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LET’S GET SMALL ADVENTURES IN MICROBIOLOGY! BIO 220 Laboratory Manual January 2013 BY STAN KIKKERT, PHD

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Page 1: Let's Get Small

LET’S GET SMALL

ADVENTURES IN MICROBIOLOGY!

BIO 220 Laboratory Manual

January 2013

BY

STAN KIKKERT, PHD

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Table of Contents

LAB TOPIC PAGE

1 Safety 3

2 Use of Micropipettes 6

3 Aseptic Technique 12

4 Streak For Isolation 13

5 Bacterial Sampling 15

6 Food Microbiology 17

7 Oxygen Tolerance and Bacterial Growth 19

8 Bacterial Growth Curve 22

9 The Microscope 25

10 Simple Stain 30

11 Gram Stain 33

12 Identification of Spore Forming Bacteria 36

13 Spore Stain 38

14 Acid fast Stain 40

15 Negative Stain 42

16 Viral Plaque Assay 44

17 Characterization of a Gram Positive Throat Isolate

47

18 Characterization of a Gram Positive Skin Isolate 50

19 IMViC Test 53

20 Nitrate Reduction 59

21 Urea Hydrolysis 62

22 Motility Test 65

23 Hemolysis on Blood Agar 68

24 Mannitol Salt Agar 69

25 Gelatin 71

26 Oxidase Test 74

27 Catalase Test 76

28 KIA Test 77

29 Transformation 80

30 Mini-prep 85

31 Restriction Endonucleases 87

32 Transposon Mutagenesis 89

33 PCR 93

34 Antibiotic Sensitivity (MIC test) 97

35 MacConkey Agar 99

36 Blood Cells 100

37 Blood Typing 102

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1. SAFETY

Background: Laboratories, even in academic settings can be dangerous environments. Laboratories may contain chemicals, high-voltage electronic equipment, open flames, glassware and live microorganisms. In order to ensure a safe and welcoming environment in the laboratory, a set of rules are followed at all times. Lab rules may vary from lab to lab, but all serve to maintain the safety and efficiency of the laboratory. It is the individual’s responsibility to learn the safety rules whenever joining a new laboratory regardless whether the lab is in an academic or industrial setting. Lab Rules: 1. All Food and Drink must remain in sealed back-packs and may never be brought out. No Food or Drink is allowed to be seen in the laboratory. 2. Closed toe shoes, lab coat and goggles are required for every laboratory. 3. Always wear vinyl gloves when working with live microorganisms. 4. No running in the laboratory. 5. Persons not on class roster are forbidden to attend laboratory. 6. Personal, non-laboratory items such as mobile phones, lipstick, newspapers, etc. must remain in a sealed back-pack at all times during laboratory. 7. Wash hands before and after every laboratory. 8. Spray table with disinfectant before and after every laboratory. 9. Bio-contaminated materials will always be disposed of properly. Items that are plastic- agar plates, microcentrifuge tubes, micropipette tips, etc. and meant to be thrown away should be placed in the Autoclave Bags located at each table. When full, the bags are sealed, autoclaved and thrown away. Sharp objects such as broken glass and razor blades are never placed in an autoclave bag. These items have their own proper receptacles.

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10. Tape is removed by student on glassware- broths, agar slants, beakers etc. and placed in the appropriate racks located in the back of the room. The racks are autoclaved and the glassware is cleaned and reused. 11. Broken glassware is to be deposited in the container designated for this purpose. Broken glassware should never be deposited into autoclave bags, ordinary trash cans or the rack where glassware is returned to the stockroom. 12. Sharp objects such as razors are deposited in a container specific for that purpose. 13. All accidents, injury or non-injury are to be reported to the instructor immediately. 14. Never handle microorganisms isolated from another person’s body. You are to work only with your own specimens. 15. Never handle or disturb equipment or experiments that are part of other courses using the laboratory facility. Safety Devices are located throughout the laboratory. All students should be aware of the location and use of these devices in the event of an emergency. Eye Wash – located near door. For use when the eyes have made contact with bacterial specimens or burning chemicals. Open eyes and place face in fountain. Press operational handle to open water valve. Emergency Shower- located beside eye wash. For use when entire body is in contact with burning chemicals. Immediately disrobe, stand under shower and pull handle to release water. Fire Extinguisher- located on the floor near eye wash. For use in case of fire. Pull pin to activate, direct nozzle at flame, pull handle to spray fire retardant. Phone- located by other safety devices. All emergency numbers are posted on phone. Emergency Exit- The second door exiting the lab continues onward as an alternative escape route in case of emergency.

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STATEMENT TO BE SIGNED BY THE STUDENT

For and in consideration of being allowed to participate in the laboratory portion of BIO 220, Microbiology, the undersigned being over the age of eighteen years, does hereby acknowledge the following: I will be working with live microorganisms in Biology 220, Microbiology. These organisms will range from nonpathogenic laboratory strains to unknown organisms isolated from the skin, nose, throat and anal region of the student. The risks of participation in this laboratory include, but are not limited to contracting disease or infection and/or injury from laboratory equipment. The risk of contracting infections and/or disease, which could be serious or even fatal, is significantly increased for individuals whose immune system (body defense) is impaired for any reason. Persons with immune system deficiencies include, but are not limited to, those individuals undergoing chemotherapy, taking immunosuppressive drugs such as corticosteroids, having diabetes, having autoimmune disease (such as lupus erythematosis or multiple sclerosis), being pregnant, and/or being HIV positive. Understanding the foregoing risks, I knowingly and consensually assume all risks involved and I waive any and all claims against Maricopa Community College District, Mesa Community College, and any agents or employees thereof which might arise from any exposure. I have been advised that any student who suspects he or she has less than normal immune function should consult a physician as to the advisability of enrolling in microbiology at this time. I have been advised of the proper laboratory techniques to use at all times while working in the microbiology laboratory in order to minimize any danger to myself and others in the laboratory and agree to employ these techniques. If a procedure or directions are not clear to me, I realize it is my responsibility to ask my instructor for clarification. Done this ___________ day of ____________ (month/year), on behalf of myself, my heirs, devisees, legatees and estate. _______________________________ _____________________________ Student signature Name of student (please print) _______________________________ ____________________________ Parent Signature (if student under 18) Parent Name (please print)

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2. Use of Micropipettes

Introduction: Micropipettes are calibrated devices used to measure (and subsequently transfer) a small volume of liquid. Micropipettes are used to transfer anywhere from 0.5 μl – 1000 μl, however a single micropipette is not accurate over this range. A molecular biologist will generally have a set of 3 micropipettes to measure 1 μl – 1000 μl range. Micropipettes work on basic principles – The device has a piston that when pushed down expels a specific volume of air out of the micropipette tip. When the piston is raised a vacuum is created that will uptake the same amount of fluid that was previously expelled as a gas. Micropipettes are sensitive instruments that are expensive to repair or purchase. Because the micropipettes are used in almost every molecular biology protocol, it is essential for the technician to treat the micropipette as the valued handheld instrument it is.

Common Micropipettes P20 Used for 1 μl - 20 μl

P200 Used for 20 μl – 200 μl P1000 Used for 100 μl – 1000 μl

Other Sizes

P2 Used for 0.5 μl – 2.0 μl P10 Used for 0.5 - 10 μl

P100 Used for 10 μl – 100 μl Because most of the reactions performed in molecular biology require measurements less than 1 ml, it is essential to be technically solid when using the micropipettes. If one does not micropipette accurately one will have a hard time performing molecular biology.

Proper micropipetting involves: Selecting the proper micropipette for the task

Adjusting the instrument to the desired volume Pipetting the fluid accurately

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Selecting the proper micropipette Each micropipette has a sticker on the end of the plunger. It will have a p____ followed by a number. The stickers are color coated. Different sized micropipettes use different sized types.

Size Range Sticker Color Tip Color

P2 0.5 -2.0 μl Silver Clear

P10 0.5 – 10.0 μl Red Clear

P20 1.0 – 20.0 μl Yellow Yellow

P200 20 -200 μl Yellow Yellow

P100 100 – 1000 μl Blue Blue

Setting the volume: The volume of the Gilson Pipeteman is adjusted by rotating the rubber knob in the center of the handle. As the knob is rotated, the numbers displayed in the window (on the handle of the micropipette) will move as well. Rotate the knob to the clockwise to decrease the number, counter clockwise to increase. Never extend the range below zero or greater than the maximum value. Doing so will damage the internal seals of the instrument. Reading the display: The numeric display is read differently, depending on the size of the micropipette. Use the tables below to identify features of various micropipettes.

P20 (set at 15.00) Range = 1.0 -20.0 μl

1 (represent 10’s) Black number

5 (represent 1’s) Black number

0 (represent 1/10’s) Red number

P200 (set at 150) Range = 20 -200 μl

1 (represent 100’s) Black number

5 (represent 10’s) Black number

0 (represent 1’s) Black number

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P1000 (Set at 800) Range = 100 -1000 μl

0 (represent 1000) Red number

8 (represent 100’s) Black number

0 (represent 10’s) Black number

How to use the micropipette:

1. Select the appropriate micropipette for the task. 2. Adjust the volume to the desired level. 3. Attach tip to micropipette the following way: gently but firmly insert the tip of

the micropipette into the wide end of the disposable tip. Disposable tips are in racks containing 62 tips/rack.

4. Lift up on the micropipette. The tip should be attached. 5. Using your thumb, press down on the piston until you feel resistance. This point

is called the first stop. By pressing to the first stop you have expelled air through the tip equal to the amount of solution you wish to measure.

6. Insert the disposable tip into the solution to be micropipetted. Never touch the actual micropipette to any fluids!

7. Slowly lift your thumb up until the piston is at its original position. Fluid will rise into the tip as the piston rises. It is important not to lift your thumb too quickly. If turbulence is created the measurement will not be accurate. You may also contaminate the actual micropipette with vapors.

8. Lift the entire micropipette above the solution. 9. Examine the tip of the micropipette for air bubbles or unexpected fluid volume. 10. Manipulate micropipette so that the tip is above the receiving tube. 11. Press down on the piston – past the first stop and onto the second point of

resistance, the second stop. By doing this you will expel the measured fluid out of the tip. Additional air will also be pushed out of the tip allowing clean transfer of the solution. (Once the solution has been transferred you may lift up on your thumb, allowing the piston to return to its original position.)

12. Eject micropipette tip into a biohazard bag. This is done by pressing down on the silver handle in the base.

Tips: Always wear gloves when micropipetting. Our fingertips contain RNases and other

compounds that may affect sensitive experiments.

Never touch pipette tips without wearing gloves.

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Never use a micropipette tip more than once! Residue fluid remaining behind will reduce accuracy of subsequent measurements as well as possibly contaminating the

micropipette itself.

Always visually verify the fluid you have removed and will transfer. You will spot errors in pipetting and learn to recognize what various volumes look like when in the tip of a micropipette.

Common errors made by students:

Not creating a firm seal between the tip and the micropipette. Air will leak through and liquid will drop from the tip without depressing the piston.

Using the wrong micropipette for the task.

Pushing past the first stop and over measuring the solution.

Raising the piston too quickly during fluid removal.

Pressing the tip against the plastic microcentrifuge tube so that fluid does not pass

when the piston is raised.

Care and Maintenance: If maintained properly, micropipettes can maintain their accuracy for a long period of time. 1. Treat the instrument gently. If the micropipette is dropped or mishandled, damage to the piston can occur. 2. Do not set the measurements less than zero or greater than the maximum value. This will damage the seals. 3. Do not contaminate the micropipette via aggressive technique. 4. Always place the micropipette back in its rack when not in use. Laying the device on its side for prolonged periods will wear down the seals.

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Class Exercise: Micropipetting Objective: Master use of the p20, p200, p1000 micropipettes. Let’s create a Data Table for the experiment we will perform.

ID Pipette Size

Volume Added

Volume Added

Volume Added

Volume Added

Volume Removed

Observations

1-p # P20 1μl 3 μl 10 μl --- 14 μl

2- p # P200 30 μl 80 μl ---- ---- 110 μl

3- p # P200 30 μl 80 μl 150 μl 200 μl 460 μl (Use p1000)

4 –p # P1000 100 μl 300 μl 800 μl ------ 1200 μl Use p200 and p1000

5 –p # P1000 100 400 900 ----- Remove 700 μl twice using p1000

Materials: Microcentrifuge tubes containing 1500 μl dye. (3 tubes per student)

Tip: Always visually verify the fluid you have removed and will transfer. You will spot errors in pipetting and learn to recognize what various volumes look like when in the

tip of a micropipette. Protocol: Each student will work independently. 1. Obtain solutions from instructor. 2. Obtain a box of yellow and blue micropipette tips. 3. Using the p20, transfer the following volumes into a microcentrifuge tube: 1 μl,

3 μl, 10 μl. 4. Once the fluid has all been transferred, adjust the p20 to 14 μl and remove all

the fluid from the microcentrifuge tube.

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5. Observe your accuracy. Check if there is air in the bottom of tip or fluid remaining in the microcentrifuge tube.

6. Repeat 3 times with the p20 using numbers of your own. 7. Repeat the entire protocol with the p200 and p1000 micropipettes.

a. For the p200 use the volumes 30 μl, 80 μl, and remove 110 μl. b. Also perform with p200 using the volumes 30 μl, 80 μl, 150 μl, 200 μl and

remove 460 μl using the p1000. c. For the p1000 use the volumes 100 μl, 300 μl, 800 μl and remove 1200

μl. d. Also perform with p1000 using the volumes 100 μl, 400 μl, 900 μl, and

remove 1400 μl.

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3. ASEPTIC TECHNIQUE

Background: Central to working in a microbiology laboratory is the ability to transfer a culture from one media to another without introducing contaminants, a process commonly known as Aseptic Technique. Various devices exist for the transfer of bacteria into a new media. The Inoculating Loop will be commonly used in this course. The Inoculating Loop is a thin wire containing a loop at one end and a handle at the other. Academic laboratories often use wire loops that must be sterilized before every use. Clinical laboratories may use sterile disposable plastic inoculating loops of varying sizes. The size of the individual loop is calibrated to equal a specific volume of broth cultures. Aseptic Transfer of Bacteria: 1. Sterilize the inoculating loop using a cigarette lighter, Bunsen burner or electric powered Bacticinerator. When the inoculating loop begins to glow, it is sterile. 2. Let the inoculating loop cool for 5 seconds before applying to bacteria culture to be transferred. If the starting media is broth; swirl the broth culture to prevent settling of bacteria at the bottom of the tube. Dip the wire loop into the culture. Upon removal a thin liquid coating of broth will contain hundreds of organisms. If the original sample is bacteria growing on agar, gently touch an edge of the loop on the colony to be transferred. Such contact will result in the transfer of thousands of bacteria to the inoculating loop. 3. Apply the loop to the media to be inoculated. If the media is a broth, insert the loop below the surface of the broth and twist the inoculating loop once. If the media is agar, gently slide the loop across the surface to be inoculated. 4. Re-sterilize the inoculating loop.

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4. STREAK FOR ISOLATION Background: Under most circumstances a researcher prefers or needs to work with bacterial cultures that are pure (not mixed.) A pure culture can be generated from a single bacterial colony isolated on agar. Bacteria, when properly spread on agar will grow into distinct colonies each resulting from a single cell. Streaking for Isolation is a common technique used to generate single colonies of bacteria. The technique involves using an inoculating loop to physically spread the bacteria out to the point where single cells are isolated and can grow into distinct colonies. Streaking for Isolation requires a certain amount of finesse and technical skill but is easily mastered. Objective: Master the Streak for Isolation technique using 2 types of bacteria and 2 different growth media. Materials: Week 1 1 TSA slant E. coli / 4 students 1 TSB broth Staphylococcus simulans / 4 students 2 TSA petri dishes /student Week 2 Unknown mixed broth / student 1 TSA petri dish /student Protocol: Week 1- Streak E. coli and S. simulans for isolation. Week 2- Skills Test! Streak unknown mixed culture for isolation. Streak For Isolation technique:

1. Sterilize inoculating loop. 2. Obtain loopful (broth) or sample (slant) and streak one quadrant of petri dish. 3. Sterilize inoculating loop. 4. Streak inoculating loop across end-portion of original streak and while streaking

second quadrant. 5. Sterilize inoculating loop.

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6. Streak inoculating loop across end-portion of 2nd streak and while streaking third quadrant.

7. Sterilize inoculating loop. 8. Streak inoculating loop across end-portion of 3nd streak and while streaking

fourth quadrant. 9. Incubate plate for 30° C for 24 hours, then store at 4° C.

Results: Draw petri dish post-incubation. Conclusions: Is technique successful? If not, what steps may be taken to improve result?

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5. BACTERIAL SAMPLING Background: Bacteria as a group possess many diverse colony colors and colony morphologies. One can estimate the different types of bacteria and their relative numbers in a particular habitat by observing the colony color and morphology of bacteria colonies grown on agar media. Scientific Question: What are the numbers and diversity of bacteria obtained from a sampled environment? Hypothesis: When two environments are sampled, each environment will support different types and numbers of bacteria depending on the conditions of that environment. Materials: 2 TSA plates per student. 4 sterile swabs per student. Protocol: 1. Each student receives 1 TSA (tryptic soy agar) Petri dish. TSA is a nonselective, non-differential media designed to grow most types of microorganisms. On the underside of the dish draw a line to split the agar dish into halves. Label each section A or B. Each table of 4 students should use the table below to designate sampling assignments. Upon leaving the lab each student should have 1 TSA agar plate taped closed and 2 sterile swabs.

A B

Student 1 Toothbrush Roommate’s toothbrush

Student 2 Dish sponge Counter or floor sponge

Student 3 Dishwasher items (dirty) Dishwasher items (clean)

Student 4 T Bird Café! T Bird Café!

2. Upon the completion of sampling, discard the swab and return the Petri dish to the laboratory. Samples will be incubated at room temperature until growth is observed, then stored at 4° C.

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Results: Each student should draw the appearance of their incubated Petri dish in their lab notebook. Each student should create a table listing diversity and raw numbers of colonies in each environment sampled by the group. The Petri dish should be sealed in parafilm and stored at 4°C for use later in the course. Each student should perform the Simple Stain (Exercise 9) on 2 or 3 interesting looking colonies. (Although the protocol for the Simple Stain is listed elsewhere, the results should be drawn or photographed and included in this section.) Conclusions/Interpretations: Each student should use the Group Data to address the Scientific Question and Hypothesis listed above. Each student should use their own data to explore the relationship (if any) between colony morphology (seen on the Petri dish) and cell appearance (determined by the Simple Stain.)

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6. FOOD MICROBIOLOGY Background: As bacteria live on almost every surface of the human body including the skin, upper respiratory, conjunctiva, urinary tract and intestinal tract, it should not seem surprising that bacteria are also closely associated with the food we eat. Since the beginnings of civilization humans have manipulated the relationship between microbes and food to make wine, kim chee, sauerkraut and a variety of other food items. In these cases the microbes have an essential role in creating the food product and often prohibit the growth of unwanted microbial contaminants. Many of these “food microbes” are often still viable at the time of ingestion. Other food products, because of their association with a variety of sources including contaminated growing water, improper processing and human handling may have bacterial contamination that is harmful to humans. The symptoms from these may range from mild gastroenteritis to organ failure and death. Chicken can contain the pathogens Salmonella and Campylobacter. Ground Beef has been associated with multiple outbreaks of E. coli strain O157:H7. Because of its multiple contact points, prepared salad mixes often contain multiple species of microbes and have been associated with outbreaks of Listeria. Potato salad is a rich media, able to support the growth of many microbes. Staphylococcus aureus food poisoning is frequently connected to potato salad. Scientific Question: What are the numbers and diversity of bacteria obtained from a sampled food product? Hypothesis: When two food products are sampled, each food product will support different types and numbers of bacteria depending on the environment. Some food products will contain higher numbers of bacteria than other products. Materials: Week 1 1 package of chicken/class 1 package ground beef/class 1 bag of salad/class 1 container potato salad/class 1 test tube containing 1 ml saline/ student 1 sterile swab / student 1 TSA plate / student

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Week 2 Gram Stain reagents Protocol: Week 1 1. Each table will perform 4 samples- chicken, beef, salad and potato salad. 2. Using 1 swab per food item, gently swab the surface or juices of the food product. 4. Remove the lid of a Petri dish and streak the surface of a TSA plate, using the technique “Streaking for Isolation” (see Exercise 7). 5. Grow plate at 30 °C for 48 hours, then store at 4° C. Week 2 1. Perform Gram Stains on the resulting bacteria. 2. Save resulting growth for to test for resistance to antibiotics. Results: Each student should draw the results on their individual petri dish. Each student should have written descriptions of all the petri dishes. Each student should perform 1 Gram stain of a specimen obtained and draw/photograph the result. The Petri dish should be sealed in parafilm and stored at 4°C for use later in the course. Conclusions/Interpretations: Answer the scientific question for both the food products sampled and grown on agar as well as the live food samples. Of the chicken/beef/salad/potato salad products, which food products seemed the most contaminated? Would these bacteria likely be present in the final food product?

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7. OXYGEN TOLERANCE AND BACTERIAL GROWTH Background: Bacteria can be grouped based on the organism’s level of oxygen tolerance. Organisms that require oxygen are termed Obligate Aerobes. Such organisms use O2 as the terminal electron acceptor in the Electron transport Chain (Aerobic Respiration). Organisms that die in the presence of oxygen are termed Obligate Anaerobes. For these organisms O2 is a toxic, highly reactive compound. ATP is generated in the absence of O2 using Anaerobic Respiration or Fermentation pathways. An Anaerobic environment can be created in the laboratory using a Gas Pak Jar. The Gas Pak Jar utilizes a foil pouch placed in the jar along with the bacterial specimens. Adding water to the foil pouch initiates a chemical reaction that consumes oxygen. An indicator strip containing Methylene Blue is added to the Gas Pak Jar before sealing. Methylene Blue is blue in the presence of O2 but turns colorless in an anaerobic environment. Upon completion of the incubation period the indicator strip must be examined to determine an anaerobic environment has been created. Organisms that prefer O2 but can survive in anaerobic environments are termed Facultative Anaerobes. For these organisms Aerobic Respiration yields the highest ATP/glucose however Anaerobic Respiration and Fermentation pathways can also support growth of the organism. Organisms that prefer a small amount of O2, but less than the 19% atmospheric level are termed Microaerophilic. Organisms that prefer a reduced amount of O2 but a higher than atmospheric level of CO2 are termed Capnophilic. Microaerophiles and Capnophiles can be cultured in a Candle Jar. A Candle Jar is literally a glass jar. Before sealing, a small burning candle is placed inside the jar. As the candle burns it consumes O2 and produces CO2. The candle burns out when the O2 falls below 5%. CO2 levels may range from 3-10%. Instead of creating a multiple oxygen-based environments, an alternative approach to studying the effects of oxygen on bacterial growth is to use a single media containing different microenvironments with varying oxygen concentrations. Thioglycollate Broth includes a sulfur-containing compound that reduces oxygen to water. Thioglycollate broth, in essence, is an anaerobic broth. However, because oxygen will constantly diffuse into the broth from the air, a portion of the broth near the surface will contain an aerobic zone. To determine the location of the aerobic zone, the dye Reazurin is included in Thioglycollate broth. Reazurin reacts with oxygen, turning the compound red. Observe the pale red oxygen layer in the Thioglycollate tubes before beginning today’s laboratory.

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Scientific Question: What is the oxygen tolerance/preference for the 4 organisms to be grown in Thioglycollate broth? Materials: 6 Thioglycollate broth/ 4 students 4 TSA Petri dishes/4 students 1 TSA slant of Escherichia coli / 4 students 1 TSA slant of Clostridium sporogenes / 4 students 1 TSA slant of Streptococcus mitis / 4 students 1 TSA slant of Micrococcus luteus / 4 students Part I- Growth in Thioglycollate Broth Protocol: 1. Utilizing sterile technique, use an inoculating loop to transfer a bacterial sample from TSA slant into Thioglycollate broth. Insert the inoculating loop completely into the anaerobic zone to ensure the microorganism is exposed to all microenvironments. Close lid tightly. 2. Incubate Escherichia coli , Clostridium sporogenes and Micrococcus luteus at 37° C for 24 hours, then store at 4° C. 3. Incubate Streptococcus mitis at 30° C for 24 hours, then store at 4° C. 4. Incubate one uninoculated thioglycate broth at 37° C for 24 hours, then store at 4° C. 5, Incubate one uninoculated thioglycate broth at 30° C for 24 hours, then store at 4° C. Results: Draw appearance of all Thioglycollate broths post-incubation. Provide a written description of each growth pattern. Conclusions/Interpretations: Address the Scientific Question in regard to all samples grown in Thioglycollate broth.

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Part II – Growth on agar under varying oxygen levels Protocol: 1. Each student works with one TSA agar plate. 4 students work as a team creating plates A, B, C, D. 2. By marking the underside of each Petri dish, divide the agar plate into 4 sections. Label each quadrant with the organism shown below. 3. Using a sterile inoculating loop, streak each quadrant with the organism indicated. 4. Grow Plate A aerobically at 30 ° C for 24 hours. 5. Grow plate B in a Candle Jar at 30 ° C for 24 hours. 6. Grow Plate C in a Gas Pak Jar at 30 ° C for 24 hours. 7. Use Plate D as a duplicate for one of the above conditions. Results: Draw appearance of all agar plates post-incubation. Conclusions/Interpretations: Provide a written description of each organism’s oxygen tolerance.

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8. Bacterial Growth Curve

Background: Bacterial Growth is defined as the process of one cell turning into two cells. Growth is measured by observing the change in living bacteria (viable count) over a time period. Generation Time describes the amount of time required for the population to double. Generation time varies between organisms. The environment can also significantly affect the Generation Time within a species. Generation Time is measured during a period of constant growth during the exponential growth stage. There are multiple ways to measure bacterial growth. One of the more common methods is an indirect measure based on the changes in turbidity. As a bacterial culture grows, the solution becomes more turbid or cloudy. As a result, less light passes through the sample i.e. more light is absorbed. The changes in light absorption can be measured using a Spectrophotometer and then plotted on semi-log paper. The resulting curve is essentially identical (except during Death Phase) to curves generated via other methods used to measure the number of viable cells. When the Log10 Viable Cells is plotted against time on Semi-Log paper, a characteristic 4-stage curve can be seen. Each stage has a distinct name and can be characterized by expression of specific genes resulting in stage specific cellular events. Please refer to lecture notes for a more thorough discussion on this topic. 4 stages and major characteristics: Lag Phase- the # of viable cells remains constant. Log Phase- the # of viable cells increases exponentially. Stationary Phase- # of new cells = # of dying cells. Death Phase- # of dying cells > # of new cells. During Death Phase the turbidity will increase as a result of increased cell lysis. A spectrophotometer-generated growth curve is less accurate during Death Phase. Objective: Inoculate a broth with bacteria and measure the resulting growth using a spectrophotometer. Scientific Question: What are the effects of Temperature and Shaking on bacterial growth?

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Materials: 1, 100 ml culture of E. coli in 500 ml Erlenmeyer flask at Stationary Phase/class 1, 100 ml culture of Staphylococcus species in 500 ml Erlenmeyer flask at Stationary Phase/class 1 spectrophotometer/ 4 students 1 spec tube/ 4 students 2 sterile test tube/ 4 students 1, 250 ml waste beaker/ 4 students 1, 5 ml glass pipette/ 4 students 1 pipette pump/ 4 students Protocol: Work in groups of 4. Each table will create a growth curve under different environmental conditions. Class Data will be analyzed.

Condition Table #

37° C in a shaking water bath

37° C in a stationary incubator

Setting up the Spectrophotometer-

1. Turn on instrument. 2. Set wavelength to 600 nm. 3. Set instrument to read Absorption.

Measuring Absorption- Before Absorption can be measured, the spectrophotometer must be calibrated to form a baseline from which all other measurements are interpreted.

1. Add 2-3 ml of uninoculated luria broth to a clean cuvette. 2. Insert cuvette into spectrophotometer. 3. Calibrate spectrophotometer so that A600 = O% 4. Transfer contents to sterile test tube in case repeat calibration is necessary.

Clean cuvette with DI H20. 5. Once the instrument is calibrated it will stay calibrated at A600 and shouldn’t

need adjustment for the remainder of the laboratory. 6. Use same cuvette to measure A600 at the time points listed below.

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Establishing a Growth Curve: 1. Obtain 100 ml Luria Broth (LB) in 500 ml Erlenmeyer flask. 2. Remove 3 ml of uninoculated Luria Broth (LB) and transfer to cuvette for calibration of spectrophotometer. 3. Inoculate 15 ml of Stationary Phase E. coli into remaining 97 ml of LB in 500 ml Erlenmeyer flask. 4. Swirl flask and immediately measure A600. (T= 0 min.) 5. Record results in Table Form in notebook. 6. Take measurements of A600 every 30 minutes.(T=30, T=60, T=90, T= 120 and T= 150.) Results: Class data should be collected on front board.

Group 1 37 / shaken 37 / stationary

A600 T = 0 min

A600 T = 30 min

A600 T = 60 min

A600 T = 90 min

A600 T = 120 min

A600 T = 150 min

Plot results on semi-log paper. Interpretations: Identify each stage of the Growth Curve reached. Answer the Scientific Question for the 3 conditions tested. Why does temperature and shaking affect bacterial growth?

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9. THE MICROSCOPE Background: Microorganisms are too small to be seen with the unaided eye. Microscopy increases the resolving power and magnification of the unaided eye making it possible to visualize microorganisms. Various types of microscopy utilize different illumination sources to view specimens (i.e. visual light, ultra violet light, laser light, electron beams). Light Microscopy uses the wavelengths found in visible light, either using natural sunlight or the illumination from an electric light bulb. In BIO 220 we use a form of light microscopy called Brightfield. In Brightfield microscopy the background is brighter than the organism. In addition to using visible light as the illumination source our microscopes are compound. Compound microscopes use 2 sets of lenses. The total magnification provided by the microscope is the power of each individual lens multiplied together. Our microscopes have a lens in the eye piece called an Ocular Lens. The ocular lens often has a 10x magnification but stronger ocular lenses are also available. Our microscopes also have a Revolving Nosepiece containing multiple Objective Lenses (5x, 10x, 40x and 100x). When viewing a specimen using the 100x objective lens the total magnification is 1000x (100x times 10x). Thus the organism visualized will appear 1000 times larger than it actually is. A light passes through the second lens of a compound microscope a Virtual Image will be generated. The virtual image is inverted (compared to the actual object) and can only be visualized with the eye. (A Real Image, by contrast can be projected onto a wall.) Thus when it appears you are seeing the left side of the slide you are actually seeing the right end and vice versa. Our microscopes are Parfocal, meaning once the specimen is in focus using one objective lens it will be in focus using all the other objective lenses. Normally ine uses the 10x objective lens to bring a specimen into focus before switching to a higher power objective. To change objective lenses, rotate the revolving nosepiece by grabbing onto the rubber ring at the base of the revolving nosepiece. NEVER CHANGE OBJECTIVE LENSES BY GRABBING AN INDIVIDUAL LENS AND REVOLVING THE NOSEPIECE! In summary, by having 4 lenses in the revolving nosepiece a Brightfield microscope can be used for multiple applications. Below is a summary of each lens and what it is used for.

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5x- Scanning Objective. The scanning objective has a large depth of field and the widest field of view. Though not used to visualize microorganisms, the scanning objective is useful to visualize large parasites or samples of tissue. 10x Objective. Commonly the 10x objective is used to bring specimens into focus. Individual bacteria may barely be seen but clusters of cells and dye are easily visualized albeit with low resolution. The 10x objective lens has a relatively large depth of field (making it easy to establish an initial focus) and a relatively large field of view. 40x Objective “High Dry”. The 40x objective lens can be used to see large unicellular organisms such as yeast and paramecium. It can also be used to see small multi-cellular organisms such as flatworms (platyheliminthes). 100x Oil Immersion Objective. The 100x objective lens is used to visualize individual bacteria. 100x objective lens has a very small numerical aperture allowing for the highest resolution using visible light. The lens has a resolution of about 5 micrometers. Because the opening of the objective lens is so small, a majority of the light will refract beyond the lens opening as the light passes through the glass slide into the air media. To prevent excessive refraction and allow for the organism to be visualized, Immersion Oil is placed on the slide when using the 100x objective. The Working Distance or the distance between the objective lens and the specimen is quite small with the 100x objective lens. Thus the immersion oil will form a bead between the glass slide and the glass of the objective lens. Because immersion oil has the same index of refraction as glass, the illumination light does not bend outward as it passes through the specimen and instead enters the objective lens allowing the specimen to be visualized. Only a small drop of immersion oil is needed to use the 100x objective lens. Before putting the microscope away, the immersion oil should be cleaned off the 110x objective lens. This can accomplished using a Q-tip and small amounts of Windex. Removing the Microscope from the cabinet 1. Reach under the microscope cover and place one hand firmly on the handle found on the backside of the arm of the microscope. 2. Slide the microscope toward you. Once its base clears the shelf of the cabinet, place your other hand under the base of the microscope. 3. Carry the microscope upright to your lab bench. 4. Gently place the microscope on your lab bench. 5. Remove cover.

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How to view a specimen: 1. Plug in the microscope. Use the On/Off switch at the back of the microscope to send power to the instrument. Slowly adjust the rheostat to increase the light intensity. 2. Rotate the arm of the Mechanical Stage Clip outward, place slide against stage clip with the slide being flush with the Stage of the microscope. 3. Let the arm of Mechanical Stage Clip revert back into position. This will secure the slide in place. 4. Use the revolving nosepiece to point the 10x objective lens at the specimen. 5. Use the Coarse Adjustment Knob to bring the specimen into focus. Note that as you adjust the focusing knob the stage moves in relation to the objective lens. 6. Once the specimen is in focus use the Fine Adjustment Knob to maximize the clarity of the image. Never adjust the coarse and fine adjustment knobs simultaneously for they are inter-geared and can be damaged from this treatment. 7. Use the revolving nosepiece to rotate the 40x objective lens downward. As the microscope is parfocal only a slight adjustment of the fine adjustment knob will be required. If you lose focus of the specimen, change back to the 10x objective lens and begin the process again. Never use the coarse adjustment knob with the 40x or 100x objective lens pointed down! The depth of field is so narrow for these lenses you will move through the focus field without seeing the specimen. It is also possible to drive the objective lens into the slide, breaking the slide and possible damaging the microscope. 8. Once the specimen is in focus with the 40x objective lens, rotate downward the 100x objective (using the revolving nosepiece.) Before the 100x objective lens is in contact with the slide add 1 drop of immersion oil to the slide. If the microscope has been properly focused, the 100x objective lens should sit squarely on the immersion oil. 9. View the specimen under 1000x. Draw 4 or 5 of the specimens as accurately as possible. 10. To view another specimen, simply slide out one slide and slide the next one in. It should be unnecessary to completely re-focus the microscope.

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Additional components of the microscope that will affect the image generated: Rheostat- The rheostat adjusts the intensity of the light source. Beginning students often have the rheostat adjusted too high. This results in eye strain and doesn’t necessarily enhance the image of the specimen being viewed. Base Diaphragm- The base diaphragm is located just above the light bulb and regulates the amount of light passing through the lens. Iris Diaphragm- The iris diaphragm is located just below the condenser and regulates the amount of light passing through the lens. Condenser- The condenser concentrates the light beam and directs it through the specimen. Make sure the condenser is rotated into its full upright position (over the light source) when viewing specimens. Use the Condenser Adjustment Knob to move the condenser. Condenser Position Adjustment Knob- The condenser position adjustment knob allows the condenser to be moved upward toward the glass slide. Normally the specimen is viewed with the condenser as close to the slide as possible. However by changing the condenser position one may affect the contrast of the specimen visualized. This can be helpful when viewing a differential stain such as the gram Stain. Base- The microscope sits on its base. Make sure there is nothing under the base when placing the microscope on the lab bench. Arm- When one carries the microscope, one hand is placed on the arm (the other under the base.) Mechanical Stage Knob- The mechanical stage knob is used to visualize other areas of the slide not in the field of view. One knob moves the slide left and right. The other knob moves the slide forward and backward.

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Putting Away the Microscope 1. Clean the stage and all objective lenses with a Q-tip and Windex. Never clean an objective lens with anything other than a Q-tip. Paper towels and other cloth are abrasive and will scratch the lens! 2. Lower the stage all the way down. 3. Using the revolving nosepiece, rotate the 5x or 10x objective lens downward. 4. Unplug the microscope. Wrap the cord around itself, then place the cord and plug into the storage area found on the backside of the arm of the microscope. 5. Cove the microscope. Using 2 hands (one on the base and one on the arm) carry the microscope upright back to the cabinet. 6. Slide the microscope into the cabinet so that the arm is facing outwards (and the ocular facing inward.)

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10. SIMPLE STAIN Background: Bacteria as a group possess a multitude of shapes or morphologies. To simplify this method of categorization, bacterial morphology can be broadly grouped into 3 basic morphologies: Bacilli or rod shaped; Cocci- sphere shaped and Spiral shaped bacteria. In this course we will focus our studies on Bacilli and Cocci. Unless microorganisms are naturally pigmented, they can be difficult to visualize using light microscopy. As a result various staining techniques have been developed. Most staining procedures utilize Aniline Dyes that are derived from benzene. The Simple Stain is a more basic technique but provides useful information nonetheless. Organisms stained in this manner reveal cell morphology and growth patterns such as chains, pairs or clusters of cells. The Simple Stain (and many other protocols) utilizes Heat Fixing the specimen before applying dye. Heat fixing involves placing the slide + organism on a heating block. Gentle heat will evaporate the solution and kill the bacteria without lysing the cells. The cells are adhered to the glass and ready to be stained. Broth grown specimens can be directly heat fixed. Agar grown specimens are more concentrated and must be emulsified in water to separate the cells before heat fixing. The Simple Stain (and many other protocols) use positively charged dye molecules that are naturally attracted to the negatively charged cytoplasm. Excess dye is washed off with water while retained within the cell. Objectives: Master the Simple Stain procedure. Characterize microorganisms based on morphology and growth patterns. Each group of 3-4 students will prepare and examine the following slides:

Slide Organism 1 Organism 2

1 E. coli Staph. simulans

2 Bacterial Sampling Result 1 Bacterial Sampling Result 2

3 Food Microbiology Result 1 Food Microbiology Result 2

4 Food Microbiology Result 3 Food Microbiology Result 4

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Materials: 1 TSA slant E. coli/ table 1 TSB broth Staphylococcus simulans/table Agar plates with cultures generated during Bacterial Sampling Lab Agar plates with cultures generated during Food Microbiology Lab Commercially prepared Yogurt Commercially prepared Kim Chee Protocol: 1. Use sterile technique and proper safety procedures when working with live microorganisms. 2. Prepare smear. Using an inoculating loop, transfer a small amount of bacterial sample to glass slide. Broth and food samples can be directly applied to the slide. For colonies obtained on agar, transfer a small portion of the bacterial colony of interest to the slide. Add a small amount of distilled water to the slide and spread sample evenly across glass surface. Note: When preparing slides it is often beneficial to have more than one organism within the field of view. When placing 2 samples on a slide follow the example in the diagram below.

3. Dry and Heat Fix the sample using heating plate. 4. When glass slide is dry, transfer slide to rack over sink.

Overlap contains both organisms

A B

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5. Apply the dye Methylene Blue to the heat fixed sample. Use sufficient amount of dye to cover the sample. 6. Leave Methylene Blue in contact with bacterial sample for 1 minute. 7. Using forceps, tilt slide so that most of the dye runs into the sink. 8. Rinse excess dye off slide with distilled water in squirt bottle. 9. Blot slide dry using blotting paper. 10. Examine the slide under 1000x magnification. Results: Drawings of Escherichia coli, Staphylococcus simulans, sporogenes specimens. Drawings of microorganisms isolated during Food Microbiology Lab should be included in Food Microbiology section of student’s lab notebook. Drawings of microorganisms isolated from Yogurt and Kim-Chee should be included in Food Microbiology section of student’s lab notebook. Interpretations/Conclusions: Interpret the morphologies of all organisms examined and any discernable growth characteristics (growth in chains, clusters, etc.). Evaluate your staining technique. If not successful, offer possible explanations for such results.

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11. GRAM STAIN

Background: The Gram Stain, developed in 1883 by Christian Gram is perhaps the most common of all staining procedures. The Gram Stain is considered a Differential Stain, i.e. it allows one to differentiate between types of organisms or sometimes between sub-cellular structures. The Gram stain differentiates organisms based on the thickness of the cell wall. Gram Positive organisms have a thick cell wall containing lipotechoic acid while gram negative organisms contain a thin cell wall surrounded by an outer membrane. Although a few are considered Gram Variable, the majority of bacteria can be classified as Gram Positive or Gram Negative. The Gram Stain relies on a series of dyes and other chemicals applied to the specimen after heat fixing. Initially the dye Crystal Violet (primary stain) is applied. All organisms, regardless whether Gram Positive or Negative will take up the positively charged Crystal Violet in the cytoplasm much like the Simple Stain method discussed earlier. After washing off excess Crystal Violet, Iodine is added to the slide. Iodine acts as a mordant by precipitating the crystal violet. Gram Positive organisms retain the dye while Gram Negative organisms (due to their thin cell wall) can be decolorized using a slightly polar solution of acetone and alcohol. Gram Negative organisms (colorless due to the effect of acetone-alcohol) are counterstained using the dye Safranin. Gram Positive organisms do not lose the Crystal Violet stain in the presence of acetone-alcohol and are thus unaffected by the addition of the counter stain. The gram Stain is a relatively easy procedure however students should be aware that over use of the decolorizer (Acetone-Alcohol) can make Gram positive organisms appear to be gram negative. In addition, Gram Positive colonies that are over 24 hours old may decolorize regardless and thus give a false negative result. Objectives: Master the Gram Staining protocol. Understand the roles of the reagents used in the Gram Stain.

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Each group of 3-4 students will prepare the following slides:

Slide # Organism 1 Organism 2

1 E. coli Staph. simulans

2 L. fermentas N. sicca

3 N. sicca Staph. simulans

4 L. fermentas E. coli

5 Food Microbiology Result 1 Food Microbiology Result 2

6 Bacterial Sampling Result 1 Bacterial Sampling Result 2

7 Bacterial Sampling Result 1 Bacterial Sampling Result 2

8 Kim Chee Yogurt

Scientific Question: What are the gram Stain results for the organisms to be tested? What are the cell morphologies of the organisms to be visualized? Materials: 1 TSA slant Lactobacillus fermentens / table 1 TSA slant Neisseria sicca / table 1 TSB broth Escherichia coli / table 1 TSB broth Staphylococcus simulans / table Protocol: Week 1 1. Use sterile technique and proper safety procedures when working with live microorganisms. 2. Prepare smear. Using an inoculating loop, transfer a small amount of bacterial sample to glass slide. Broth and food samples can be directly applied to the slide. For colonies obtained on agar, transfer a small portion of the bacterial colony of interest to the slide. Add a small amount of distilled water to the slide and spread sample evenly across glass surface. 3. Dry and Heat Fix the sample using heating plate. 4. When glass slide is dry, transfer slide to rack over sink. 5. Apply the dye Crystal Violet to the heat fixed sample. Use sufficient amount of dye to cover the sample. 6. Leave Crystal Violet in contact with bacterial sample for 30 seconds. 7. Using forceps, tilt slide so that most of the dye runs into the sink. 8. Rinse excess dye off slide with distilled water in squirt bottle. 9. Add Iodine to the slide. Let sit for 30 seconds. 10. Rinse excess iodine off slide. 11. Apply Acetone-Alcohol to slide. Let sit 5 seconds.

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12. Rinse excess Acetone-Alcohol off slide. 13. Apply Safranin (counterstain). Let sit for 30 seconds. 14. Rinse excess Safranin off slide. 15. Blot slide dry using blotting paper. 16. Examine the slide under 1000x magnification. Week 2 1. Perform Gram Stain on unknown mixed culture. 2. Identify specimens based on Gram Stain result and bacterial morphology. Results: Drawings of Lactobacillus fermentens, Neisseria sicca, Escherichia coli and Staphylococcus Simulans. Interpretations/Conclusions: Answer the scientific question stated above. Evaluate your staining technique. If not successful, offer possible explanations for such results.

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12. Isolation of Spore Forming Bacteria Background: Endospores are inert forms of a bacterium able to withstand harsh environments and still retain viability under more favorable conditions. Certain species of Gram Positive bacteria form spores. Several Bacillus and Clostridium species are well characterized however many other species of spore formers exist. In the laboratory there are multiple ways to create an environment that kills vegetative cells without affecting any spores present. Such environments can be used to screen large bacterial populations and select for the presence of spores. The spores are identified by their ability to later germinate into vegetative cells. In this laboratory bacterial samples will be subjugated to heat and detergent to create an environment selecting against vegetative cells. In this type of experiment, a known spore former serves as a positive control and a non spore-forming bacteria serves as the negative control. Objective: To screen various natural environments for the presence of viable endospores. Scientific Question: What is the number of colony forming units (cfu) in a gram of sampled environment? Materials: 2 jars of soil obtained from various environments/class 1 TSA slant Bacillus subtilis/ table 1 TSA slant Escherichia coli/ table 4 TSA perti dishes / table 4 screw cap tubes containing 1ml 1% SDS solution 1 water bath at 90 ° C / class 4 250-ml beakers containing 50 ml ethanol /class 4 glass cell spreaders /class 4 1.5 ml screw cap tubes/ table 8 Pasteur pipettes / table 1 tube containing 5ml luria broth / table

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Protocol: Week 1: Each student should perform the protocol on 1 bacterial sample. Group data will be compiled in Results section. 1. Remove “a pinch” (or 5 loopfuls of control bacteria) and transfer to screw cap or glass test tube containing 1 ml 1% SDS solution. 2. Close lid and transfer to 90 ° C water bath. 3. Incubate sample for 10 minutes at 90 ° C. 4. Remove SDS solution using a Pasteur pipette and transfer to 1.5 ml screw cap tube. 5. Spin sample on tabletop centrifuge for 1 min. 6. Remove the majority of SDS solution using a Pasteur pipette and discard. Be careful not to disturb any cellular debris pelleted on the bottom of the tube. 7. Using a Pasteur pipette add 1 ml luria broth to screw cap tube. 8. Mix luria broth with cellular debris in tube. 9. Transfer contents to TSA petri dish. 10. Spread contents across agar using cell spreader sterilized with ethanol and flame. (See instructor demo before performing this step!) 11. Incubate petri dishes at room temperature for 1 week. Week 2: Perform Spore Stain on colonies from each agar plate to verify presence of Endospores. Results:

Sample Number of colonies

E. coli (Negative Control)

B. subtilis (Positive Control)

Soil sample A

Soil sample B

Drawings of Spore Stains of organisms obtained from soil. Interpretations/Conclusions: Evaluate the success of the technique in selecting for Endospores. Evaluate each organism/environment’s ability to support spore formation. Discuss shortcomings in this experimental design that may result in lower reported values of CFU/gram soil than actually exist in the sample. Answer the scientific question stated above.

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13. SPORE STAIN Background: The Spore Stain is a differential staining procedure used to differentiate between endospores and vegetative cells. The technique relies on prolonged contact to drive in the primary stain Malachite Green into the spores. De-ionized water rinses the primary stain off the vegetative cells but the dye is retained by the endospores. Safranin acts as a counterstain, staining vegetative cells pink. In this lab we will work with the same control organisms used in the previous lab and attempt to Identify endospores in organisms isolated from soil. Objective: Master the Spore Stain. Investigate the presence of endospores from bacteria isolated from soil. Scientific Question: Are spores present in bacterial samples isolated from soil? Materials: 1 TSA slant Bacillus subtilis/ table 7.5% Malachite Green Distilled water Safranin Petri dishes containing specimens from previous lab identifying spore forming bacteria. Protocol: Each student should perform two spore stains individually. 1. Use sterile technique and proper safety procedures when working with live microorganisms. 2. Prepare smear. Using an inoculating loop, transfer a small amount of bacterial sample to glass slide. Broth and food samples can be directly applied to the slide. For colonies obtained on agar, transfer a small portion of the bacterial colony of interest to the slide. Add a small amount of distilled water to the slide and spread sample evenly across glass surface. 3. Dry and Heat Fix the sample using heating plate. 4. Leaving slide on heating plate, add 7.5% Malachite Green to smear area.

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5. Let heating plate slowly dry off Malachite Green and drive dye into Endospores. 6. When dye appears dry, transfer slide to rack over sink. 7. Rinse off excess Malachite Green with distilled water. 8. Leaving slide on rack over sink, add the dye Safranin to sample. 9. Let sit for 5 min. 10. Rinse off excess Safranin with distilled water. 11. Blot dry and visualize under 1000 magnification. Results: Drawings of soil specimens obtained from previous lab. Drawings of control organisms. Interpretations/Conclusions: Evaluate the success of the technique in differentiating between endospores and vegetative cells. Address the scientific question stated above.

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14. ACID FAST STAIN Background: Some organisms, resistant to traditional Gram Staining procedures, have forced the development of alternative staining procedures. The Acid Fast Stain was developed specifically to visualize the genus Mycobacterium. It can also be used to visualize Nocardia species. Certain members of these genera are pathogenic to humans. Mycobacterium tuberculosis causes tuberculosis, a disease of the lower respiratory tract. The cell wall of Mycobacterium species contains mycolic acid, a waxy lipid that repels most dyes. However once the organism picks up the primary stain (Carbol Fuschin) it is very difficult to decolorize. Acid Fast organisms resist decolorizing in the presence of polar Acid-Alcohol (3% HCl, 97 % Ethanol). Non-Acid Fast organisms are easily decolorized by acid alcohol and pick up the counterstain (Methylene Blue). At times the positive control for this laboratory is heat fixed sputum samples of Mycobacterium smegmatis. The organism, being heat fixed, is no longer viable. The negative control (Staphylococcus simulans) is added directly on top of the heat fixed Mycobacterium smegmatis slide. Upon completion of the staining procedure, both Acid Fast rods and Non Acid Fast cocci are present in the field of view. Other times the control uses a nonpathogenic culture of Mycobacterium. When this control is used, the slide is prepared as Objective: To master the Acid Fast protocol and interpret results. Materials: 1 TSA slant nonpathogenic Mycobacterium 1/ student 1 TSB broth of Staphylococcus simulans / table or E.coli Carbol Fuschin De-ionized water Methylene Blue Small square of filter paper Acid Alcohol decolorizer

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Protocol: 1. Obtain a heat fixed slide of a sputum sample containing Mycobacterium smegmatis or apply a lab strain of Mycobacterium to a glass slide and proceed to step 2. 2. Add directly to the Mycobacterium smear 10 loopfuls of broth containing Staphylococcus simulans. 3. Dry/Heat Fix specimen by placing slide on heating plate. 4. When fixed, transfer slide to staining rack over sink. 5. Cover specimen with Carbol Fuschin 6. Apply filter paper to area containing bacteria + Carbol Fuschin. Add more dye until filter paper is saturated. 7. Wait 10:00. Remove filter paper using forceps. 8. Rinse slide with de-ionized water to remove excess remove excess Carbol Fuschin. 9. Place slide in contact with decolorizer Acid-Alcohol for 10 seconds. 10. Tilt slide and continue to rinse with acid-alcohol to remove remaining Carbol Fuschin. 11. Rinse slide with de-ionized water to remove excess remove decolorizer. 12. Without removing slide from staining rack, apply counterstain Methylene Blue. 13. Let sit for 2 minutes. 14. Rinse slide with de-ionized water to remove excess Methylene Blue. 15. Blot dry. 16. Visualize under 1000 magnification. Results: Draw organisms after staining. Interpretations/Conclusions: Characterize the tested organisms in terms of Acid Fastness.

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15. NEGATIVE STAIN

Background: Negative Staining procedures are unique from most other protocols in that instead of the organism being stained and the background clear, the background is stained and the organism remains clear. Negative Staining methods are most useful when the organism possesses a glycohalyx- a capsule or slime layer that accentuates the region where dye is repelled by the organism. In this laboratory we will visualize the same organism, Enterobacter aerogenes, using two different Negative Staining procedures. In the classic Negative Stain, the negatively charged dye Nigrosin is repelled by the glycohalyx and negatively charged cytoplasm. In Anthony’s Method to stain capsules, negatively charged 1% Crystal Violet stains everything but the capsule. Crystal Violet enters the cytoplasm and also remains in the background. The addition of Copper Sulfate washes Crystal Violet out of the glycohalyx leaving it unstained. Objective: Master the classic Negative Stain and Anthony’s Method for Capsules. Scientific Question: What are the similarities and differences between the images of Enterobacter aerogenes created by the two methods? Materials: 1 litmus milk broth containing Enterobacter aerogenes / table Nigrosin 1% Crystal Violet Copper Sulfate solution

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Protocol: Negative Stain- 1. Apply 10 loopfuls of Enterobacter aerogenes to far end of slide. 2. Add enough Nigrosin to cover specimen. 3. Using another slide, spread specimen across slide. (See Instructor demonstration.) 4. Cover with glass cover- slip. 5. Visualize under 1000 magnification. Anthony’s Method- 1. Apply 10 loopfuls Enterobacter aerogenes to center of slide. 2. Air Dry. (Do Not Heat Fix slide!) 3. Apply 1% Crystal Viotel to specimen. 4. Let sit 2 minutes. 5. Rinse slide with Copper Sulfate to remove excess Crystal Violet. 6. Blot gently. 7. Visualize under 1000 magnification. Results: Drawings of Enterobacter aerogenes after both staining methods. Interpretations/Conclusions: Address the scientific question stated above.

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16. VIRAL PLAQUE ASSAY Background: Bacteriophages are a group of viruses that infect prokaryotic cells. Possessing a small genome and short life cycle, bacteriophages have long served as model organisms in the study of cell and molecular biology. Bacteriophage T4 undergoes the Lytic life cycle where the bacterial cell, originally infected with a single bacteriophage, bursts to release hundreds or thousands of progeny bacteriophage. The newly releases bacteriophage will infect surrounding cells and repeat the cycle eventually resulting in a clear zone of lysis in a lawn of E. coli cells. The zone is referred to as a plaque. The single phage that initiated the formation of the plaque is called a Plaque Forming Unit (PFU). The concentration of a bacteriophage solution is measured in Plaque Forming Units (PFU)/ ml. In a pure sample the phage titer (PFU/ml) may be extremely concentrated, resulting in overlapping plaques that are difficult to measure directly. In such cases the phage sample is treated to a series of dilutions to create samples within the range of measurable concentrations. Serial Dilutions are performed in many laboratory settings when a sample is too concentrated to be accurately analyzed. Successful serial dilution technique requires accurate measurement skills and the proper mixing of samples before creating the next dilution. See Instructor demonstration of Serial Dilution. In this laboratory we will use Luria broth to make serial dilutions of a T4 phage stock solution. Once created, the phage dilutions will be mixed with an excess of susceptible E. coli to initiate a bacteriophage infection. The phage + E. coli mixture will be added to Top Agar and poured across petri plates containing Bottom Agar. See Instructor demonstration on the use of Top Agar. Objective: Master Serial Dilution technique. Infect E. coli strain B with phage T4. Determine the phage titer (PFU/ml). Scientific Question: What is the titer of the T4 phage solution?

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Materials: 3 tubes of 9.9 ml Luria Broth/table 2 tubes containing 9.0 ml Luria Broth /table 1 tube T4 phage stock solution / table 5 1.5 ml screw cap tubes / table 1 P1000 micropipette + tips / table 1 TSB broth containing E. coli strain B / table 5 petri plates containing Luria Agar / table 1 50 ° water bath/class 5 tubes of 5.0 ml Top Agar/ table Protocol: Phage Dilution- 1. Set up 5 tubes (labeled 1-5). See Table.

Sample ID (Phage Dilution)

Contains _____ ml luria broth

Receives Resulting Phage Dilution PFU/ml

Tube 1 9.9 0.1 ml Phage stock solu.

10-2

Tube 2 9.9 0.1 ml Tube 1 10-4

Tube 3 9.9 0.1 ml Tube 2 10-6

Tube 4 9.0 1.0 ml Tube 3 10-7

Tube 5 9.0 1.0 ml Tube 4 10-8

2. Perform Serial Dilution of T4 Phage solution. See above Table. Mixing Phage + E. coli- Perform the following protocol for all 5 dilutions. Label each petri plate with the phage dilution used. 1. Aliquot 100 μl E. coli strain B + 100 μl phage dilution into a screw cap tube. 2. Let contents incubate for 5 minutes at room temperature.

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3. Transfer contents to 5 ml Top Agar obtained from 50° water bath. Roll tube between hands to mix thoroughly. 4. Pour Top Agar across petri plate containing Bottom Agar. Top Agar will solidify as it cools. 5. Without inverting plate, incubate at 37° for 24 hours. Store at 4°. Results: Count plaques on each plate. Observe texture of plaques and cells at each dilution.

ID Phage Dilution

Phage Dilution PFU/ml

Number of plaques generated by 0.1 ml phage

PFU/ml

Tube 1 10-2

Tube 2 10-4

Tube 3 10-6

Tube 4 10-7

Tube 5 10-8

Interpretations/Conclusions: Address the differences in plaque appearance for each petri plate. Determine PFU/ml of T4 solution.

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17. Characterization of a Gram Positive Throat Isolate Background: Normal Flora of the Upper Respiratory tract includes organisms of the genera Staphylococcus, Streptococcus, Corynebacterium and Neisseria. In this laboratory we will perform a throat swab and select for the isolation of a Gram Positive organism by growing the isolate on CNA agar. CNA agar contains Colistin and Nalidixic Acid. Colistin disrupts the cell membrane while Nalidixic Acid inhibits supercoiling of newly synthesized DNA of Gram Negative organisms. The isolated Gram Positive organism will be characterized using various biochemical tests. Objective: Isolate and characterize a Gram Positive organism from throat region. Scientific Question: What is the identity of the organism isolated? Materials: Week 1 1 Sterile Swab / student 1 CNA petri plate / student Week 2 1 TSA slant / student 1 TSB broth / student Gram Staining Reagents Week 3 1 Blood Agar petri plate / student 1 Mannitol Salt Agar (MSA) / student Week 4 H2O2 in dropper bottles 2 Mueller-Hinton agar plates / student

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Protocol: The Experimental Flowchart listed below details the order of the characterization process. The majority of the protocols (and how to interpret the results of tests) used in this laboratory are discussed in detail in other sections of the Lab Manual. Protocols specific to this laboratory are discussed below. For all tests and protocols isolate will be grown at 30°C for 24 hours under microaerophilic conditions unless stated otherwise. Experimental Flowchart: Characterization of Gram Positive Throat Isolate

Week Task

1 Streak throat swab across CNA petri plate.

2 Pick single Colony and inoculate TSA, TSB. Perform Gram Stain.

3 Streak on Blood Agar. Streak on Mannitol Salt Agar (MSA).

4 Antibiotic Sensitivity. Catalase Test Results: Blood Agar. MSA.

5 Results: Antibiotic Sensitivity

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Results: In addition to drawings and other descriptions of the tests performed, package all data in a central table such as the one listed below.

Test Result

Colony Color and Appearance

Gram Stain

Growth on Blood Agar

Mannitol Fermentation

Salt Tolerance

Catalase Test

Kirby Bauer- Erythromycin

Kirby Bauer- Gentamycin

Kirby Bauer- Chloramphenicol

Kirby Bauer- Bacitracin

Kirby Bauer- Ampicillin

Kirby Bauer- Vancomycin

Kirby Bauer- Oxacillin

Kirby Bauer- Tetracycline

Interpretations/Conclusions: Using Bergey’s Manual of Bacteriology, is it possible to identify the unknown isolate? If the organism displayed resistance to antibiotics, discuss how this resistance may have been acquired.

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18. CHARACTERIZATION OF A GRAM POSITIVE SKIN ISOLATE Background: Normal Flora of the skin includes organisms of the genera Staphylococcus, Actinobacter, Corynebacterium and Propionibacterium. In this laboratory we will perform a skin swab and select for the isolation of a Gram Positive organism by growing the isolate on CNA agar. CNA agar contains Colistin and Nalidixic Acid. Colistin disrupts the cell membrane while Nalidixic Acid inhibits supercoiling of newly synthesized DNA in Gram Negative organisms. The isolated Gram Positive organism will be characterized using various biochemical tests. Objective: Isolate and characterize a Gram Positive organism from skin region. Scientific Question: What is the identity of the organism isolated? Materials: Week 1 1 Sterile Swab / student 1 CNA petri plate / student Week 2 1 TSA slant / student 1 TSB broth / student Gram Staining Reagents Week 3 1 Blood Agar petri plate / student 1 Mannitol Salt Agar (MSA) / student Week 4 H2O2 in dropper bottles 2 Mueller-Hinton agar plates / student

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Protocol: The Experimental Flowchart listed below details the order of the characterization process. The majority of the protocols (and how to interpret the results of tests) used in this laboratory are discussed in detail in other sections of the Lab Manual. Protocols specific to this laboratory are discussed below. For all tests and protocols isolate will be grown at 30°C for 24 hours under aerobic conditions unless stated otherwise. Experimental Flowchart: Characterization of Gram Positive Skin Isolate

Week Task

1 Streak throat swab across CNA petri plate.

2 Pick single Colony and inoculate TSA, TSB. Perform Gram Stain.

3 Streak on Blood Agar. Streak on Mannitol Salt Agar (MSA).

4 Antibiotic Sensitivity. Catalase Test Results: Blood Agar. MSA.

5 Results: Antibiotic Sensitivity

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Results: In addition to drawings and other descriptions of the tests performed, package all data in a central table such as the one listed below.

Test Result

Colony Color and Appearance

Gram Stain

Growth on Blood Agar

Mannitol Fermentation

Salt Tolerance

Catalase Test

Kirby Bauer- Erythromycin

Kirby Bauer- Gentamycin

Kirby Bauer- Chloramphenicol

Kirby Bauer- Bacitracin

Kirby Bauer- Ampicillin

Kirby Bauer- Vancomycin

Kirby Bauer- Oxacillin

Kirby Bauer- Tetracycline

Interpretations/Conclusions: Using Bergey’s Manual of Bacteriology, is it possible to identify the unknown isolate? If the organism displayed resistance to antibiotics, discuss how this resistance may have been acquired.

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19. IMViC TEST Background: The IMViC is a series of biochemical tests designed to separate Escherichia from Enterobacter or Klebsiella. While species of Enterobacter are generally nonpathogenic, strains of Escherichia and Klebsiella may be pathogenic. It is important to be able to quickly differentiate between these gram negative bacilli. The IMViC test uses 3 different media to perform 4 different analyses. The tests of the IMViC are outlined below. Indole- The Indole Test examines whether or not an organism can break down the amino acid tryptophan. If the organism contains the endoenzyme Tryptophanase, tryptophan will be broken down to Indole, Pyruvate and NH4

+. To facilitate the process, the Indole test is performed using Tryptone broth containing high levels of tryptophan. To test for the presence of Indole, Kovac’s reagent is added to an overnight culture of Tryptone Broth. Kovac’s Reagent is a nonpolar compound with a slight yellow tinge. In the presence of Indole, Kovac’s reagent chemically converts into a pinkish-red substance indicating a positive Indole Test. Methyl Red- Vogue Proskauer- The MR-VP tests examine whether acidic or non-acidic end products are created during glucose fermentation. In the MR-VP tests, a single tube of glucose broth is inoculated with the organism. After the organism has grown overnight, the sample is split into 2 test tubes. In one tube the Methyl Red Test is performed. In the other tube the Vogues-Proskauer Test is performed.

To perform the Methyl red Test, the pH indicator methyl red is added to the half-sample. If acidic end products are present, Methyl Red will detect the reduced pH and turn the entire sample red. If fermentation of glucose produces non-acidic end products, addition of Methyl Red will produce no change in the sample. To perform the Vogues-Proskauer Test, the reagents Barrit’s A (alpha napthol) and Barrit’s B (potassium hydroxide) are added to the half-sample. The reagents react with an intermediate in the non-acid fermentation pathway, acetyl methyl carbinol to produce a dark burgundy color. The dark burgundy color which takes 10-20 minutes to develop is indicative of a positive VP Test. Citrate- The Citrate Test examines whether an organism can use Citrate as its sole source of carbon. In the Citrate test, an organism is grown on a Citrate Slant where Sodium Citrate is the only source of carbon. In addition, the slant contains the pH indicator Bromthymol Blue which turns blue in a basic environment. If the organism can utilize Citrate as the sole carbon source, the bacterium can directly import citrate into the cell. The Citrate anion is co-transported with H+ across the membrane. As citrate is imported into the cell, the pH of the surrounding media begins to rise causing the agar to turn from green to Prussian Blue in appearance.

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Objective: Master the IMViC test. Scientific Question: How are Escherichia from Enterobacter differentiated by using the IMViC test? Materials: Week 1 1 TSA slant Escherichia coli / table 1 TSA slant Enterobacter aerogenes / table 2 Tryptone broth / student 2 MR-VP broth / student 2 Simmon’s Citrate slant / student Week 2 Kovac’s Reagent Barrit’s A and Barrit’s B Protocol: Indole Week 1 1. Using sterile technique, inoculate test organism into Tryptone broth. 2. Incubate at 37°C for 24 hours. 3. Store at 4°C. Indole Week 2 1. Add 10-15 drops of Kovac’s Reagent to culture. 2. View color of alcohol layer on surface to interpret result. MR-VP Week 1 1. Using sterile technique, inoculate test organism into MR-VP broth. 2. Incubate at 37°C for 24 hours. 3. Store at 4°C. MR-VP Week 2 1. Transfer half of grown sample to a sterile test tube. 2. To the first tube, add 10 drops of Methyl Red, swirl. 3. Examine tube for color change indicating presence of acid. 4. To the second tube add 10 drops each of Barrit’s A and Barrit’s B, swirl. 5. Let sit at room temperature for 10-20 minutes. 6. Examine the tube for the development of a burgundy color indicating the presence of non-acidic fermentation end products. Citrate Week 1

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1. Using sterile technique, streak test organism across surface of Citrate slant. 2. Incubate at 37°C for 24 hours. 3. Store at 4°C. Citrate Week 2 1. Examine tube for growth and report color of agar. Results: Color of broths should be noted before and after incubation. Note appearance of broth after test is performed (if applicable.) Interpretations/Conclusions: Summarize interpretations of IMViC Tests in table form.

Test E. coli Enterobacter aerogenes

Indole

MR

VP

Citrate

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Results- Citrate Test

+ -

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20. NITRATE REDUCTION Background: Nitrogen reduction is a form of anaerobic respiration where inorganic Nitrogen compounds act as the terminal electron acceptor in the Electron Transport Chain. The Nitrate Test evaluates whether or not an organism can reduce Nitrate (NO3

2-) to Nitrite (NO2

-) or even further reduced nitrogen compounds such as NH3 or N2 gas. The reaction NO3

2- + 2e- + 2H+ -> NO2- + H2O is catalyzed by the membrane bound

enzyme Nitrate Reductase. To activate expression of the genes involved in nitrogen reduction, the Nitrate Test is performed in Nitrate Broth containing a high concentration of NO3

2-. The Nitrate Test uses Alpha-Napthylamine and Sulfanilic Acid to test for the presence of Nitrite (NO2

-). Nitrite will react with these compounds to produce a red color throughout the tube, indicating the presence of the enzyme Nitrate Reductase.

The absence of red color upon addition of Alpha-Napthylamine and Sulfanilic Acid, however, does not necessarily indicate a negative Nitrate Test. As Alpha-Napthylamine and Sulfanilic Acid react only with Nitrite, further reduced forms of nitrogen (NH3 or N2 gas) will give the appearance of a false negative result. To determine whether a more reduced form of nitrogen is present, powdered zinc is added to the sample. Powdered zinc chemically converts Nitrate (NO3

2-) to Nitrite (NO2-). If

nitrate is still present in the broth, it will be converted by the zinc to nitrite where upon it will react with the already added Alpha-Napthylamine and Sulfanilic Acid and turn the solution red. In this case, after the addition of zinc, the appearance of a red color in the broth is a negative test result. However, if upon the addition of zinc, no color change occurs, the nitrogen has already been converted by the bacterium to a more reduced form such as NH3 or N2 gas. In this case, the lack of a red color indicates a positive Nitrate Test. Objective: Master the Nitrate test. Learn how to interpret results properly. Scientific Question: Is the tested organism capable of Nitrate Reduction? What form of Nitrogen is generated by the organism when grown in Nitrate Broth? Materials: Week 1 8 Nitrate broth /table 1 TSB broth Escherichia coli /table 1 TSB broth Enterobacter aerogenes /table 1 TSB broth Pseudomonas aeruginosa /table 1 TSB broth Pseudomonas fluorencens /table 1 Nitrate Broth/ student 1 TSB broth containing unknown organism

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Week 2 Alpha-Napthylamine Sulfanilic Acid Powdered Zinc Protocol: Week 1 1. Using sterile technique, inoculate test organism into Nitrate broth. 2. Incubate most test organisms at 37°C for 24 hours. (Incubate P. fluorencens at 30°C for 24 hours.) 3. Store at 4°C. Week 2 1. Add 10 drops of Alpha-Napthylamine and 10 drops of Sulfanilic Acid to sample. 2. Record appearance of broth. 3. If no color change, add a minute amount of powdered zinc to sample. 4. Record appearance of broth. Results:

Organism Color after addition of Alpha-Napthylamine and Sulfanilic Acid

Color after addition of zinc (if necessary)

Nitrate Test Result (+/-)

Uninoculated Nitrate Broth

Escherichia coli

Enterobacter aerogenes

Pseudomonas aeruginosa

Pseudomonas fluorencens

Interpretations/Conclusions: Address the Scientific Question stated above.

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21. UREA HYDROLYSIS Background: Bacteria possessing the exoenzyme Urease have the ability to break down urea into carbon dioxide and ammonia. These end products may serve as carbon and nitrogen sources for the organism, in addition to raising the pH of the surrounding media. The Urea Test is used to differentiate between nonpathogenic, non-lactose fermenters such as Proteus from potential pathogenic non-lactose fermenters such as Salmonella and Shigella. The Urea test relies on the pH increase caused by the generation of NH4

+ during urea hydrolysis. The pH indicator dye Phenol Red is added to Urea Broth. Phenol Red (and the subsequent Urea Broth) is salmon color at pH =6.8 but turns cerise when the pH increases beyond 8.1. The color change from salmon to cerise indicates a positive urea test. Objective: Master the Urea Test. Scientific Question: Does the tested organism have the ability to hydrolyze urea? Materials: 1 TSB broth Proteus vulgaris / table 1 TSB broth Salmonella / table 2 Urea broth / table Protocol: Week 1 1. Inoculate Urea broth with test organism. 2. Incubate at 37°C for 24 hours. 3. Store at 4°C. Week 2 1. Observe appearance of tube. Evaluate organism’s ability to hydrolyze urea. Results: Include drawings of urea Broth pre- and post incubation for each organism tested. Interpretations/Conclusions: Address the Scientific Question stated above.

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+ -

Urea Test - Results

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22. MOTILITY TEST Background: The Motility Test offers a quick and easy way to evaluate the motility of an organism. An Inoculating Stab is used to introduce a bacterial species into a tube of Motility Agar. If the organism is motile, it will grow outward, away from the line created by stabbing the agar when the bacteria is introduced (Stab Line). Often motile organisms will also grow across the surface of the agar and then back into the agar from the surface. (See demonstration.) Temperature may of concern when performing the Motility Test. Proteus vulgaris, for example, is motile at 30 °C but loses its flagella at 37 °C. Objective: Master Motility Test Scientific Question: What is the Motility designation for each organism tested? Materials: Week 1 1 TSA slant Escherichia coli / table 1 TSA slant Staphylococcus simulans / table 2 Motility Agar Tubes / student Week 2 1 TSA slant Unknown / student 1 Motility Agar Tube / student Protocol: 1. Using a sterile Inoculating Stab, pick a portion of a bacterial colony with tip of wire. 2. Spear Inoculating Stab 1-2 inches into Motility Agar Tubes. 3. Incubate at 37 °C for 24 hours under aerobic conditions. Results: Draw pictures of raw data.

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Interpretations/Conclusions: Answer Scientific Question stated above.

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Growth beyond the stab line

Growth from the surface into the media

+ -

Growth along the stab line only

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23. HEMOLYSIS ON BLOOD AGAR Background: Blood Agar is a commercial preparation containing intact red blood cells from sheep. Blood Agar is a Differential media, allowing one to differentiate organisms based on hemolysis, the ability to lyse blood cells. There are three basic patterns of hemolysis- Alpha (α), Beta (β) and Gamma (γ). Alpha Hemolysis- Partial lysis of red blood cells produces a greenish halo surrounding bacterial colony. Beta Hemolysis- Complete lysis of red blood cells results in a clear area surrounding the bacterial colony to be tested. Gamma Hemolysis- No lysis of red blood cells. Objective: Observe patterns of hemolysis generated by test organisms. Scientific Question: What pattern of hemolysis is produced by test organism? Materials: 1 TSB broth Strep Group C / table 1 TSB broth Streptococcus mitis / table 1 TSB broth Staphylococcus simulans /table 3 Blood Agar Petri dish/ table Protocol: Week 1 1. Streak organism across blood agar. 2. Incubate in Candle Jar at 30 °C for 48 hours. 3. Transfer to 4 °C. Week 2 1. Observe growth and hemolysis pattern. Results: Include drawings of hemolysis patterns generated by all organisms tested. Interpretations/Conclusions: Address Scientific Question stated above.

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24. MSA- Mannitol Salt Agar Background: Mannitol Salt Agar (MSA) is a Selective and Differential media. MSA contains Mannitol, 7.5 % NaCl and the pH indicator Phenol Red. Organisms able to ferment mannitol will reduce the pH of the surrounding media, causing MSA to turn yellow. However, MSA also selects for organisms capable of growing in a high salt environment. Thus, if a tested organism is sensitive to high salt one is unable to determine the organism’s ability to ferment mannitol. MSA is used frequently to differentiate between species in the family Micrococcaceae. Staphylococcus aureus is a potential pathogen able to ferment mannitol in a high salt environment. Objective: Master use of Mannitol Salt Agar Scientific Question: What is the salt tolerance of the organisms tested? Are the salt tolerant organisms able to ferment mannitol? Materials: 1 TSA slant Staphylococcus simulans/ table 1 TSA slant Staphylococcus epidermis/ table 1 MSA Petri plate /table Protocol: Week 1 1. Draw a line on bottom of Petri dish, splitting plate in halves. 2. Streak each organism to be tested across half section of MSA plate. 2. Incubate at 30 °C for 48 hours. 3. Transfer to 4 °C. Week 2 1. Observe growth and fermentation patterns.

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Results: Include drawings of all data. Interpretations/Conclusions: Address Scientific Question stated above.

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25. Gelatin Background: The Gelatin test is a simple method for determining whether or not an organism has the ability to produce the exoenzyme Gelatinase. Gelatin is a solid at temperatures below 28°C however turns to liquid at higher temperatures. Organisms that produce Gelatinase will hydrolyze Gelatin into sub-units that will remain in liquid form even at temperatures below 28°C. Gelatin is a component of connective tissue and the extracellular matrix. An organism’s ability to break down gelatin may enhance its pathogenicity. In this laboratory, an organism is incubated for several days at 30°C in Gelatin agar. During the incubation process the Gelatin may liquefy regardless whether the enzyme Gelatinase is produced. To avoid false-positive results, the Gelatin Agar Tube is stored at 4°C for 30 minutes before the contents examined. Hydrolyzed gelatin will remain liquid at 4 °C, indicating a positive Gelatin test. Gelatin that has not been broken down will solidify 4 °C. Objective: Master the Gelatin Test Scientific Question: Does the tested organism produce Gelatinase? Materials: 1 TSB broth Escherichia coli /table 1 TSB broth Proteus vulgaris / table 1 TSB broth Enterobacter aerogenes / table 1 TSB broth pseudomonas fluorencens / table 4 Tubes Gelatin Agar / table Protocol: Week 1 1. Inoculate 100 μl of specimen into Gelatin Agar Tube. 2. Use Inoculating Stab to make repeated stabs into Gelatin Agar. 3. Incubate at 30°C for 7 days. Week 2 1. Store Gelatin Agar at 4°C for 30 minutes.

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2. Examine consistency of agar. Results: Include drawings of Gelatin tubes before and after incubation. Interpretations/Conclusions: Address the Scientific Question stated above.

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26. OXIDASE TEST Background: The Oxidase Test tests for the presence of the enzyme Cytochrome c, an enzyme complex involved in the Electron Transport Chain of certain organisms. Often the Oxidase test is used in clinical settings to test for the presence of Pseudomonas aeruginosa, an opportunistic pathogen implicated in various nosocomial infections. Bergey’s Manual of Determinative Bacteriology provides additional classification information using the Oxidase test. The Oxidase test utilizes a commercially prepared strip containing a compound that reacts with Cytochrome c. Reaction between the compound and Cytochrome c results in the formation of a purple color on the strip. Although the Oxidase Test is technically simple, care must be used when interpreting the results. It is important to use a slant culture to ensure a large enough sample will be in contact with the Oxi-Strip. Sometimes a Oxidase positive organism will only produce the purple color on the swab used to rub the Oxi-strip and not on the strip itself. In addition, some organisms will generate a purple color on the Oxi-Strip 5 minutes after performing the test. If the purple color does not develop instantly, consider the result a negative Oxidase test. Objective: Master the Oxidase Test Scientific Question: Does the tested organism possess the enzyme Cytochrome c? Materials: 2 Oxi-Strips/table 4 Sterile Swabs / table 1 empty perti dish (no agar) /table 1 TSA slant of Alcaligenes faecalis 1 TSA slant of Pseudomonas fluorencens 1 TSA slant of Escherichia coli 1 TSA slant of Pseudomonas aeruginosa

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Protocol: Perform 3 tests on a single Oxi-Strip! 1. Place an Oxi-Strip in the bottom (or lid) of an empty Petri plate. 2. Add a small amount of de-ionized water to the strip. 3. Use a sterile swab to sample a large swath of TSA slant of tested organism. 4. Rub swab + organism on a small section of Oxi-Strip. 5. Immediately look for the appearance of purple color. 6. Repeat procedure with another organism to be tested, using clean area of same Oxi-Strip. 7. Repeat procedure with 3rd organism to be tested, using clean area of same Oxi-Strip. Results: Describe color of Oxi-Strip following test procedure. Interpretations/Conclusions: Address the Scientific Question stated above.

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27. CATALASE TEST Background: Hydrogen Peroxide (H2O2) is a compound with antimicrobial properties. H2O2 is not stable and breaks down into oxygen free radicals. H2O2 is found in the lysosome of Neutrophils and other phagocytic cells of the immune system. It is also used as a topical treatment against infection. Some aerobic organisms produce the enzyme catalase. Catalase converts H2O2 into harmless end products oxygen and water. The Catalase test is largely used to differentiate between Staphylococcus and Streptococcus species. Applying 3% H2O2 to Catalase Positive organism will result in the immediate formation of oxygen bubbles. Objective: Master the Catalase Test. Scientific Question: Does the tested organism produce catalase? Materials: Glass Slides Dropper bottles containing 3% H2O2 1 TSA slant Staphylococcus Simulans /table 1 TSA slant Strep Group C /table Protocol: 1. Use an inoculating loop to transfer a small amount of colony to glass slide. 2. Add 2-3 drops of 3% H2O2. 3. Examine sample for the presence of oxygen bubbles. Results: Draw and describe the glass plates after the addition of 3% H2O2. Interpretations/Conclusions: Address the Scientific Question stated above.

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28. KIA TEST Background: The KIA Test is often used as a compliment to the IMViC in identifying Gram negative enteric pathogens. KIA agar (Klinger’s Iron Agar) contains a mixture of 1% lactose, 0.1 % glucose, Ferrous Sulfate and the pH indicator dye Phenol Red. KIA Test allows to simultaneously assay for glucose and lactose fermentation, as well as the production of H2S gas. In the KIA test, the organism to be tested is stabbed into the agar and struck across the surface of the slant before incubation. Acid production during glucose fermentation will cause the Phenol Red pH indicator to turn the agar at the butt of the slant yellow. Lactose fermentation will cause the Phenol Red pH indicator to turn the agar at the top of the slant yellow. H2S gas produced during fermentation will react with the Ferrous Sulfate (FeSO4) present in the agar to form Ferric Sulfate (Fe(SO4)3). Ferric Sulfate forms a black precipitate in the KIA agar. Other gas production may result in cracks or bubbles in the KIA agar. Objective: Master the KIA test. Scientific Question: What does incubation in KIA agar tell us about the metabolic capabilities of the tested organism? Materials: 4 KIA agar slants / table 1 TSA slant of Alcaligenes faecalis 1 TSA slant of Proteus vulgaris 1 TSA slant of Escherichia coli 1 TSA slant of Enterobacter aerogenes

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Protocol: Week 1 1. Using an inoculating stab, inoculate KIA agar slant in the following manner: Stab organism into agar then streak stab tool across surface of slant. 2. Incubate at 37°C for 24 hours. 3. Store at 4°C. Week 2 1. Examine color and appearance of KIA agar for results. Results: Include drawings of KIA agar before and after incubation. Interpretations/Conclusions: Address the Scientific Question listed above.

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29. Transformation

Background: Transformation is the process by which bacteria cells “pick up” or accept foreign DNA from the environment. If the bacteria are able to maintain the exogenous DNA (either as a self-replicating plasmid or by adding it to the chromosome through recombination) the bacteria is said to be transformed. This terminology is derived from Frederick Griffiths famous 1928 experiment involving Rough and Smooth strains of Streptococci. Bacteria that can acquire foreign DNA naturally and said to be competent for transformation. In the lab bacteria such as E. coli that are not naturally competent can be made so through various methodologies. In molecular biology competent E. coli are of often transformed with a self-replicating plasmid, but other vectors exist as well. Common methods for introducing a plasmid vector into E. coli include Electroporation (using an electric current to drive the DNA into the cell) and a Heat Shock method that is discussed below. The preparation of electro-competent and chemi-competent E. coli is done using different protocols. One should never use chemi-competent cells for electroporation. Electroporation has its advantages over heat Shock transformation in that it generates a higher transformation efficiency. However for many cloning applications chemi-competent cells work fine. Plasmid Pglo: Pglo is a plasmid (5371 base pairs) developed by BioRad corporation. The plasmid contains an E. coli origin of replication, the gene for Beta Lactamase driven by a constitutive promoter (meaning the gene will always be expressed.) The enzyme Beta Lactamase cleaves the beta lactam ring of ampicillin, rendering the antibiotic ineffective. The plasmid also contains a gene encoding for Green Fluorescent Protein (GFP). This gene was originally isolated from the jelly fish Aequorea victoria. The gene for GFP is driven by an inducible promoter pBAD. The pBAD promoter is natively part of the arabinose operon in E. coli. (The genes in the native arabinose operon encode for enzymes involved in the catabolism of arabinose.) The regulatory protein AraC binds the pBAD promoter. In the absence of arabinose AraC adopts a conformation that allows the DNA binding protein to act as a repressor of transcription. In the presence of arabinose the protein undergoes an allosteric activation and now functions as a transcription factor, recruiting RNA Polymerase to the

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pBAD promoter. A gene encoding for the protein AraC is also found on the pGLO plasmid. The gene is driven by a constitutive promoter. For this lab each pair of students will perform 2 transformations with plasmid pGlo as well as one set of controls (Control A, Control B and Control C.) Materials: Styrofoam ice buckets (filled to rim): 1 per 2 students 250 ml beaker containing ethanol and cell spreaders: 1 per 2 students Chemi-competent E. coli: 50 μl per student. Tubes of cells should be thawing on ice when lab begins. (For future labs, cells should be prepared in tubes containing 100 μl cells per tube so that each pair of students has their own tube.) Plasmid pGlo: 2-5 μl per student, depending on the concentration of the DNA prep. Tubes of DNA should be thawing on ice when lab begins. (For future labs create a set of tubes with excess DNA in each tube. Samples can be reused between BIO 220 and BIO 244 and possibly 212 series courses. Create separate preps of pGlo for BIO 100 and BIO 205 courses. If stored in -80° C, DNA should be good for at least one academic year.) Microcentrifuge tubes: 1 per student Microcentrifuge tube racks: 1 per student Test tubes containing 1 ml- 2ml Luria Both (no ampicillin): 1 per student Luria Agar plates containing ampicillin (100μg/ml agar) and L-Arabinose (0.2 g/100 ml agar): 1 per student Luria Agar plates (no ampicillin, no L-arabinose): 1 per 2 students Dry Bath set at 37° C - 42° C. Bath should be at temperature before lab begins. Micropipettes (p1000, p200, p20) and tips will be provided by instructor.

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Protocol: 1. Thaw chemi-competent cells on ice. (If cells have previously been thawed and re-frozen, make sure cells are suspended uniformly and not settled on the bottom.) 2. Add 50 μl chemi-competent cells to a sterile microcentrifuge tube already chilled on ice. 3. Add into the cells 5 μl plasmid Pglo DNA (1-pg – 100 ng is the optimal range for supercoiled plasmids). 4. Let cells and DNA incubate together on ice for 30 minutes. 5. Heat Shock: Place cells in 42° C dry bath for 30 seconds. (Other protocols call for 37° heat shock for 3 minutes). 6. Place tubes immediately on ice for 5 minutes. 7. Add 600 μl luria broth to sample. 8. Incubate at 37° C for 30 minutes in a shaking incubator. 9. Plate cells on Luria Agar (containing Ampicillin and L- arabinose). 10. Incubate plate upside down, overnight at 37° C. 11. Week 2: Record data. Observe plates under ultraviolet light. Protocol for Control A: (Cells without plasmid) The Control A protocol is performed as above with slight modifications: Step 3 is eliminated. Control A is plated on luria agar containing ampicillin and L-arabinose. What is the purpose of Control A? Protocol for Control B: (Cells without plasmid plated on luria agar lacking ampicillin and L-arabinose.) The Control B protocol is performed as above with slight modifications: Step 3 is eliminated. Control B is plated on luria agar lacking both ampicillin and L-arabinose. What is the purpose of Control B?

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Protocol for Control C: (Cells +pGlo plated on luria agar + ampicillin but lacking L-arabinose.) The Control A protocol is performed as above with slight modifications: Step 9 is modified- cells are plated on luria agar + ampicillin plates that lack L- arabinose. What is the purpose of control C? Results: Record appearance of plates post incubation. Record relative size of colonies, distribution across plate surface. Phenotype of bacteria under ultraviolet light. Conclusions/ Interpretations: Evaluate the success of carrying out the transformation protocol. Interpret the data on Control plates. Notes on steps: 1. If cells are thawed at room temperature or hand-thawed a decrease in transformation efficiency will result. 2. Although technically the order (DNA or cells in the tube) doesn’t matter, it is preferable to add the larger volume component first. 3. To facilitate diffusion of the DNA among the cells, some molecular biologists will gently move the micropipette tip around in the cells as the DNA is being added. 4. Incubating cells + DNA on ice allows for the negatively charged DNA to coordinate with the positively charged calcium ions on the surface of the cell. 5. The Heat Shock step expands the pores in the plasma membrane created by the calcium ions. Opening of the pores seems to drive the DNA inside the cells. If the pores are left open too long the cytoplasm will leak from the cells and they will die. 6. Placing the cells on ice closes the pores, trapping the DNA inside. 7. Luria broth provides nutrients for the cells. 8. Incubation of the cells + luria broth allows for expression of the gene conferring antibiotic resistance. It is essential that this protein is synthesized before plating the cells on agar containing the antibiotic.

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9. Plating the cells allows for single cell transformants to be isolated. 10. Incubation allows for the single cell transformants to grow into a colony as well as providing selective pressure to kill the cells that did not acquire the plasmid. L-arabinose acts as an inducer of pBAD, allowing expression of the gene for Green Flourescent Protein.

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30. Mini-Prep Background Because of the ease by which it can be transformed, E. coli is often used to harbor and amplify certain man-made plasmids. Various methodologies have been developed to isolated plasmid DNA from bacteria. The Alkaline Lysis method utilizes a series of 3 solutions- Lysis I, Lysis II and Lysis III. The roles for each solution in the protocol are listed below. Protocol- Alkaline Lysis Mini-Prep of plasmid DNA Protocol is largely performed in 1.5 ml microcentrifuge tubes and table top centrifuges at room temperature and 4 ° C.

1. Transfer overnight culture (grown in Luria Broth + 100 μg/ml Ampicillin) to 1.5 ml microcentrifuge tube.

2. Spin cells for 20 seconds at room temperature. 3. Remove supernatant and discard. 4. Resuspend cellular pellet in 100 μl Lysis I solution. 5. Add 200 μl Lysis II, invert tube 5 times, and place cells on ice 5 minutes. 6. Add 150 μl Lysis III, invert tube 5 times, and place tube on ice 5 minutes. 7. Spin for 5 minutes at room temperature. 8. Transfer DNA containing aqueous solution to a new microcentrifuge tube. 9. Add 500 μl of Isopropanol. Mix thoroughly, place on ice 10 minutes. 10. Spin 12 minutes at 4 ° C. 11. Remove and discard alcohol supernatant being careful not to disturb the DNA/salt

pellet at the bottom of the tube. 12. Dry pellet using speed vacuum or leave open and let air dry for 1 week. 13. Re-suspend pellet in 50 μl H20. 14. Store at -20° C.

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Role of Reagents- Alkaline Lysis mini-prep of plasmid DNA. 1. Lysis I Glucose- maintains osmotic balance. Cells do not prematurely lyse. Tris – buffering agent for pH stability. EDTA- chelates divalent metal ions (Mg++, Ca++) which are required for certain enzymes to be active. Removal of the ions will prevent nucleases from degrading the DNA. 2. Lysis II- NaOH- basic environment lyses cells and denatures double stranded DNA. SDS- detergent disrupts phospholipid bilayer of membrane. 3. Lysis III K+ Acetate – neutralizes the pH. Double stranded DNA reanneals. Raising pH causes cellular debris to precipitate out of solution. Chromosomal DNA remains membrane bound and will precipitate along with cellular debris.

4. Isopropanol - DNA is insoluble in equal volume Isopropanol and can be separated from aqueous solution.

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31. Restriction Endonucleases Background: Restriction endonucleases or restriction enzymes are naturally occurring enzymes produced by bacteria as a defense against viral infections. Restriction enzymes cleave DNA in a site specific manner. Here is a link to detailed descriptions of restriction enzymes:

http://en.wikipedia.org/wiki/Restriction_enzyme Today restriction enzymes are mass produced and commercially available from many biotech firms. Restriction enzymes allow molecular biologists to cut DNA at a specific sequence and generate a predictable end product. Because of the ease by which DNA can be manipulated, restriction enzymes are an integral part of any cloning procedure. New England Biolabs has been selling restriction enzymes for a several decades. Her is a link to an overview of restriction enzymes provided by NEB:

http://www.neb.com/nebecomm/tech_reference/restriction_enzymes/overview.asp Objectives: Digest plasmid pGlo with restriction endonuclease BamH1 to remove the green fluorescent protein (gfp) gene. Resulting DNA will be characterized by electrophoresis. (We are basing this experiment on a map of pGLO that has not been verified and are expecting to see 2 DNA fragments; 900 base pairs (gfp gene) and 4471 base pairs (remainder of the pGlo plasmid). Digest Protocol: 5 μl DNA 2 μl NEB 2 (10x buffer) 1 μl restriction enzyme BamH1 (NEB 5 units/ μl) 12 μl H20 Incubate sample at 37°C for 1 hour. Add 6 μl 6x loading dye to sample. Load sample in 0.7% agarose gel.

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Preparation of 0.7% agarose gel: Weigh out 0.7 g agarose. Transfer to 500 ml Erlenmeyer flask. Add 100 ml 1x SB (Sodium Borate) buffer. (100 ml makes 2 gels) Stopper flask with paper towel and place in microwave. Heat flask until agarose is completely dissolved. (2-3 minutes) Let flask cool. Add 5 μl Sybr-Safe DNA stain to the agarose – swirl flask Pour contents in casting tray. (See instructor demo.) Insert well-comb before agarose solidifies. Results: Sample data table for electrophoresis

Lane Sample # ID Volume

1 1 kb base pair ladder 5 μl total

2 original mini prep

pGLO uncut 5 μl DNA + 2 μl 6x dye

3 Digested plasmid

pGLO + BamH1 Entire reaction + 6 μl 6x dye

Photograph gel. Conclusions: Did the restriction digest cut to completion? What observations can be made about uncut verses cut plasmid DNA?

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32. Transposon Mutagenesis

Overview: Mate A. radiobacter K84 and E. coli BW20767/pRL27 to allow transposition of K84 to occur, select for transposed strains on Km50 Rif100 plates. Screen for auxotrophic mutants by growing Kanamycin + Rifampin resistant cells on minimal media. Background Information:

1. Host Strain - Agrobacterium radiobacter K84 a. A biocontrol strain – produces agrocin 84 which inhibits tumor formation

when K84 is mixed with some tumor forming Agrobacterium strains. b. The genome is composed of 5 replicons, the entirety of which have been

sequenced. Sequence can be viewed at “Agrobacterium.org”. Chromosome 1 Chromosome 2 Plasmid 1 (388k) Plasmid 2 (185k) Plasmid 3 (44k)

2. Donor Strain - E. coli BW20767/pRL27 (see appendix for map of pRL27 Vector) a. plasmid carries transposon delimited by inverted repeats b. transposable element contains an origin of replication so that it can be recovered

and maintained epigenetically after transposition c. transposase resides outside of transposable element; once transposition has

occurred, the transposon is “fixed” into the genome at that location and can no longer move.

Protocols:

1. Growth of host and donor strains. (To be done preceding Lab I by Staff.)

a. Grow a 3 ml overnight culture (18-24 hrs) of A. rhizogenes K84Rif in MG/L broth (no antibiotic is needed because the Rif resistance is chromosomal), shaking at 225 rpm at 28°C.

b. Grow a 3 ml overnight culture (18-24 hrs) of E. coli BW20767 (pRL27) in LB containing 50μg/ml Km, shaking at 225 rpm at 37°C. (The plasmid is readily lost in the absence of Km, so be sure to include it to provide selection pressure for maintenance of the plasmid).

2. Lab I - Bacterial Mating for Agrobacterium radiobacter K84Rif with Escherichia coli

BW20767/pRL27. a. Pipet 1ml of each culture (K84Rif recipient and BW20767/pRL27 donor

strains) into separate sterile microfuge tubes and pellet the cells using a

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tabletop microfuge-spin at 13,000 rpm for 30 seconds. Discard the supernatant into a biohazard bucket.

b. Resuspend resulting pellets in 400 μl each of MG/L broth. (This step eliminates residual antibiotic that would be toxic to the host A. radiobacter K84 strain and concentrates the cells for efficient mating). Combine the 2 cultures.

c. Pipet the cell suspension onto 4 places of an MG/L plate (leave as 4 puddles, do not spread). Let the plate sit on the bench for 15-20 minutes. Afterward

3. Lab II - Selection of RL27 Transposon Mutants a. Scoop up the mating puddle using a sterile inoculating loop and resuspend in

1000 μl MG/L broth. Plate 330 μl onto each of 3 MG/L plates containing Rif100 Km50. This step selects for host cells (Rif resistant) which have acquired an integrated transposon (Km resistant), i.e. mutant strains. Incubate the plates for 1-colonies when transposition has occurred efficiently.

4. Lab III- Identification of Auxotrophic Mutants. Select individual colonies with a

sterile toothpick and inoculate onto an MG/L Rif100 Km50 plate in a 36 well grid pattern and on a minimal media agar plate in a 36 well grid pattern. Grow both plates overnight at 28C. It is important to plate a particular mutant in the same square of the grid (i.e. A1) regardless of the type of media used.

5. Lab IV - Identification of Amino Acid Auxotrophs. All colonies plated on MG/L Rif100 Km50 grid plates will grow, however not all organisms will be able to grow on minimal media. Organisms that are unable to grow on minimal media are auxotrophs –mutants deficient in the synthesis of an essential metabolite. To further characterize and identify amino acid auxotrophs The colonies that grew on MGL but not minimal media will be picked and replica plated on a series of 36 well grid plates: MGL, minimal media + polar uncharged amino acids, minimal media + nonpolar amino acids, minimal media + acidic amino acids, and minimal media + basic amino acids. An amino acid auxotroph should show growth on the MGL plate as well as one of the minimal media plates (with added amino acids). Such growth will not identify which amino acid the mutant cannot synthesize, however it will narrow the group to a particular class of amino acids. Further experimentation performed by independent studies students will further characterize the mutants.

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Tech Prep- make all reagents available by Lab 1. Instructor will access reagents from refrigerators. Turbid culture of Agrobacterium radiobacter strain k84 Rif in 2ml MG/L broth (no antibiotics). 1/ student Turbid culture of E. coli BW20767/pRL27 in 2ml luria broth (kanamycin 50 μg/ml). 1/student 5 ml tube of 0.9% NaCl. 1/Student (Prepare in short tubes). MG/L agar plate (no antibiotic) 1/Student

tetA promoteroriT

transposase

Km Resistance (aph)

tpnRL 17-1

tpnRL 13-2

R6K Ori

Tn5 RL27 IR

Tn5 IR

Transposon RL27

pRL27 Transposon

4100bp

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MG/L agar plate (Rifampin 100 μg/ml , kanamycin 50 μg/ml) 3/Student 5 ml tube of MG/L broth (no antibiotics) 1/ Student MGL agar plate (Rifampin 100 μg/ml , kanamycin 50 μg/ml) grid plates at least 3 /Student Minimal media agar plate (Rifampin 100 μg/ml , kanamycin 50 μg/ml) grid plates at least 3 /Student Controls: Positive Control: 5 ml culture MG/L broth 1/ 4 Students any of the A. radiobacter k84 (RifR, KanR) generated in previous BIO 220 labs. Negative Control: 5 ml culture MG/L broth 1/ 4 Students A. radiobacter k84 (RifR, KanR) auxotrophic mutants created by Slater

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33. Polymerase Chain Reaction Background: Polymerase Chain Reaction (PCR) is in essence an application of DNA replication. Original PCR methods were used solely to amplify stretches of DNA. Today PCR is used for a variety of applications including: amplification of DNA, site-directed mutagenesis, DNA sequencing, creating gene fusions and deletions, genotyping and forensic applications. PCR is a synthesis reaction and thus contains all the compounds required to synthesize DNA in addition to a thermal stable DNA Polymerase which is also required. What makes PCR unique compared to many other synthesis reactions is a thermal cycle (the reaction takes place in 3 stages involving 3 different temperatures). PCR is also unique in that the thermal cycler is repeated, generally 20-30 times during the course of a reaction. Here is a link to more information on PCR: http://en.wikipedia.org/wiki/Polymerase_chain_reaction#PCR_principles_and_procedure

Tavi has a website dedicated to PCR http://www.med.yale.edu/genetics/ward/tavi/PCR.html Fermentas corporation also has website dedicated to PCR http://www.fermentas.com/techinfo/pcr/dnaamplprotocol.htm Steps of a PCR Thermal Cycle: Initialization 94° C 5 minutes During this step the DNA template is denatured into single strands. Many PCR protocols with short DNA templates don’t use the initialization step, however if you are performing PCR using genomic DNA as a template it might be a good idea. Denature 94° C, 30 seconds During this step double stranded DNA is melted into single strands. Anneal 55° C, 30 – 60 seconds During this step the primers will bind to the target sequence. Once this has taken place the Taq Polymerase will bind the primer:template duplex and extend off the 3’ end of the primer, synthesizing the compliment of the template strand DNA.

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Extension 72° C, 30-90 seconds During the extension step Taq Polymerase continues to synthesize the compliment of the template strand. The increased temperature of the extension step ensures that primers hybridize to the template only during the anneal stage. Thus all synthesis occurs in a coordinated manner. Cycle is repeated 25-30 times Final Extension 72° C, 5:00 The final extension is optional, and generally used when amplifying especially long DNA sequences such as a viral genome. The final extension allows any PCR products whose synthesis was not complete by the end of the extension step to be “cleaned up”. 4° Hold (useful for overnight experiments) Often this step is programmed in at the end of the thermal cycles. The PCR product is kept cool and will not evaporate while sitting in the thermal cycler. These steps are listed as generalities. Many times PCR reactions that are novel may have to play with experimental conditions to optimize the reaction for a specific set of primers and DNA template. Components of a Typical PCR Reaction: 100 ng DNA Template 0.4 µM Upstream Primer 0.4 µM Downstream Primer 200 µM of each dNTP 1 unit Taq Polymerase 10x PCR Buffer (containing MgCl2) Ultra pure H20 General Example of a PCR Reaction 1 µl DNA template 1.5 µl Primer 1 [15 µM starting concentration] 1.5 µl Primer 2 [15 µM starting concentration] 1.0 µl 10mM dNTP’s 1.0 µl Taq Polymerase (2.5 U/µl) 5 µl 10x PCR buffer (containing MgCl2) 39.5 µl ultra pure H20

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Objective: Amplify the DNA Ligase from bacteriophage T4 using PCR. The intended product is approximately 1464 base pairs. 1 PCR reaction/student. Materials – PCR reaction: 1 µl DNA template (bacteriophage T4) 1.5 µl Primer T4 forward [15 µM starting concentration] 1.5 µl Primer T4 reverse [15 µM starting concentration] 1.0 µl 10mM dNTP’s 1.0 µl Taq Polymerase (2.5 U/µl) 5 µl 10x PCR buffer (containing MgCl2) 39.0 µl ultra pure H20 Protocol: Prepare PCR reactions. Load into thermal cycler using the following parameters: 94° C, 1:00 55° C, 30 seconds 74° C, 1:00 30 cycles 4° C hold Preparation of 0.7% agarose gel: Weigh out 0.7 g agarose. Transfer to 500 ml Erlenmeyer flask. Add 100 ml 1x SB (Sodium Borate) buffer. (100 ml makes 2 gels) Stopper flask with paper towel and place in microwave. Heat flask until agarose is completely dissolved. (2-3 minutes) Let flask cool. Add 5 μl Sybr-Safe DNA stain to the agarose – swirl flask Pour contents in casting tray. (See instructor demo.) Insert well-comb before agarose solidifies. Results: (Week 2) Samples will be visualized on a 0.7% Agarose gel Photograph results using the gel documentation system. Conclusion: Was the PCR successful? What modifications may you want to try to optimize the reaction?

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Tech Prep: Week 1: PCR solution set. 1 set of tubes/group of 4 students.

Solution Volume

Bacteriophage T4 5 µl

10 mM dNTP’s 5 µl

10x PCR buffer (+Mg++) 25 µl

15 µM T4 forward primer 8 µl

15 µM T4 reverse primer 8 µl

H20 250 µl

Taq Polymerase* Instructor will distribute

Week 2: 0.7 % agarose melted in 1x SB buffer (enough for 5 small gels) 1x SB – enough to run all gels

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34. Antibiotic Sensitivity (MIC Test) Background: Antibiotic sensitivity is of primary concern to clinical microbiologists and health care professionals. Identifying the causative agent of a microbial infection, and then identifying what resistance, if any, the organism has to antibiotics will dictate the initial steps of treatment. One way to determine bacterial resistance to a wide range of antibiotics is to perform the MIC test (Minimum Inhibitory Concentration). In essence, an organism is struck across Mueller Hinton agar to create a confluent layer across the agar surface. Next, paper discs containing a particular antibiotic are applied to the surface of the agar. The organism is grown overnight and the results are determined. If an organism is resistant to an organism its growth will not be inhibited by the antibiotic – it will grow directly up to the paper disc. If the organism is sensitive to the antibiotic, a zone of inhibition (clearing area) will be observed. The diameter of the zone of inhibition can be measured and then a designation; resistant, sensitive or middle state can be applied. (Different antibiotics will diffuse through the media at different rates, thus no one size of a zone of inhibition is appropriate for all antibiotics.) Brock’s 12th edition Chapter 32 has a section on the MIC test. Below is a link to more information on the MIC test: http://en.wikipedia.org/wiki/Minimum_inhibitory_concentration Objective: The MIC test will be performed on skin and throat isolates to determine antibiotic resistance of the organism. Antibiotic resistance of other organisms isolated during the course of the semester may also be tested. Scientific Question: Is the organism(s) tested resistance to antibiotics? If so, which ones? Which antibiotics is the organism(s) sensitive to? Materials: Overnight TSB culture of the organism to be tested. 1 large Mueller Hinton agar plate per organism tested. 1 sterile swab per agar plate.

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Protocol: 1. Insert a sterile swab into an overnight broth of the organism to be tested. 2. Streak across the plate covering the entire surface. (Your instructor will demonstrate the appropriate technique for this protocol.) 3. Using the antibiotic dispenser, apply antibiotics to the surface of the agar plate. (One large Mueller Hinton plate can accommodate at least 6 antibiotic discs.) Your instructor will announce which antibiotics will be used. 4. Tap the surface of the antibiotic disc with a sterile tweezers. One small tap will ensure that the disc will stay adhered to the agar surface. 5. Incubate overnight at 37 degrees C. 6. Post incubation, measure the zone of inhibition surrounding each antibiotic disc and the bacterial growth. 7. Using the manufacturer’s guidelines, determine if the organism is resistant, sensitive or intermediate. Results: 1. Draw or photograph results. 2. Measure the zone of inhibition for each antibiotic. 3. Determine if the organism is resistant, sensitive or intermediate. Conclusions: Address the scientific question stated above.

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35. MacConkey Agar Background: MacConkey agar is an example of selective and differential media. MacConkey agar contains bile salts and the dye crystal violet which together inhibits the growth of most gram Positive bacteria. The media also contains the dye neutral, lactose and peptone. Gram Negative organisms that ferment lactose will be produce acid as an end product of the fermentation pathway. The acid lowers the pH of the media, causing a red dye to precipitate out of the media, making the colonies appear reddish. If the Gram Negative organism is unable to ferment lactose, the neutral red dye found in the media pre- incubation will appear “washed out” or almost colorless once the incubation period is complete.

Objective: Grow various microbes on MacConkey agar to demonstrate its selective and differential properties. Scientific Question: Are the organisms tested Gram negative? Do they have the ability to ferment lactose? Materials: Stock cultures of:

Escherichia coli Alcaligenes faecalis Staphylococcus simulans or carnosus Pseudomonas fluorencens 2 plates MacConkey agar/ 4 students Protocol: 1. Using a marker on the back side of the petri dish, divide the agar plate in half. 2. Streak each organism on one half of an agar plate. 3. Incubate at 37 degrees overnight. 4. Read results the following week. Results: Draw and describe growth of microorganisms, color of agar. Interpretations/Conclusions:

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Address the Scientific Question stated above.

36. Blood Cells Background: Human blood contains a variety of cell types, each performing specific roles in human health. Here are a few links describing blood cells in greater detail:

http://www.funsci.com/fun3_en/blood/blood.htm

http://en.wikipedia.org/wiki/Red_blood_cell

http://en.wikipedia.org/wiki/White_blood_cell

Listed below are the types of cells found in human blood. Red Blood Cells Function to transport oxygen from the lungs to the tissues. Red blood cells lack a nucleus and have an hour glass shape that makes them appear hollow under a light microscope. Red blood cells are the most abundant cell found in blood. Generally the cells appear red though they may take on a blue tinge depending on the staining procedures employed. Platelets Platelets function in blood clotting. The cells are much smaller than red blood cells, irregularly shaped and staining light blue. The nuclear material in platelets will stain darker blue. Monocytes Precursors to Macrophages and Dendritic Cells, Monocytes are generally larger cells that stain light blue. Monocytes may be distinguished by the cleft shape in the nucleus (although this is not always present). Lymphocytes The majority of lymphocytes found in the blood are T cells and B cells. Lymphocytes often appear smaller than red blood cells. The nucleus of lymphocytes may appear to be as large as the entire cell. Occasionally a small crescent of cytoplasm can be seen. Eosinophils Eosinophils are involved in inflammatory and allergic responses as well as cases involving infections from multicellular parasites. Eosinophils have a bilobular nucleus that stains blue, and granules in the cytoplasm that are acidic and stain red. Basophils

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Basophils participate in an allergic response. The cells possess a bi-lobed or tri-lobed nucleus that stains light blue. Often the nucleus is obscured by basic granules that stain darker blue. The granules may appear differently from one basophil to another. (During an immune response the granules will swell and migrate towards the cell surface.) Neutrophils Neutrophils are the most common leukocyte (or white blood cell). The cells have a tri-lobed nucleus that stains light blue. Often the cytoplasm appears clear though small granules may be present. Neutrophils participate in an immune response as nonspecific phagocytes. Objective: Identify the various types of blood cells found in a commercially prepared slide of human blood. Materials: Commercially prepared slide of human blood. 1/student Microscope. 1/student Protocol: View specimens under 40x and 100x objective lenses. Draw or photograph the various cell types. Results: Drawings or photographs of the various cells types.

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37. Blood Typing Background: Blood Typing is a process to differentiate blood cells by the presence or absence of antigens found on the surface of the red blood cell. Here is a link with detailed information on blood typing:

http://en.wikipedia.org/wiki/Blood_type Objective: Using a commercial kit, students will determine the ABO and Rhesus (Rh) type of their blood. Scientific Question: What is the ABO and Rh types of my blood? What types of people could use my blood in a transfusion? What types of blood could I receive in a transfusion.