as too numerous to count (TNTC). Plates that have less than 30 plaques cannot be counted and referred as too few to count (TFTC). 12. If the plate you have chosen to count has closer to 30 plaques, you may count the entire plate. If the plate you have chosen to count has closer to 300 plaques, you will need to use the ten square method to estimate the total number of plaques on the plate.

13. Calculate the number of plaques per ml of original sample by using the following formula.

a. Number of plaques/ml= number of plaques X dilution factor (reciprocal of dilution) OR b. Number of plaques/ml= number of plaques from 10 squares X πr2 X dilution factor 10

14. Record your bacteriophage counts/ml of sample in the table.

PFUs/ml Pla t e # Dilution Factor Number of PFU’s of sample

10-4 10-5 10-6

10-7 -8 10

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1ml 1ml 1ml 1ml 1 ml 1ml 1ml 1ml 1ml

10-1 10-2 10-3 10-4 10-5 10-6 10-7 10-8 10-9 Phage

Tryptone broth 1ml 1ml 1ml 1ml 1ml 1m

Tryptone soft-agar:

Add two drops of E.coli culture

Tryptone hard-agar

104 105 106 107 108

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PlaquePlaq ue s

Bacteria (Lawn of growth)

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QUANTITATIVE ANALYSIS OF WATER: MEMBRANE FILTER METHOD

Membrane filters capable of trapping bacteria larger than 0.45 microns are frequently used for analysis of water samples. A water sample is passed through the filter using a vacuum suctioning device. The filter is then transferred to a sterile Petri dish containing an absorbent pad saturated with a selective and differential liquid medium used for detecting coliforms, including E.coli. The plate and filter are incubated, and following incubation, the colonies are counted. Membrane filtration analysis is used in state public health labs throughout the US for quantifying bacteria associated with fecal pollution in drinking, recreational, and shellfish harvesting waters.

Coliforms, fecal coliforms, and E.coli are three types of indicator bacteria that are often used to determine bacterial water quality. Fecal coliforms are a sub-set of coliforms, and E.coli is the most well-known example of the fecal coliform group. It is important to note that these organisms are generally not pathogens, but are “indicators” of fecal pollution which may contain pathogens harmful to humans.

Coliforms refer to a group of bacteria with the following characteristics: Gram-negative, rod-shaped, facultative anaerobic, produce gas from glucose (and other sugars), and ferment lactose to acid and gas within 48 hrs at 35ºC. Coliforms have historically been used as an indicator for fecal pollution, and thus as an indicator of potential pathogens (e.g., enteric viruses). However, it is now understood that these organisms are found in other environments (e.g., soils, vegetation) and not just in the digestive tracts of animals. Consequently, coliforms may indicate that there is contamination, but not necessarily from a fecal source. EPA has standards for total coliforms in drinking water supplies, but no longer recommends total coliforms for recreational/surface waters (e.g., streams and lakes).

Fecal coliforms, such as E.coli, are a subset of the coliform group and have the ability to grow at higher temperatures (44.5 C) than coliforms. Fecal coliforms are found in the digestive tracts of warm and cold blooded animals and are considered more directly associated with fecal pollution compared to coliforms. The presence of fecal coliforms in a water body indicates fecal contamination (e.g., sewage overflow) and a potential public health risk. A 1986 EPA study “Ambient Water Quality Criteria for Bacteria” found that E.coli was more closely associated with illness in freshwater than fecal coliforms. Subsequently, some states including Kentucky, have adopted E.coli as their bacteria indicator of choice for determining health risk in surface waters. Water quality standards may very between States, but must be at least as stringent as Federal standards.

The Federal criteria for coliforms, fecal coliforms, and E.coli are shown below.

Maximum Permissible Total Coliform, Fecal Coliform, and E.coli Counts/100 ml

Water Use Total Coliform Fecal Coliform E.coli Municipal drinking 0 0 0 water Waters used for 70 14 14 shellfishing Recreational waters N/A 200 126

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Procedure:

Label1. Labe fourl f 90our ml 90 sterile ml ster waterile w bottlesater bot tles with the following dilutions: 10-1, 10-2, 10-3, and 10-4. Label five small Petri with the following dilutions: 10-1, -1 -2 -3 -4 10-2,dishe 10-3,s as f andollow 10-4.s: un Labeldilut efived, 10 small, 10 , 10 , and 10 . Petri dishes as follows: undiluted, 10-1,2. U si10-2,ng st 10-3,erile fandorce 10-4.ps di pped in alcohol and flamed, add a sterile absorbent pad to all five dishes.

With3. Wi a tsterileh a ster pipette,ile pip etasepticallyte, aseptica addlly add 2 ml of M-endo broth to each. This is the selective and differential media 2 fmlor ofdet M-endoection o brothf coli ftoor each.ms, incl Thisud iisng E.coli. the selective and differential media for detection of coliforms, including E.coli.Using4. Usi ng10 10ml msterilel ster pipettes,ile pipett esperform, perfor am a serial dilution as follows: transfer 10 ml of the original water sample into serial dilution as follows: transfer 10 mlthe of fi rthest bot originaltle and water mix samplewell. U siintong theanot her pipette, transfer 10 ml from bottle #1 into bottle 2 and mix well. Using firstanot bottleher pi peandtte, mix tran well.sfer Using10 ml another from bot tle #2 into bottle #3 and mix well. Finally, using another pipette, transfer 10 pipette,ml from transfer bottle #3 10 i ntmlo from bottl bottlee #4. #1 into bottle 2 and mix well. Using another pipette, transfer 10 ml from bottleAssemble5. Ass #2em intobl thee tbottle hevacuum vac #3uum apparatusand appa mixr awell.tus as as shown. shown. Start with the highest water sample dilution (10-4). Using a sterile Finally,Startpipet twithe, usingpl theace anotherhighest20 ml of waterpipette, wate sampler i transfernto the f unnel and start the vacuum. When the sample has filtered, run through 10dilution ml from (10-4). bottle Using #3 into a sterile bottle #4. pipette,another place20 m l20 of mlster ofil ewater water into to rtheinse out the funnel. funnel and start the vacuum. When the Disconnectsample6. Disc ohasnnec the filtered,t tapparatus,he appa runr atthroughus and, and with another wi th st erilized forceps, carefully remove the membrane filter and place it on top of 20 ml of sterile water to rinse out the sterilizedthe pad i nforceps, the Pet rcarefullyi dish lab removeeled 10 -the4. membranefunnel. filter and place it on top of the pad in the Petri dish labeled 10-4. 7. Repeat steps #5 and 6 for the other dilutions and for the undiluted sample.

8. DoDo not not invert inver tplates! plates! Place Place in in the the incubator for 24 hours at 37C. incubator for 24 hours at 37 decrees Celsius.

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Water Quality Results

Using a colony counter, choose plates that have between 30 and 300 colonies to count. Designate plates with fewer than 30 colonies too few to count (TFTC) and plates with more than 300 colonies as too numerous to count (TNTC). Determine the numbers of organisms per ml in the original sample by multiplying the number of colonies counted on the plate by the dilution factor for that plate. Divide the number by the amount filtered and multiply it by the total volume.

Example: 50 ml water was filtered from a 100 ml sample

45 colonies counted on 10-3 plate = 45 x 103 x 100 = 9.0 x 104 CFU/100ml 50

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STUDY QUESTIONS

1. For the water sample you tested, would that water be safe to drink? ______Why or why not? to swim or water-ski in? ______Why or why not? to fish for shrimp or oysters? ______Why or why not?______

2. Why would it be acceptable to eat fish such as bass or trout but not shellfish such as shrimp or oysters from contaminated waters? ______

3. List four (4) pathogenic microbes that can be transmitted by contaminated water. ______

4. After performing a membrane filtration procedure, you count 137 colonies with a green metallic sheen from a 10-3 dilution of 20 ml of filtered water; the original sample is 100ml. What is the level of contamination in the original water sample? Show your math. ______

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QUANTITATIVE ANALYSIS OF WATER: IDEXX Quanti-Tray 2000 using Colilert

(IDEXX product information and procedures were obtained from the IDEXX website at http://www.idexx.com/view/xhtml/en_us/water/colilert.jsf)

IDEXX’s Colilert and Quanti -Tray 2000 test for total coliforms and E.coli. Colilert is a type of selective and differential media that contains substrates u pon which enzymes of both total coliforms and E.coli react. Total coliforms have the enzyme β-galactosidase and act upon the substrate ONPG to turn the media yellow. E.coli has the enzymes β-galactosidase and β-glucuronidase. As mentioned above, the fir st enzyme will turn the media yellow; the 2 nd enzyme, β-glucuronidase, cleaves the substrate MUG to form the fluorescent product, 4 -methyl- umbelliferone. Since most noncoliforms do not have these enzymes, they are unable to grow and interfere with the test results. The noncoliforms that do have these enzymes are selectively suppressed by reagents within the Colilert media.

Unlike membrane filtration, Quanti-Tray uses most probable number (MPN) to quantify total coliforms and E.coli. The MPN method of quantification is a statistical determination of bacteria numbers (usually coliforms) per 100 ml of water or 100g of food. An MPN table is used to determine the MPN value and should correlate closely with CFU counts of membrane filtration. For example, if there were 40 yellow large wells and 15 yellow small wells, using the Quanti-Tray 2000 MPN table, the MPN value would equal 112.4 total coliforms per 100 ml. This number would equal approximately 112 CFU/100 ml with membrane filtration. The MPN table is used the same way with fluorescent wells to quantify E.coli. Remember, if a sample is diluted, to follow the equation above for membrane filtration to determine the final MPN Value.

Colilert with Quanti-Tray is known for being easy, rapid, accurate, and cost effective. The Kentucky Department of Environmental Protection (KDEP) uses this method for water quality analysis to detect total coliforms and E.coli in surface waters.

IDEXX Colilert and Quanti-Tray 2000 Procedure

1. Label one 90 ml bottle as 10-3

2. Transfer 10 ml of the 10-2 dilution used during membrane filtration into a new 90 ml 10-3 dilution bottle.

3. Aseptically transfer the water from the dilution bottle into the IDEXX sampling bottle.

4. Add 1 pack of Colilert reagent to the sample bottle and mix well

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5. Open Quanti-Tray

6. Pour reagents/sample mixture from bottle into tray. Avoid tray/bottle contact

7. Tap the wells 2-3 times to release any air bubbles and allow the foam to settle

8. Place the sample-filled Quanti-Tray onto the Quanti-Tray rubber insert of the sealer with the well side (plastic side) facing down

9. Ensure that the large cutout of the rubber insert is facing away from the sealer

10. Gently slide the rubber insert with tray into the sealer until the motor grabs the rubber insert and begins to draw it into the sealer

11. In approximately 15 seconds, the tray will be sealed and partially ejected from the rear of the sealer. Remove the rubber insert and tray from the rear of the sealer.

12. If a misaligned tray is accidentally fed into the sealer, press and hold the reverse button. However, do not reverse the motor once the rubber insert has been drawn fully into the input slot!

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13. PlacePlace sealedsealed traystrays inin thethe 3737 degreesC incubat or with white side up for 24 hours Celsius incubator with white side up for 24 hours For the Follow-Up Lab

a. Remove trays from incubator b. Add up the total # large yellow wells c. Add up the total # of small yellow wells d. Use the MPN table to calculate MPN of total coliforms per 100 ml e. Place each tray under the UV light cabinet f. Using a sharpie, mark both large and small wells that exhibit fluorescence; these are E.coli g. Follow procedure #b-d to determine the MPN value of E.coli per 100 ml

STUDY QUESTIONS

1. Did the total coliform numbers from the membrane filtration method correlate with the MPN value you obtained using Quanti-Tray 2000? If not, why do you think a discrepancy was observed?______

2. What was the MPN value for E.coli in your sample? Would the results be acceptable for Drinking water?______shellfish harvesting waters?______recreational waters?______

3. Compare and contrast membrane filtration with Colilert/Quanti-Tray ______

4. What would be possible sources of fecal contamination at a neighborhood pond?______

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IDENTIFICATION OF FUNGI

Objectives: After completion of this part of the laboratory exercise, you should be able to:

1. Describe and compare colony morphology of yeasts vs. molds vs. bacteria.

2. Describe and compare the microscopic appearance of yeasts vs. molds vs. bacteria.

3. Describe and diagram the microscopic features and sporulation of Aspergillus, Penicillium, Rhizopus, Saccharomyces, Candida, Histoplasma capsulatum, Cryptococcus neoformans, Sporothrix schenckii, and Trichophyton.

Before beginning this laboratory exercise, read the sections on Fungi in your textbook. Also, refer to the table of Medically Important Fungi at the back of this lab manual.

Materials:

1. Sabouraud's agar culture Saccharomyces yeast 2. Sabouraud’s agar cultures of Rhizopus, Aspergillus, and Penicillium molds 3. culture of bacteria for comparison purposes 4. sterile water for preparation of wet mount 5. sterile transpettes 6. microscope slides 7. cover slips 8. lactophenol cotton blue stain 9. prepared demonstration slides of various molds and yeasts 10. microscope 11. lens paper and cleaner

Procedure #1: Comparisons of colony (macroscopic) characteristics of molds and yeasts and bacteria

1. Examine the culture plates of Aspergillus, Penicillium, and Rhizopus molds provided by the instructor. Describe the colonies of these molds. Observe the aerial mycelium and the vegetative mycelium.

2. Examine the culture plates of Saccharomyces and Candida (yeasts). Describe the colonies of the yeasts.

3. Examine the culture of bacterial colonies and compare them to the colonies of the molds and yeasts.

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A. Rhizopus (mold)

B. Aspergillus (mold)

C. Penicillium (mold)

D. Candida (yeast)

E. Saccharomyces (yeast)

F. Bacterial Colonies

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Procedure #2: Comparison of microscopic characteristics of molds and yeasts

1. Using the Saccharomyces (“bakers” or “brewers” yeast) cultures provided, prepare a wet mount of yeast cells stained with lactophenol cotton blue. Examine the wet mount under low power and high power magnification. Draw a few cells and describe their morphology. Look for the budding (blastoconidia) (NOTE: the descriptive terms used to describe bacterial morphology, such as streptococcus, do not apply to fungi.)

2. Examine the prepared demonstration slides of the various molds and yeasts provided by the instructor. Draw the structures that you see for each fungus and describe their morphology. Look for hyphae and sporulation. Describe the disease processes caused by each.

For the medically important mycoses, refer to the chart in the back of your lab manual entitled “Summary of Significant Characteristics of Medically Important Fungi.”

You will be required to: 1. Identify the fungus and specific structures microscopically 2. Describe the mode of infection and disease process caused by each fungus. 3. Identify the type of specimen required for identification

Saprophytic fungi:

Saccharomyces under high power and oil immersion magnification (40X, 100X)

Rhizopus (40X)

Penicillium (40X)

Cutaneous mycoses:

Trichophyton (100X)

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Systemic mycoses:

Histoplasma capsulatum (100X)

Mold form

Yeast form

Cryptococcus neoformans (100X)

Subcutaneous mycoses:

Sporothrix schenckii (100X)

Opportunistic mycoses:

Candida albicans (100X)

Aspergillus (40X)

Pneumocystis carinii cysts in lung tissue (100X)

Procedure #3: Preparation of a microculture/slide culture for studying molds

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Sometimes, to be able to view the type of sporulation produced by a mold in order to identify that mold, it is necessary to view the mold growing in its intact, undisturbed state. This is achieved by setting up a microculture or slide culture. A slide culture is a miniature version of a culture, using a small piece of agar placed on a glass microscope slide. After the slide culture has grown the mold, the slide can be placed directly onto a microscope and observed for growth. The mold can then be seen in its living, undisturbed, growing state and identified.

Materials: teasing needle forceps Sabouraud dextrose agar small test tube (“cookie cutter”) wrapped, sterilized slide culture package tube of sterile water culture plate of a mold

Procedure:

1. Unwrap the slide culture package, but DO NOT remove the lid of the Petri dish until you are ready. The contents are sterile.

2. Using sterile forceps, assemble the contents of the slide culture dish (as shown by the instructor) so that the glass microscope slide and one of the cover slips is sitting on top of the wooden sticks.

3. Briefly flame the mouth of the small test tube in the Bunsen burner, let it cool, and then use it to cut out a small circle of “Sab” agar from the agar plate.

4. Aseptically remove the agar circle and transfer it to the center of the microscope slide inside the slide culture dish. Close the lid when you are done.

5. Using the teasing needle and aseptic technique, cut a very tiny piece of mold from the mold culture provided and touch it to the top edge of the side of the agar circle. Repeat the procedure on the opposite side of the circle. Replace the lid immediately afterwards.

6. With sterile forceps, place a cover slip on top of the agar circle. Tap it into place gently.

7. Pour a small amount of sterile water into the bottom of the dish so that the filter paper is well soaked, but not swimming. This will keep the chamber moist for the mold to grow properly.

8. Incubate the slide culture dish at room temperature in your drawer until the next lab period or for one week.

9. After the incubation period, remove the microscope slide with its agar circle and cover slip. Gently wipe off any moisture that may have accumulated on the bottom of the slide. Place the entire set-up on the microscope stage inbetween the clips.

10. Using low power (10X), focus on the outer edge of the agar circle where the mold is growing. You should be able to see hyphae and spores. If you can do so without hitting the cover slip, carefully

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switch to high power (40X) to see more detail. (NOTE: everything will be in black and white, since this is not a stained preparation)

11. After viewing the slide culture in its intact state, remove it from the microscope.

12. Obtain a clean microscope slide and place 1-2 drops of lactophenol cotton blue (LCB) stain in the center. Remove the cover slip from the top of the agar circle. The underside of the cover slip will have a circular imprint of mold growth. Place this side down into the LCB stain.

13. Take the microscope slide with the agar circle left on it and hold it over the container of disinfectant. Using the forceps, remove the agar circle and drop it into the disinfectant. Remaining will be the microscope slide with a circular imprint of mold growth. Place 1-2 drops of LCB stain onto this imprint and cover it with a clean cover slip.

14. You will now have two LCB-stained preparations of the mold. Observe both slides under low power (10X) and high power (40X). Sketch the fungal structures that you see, labeling both the hyphae and type of sporulation observed using proper terminology.

Observations:

Procedure #4: Identification of yeast species using the API C AUX system

API 20C AUX is a system for precise identification of frequently encountered yeasts. The system consists of 20 cupules containing dehydrated substrates. The yeast will grow only if it is capable of utilizing that substrate as its sole carbon source.

The reactions are read by comparing them to a control cupule. Identification is obtained by looking up the resulting profile number in the API Index.

1. Set up an incubation tray. Dispense distilled water into the bottom of the tray to provide humid atmosphere. 2. Record the specimen number on the end flap. 3. Open a pouch and remove an API strip from the pouch. Place the strip in to the tray on top of the water- filled wells. 4. Open an ampule of API Suspension Medium of API 8.5 % NaCl medium. Using a sterile pipette, pick up a yeast colony either by suction or by repeated touching. Transfer aseptically to the suspension, creating a turbidity equal to a #2 McFarland standard. 5. Open an ampule of API 20 Medium. Transfer 100 µl of the suspension prepared in step #4. 6. Using a pipette, fill the cupules with the suspension from step #5. Avoid bubbles. 7. Place the lid on the tray and incubate at 300C for 48-72 hours.

Reading the Strip:

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1. After 48-72 hours, compare growth in each cupule to the “0” cupule which is used as a negative control. 2. A cupule more turbid than the control indicates a positive reaction.

Morphology Test:

1. Determine the presence or absence of hyphae or pseudohyphae using Rice Agar Tween (RAT) medium. 2. This is considered to be the 21st test of the strip. It is recorded as positive if either hyphae or pseudohyphae are present. *(NOTE: We will not be performing this test, so the results will be recorded for you on the end of the API strip. Be sure to record this as either + or – on your results form in the box at the end.)

Interpretation:

Identification is obtained with a numerical profile. To determine the profile number, the test results are divided into groups of three on the results sheet (see next page). A value of 1, 2, or 4 is assigned to each of the three test in the group. By adding the numbers corresponding to the positive results within each group, a 7 digit numerical profile is obtained.

Look up the numerical profile in the API C AUX Index or on https://apiweb.biomerieux.com. (user ID and password required)

RESULTS

Unknown yeast #1 API profile # = ______

Identification: ______

Unknown yeast #2 API profile # = ______

Identification: ______

Unknown yeast #3 API profile # = ______

Identification: ______

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Study Questions:

1. Are these yeast species normal microbiota or are they pathogenic? Under what circumstances could they become pathogenic?

______

______

2. What is the difference between hyphae and pseudohyphae? Which are usually seen with yeasts?

______

______

3. How do yeasts reproduce? ______

______

4. What type of tests are those used in the API C AUX for identifying yeast species?

______

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IDENTIFICATION OF PROTOZOA

Before beginning this laboratory exercise, read the sections on Protozoa in your textbook. Also, refer to the table of entitled “Summary of Significant Characteristics of Parasitic Protozoa” at the back of this lab manual.

Draw and describe the microscopic appearance of the following protozoans. Describe the disease process caused by each. You will be required to: 1. Identify the protozoan microscopically 2. Describe the mode of transmission and disease process caused by each protozoan. 3. Identify the type of specimen required for identification

A. Entamoeba histolytica cysts and trophozoites in feces (oil-immersion)

B. Giardia lamblia cysts and trophozoites in feces (oil-immersion)

C. Trichomonas vaginalis trophozoites from vaginal exudate (oil-immersion)

D. Trypanosoma species hemoflagellate (blood smear on oil-immersion)

E. Plasmodium species trophozoites (merozoites) (blood smear on oil-immersion)

F. Toxoplasma gondii trophozoites in tissue (oil immersion)

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ARTHROPOD VECTORS

Before beginning this laboratory exercise, read the section in your textbook on arthropod vectors. Also refer to the table on Medically Important Arthropod Vectors at the end of this lab manual.

Examine each vector under the dissecting microscope. You will be required to: 1. recognize the vector 2. name the microorganism the insect vector transmits 3. name and describe the disease process the microbe causes in humans

Anopheles (mosquito)

Xenopsylla (rat flea)

Aedes (mosquito)

Pediculus (louse)

Culex (mosquito)

Triatoma (kissing bug)

Dermacentor (tick)

Glossina (tsetse fly)

Ixodes (tick)

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IDENTIFICATION OF HELMINTHS

Before beginning this laboratory exercise, read the section on helminths in your textbook. Also, refer to the table of Medically Important Helminths at the end of this lab manual.

Draw and describe the microscopic appearance of the following helminths. You will be required to: 1. Identify the helminth microscopically 2. Describe the mode of transmission and disease process caused by each helminth. 3. Identify the type of specimen required for identification

A. Trichinella spiralis cysts in muscle tissue (10X-40X)

B. Schistosoma cercaria (10X-40X) and ova (10X-40X)

C. Strongyloides larva (10X-40X)

D. Ascaris lumbricoides ova (10X-40X)

E. Trichuris trichiura ova (10X-40X)

F. Enterobius vermicularis ova (10X-40X)

G. Necator americanus ova (10X-40X)

H. Taenia sp. ova (10X-40X) and proglottid

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HELMINTH UNKNOWNS

Using the prepared fecal specimens supplied by the instructor, make a wet mount of each, and examine under low (10X) and high (40X) power for the presence of ova or larvae. Write the scientific name of the parasite on the results sheet, and have it checked by the lab instructor before you leave.

A.

B.

C.

D.

E.

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FOOD AND WATER MICROBIOLOGY

Food and water are often contaminated with microorganisms and can be an important inanimate vector for the transmission of disease. These microbial contaminants come from a variety of sources such as: 1. soil 2. equipment and utensils used in the preparation of food or water 3. fecal contamination from animals or humans 4. food handlers Some examples of pathogenic microbes that have been found to transmit disease through contaminated food or water are Salmonella, E .coli 0157:H7, Staphylococcus aureus, Hepatitis virus, typhoid, cholera, and numerous others, including various parasitic protozoa and helminths. By demonstrating the presence and number of microorganisms in food or water, the quality of a particular substance can be evaluated. Even if the microbes are not specifically identified, a high microbe count strongly suggests the possible presence of pathogens. Even if a sample has a low count, such as drinking water, it may not be safe for consumption. The following procedures are designed to determine the quality of a sample of food or water.

THE METHYLENE BLUE REDUCTASE TEST

In a sample of milk containing large number of microbes, the microbes will actively metabolize the oxygen present in the milk. As the oxygen is used up by the microbes, the environment in the milk will become more anaerobic. This change from an aerobic to an anaerobic environment can be demonstrated by the addition of an indicator, methylene blue. In its oxidized (aerobic) form, the dye is blue. When the oxygen becomes metabolized by the microbes in the milk, the methylene blue dye will be reduced and will lose its blue color to become colorless. The rate at which this color change occurs correlates with the amount of microbial contamination in the milk. A highly contaminated sample will become reduced in less than thirty minutes. A very slightly contaminated sample of milk may require over 6-8 hours to become reduced. This determination is made according to the following guidelines:

1. very poor quality (highly contaminated) = reduction within 30 minutes 2. poor quality = reduction takes between 30 minutes and 2 hours 3. fair quality = reduction takes between 2 and 6 hours 4. good quality = reduction takes 6-8 hours or longer

Procedure:

1. You will be given two milk samples, one of which has been pasteurized, and one of which is "raw" unpasteurized milk that is highly contaminated with bacteria.

2. Using a sterile pipette, add 1 ml of methylene blue dye to each test tube.

3. Mix the milk and dye, and place in a 37 degree C waterbath. Record the starting time.

4. Observe the milk samples every 30 minutes until the milk turns white. Record this time. Based on your observations, determine and record in the chart the quality of each sample as very poor, poor, fair, or good.

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Sample A Sample B

Reduction time

Quality of milk sample

STUDY QUESTIONS:

1. Which milk sample was pasteurized? ______Which sample was "raw" milk?______

2. List four pathogens that have historically been found in raw, unpasteurized milk.

a. ______

b. ______

c. ______

d. ______

3. Explain the function of the methylene blue dye in this experiment.

4. Describe the process used to pasteurize milk and other food substances.

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MICROBIOLOGICAL ANALYSIS OF HAMBURGER: BACTERIAL COUNT

In this procedure, several dilutions will be made of ground beef in sterile water, which will then be plated onto EMB (eosin methylene blue) agar and onto BHI (brain heart infusion) agar. EMB is a selective and differential agar used for the detection of fecal coliforms including E. coli. BHI is commonly used for making pour plates for the counting of bacteria. The amount of bacteria from the hamburger on these plates will be counted, and a determination will be made as to the quality of the hamburger.

Procedure:

1. You will be provided with two hamburger samples, sample A and sample B. Weigh out 20 grams of each hamburger sample, using a plastic weighing "boat" for each sample.

2. Place one of the samples in a blender with 180 ml of sterile, distilled water. Blend for 5 minutes. You have now created a 10-1 suspension of hamburger. While you are waiting, label one EMB plate for each sample with 10-1 and three empty Petri plates for each sample with the following dilutions: 10-2, 10-3, and 10-4. (You should end up with eight Petri dishes: 2 EMB's and 6 empties)

3. Using a sterile pipette, transfer 0.l ml of this suspension onto the 10-1 labeled EMB plate. Spread the sample out onto the plate using the streak plate technique.

4. Using the same pipette, transfer 0.l ml of this suspension into the empty Petri dish labeled 10-2.

5. Using the same pipette, transfer 1 ml of the suspension into a bottle containing 99 ml of sterile, distilled water. You have now created a 10-3 dilution. Label the bottle.

6. Using another pipette, transfer 1 ml of this 10-3 dilution from the bottle into the empty Petri dish labeled 10-3.

7. Using the same pipette, transfer 0.l ml of this 10-3 dilution from the bottle into the empty Petri dish labeled 10-4.

8. Repeat steps #2-7 for the second hamburger sample.

9. Quickly pour the contents of one tube of molten BHI agar (keep in the water bath until you are ready to pour!) into each empty Petri dish. Gently swirl the agar around to mix and cover the bottom of the dish. Let the plates set until hardened.

10. Invert and incubate all plates at 37oC for 24-48 hours

11. Using a colony counter, choose plates that have between 30 and 300 colonies to count. Designate plates with fewer than 30 colonies too few to count (TFTC) and plates with more than 300 colonies as too numerous to count (TNTC).

12. Determine the numbers of organisms per ml in the original sample by multiplying the number of colonies counted on the plate by the dilution factor for that plate. Example: 50 colonies counted on 10-3 plate = 50 x 103 = 5.0 x 104 CFUs/ml

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Sample EMB BHI plate counts CFU's/ml Type of colonies observed 10-2 10-3 10-4___

Hamburger A Hamburger A______

Hamburger B Hamburger B______

Weigh 20 g of sample

Blend food with 180 ml 11 ml m l (from the blender) Sterile water

99 ml dH20

10-3 10-1

Streak for 0.1 ml 1 ml 0.10.1 ml m l(from the bottle) isolation 0.1 ml (from the blender)

Petri10- 1dish 1. 10-1 EMB Petri10-2 dish 2. 10-2 BHI Petri10-3 dish 3. 10-3 BHI Petri10-4 dish 4.10-4 BHI Plate EMB BHI BHI BHI

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STUDY QUESTIONS

1. Which hamburger sample contained E.coli? How did you know this? ______

______

2. Explain why E.coli colonies produce a green, metallic sheen on eosin methylene blue agar. ______

______

______

3. How do coliform bacteria like E.coli get into foods like hamburger? List several possible ways foods may become contaminated with enteric organisms. ______

______

______

______

4. Would the hamburger sample contaminated with E.coli be safe to eat? Why or why not? ______

______

5. In recent years, a pathogenic strain of E.coli has appeared that has been designated E.coli 0157:H7. How would it be determined whether a sample of ground meat contained E.coli 0157:H7 rather than any of the other strains of normal microbiota E.coli? Explain the methodology______

______

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Antibiotic Susceptibility Testing

Objectives: After completion of this laboratory exercise, the student will be able to:

1. Demonstrate the activity of certain antibiotics against certain microbes. 2. Show the antibiotic susceptibility patterns of microorganisms that cause human infections. 3. Explain the importance of susceptibility testing in clinical microbiology.

As antibiotics have been used to treat infections over the years, resistant strains of bacteria have developed. The development of resistance to an antibiotic involves these processes:

1. Genetic mutation: natural selection operates to promote "survival of the fittest": survival of new mutant strains that are resistant to the effects of a particular drug with the old, sensitive bacteria being killed off by the antibiotic.

2. Transfer of a plasmid (the R factor) to the bacterial cell. The plasmid contains a gene or group of genes causing resistance to an antibiotic. This transfer occurs when resistant bacteria (carrying an R factor) come in contact with sensitive bacteria (do not have a R factor).

In order to choose the proper antibiotic for therapy it is important not only to identify the causative bacterium but to test it for its susceptibility to a variety of antibiotics. The variety of antibiotics to which a given organism is susceptible or resistant is called its antibiotic susceptibility pattern. This susceptibility is based on the genetic characteristics of each individual species of microorganism.

Among the variety of tests that are available, the disk-diffusion method (Kirby-Bauer test) is probably the simplest to perform and interpret. Discs of filter paper are impregnated with antibiotic solutions in the same range of concentrations obtainable in the human body. These are placed on an agar plate that has been uniformly inoculated with the organism to be tested. The test organisms grow in a smooth "lawn" of growth on the plate except in a clear round zone around each antibiotic disc which inhibits the growth of the organism. This zone indicates the susceptibility of the organism. Bacteria resistant to an antibiotic show little or no inhibition.

You will perform antibiotic susceptibility tests on different bacterial species to a variety of antibiotics. The microorganisms used in this exercise are common (such as S. aureus) and the antibiotics used here have been selected because they are widely used. They are not necessarily the most appropriate therapeutic choice.

Materials:

1. 24-hour broth cultures of Staphylococcus aureus (gram-positive coccus) Escherichia coli (gram-negative bacillus)

2. antibiotic discs 3. forceps 4. Mueller-Hinton agar plates 5. sterile cotton swabs 6. MacFarland Standard or spectrophotometer 7. sterile pipettes 8. sterile TSB

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Procedure:

1. Using the 24-hour broth culture you were assigned, standardize the inoculum by either comparing the turbidity (cloudiness) of your culture to the MacFarland Standard provided, or use a spectrophotometer. If the culture is too turbid ,use sterile TSB and a sterile pipette to dilute it until it is the same turbidity as the standard.

2. Using a sterile swab, dip the swab into the standardized bacterial suspension. Spread the bacteria out over the surface of a Mueller-Hinton agar plate to create a solid “lawn” of bacteria.

3. Using a lab marker, divide the bottom of the Mueller-Hinton agar plate into four quadrants.

4. Choose four antibiotic discs to test. Make sure that you are using the appropriate type antibiotics for the microorganism you were assigned. For example, penicillin is used to test with S.aureus, a gram-positive bacterium. It should not be used to test with E.coli, a gram-negative bacterium.

5. Label the bottom of each quadrant of the petri dish with the abbreviation (code) of the antibiotic being tested. Also put your name, date, and class section.

6. Dip the forceps into a bottle of alcohol and then hold the forceps in the flame of your Bunsen burner until the alcohol has burned off. This will sterilize the forceps. Allow them to cool before using.

7. With the sterile forceps, remove an antibiotic disc aseptically from its container and place it gently on the surface of the agar in the center of the section labeled for that disc.

8. Tap the disc gently onto the surface of the agar so that it will not fall off when the plate is inverted in the incubator.

9. Reflame the forceps.

10. Place the other disc onto the agar in the same manner. Flame the forceps in between each use. DO NOT CONTAMINATE THE ANTIBIOTIC VIALS!

11. Invert the plate and incubate at 370C for 24 hours.

12. After incubation, examine the plate for a zone of inhibition. Using a ruler marked in millimeters (mm), measure the diameter of each zone. Be sure to make a note of any colonies growing inside the zone of inhibition. These are called “satellite colonies” and indicate the development of a resistant mutation.

Look up each zone measurement in the interpretation chart provided. Record the measurement in the appropriate box on the sample report sheet. (R= resistant, I=intermediate, S=sensitive). Zones containing satellite colonies should be recorded as “R” (resistant.)

84

86

ZONE SIZE INTERPRETATION CHART

INHIBITION ZONE DIAMETER IN MM ANTIMICROBIAL DRUG OR ANTIBIOTIC TESTED RESISTANT INTERMEDIATE SENSITIVE mm or less mm mm or more Ampicillin (AM) gram negatives 13 14-16 17 Staphylococci 28 29

Carbenicillin (CB) gram negatives 19 20-22 23 Pseudomonas 13 14-16 17

Cephalothin (CR) 14 15-17 18 Ceftiofur (XNL) Ceftazidime (CAZ) Cefmetazole (CMZ)

Clindamycin/Lincomycin 14 15-20 21 (CC/L)

Erythromycin (E) 13 14-22 23

Gentamycin (GM) 12 13-14 15

Kanamycin (K) 13 14-17 18

Neomycin (N) 12 13-16 17

Nitrofurantoin (FD or F/M) 14 15-16 17

Penicillin (P) Staphylococci only 28 29

Tetracycline (Te) 14 15-18 19

Ticarcillin (TIC) gram negatives only 14 15-19 20

Tobramycin (NN) 12 13-14 15

Trimethoprim/ Sulfamethoxazole (SXT) 10 11-15 16

87

Physical Control of Microbial Growth Ultraviolet Light

Objectives: After completing this laboratory exercise, the student will be able to:

1. observe the effects of exposure to ultraviolet radiation as a mutagenic agent. 2. describe the variables that must be controlled in order to use UV irradiation as an effective disinfecting agent

Ultraviolet light is often toxic to bacteria. The DNA in the bacterial cell is distorted by the formation of thymine dimers. The mutation that results can take several forms. A different protein may be formed, such as a change in an enzyme, which can produce a different trait. A protein or enzyme may be destroyed, so that it no longer functions and the trait is lost. The most serious result is death of the organism due to a lethal mutation. When attempting to achieve disinfection or sterilization, obviously a lethal mutation is desirable.

The advantages of using UV light for sterilization are its ease of application, and its rapid effect. Therefore, it is widely used in clinical applications such as in hospital operating rooms over instrument trays. However, UV light also has some disadvantages. UV light has very little penetrating power, so that unless the microorganisms are directly exposed to the UV light, they will not be killed. Glass, plastic, and dust can block the penetration of UV light. In addition, UV light can burn the skin and eyes, and must be carefully used around human contact areas.

UV light is also not as effective against bacterial endospores, and some species of bacteria can recover from the damage imposed by UV light. This process is called photoreactivation, and can take place if the bacteria are reexposed to light.

Sunlight, since it contains UV light, is harmful to bacteria. That is why drying clothes on a line outside in the sun is more beneficial than using a clothes dryer, and the clothes smell fresher.

Other types of radiation, such as gamma rays, have more energy and greater penetrating power. For example, gamma radiation is used in sterilizing medical supplies. However, they are more dangerous to use than UV light because normal tissue can be damaged.

Materials:

1. 24-hour TSB culture of Serratia marcescens 2. four (4) TSA plates 3. small, sterile tube of TSB 4. sterile pipettes 5. 3 x 5" cards and tape 6. safety glasses 7. ultraviolet lamp 8. sterile cotton swabs

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Procedure:

1. Transfer two (2) drops of a 24-hour broth culture of Serratia marcescens with a sterile pipette to a small tube of sterile TSB. Mix by tapping the tube gently with your fingers.

2. Spread this diluted broth culture over the entire surface of a TSA plate, using streaks back and forth across the entire plate with a cotton swab. Repeat with 3 more TSA plates.

3. Label the lids and bottoms "5 sec.", "30sec.", "1 min." and “5 min.” This indicates the exposure time to be used.

4. Put on safety glasses. If you already wear glasses, use the safety glasses designed to go over your own. These glasses have special UV protective coatings to protect your eyes.

5. Remove the lids of the Petri dishes. Tape an index card over one-half of the agar plate.

6. Have a partner time the exposure. Expose each plate for the time specified (30 sec., 1 min., 5 min. and 10 min.)

7. After exposure, remove the cards, replace the lids, invert the plates, and incubate in the dark at 25C (in your lab drawer) to prevent the possibility of photoreactivation until the next lab period.

8. During the next lab period, examine the plates and record your results.

Results: 0 = no growth 1+ = a few colonies 2+ = moderate # of colonies 3+ = heavy growth (solid)

5 seconds 30 seconds 1 minute 5 m5i Minutesnutes Lamp # covered exposed covered exposed covered exposed covered exposed

89

STUDY QUESTIONS

1. Is UV irradiation effective in controlling microbial growth, according to your results?

______

2. What length of time gave the most killing, using UV irradiation?

______

3. What factors could have affected the outcome of UV treatment? (What variables do you have to control in order for UV light to be an effective killing agent?)

______

______

______

4. What mechanism is responsible for the killing of microbes by UV irradiation?

______

______

______

5. List three (3) practical applications for the use of UV light.

______

______

______

6. Why did you incubate your plates in your drawers at room temperature?

______

______

______

______

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STUDY QUESTIONS Physical Methods of Microbial Control

1. Give an example of a medical or laboratory use of each of the following to control microbial growth:

incineration ______

______

pasteurization ______

______

autoclaving ______

______

filtration ______

______

osmotic pressure ______

______

desiccation ______

______

lyophilization______

______

7. For each of the following items, choose the best or most practical method of controlling microbes:

plastic Petri dishes, test tubes, or pipettes packaged inside a plastic wrapping

canned fruits or vegetables

inoculating loop or needle

milk

water inside of a glass container with a screw-cap

beef jerky

a used, soiled paper lab coat

bacteria to be sent through the mail

91

Chemical Control of Microbial Growth Filter Paper Disc Method for Evaluating Antiseptics

The filter paper disc method is a simple method for evaluating the effectiveness of an antiseptic. In this method, a disc of filter paper is soaked with the antiseptic and placed on a nutrient agar plate that has been streaked with a particular type of organism. The plate is then incubated for 24 hours. If the antiseptic is inhibitory, a clear zone of inhibition will surround the disc. The size of the zone is related to the effectiveness of the antiseptic, and therefore can be measured and compared to other substances. In this exercise we will measure the relative effectiveness of various antiseptics against a common inhabitant of the skin and respiratory tract, and a potential pathogen, Staphylococcus aureus.

Materials:

1. four (4) antiseptics 2. sterile filter paper discs 3. TSA plate 4. forceps 5. paper towel 6. a 24-hour broth culture of Staphylococcus aureus 7. sterile cotton swabs

Procedure:

1. With a marker, divide the bottom of the TSA plate into four quadrants and label them with the names of the antiseptics to be used.

2. Label the plate with the name of the organism tested, your initials and section number, and the date.

3. Take a sterile cotton swab and carefully insert the swab using aseptic technique into the 24-hour broth culture that you are assigned. Press the swab against the walls of the tube to remove excess liquid. Streak this swab thoroughly across the entire surface of the TSA plate, making sure that there are no uncovered areas.

4. With sterile forceps, remove one of the paper discs provided and dip it into the antiseptic solution.

5. Blot off any excess liquid on the paper towel. Then place the disc in the center of the quadrant labeled for that particular antiseptic. Tap it gently into place so that it will stick to the surface of the agar. DO NOT PRESS IT INTO THE AGAR.

6. Repeat the procedure for the other three antiseptics.

7. Invert the plate and incubate it at 37C for 24 hours.

8. After incubation, measure the zones of inhibition surrounding each disc. Use the ruler marked off in millimeters, and record the zone sizes in millimeters, not centimeters or inches. Measure the complete diameter of the zone, from one side of the circle to the other. (This will include the paper disc.)

9. Also note in your results if there are any colonies growing inside the zone of inhibition. These are called “satellite colonies” and indicate a resistant mutation has occurred.

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RESULTS:

Organism tested:______

Antiseptic Zone of inhibition (mm) Satellite colonies (yes/no)

Study Questions:

1. Which antiseptics were the most effective against this strain of Staphylococcus aureus? How do you know this?

______

______

2. Which antiseptics were the least effective against this strain of Staphylococcus aureus? How do you know this?

______

______

3. Explain “satellite” growth. What does this mean in terms of the effectiveness of the antiseptic?

______

______

4. Match the following items to the correct category of antiseptic/disinfectant to which they belong:

___ dishwashing detergent a. phenolics/bisphenols ___Triclosan b. aldehydes ___ povidone-iodine c. surfactants ___ powders/ointments for athletes’ foot d. halogens that contain zinc e. alcohols ___ Lysol ab. organic acids ___ bleach ae. heavy metals ___ shampoo bd. chlorhexidine ___ prepackaged toaster treats cd. oxidizing agent de. quaternary ammonium compound (“quat”)

5. What is a “tincture”? ______

6. What is the only category of disinfectants that can also sterilize?______

7. Define “surfactant”.______

93

IMMUNOLOGICAL TESTS FOR IDENTIFICATION OF MICROORGANISMS AND INFECTIOUS DISEASES

An often used and alternative method for identification of microbes and the diseases they cause is to identify them by their antigenic structure, or by the antibodies that are produced against them. Antigens are molecular markers that are part of the structure of the microbes themselves. When the body is exposed to these antigens, serum proteins called antibodies (“immunoglobulins”) are usually produced that will specifically react with these microbial antigens in an attempt to eliminate them. Serum or solutions containing antibodies are called antisera. These antigen-antibody reactions are very specific; that is to say, for example, that antibodies produced against S.aureus will only react with S. aureus and not with other microbial species.

LATEX AGGLUTINATION TEST FOR IDENTIFICATION OF STAPHYLOCOCCUS AUREUS

The latex agglutination procedure is used for the rapid identification of S. aureus utilizing the detection of protein A in the cell wall. The coagulase test has long been recognized as the principle aid in the identification of S. aureus. This test takes a minimum of four hours to perform, and sometimes as long as 24 hours to become positive. S. aureus can be differentiated by a rapid slide agglutination procedure using latex particles coated with antibody. When bacteria resembling S. aureus are mixed with this S.aureus antiserum, agglutination of the cells (clumping) that is visible to the naked eye will occur.

MATERIALS latex reagent (antiserum = latex particles coated with antibody) disposable reaction cards and disposable stirring sticks culture of suspected S.aureus

PROCEDURE

Step 1. Add one drop of latex antiserum to a circle on the test card.

Step 2. Using a plastic or wood stirring stick, mix at least 3-5 colonies of suspected S. aureus in the latex antiserum to achieve an even, heavy suspension. Discard the stick in disinfectant.

Step 3. Continue stirring with the stick for 30 seconds and observe for clumping. Discard the used stick and card in disinfectant.

Results: unknownunknown AA ______unkunknownnown BB ______unkunknownnown CC ______

STUDY QUESTIONS:

1. What is the purpose of the latex particles? ______

2. Define “antigen.”______

3. Define “antibody.”______

4. Define “agglutination.”______

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INDIRECT ELISA TEST FOR IDENTIFICATION OF HIV ANTIBODIES

The ELISA test ("enzyme-linked immunosorbent assay") is a screening test that is currently used to detect the presence of antibodies to HIV. The procedure is done by placing a drop of blood on a piece of clean filter paper. The sample is placed in a microtiter well that has previously been coated with HIV antigens and allowed to incubate. Any HIV antibodies present in the blood sample will then bind to the antigens on the surface of the well. The well is then rinsed to wash away any unbound antibodies.

The next step is to "visualize" the presence of antigen-antibody complexes attached to the well. This is done by adding a solution of antibodies designed to attach to human immunoglobulins. These "anti-human IgG antibodies" will attach to the HIV antibodies that are already bound to the HIV antigens on the well surface. These secondary antibodies have been tagged with an enzyme and are therefore called "conjugated antibodies". If there are any HIV antigen-antibody complexes present in the well, the conjugated antibodies will attach to them, creating a "sandwich" with the HIV antibodies in the middle (HIV antigen - HIV antibody - conjugated antibody). The well is then rinsed again; if there are no HIV antibodies in the patient's sample, the conjugated antibodies will be washed away.

The last step is to add a "substrate-chromagen" to the well. This substrate will undergo a chemical reaction when it comes in contact with its enzyme, and will change color. If the patient has HIV antibodies, the HIV antigen-HIV antibody complex will be detected when this substrate is added.

NOTE: This test kit that you will be using is a simulation. This kit contains no blood or blood products or HIV. However, as with any chemicals, care should be taken when handling any of the reagents.

PROCEDURE:

1. Obtain a plastic microtiter plate. You will use only the rows labeled with the letter of the serum samples you are to test.

2. Obtain one microtiter pipette for each serum sample. Label each pipette with the letter of the sample.

3. Place six (6) drops of serum in the first two wells of the row labeled for that sample.

4. Obtain another pipette and label it for distilled water. Skip the first well and add six (6) drops of distilled water to wells #2 thru 7 in each row. Since there is undiluted serum in well #1, this is commonly referred to as the undiluted sample or 1:1 dilution. Since there is an equal amount of water and serum in well #2, this is commonly called a 1:2 dilution.

5. Using the appropriate serum sample pipette labeled for each row, mix the sample in the second well by gently sucking the solution up and down into the pipette. Then suck the contents of well #2 into the pipette and transfer only six (6) drops to well #3. Squirt the remaining solution in the pipette back into well #2.

6. Using the same pipette, mix the contents of well #3 and then transfer six (6) drops to well #4. Return the remaining solution back into well #3.

7. Continue this serial dilution process until you reach the eighth well. The dilution of antibody in well #7 is 1:64.

8. Let the plate sit undisturbed for 10 minutes to allow any antibodies in the serum to react with the antigen in the wells.

9. Label a clean pipette “conjugate”. Add two drops of conjugate to wells #1-7. This simulates the addition of the conjugated antibody-enzyme in the actual ELISA test 95

10. Let the plate sit undisturbed for 5 minutes to allow the conjugate to adhere to any antigen- antibody complexes in the well.

11. Label a clean pipette “chromogen”. Add three drops of chromogen to each well. This simulates the addition of the substrate-chromogen in the actual ELISA test.

12. Observe the color change that occurs in each well. A light yellow or clear color is a negative test result. A reddish color is a positive test for HIV antibodies.

CLINICAL LAB REPORT

Date: ______Technologist: ______

Test: Enzyme Linked Immunosorbent Assay (ELISA) for detection of antibodies to the Human Immunodeficiency Virus (HIV)

Patient A ______(positive or negative?)

Patient B ______

Patient C ______

Patient D ______

Patient E ______

Patient F ______

Patient G ______

Patient H ______

96

STUDY QUESTIONS:

After1. A readingfter read theing biographicalthe biographi sketchescal sketche s for each of the above patients, which ones did you predict would be forposi eachtive? of Whythe above or why patients, not? Whi whichch behaviors are considered to be of the highest risk for HIV infection? onesWhi cdidh a ryoue the predict lowes wouldt risk? be______positive?______Why or why not? Which behaviors are considered to be of the highest risk for HIV______infection?______Which______are the______lowest ______risk? ______Define______2. Def “antibodyine “antibody titer”.______titer”.______What3. Wha wast w theas antibodythe antibo titerdy tofite yourr of y our patient? ______patient? ______Does______4. D oesthe tantibodyhe anti body titer tmakeiter m ak e a difference in the prognosis of your patient with HIV? ______difference in the prognosis of your patient______with ______HIV? ______Why5. Why is this is testthis calledtest ca lanled “indirect” an “indi rect” test for HIV? ______test for HIV? ______

97

IMMUNODIFFUSION

In this exercise you will study a technique of immunology called immunodiffusion. Each team should obtain a dropper and a petri dish containing agar. Become familiar with the following procedures before beginning the exercises.

Making the Wells in the Agar

Set the petri dish right side up over the pattern below. Remove the dish cover and hold the dropper vertically over one of the circles on the pattern. Squeeze the dropper bulb and gently touch the tip to the surface of the agar. While releasing the bulb, push the pipette tip down through the agar to the bottom of the dish. Lift the pipette vertically; this should leave a straight-walled well in the agar.

CAUTION: As the bulb is released, a vacuum pull is exerted on the agar. If this vacuum is not maintained while pushing the pipette in to the agar, hairline fractures can develop in the well which will interfere with the results.

Filling the Wells

Each vial of antigen or antibody has a dropper tip. Draw a small amount of solution into the dropper, avoiding air bubbles. Wipe the dropper tip on the inside edge of the vial to remove any excess solution from the tip. Insert the dropper tip to the bottom of the appropriate well and slowly eject solution until the well is filled to, but not above, the surface of the agar. Either underfilling or overfilling a well may cause poor results. Immediately return the dropper to its vial.

CAUTION: Do no exchange the droppers or use the dropper of one vial for solution in another vial.

EXERCISE 1: ANTIBODY-ANTIGEN REACTION IN AGAR

Place petri dish section over template. Make the four wells indicated and then fill them as listed below. Replace the cover and set the dish at room temperature. Observe your results 16 to 48 hours later. The results are best viewed by holding the dish (without its cover) vertically between the fact and light source and then moving the dish to the side until all glare vanishes. Diagram the results on template below.

Well Solution

1 Bovine Albumin (antigen) 2 Anti-horse Albumin (antibody) 3 Anti-bovine Albumin (antibody) 4 Anti-swine Albumin (antibody)

EXERCISE 2: IDENTIFICATION OF AN UNKNOWN

98

Set petri dish section over template and make the indicated four wells. Your instructor will prepare an extract for testing. Fill the wells as listed below. Replace the cover and set the dish at room temperature. Observe and record your results 16 to 48 hours later. Diagram the results on template below.

Well Solution

1 “Mystery Meat” Extract (antigen) 2 Anti-horse Albumin (antibody) 3 Anti-bovine Albumin (antibody) 4 Anti-swine Albumin (antibody)

QUESTIONS

1. Which serum functions as the antigen in Exercise 1? ______Exercise 2? ______

2. Which antiserum reacted with the antigen in Exercise 1? ______Exercise 2? ______

3. According to your test results from Exercise 2, what did the unknown extract contain? ______

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Colony Morphology

Shape Surface -Smooth, Shiny Circular -Rough -Wrinkled -Dry, Powdery Irregular -Mucoid

Punctiform … Elevation

Flat

Rhizoid (branched Raised like roots) Convex

Umbonate Filamentous Pulvinate (very convex) Margin Entire (smooth)

Curled

Undulate (wavy)

Lobate (lobed)

Fillamentous

100

SUMMARY OF MEDICALLY IMPORTANT ARTHROPOD VECTORS

Scientific name Type of pathogenic Scientific name of Disease of vector microbe transmitted pathogen Process ______

Anopheles mosquito protozoan Plasmodium malaria

Aedes mosquito viruses arboviruses dengue fever, yellow fever

Culex mosquito viruses arboviruses encephalitis

Dermacentor tick bacteria Rickettsia Rocky Mountain spotted fever

Ixodes tick bacteria Borrelia Lyme disease

Glossinia tsetse fly protozoan Trypanosoma African trypanosomiasis (sleeping sickness)

Triatoma kissing bug protozoan Trypanosoma Chagas’ disease

Pediculus louse (lice) bacteria Rickettsia epidemic typhus

Xenopsylla rat flea bacteria Yersinia pestis plague

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SUMMARY OF SIGNIFICANT CHARACTERISTICS OF MEDICALLY IMPORTANT FUNGI *slides available in lab

Classification Scientific or Common Type of Portal of Entry or Mode Disease or Condition in Specimen of Choice Name Sporulation of Transmission Humans for Identification Systemic mycoses: *Histoplasma dimorphic fungus respiratory inhalation histoplasmosis sputum/lung tissue capsulatum 25C=mold of spores (tuberculated macroconidia) 370C=yeast

*Cryptococcus budding yeast with respiratory inhalation of cryptococcal meningitis CSF neoformans capsule spores

Subcutaneous *Sporothrix schenckii subcutaneous implantation sporotrichosis exudate from draining mycoses: of spores lesion

Opportunistic *Pneumocystis carinii cysts respiratory opportunist pneumonia sputum, lung tissue Mycoses: *Aspergillus conidiospores respiratory, brain aspergillosis sputum, tissue

*Candida albicans budding yeast normal microbiota vaginitis, thrush throat or vaginal swab (blastoconidia)

Cutaneous *Trichophyton microconidia and cutaneous contact or tinea pedis skin or nail Mycoses: macroconidia contaminated fomites

102

SUMMARY OF SIGNIFICANT CHARACTERISTICS OF PARASITIC PROTOZOANS/ALGAE *slides available in lab

CLASSIFICATION PARASITIC PORTAL OF ENTRY DISEASE OR SPECIMEN OF by means of REPRESENTATIVE OR MODE OF CONDITION IN CHOICE FOR locomotion ENTRY HUMANS IDENTIFICATION

Amoebas *Entamoeba histolytica ingestion of cysts amoebic dysentery fresh stool (pseudopods)

Flagellates *Trichomonas sexual contact vulvovaginitis vaginal or urethral vaginalis discharge

*Giardia lamblia ingestion of cysts enteritis and diarrhea fresh stool “backpacker’s disease”

Hemoflagellate *Trypanosoma species bite of insect vector African sleeping blood smear (tsetse fly or kissing sickness/S. American bug) Chagas’ disease

Nonmotile obligate *Plasmodium species bite of insect vector malaria blood smear Intracellular parasite (Anopheles mosquito)

*Toxoplasma gondii ingestion or inhalation toxoplasmosis tissue culture of oocysts (cat feces) serologic tests

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A SUMMARY OF THE PARASITIC HELMINTHS

PARASITE DISEASE INFECTIVE OR DIAGNOSTIC INFECTIVE STAGE/MODE OF STAGE TRANSMISSION PLATYHELMINTHS (flatworms)

TREMATODA (flukes) schistosomiasis: liver damage, ova in feces; elongated, with a free-swimming cercaria in fecally Schistosoma mansoni dysentery single, lateral spine contaminated water penetrate skin cercaria (larvae) with forked tail

CESTODA (tapeworms) Taenia species intestinal involvement ova or proglottids in feces ingestion of cysticerus or ova in undercooked beef, pork, or fish

(see next page for Nematoda:roundworms)

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NEMATODA (roundworms):

Ascaris lumbricoides (roundworm) intestinal or lung involvement ova in feces; (oval with thick, ingestion of ova; often in fecally course, bumpy outer shell) contaminated water or food

Trichuris trichiura (whipworm) intestinal ova in feces; (lemon-shaped with same as Ascaris bipolar knobs)

Enterobius vermicularis (pinworm) intestinal ova from perianal region by Scotch ingestion or inhalation of ova tape method (asymmetrical oval shape with well-formed larva)

Necator americanus (hookworm) intestinal ova in feces; (rounded with single, larvae in fecally contaminated soil thin, transparent shell; larvae not burrow through skin of bare feet usually seen in feces) OR ingestion of ova

Strongyloides stercoralis similar to hookworm microscopic larva in feces; ova not larvae in fecally contaminated soil found in feces burrow through skin

Trichinella spiralis trichinosis muscle biopsy for encysted larvae; ingestion of larvae in undercooked muscle involvement serologic tests pork or other meat

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