Microbiology Lab Contents

Transcription

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2 Contents GENERAL DIRECTIVES... 4 GRADING SCHEME... 5 SCHEDULE... 6 LAB N O DILUTIONS AND CONCENTRATIONS... 7 PERCENTAGE... 7 MOLARITY... 9 WEIGHT/VOLUME... 9 RATIOS... 9 DILUTIONS EXERCISE 1.0: GENERATING A STANDARD CURVE AND DETERMINING AN UNKNOWN CONCENTRATION OF METHYLENE BLUE (Groups of 2) DIFFUSION OSMOSIS TONICITY OSMOLARITY OSMOLARITY VS TONICITY EXERCISE 1.1: DIFFUSION, OSMOSIS AND TONICITY IN RED BLOOD CELLS (Groups of 2) MICROBIAL GROWTH IN THE LAB MICROBIOLOGICAL MEDIA INOCULATING SOLID MEDIA: SPREADING EXERCISE 1.2: VIABLE COUNTS OF A SOIL SAMPLE (Groups of 2) EXERCISE 1.3: SAMPLING OF YOUR ENVIRONMENT (EACH PERSON)... Error! Bookmark not defined. LAB N O COLIFORMS MOLECULAR TECHNIQUE TO EVALUATE THE PRESENCE OF MICROBES EXERCISE 2.0: DETECTION OF COLIFORMS BY PCR (Groups of 2) AGAROSE GEL MIGRATION (Groups of 4) MOST PROBABLE COUNTS EXERCISE 2.1: MPN OF COLIFORMS IN YOUR ENVIRONMENT (Groups of 2) EXERCISE 2.2: BACTERIAL COUNTS OF A SOIL SAMPLE (GROUPS OF 2) EXERCISE 2.3: COUNTS OF ACTINOMYCETES IN SOIL (GROUPS OF 2) FUNGI EXERCISE 2.4: COUNTS OF FUNGI IN SOIL (Group of 2) INOCULATING SOLID MEDIA: STREAKING FOR SINGLE COLONIES EXERCISE 2.5: STREAKING FOR SINGLE COLONIES (Individually) LAB N O EXERCISE 3.0: MPN OF COLIFORMS IN YOUR ENVIRONMENT (GROUPS OF 2) DIRECT COUNTS (HEMOCYTOMETER SLIDE) EXERCISE 3.1: DIRECT COUNT OF A YEAST SUSPENSION (Groups of 2) VIEWING MICROORGANISMS MACROSCOPIC VISUALIZATION COLONY MORPHOLOGIES EXERCISE 3.2: COLONY MORPHOLOGIES (Groups of 2) MICROSCOPIC VISUALIZATION EXERCISE 3.3: MICROSCOPIC VISUALIZATION OF BACTERIA SIMPLE STAINS (Groups of 2) BACTERIAL CELL MORPHOLOGIES

3 LAB N O MICROSCOPIC VISUALIZATION - GRAM STAINING EXERCISE 4.0: GRAM STAINING (Groups of 2) MICROSCOPIC VISUALIZATION - ACID-FAST STAINING EXERCISE 4.1: ACID FAST STAINING (Groups of 2) EXERCISE 4.2: MICROSCOPIC VISUALIZATION SPORE STAINING LAB N O GROWTH OF BACTERIA GROWTH CURVE EXERCISE 5.0: E.COLI GROWTH CURVE (Groups of 2) YEAST FERMENTATION EXERCISE 5.1: YEAST FERMENTATION BIOASSAY (Groups of 2) DEATH KINETICS EXERCISE 5.2: SUSCEPTIBILITY TO AND MUTAGENESIS BY ULTRA VIOLETS (GROUPS OF 2) Error! Bookmark not defined. EXERCISE 5.3: DIFFERENTIAL STAINS AND STERILIZATION (Groups of 2) LAB N O CONTROL OF MICROBIAL GROWTH - ANTIBIOTICS KIRBY-BAUER DISC DIFFUSION METHOD EXERCISE 6.0: KIRBY-BAUER ASSAY (Groups of 2) E-TEST EXERCISE 6.1: SENSITIVITY OF S. FAECALIS TO VANCOMYCIN (Groups of 2) DETERMINING THE THERAPEUTIC DOSE EXERCISE 6.2: DETERMINING MICs (Groups of 2) LAB N O CONTROL OF MICROBIAL GROWTH (CONT D) EXERCISE 7.0: KIRBY BAUER DIFFUSION ASSAY EXERCISE 7.1: SENSITIVITY OF S. FAECALIS TO VANCOMYCIN - E-TEST (Groups of 2) BACTERIAL METABOLISM AND DIFFERENTIAL TESTS UTILIZATION OF COMPLEX CARBON SOURCES: EXOCELLULAR ENZYMES EXERCISE 7.2: DEGRADATION OF COMPLEX CARBON SOURCES (Groups of 2) SUGAR METABOLISM PHENOL RED BROTH EXERCISE 7.3: METABOLISM IN PHENOL RED BROTH (Groups of 2) GROWTH IN TSI MEDIUM (TRIPLE SUGAR IRON) EXERCISE 7.4: GROWTH IN TSI MEDIUM (Groups of 2) USE OF CITRATE AS A CARBON SOURCE EXERCISE 7.5 GROWTH ON SIMMON S CITRATE SLANT (Groups of 2) UREA METABOLISM EXERCISE 7.6: GROWTH ON UREA SLANT (Groups of 2) DECARBOXYLASES AND DEAMINASES EXERCISE 7.7: DECARBOXYLASE AND DEAMINASE ASSAYS (Groups of 2) SIM: PRODUCTION OF HYDROGEN SULFIDE, INDOLE AND MOTILITY EXERCISE 7.8: SIM TEST (Groups of 2) NITRATE AND NITRITE REDUCTION EXERCISE 7.9: NITRATE REDUCTION ASSAY (Groups of 2) ENTEROPLURI TEST: ENTEROBACTERIACEAE SYSTEM EXERCISE 7.10: ENTEROPLURI TEST (Groups of 2) THE STREPTOCOCCI AND THE STAPHYLOCCOCI BLOOD HEMOLYSIS EXERCISE 7.11: THROAT SAMPLING ON BLOOD AGAR PLATES (Groups of 2)

4 LAB N O EXERCISE 8.0: DEGRADATION OF COMPLEX CARBON SOURCES (Groups of 2) EXERCISE 8.1: METABOLISM IN PHENOL RED BROTH (Groups of 2) EXERCISE 8.2: GROWTH IN TSI (Groups of 2) GLUCOSE FERMENTATION: PRODUCTION OF MIXED ACIDS OR ACETOIN EXERCISE 8.3: METHYL RED - VOGUES-PROSKAUER TEST (MRVP) (Groups of 2) EXERCISE 8.4 GROWTH ON SIMMON S CITRATE AGAR SLANT (Groups of 2) EXERCISE 8.5: GROWTH ON UREA SLANT (Groups of 2) EXERCISE 8.6: DECARBOXYLASE AND DEAMINASE ASSAYS (Groups of 2) EXERCISE 8.7: SIM TEST (Groups of 2) EXERCISE 8.8: NITRATE REDUCTION ASSAY (Groups of 2) EXERCISE 8.9: ENTEROPLURI TEST (Groups of 2) EXERCISE 8.10: BLOOD HEMOLYSIS (Groups of 2) EXERCISE 8.11: CATALASE (Groups of 2) BILE-ESCULIN BACITRACIN, OPTOCHIN AND NOVOBIOCIN SENSITIVITY MANNITOL + SALTS AGAR TELLURITE AGAR OR BAIRD PARKER AGAR PYR TEST METRIC UNITS GROWTH MEDIA COMPOSITION

5 GENERAL DIRECTIVES 1. Attendance in lab is mandatory. Please be on time. 2. Shoes and appropriate dress must be worn at all times. Secure long hair. 3. Wear a lab coat they are easier to sterilize than your clothing, should you spill a culture or staining reagents. 4. Leave outerwear, backpacks, and any other extraneous materials in the lockers outside of the lab. 5. Be careful with Bunsen burners keep them away from microscopes, paper, ethanol, and watch your hair. Never leave the flame unattended. 6. Always place used pipettes, swabs, and other materials in the biohazard bags provided so that they can be autoclaved and disposed of properly. Do NOT throw trash in the autoclave bag. 7. No eating or drinking in lab. 8. Never lick your fingers, or put your fingers in your mouth. 9. Treat every organism as a potential pathogen. 10. Treat spilled cultures with disinfectant before cleaning them up. Cover the spill with a paper towel. Spray the paper towel with disinfectant until the towel is soaking wet. Let this sit for 10 minutes. Wearing gloves pick up the paper towels and discard in the autoclave bag. Ask the instructor or T.A. for help as soon as the spill occurs. 11. Remember to wipe the oil off the lenses before putting the microscope away. 12. No radios, MP3 players, or CD players in the lab. 13. No use of cell phones or texting in the lab. 14. Notify the T.A. or instructor of any accident, no matter how minor. 15. At the beginning of each lab period, clean your bench with disinfectant. Clean it again at the end of lab. 16. WASH YOUR HANDS before beginning the lab exercises. WASH YOUR HANDS before leaving the lab, even if it s only for a break. Material you MUST have to work in the microbiology lab: A lab coat A thin tipped permanent, preferably black, marker for labelling. A note book to record your results. Any type is acceptable. Do not waste your money. A USB key to save your pictures Optional but strongly recommended: Do not wear contact lenses in the lab. They can be quite hazardous. Notify the instructor of any safety or medical concerns so that appropriate accommodations can be taken. For example, allergies, diabetes, hypoglycemia, epilepsy, exposed wounds, color blindness, etc.. Notify the instructor of any special needs you may require so that appropriate accommodations can be taken. For example, if you write your exams with SASS. 4

6 Quiz GRADING SCHEME 2 bonus points for 100% on 4/8 quizzes Assignments 20% Midterm Exam 30% Practical Final Exam 10% Final Exam 40% Quizzes: At the beginning of each lab, a 10 minute quiz consisting of one to two questions will be given. Your performance on these cannot have a negative impact on your final grade. However, if you obtain 100% on at least 4 of the 8 quizzes you will be granted 2 bonus points on your final grade. Assignments: This lab includes 4 assignments on the theory of the experiments performed and the analysis of the results obtained. These assignments may be submitted either individually or in groups of two (you and your lab partner). A 10%/day penalty will be imposed on late assignments. (Weekends will be considered as one day) All assignments are due on the indicated date (See schedule on page 6) before you leave the lab for the day. Midterm: A midterm exam will be given during lab hours at the date specified in the schedule on page 6. The midterm will consist of 30 short answers and multiple choice questions given over a 2 hour period. You will be allowed to bring a single one sided cheat sheet (8 1/2 X 11 in.), scrap paper, and calculators. Practical final exam: For this exam, you will have to come on an individual basis for a 2.5 hour period to perform techniques commonly used during the semester. The tasks you will have to perform include a Gram stain and the identification of an unknown, a streaking for single colonies, a viable count and a direct count. This is an open book exam and you are therefore allowed any printed resource. Theoretical final exam: A final exam, which is cumulative, will consist of 40 short answer and multiple choice questions given over a 3 hour period. You will be allowed to bring a single one sided cheat sheet (8 1/2 X 11 in.), scrap paper, and calculators. 5

7 SCHEDULE Date Due dates Lab 1 Sept. 11 (sec. A) or 13 (sec. B) Lab 2 Sept. 18 (sec. A) or 20 (sec. B) Assignment 1 Lab 3 Sept. 25 (sec. A) or 27 (sec. B) Lab 4 Oct. 2 (sec. A) or 4 (sec. B) Assignment 2 Thanksgiving Oct NO LABS Midterm exam Oct. 16 (sec. A) or 18 (sec. B) Study break Oct Lab 5 Oct. 30 (sec. A) or Nov. 1 (sec. B) Lab 6 Nov. 6 (sec. A) or 8 (sec. B) Assignment 3 Lab 7 Nov. 13 (sec. A) or 15 (sec. B) Lab 8 Nov. 20 (sec. A) or 22 (sec. B) Practical exam Nov. 27 (sec.a) or 29 (sec. B) Assignment 4 Theoretical final exam Final exam period 6

8 LAB N O 1 DILUTIONS AND CONCENTRATIONS One very important property of solutions that must be addressed is concentration. Concentration generally refers to the amount of solute contained in a certain amount of solution. To deal with concentration you must keep in mind the distinctions between solute, solvent and solution. Because varying amounts of solute can be dissolved in a solution, concentration is a variable property and we often need to have a numerical way of indicating how concentrated a solution happens to be. Over the years a variety of ways have been developed for calculating and expressing the concentration of solutions. That can be done with percentages using measurements of weight (mass) or volume or both. It can also be done using measurements that more closely relate to ways that chemicals react with one another (moles). In the pages that follow, several concentration types will be presented. They include volume percent, weight percent, weight/volume percent, molarity (the workhorse of chemical concentrations), and weight/volume. You will get experience with more than one way of establishing the concentration of solutions. You can prepare a solution from scratch and measure each of the components that go into the solution. You can prepare a solution by diluting an existing solution. PERCENTAGE The use of percentages is a common way of expressing the concentration of a solution. It is a straightforward approach that refers to the amount of a component Per 100. Percentages can be calculated using volumes as well as weights, or even both together. One way of expressing concentrations, with which you might be familiar, is by volume percent. Another is by weight percent. Still another is a hybrid called weight/volume percent. Volume percent is usually used when the solution is made by mixing two liquids. For example, rubbing alcohol is generally 70% by volume isopropyl alcohol. That means that 100 ml of solution contains 70 ml of isopropyl alcohol. That also means that a liter (or 1000 ml) of this solution has 700 ml of isopropyl alcohol plus enough water to bring it up a total volume of 1 liter, or 1000 ml. Volume percent = volume of solute volume of solution x 100 7

9 Weight Percent is a way of expressing the concentration of a solution as the weight of solute/ weight of solution. Weight percent = weight of solute weight of solution x 100 As an example, let's consider a 12% by weight sodium chloride solution. Such a solution would have 12 grams of sodium chloride for every 100 grams of solution. To make such a solution, you could weigh out 12 grams of sodium chloride, and then add 88 grams of water, so that the total mass for the solution is 100 grams. Since mass is conserved, the masses of the components of the solution, the solute and the solvent, will add up to the total mass of the solution. To calculate the mass percent or weight percent of a solution, you must divide the mass of the solute by the mass of the solution (both the solute and the solvent together) and then multiply by 100 to change it into percent. Percentage weight/volume is a variation which expresses the amount of solute in grams but measures the amount of solution in milliliters. An example would be a 5 % (w/v) NaCl solution. It contains 5 g of NaCl for every 100 ml of solution. Volume percent = 12 % NaCl solution = 12 g NaCl (12 g NaCl + 88 g water) 12 g NaCl 100 g solution = 12% NaCl solution weight of solute (in g) volume of solution (in ml) x 100 This is the most common way that percentage solutions are expressed in this lab course. 8

10 MOLARITY Another way of expressing concentrations is called molarity. Molarity is the number of moles of solute dissolved in one liter of solution. The units, therefore are moles per liter, specifically it's moles of solute per liter of solution. Molarity = moles of solute liter of solution Rather than writing out moles per liter, these units are abbreviated as M. So when you see M it stands for molarity, and it means moles per liter (not just moles). You must be very careful to distinguish between moles and molarity. "Moles" measures the amount or quantity of material you have; "molarity" measures the concentration of that material. So when you're given a problem or some information that says the concentration of the solution is 0.1 M that means that it has 0.1 moles for every liter of solution; it does not mean that it is 0.1 moles. WEIGHT/VOLUME This means of expressing concentrations is very similar to that of percentages and is one of the most popular ways used by biologists. In contrast to percent, the concentration is expressed as a mass per any volume the user wishes to use. Most commonly, these concentrations are expressed per one measuring unit. For example per 1 ml, 1 µl or 1 L, etc. Essentially these expressions represent the mass of solute present in a given amount of solution. For example a solution at a concentration of 1mg/mL contains 1mg of solute in 1 ml of solution. RATIOS All the ways described above to express concentrations are done as a function of the total volume of the solution which is the volume of the solvent and that of the solute. A common method used by many microbiologists and chemists to express concentrations are ratios. In this case, the relationship between the solvent and the solute is expressed independently of one another. For example, we could say that the ratio between a solute and its solvent is 2:1. This indicates that for two parts of the solute there is one part of solvent. Thus three parts total of solution. 9

11 DILUTIONS The preparation of dilutions is essential in all fields of science as well as in everyday life. Dilutions are used to precisely reduce the concentration of elements, either chemical or alive, within a solution. For example, if you wished to reduce the concentration of fat in 3.5% milk to 0.35% you would have to perform a 10-fold dilution. To comprehend how dilutions are prepared, you must grasp the following three concepts: Concentration, dilution factor, and the dilution. The dilution factor represents the multiple by which an initial concentration must be divided by in order to obtain the desired final concentration. For example, if a solution contains 30g of caffeine per liter of solution and you wish to reduce the caffeine concentration to 0.3 g/l, then you will have to divide the initial concentration by 100, which represents the dilution factor. You can use the following formula in order to determine a dilution factor. Dilution Factor = Initial Concentration Final Concentration The dilution represents the fraction of the component being investigated. For example, in the previous problem a dilution of 1/100 was prepared. The dilution is expressed as a fraction of 1 over the dilution factor. In order to properly setup dilutions you must learn to properly use pipettes. Here are some general guidelines: Choose the pipette whose capacity is closest to the volume you wish to measure. For instance, to measure 0.1mL it is best to use a 1.0 ml pipette rather than a 10 ml pipette. Minimize the number of pipetting s done to minimize the chances of error. For instance if you wish to dispense 1 ml in ten tubes, it is best to pipette 10 ml once and dispense 1 ml ten times rather than pipetting 1 ml ten times and dispensing ten times. Change pipettes for each different solution or dilution. 10

12 Performing serial transfers: When performing serial transfers as in serial dilutions use a different pipette for each dilution. Follow the following directives: Going from a more concentrated to a less concentrated solution: Pipette the desired volume from the source and then dispense into the new tube ( A in the picture below). Rinse by pipetting up and down several times (In A in the picture below). Using a new pipette, pipette the desired volume from this tube ( A ) and dispense into the next tube ( B ). Repeat the process from A to B with the new pipette. Source A B C Dispense and rinse pipette in this tube before changing pipettes for the uptake of the desired volume from this new solution. Going from a less concentrated to a more concentrated solution: In this instance, pipetting is even easier. Just use the same pipette to transfer the desired volume from a lower concentration solution to one of higher concentration. No equilibration or rinsing is required in this case. Calculation of serial dilutions Dilutions essentially represent fractions and thus follow the same mathematical principals. That being said, the dilution (or the fraction) indicates what fraction of the total is represented by the compound being diluted. Ex. You wish to dilute a solution by a factor of 4. To do so the fraction desired is therefore 1/4; i.e. a quarter of the total volume must be represented by whatever is being diluted. Therefore, two fractions which are equal, for example 2/4 and 4/8 represent the same dilutions or dilution factors. 11

13 Preparing dilutions: The things you must determine before preparing dilutions are what final total volume you want, what is the dilution factor desired, and what the final concentration you want is (if this is known). For example, I want a final volume of 50 ml and a dilution factor of 4X. The fraction desired is thus 1/4 where the denominator represents the total. Since I want a final volume of 50 ml, 1/4 of the 50 must represent the compound being diluted; thus What this means is that 1/4 = 12.5/50. Therefore to prepare this dilution you would add 12.5 ml of the solution to be diluted in (50 ml ml = 37.5 ml) of solvent. You can use the following formula to determine the volume of the stock solution to dilute if you know the final concentration that you wish to obtain: Concentration you want Concentration you have X Final volume wanted = Volume of stock solution to be added to the mixture Serial dilutions are simply sequential dilutions where the stock solution used for each dilution represents the previous dilution. The final dilution for the series is the product of each individual dilution. Final Dil. = Dil.1 X Dil. 2 X Dil. 3 etc. 12

14 Schedule 1-2 pm Lecture 2 3:30 pm Exercise 1.0 3:30 5 pm Exercise :30 pm Exercise 1.2 EXERCISE 1.0: GENERATING A STANDARD CURVE AND DETERMINING AN UNKNOWN CONCENTRATION OF METHYLENE BLUE (Groups of 2) Materials Methylene blue solution (0.26% m/v, M.W. 320g/mole) Methylene blue solution of unknown concentration 100 ml of water Test tubes Method 1. From the stock solution of methylene blue, prepare 5 ml of a methylene blue solution at a final concentration of 0.4 mm. Label this tube N o 1. Make sure to write down how this solution was prepared. 2. Add to a test tube, labelled N o 2, 1.2 ml of water and 4.8 ml of solution N o Add to a test tube, labelled N o 3, 3.0 ml of water and 2.5 ml of solution N o Add to a test tube, labelled N o 4, 1.5 ml of water and 2.0 ml of solution N o Add to a test tube, labelled N o 5, 0.5 ml of water and 0.8 ml of solution N o Add to a test tube, labelled N o 6, 1.0 ml of water and 1.5 ml of solution N o From the stock solution of methylene blue of unknown concentration, prepare dilutions in a final volume of 5 ml of 1/4.5 and of 1/8. Label these test tubes UNK 1 and UNK 2 respectively. Make sure to write down how these dilutions were prepared. 8. Transfer 0.1 ml of each of the solutions to the wells of a 96 well plate as indicated below. 9. Obtain the absorbance readings at 550 nm. Layout of 96 well plate (one plate/table) Water Blank Soln. N o 1 Soln. N o 2 Soln. N o 3 Soln. N o 4 Soln. N o 5 Soln. N o 6 UNK 1 UNK 2 Group 1 Water Blank Soln. N o 1 Soln. N o 2 Soln. N o 3 Soln. N o 4 Soln. N o 5 Soln. N o 6 UNK 1 UNK 2 Group 2 Water Blank Soln. N o 1 Soln. N o 2 Soln. N o 3 Soln. N o 4 Soln. N o 5 Soln. N o 6 UNK 1 UNK 2 Group 3 Water Blank Soln. N o 1 Soln. N o 2 Soln. N o 3 Soln. N o 4 Soln. N o 5 Soln. N o 6 UNK 1 UNK 2 Group 4 13

15 DIFFUSION The internal environment of any organism consists mainly of water-based solutions. Many solutes may be dissolved in these solutions. Since the movement of compounds across cell membranes is strongly influenced by the differences in concentrations and by the permeability of the lipid bilayer, it is important to understand how differences in concentration of a solute influence passive membrane transport. The particles in a solution are generally free to move randomly throughout the volume of a solution. If there is a difference in the concentration of a solute between a region of a solution and another, the substance will tend to spread to the area where it is more concentrated to the location where it is less concentrated until such time that the distribution of the compound is uniform throughout the volume of the solution. Thus the net distribution occurs from a high concentration area to a low concentration area, until an equilibrium state is reached. The simple diffusion represents any non-assisted movement of any compound down the concentration gradient. In the case of cells, solutes which can readily cross the lipid bilayer (such as small uncharged molecules) are transported across the cell membrane by simple diffusion. OSMOSIS The concentration of water in a solution is inversely proportional to the solute concentration - the higher is the total concentration of solutes in a solution the smaller is the number of water molecules per unit volume of the solution. By this fact, the water will diffuse from the area with the lowest solute concentration to the area with the higher concentration of solutes. Thus, the diffusion of water in or out of the cell is driven by the differences in the total concentration of solutes that cannot cross the plasma membrane. This movement of the water by diffusion is referred to as osmosis. If water flows from one solution to another, the volume of the second solution will tend to increase, while that of the first will decrease. In the case of a cell, this change in volume will have the effect of forcing the membrane to stretch to accommodate the increase in volume, causing a pressure increase within the cell. The osmotic pressure is therefore directly related to the total concentration of non-permeable solutes of a solution. TONICITY Living cells have the potential to make or lose water by osmosis relative to the extracellular environment. The net movement of water into or out of the cell is caused by differences in the concentrations of impermeable solutes to the cell. Thus the effect that an extracellular solution has on the osmotic movement of water into or out of the cell is described by the tonicity of the extracellular fluid. For example, if the concentrations of impermeable solutes in the intracellular and extracellular fluids are the same, no net osmosis will occur. In this case the extracellular solution is said to be isotonic ("equal tonicity"). If a cell is placed in a solution with a greater concentration of impermeable solutes relative to the intracellular fluid (e.g., human blood cells in seawater), water will flow out of the cell and into the extracellular fluid, causing the cell to shrink and crenate. In this case, the extracellular fluid is said to be hypertonic ("greater tonicity"). Conversely, if a cell is placed in a solution with a concentration of impermeable solutes which is lower (e.g. distilled water) water will flow into the cell, causing it to swell to the 14

16 point where the cell may undergo lysis. In this situation, the extracellular fluid is said to be hypotonic. OSMOLARITY Osmosis is driven by differences in the relationship between solutes and solvents that exist across a semipermeable membrane. The total solute concentration of a solution (and thus the osmotic concentration) can be quantified by the osmolarity of the solution. Osmolarity is the ratio of total moles of solute particles per liter of solution. The unit for the concentration measurement is osmolality (OsM), where 1 OsM is equal to one mole total of solute particles per liter of solution. If the solution contains a single solute that cannot dissociate, such as glucose, the osmolarity of the solution is equal to the molarity of the solution. However, if the solution contains an ionic solute that can dissociate, such as NaCl, this will have a considerable influence on the osmotic concentration. E.g. NaCl readily dissociates in water into Na + and Cl-. Thus, for each mole of NaCl in a solution, there will be two moles of solute particles (1 mole of Na + and 1 mole of Cl). Thus the osmolarity of a solution containing 1 mole of NaCl represent a solution of 2 osmoles. OSMOLARITY Vs TONICITY In contrast to osmolarity, tonicity is only influenced by the concentrations of impermeable solutes. For example, a glucose solution of 300 mm, a urea solution of 300 mm, and a NaCl solution of 150 mm all have the same osmolarity; 300 mosm. But if a cell with an internal concentration of 300 mosm was placed in each of these solutions, it would behave very differently. In a 150 mm NaCl solution, the cell would be isotonic and iso-osmotic with respect to its environment. The osmotic pressure would be equal on both sides of the cell and it would maintain the same volume. However, in the case of urea, which is highly permeable through most membranes, the extracellular environment would be iso-osmotic, but hypotonic in relation to the interior of the cell. Consequently, the cell would quickly swell as a result of the rapid entry of urea and water. 15

17 EXERCISE 1.1: DIFFUSION, OSMOSIS AND TONICITY IN RED BLOOD CELLS (Groups of 2) Materials Sheep s blood- 3 ml 50% (v/v) Glycerol (MW: 92g/mole) 1 M NaCl (MW: 58g/mole) 50% (m/v) Sucrose (MW: 342g/mole) 100 ml of water Test tubes Method 1. Prepare three series of 6 test tubes; one series for each solute. 2. For each series, prepare from the stock solutions 2 ml solutions representing the following solute concentrations: Series 1: Sucrose: 0.32 M, 0.25 M, 0.15M, 0.08M, 0.04M and 0.0M Series 2: NaCl: 0.16 M, 0.12 M, 0.08M, 0.04M, 0.02M and 0.0M Series 3: Glycerol : 0.5 M, 0.4 M, 0.3 M, 0.2 M, 0.1 M and 0.0M 3. Add to each test tube, 0.1 ml of sheep s blood and mix. 4. Prepare a positive control by adding 2 ml of water to 0.1mL of sheep s blood and mix. 5. Wait for 15 to 20 minutes. 6. Mix well each tube and transfer 1 ml to labelled microcentrifuge tubes. 7. Centrifuge for 30 seconds à maximum speed in a microcentrifuge 8. Transfer 0.2 ml from each tube to the wells of a 96 well plate. 9. Have your plate read at a wavelength of 540 nm. Calculate the percentage of hemolysis under each condition: % hemolysis = 100 X [(Absorbance of sample) (Absorbance of positive control)] 16

18 MICROBIAL GROWTH IN THE LAB In their natural setting, not only are the number of microorganisms relatively low, but in addition several different species live together. For example, the tips of your fingers are probably covered by at least ten different species of bacteria in relatively low numbers (between a few hundred to several thousands). In addition, optimal conditions required for the growth of a given species (for example its nutritional requirements, temperature, ph, etc.) are very diverse. By keeping in mind these characteristics; microbiologists have developed several different strategies to enable the study of microorganisms. MICROBIOLOGICAL MEDIA It is often impractical to study a microorganism in its natural environment. For instance, if one wanted to investigate the effects of newly developed antibiotics on a given bacterial pathogen it would be unethical to do this on humans themselves. Furthermore, since bacterial populations are heterogeneous, the effect on a single isolated species would be difficult to interpret. Consequently, methods are necessary to grow bacteria in culture in a laboratory environment. This is achieved by using a variety of different media, which can be synthetic or non-synthetic in nature. Growth media are available in liquid, semi-solid, or solid form. Semi-solid and solid media are obtained by the addition of different concentrations of a solidifying agent. The most common solidifying agent used in microbiology is agar; a polysaccharide derived from seaweed. Agar possesses several advantageous characteristics. 1) It can be easily liquefied by boiling, and can be maintained in its molten form at temperatures as low as 45 o C. 2) Agar solidifies at temperatures below 45 degrees and finally 3) most bacteria do not digest agar. Another solidifying agent that is less commonly used is gelatin. Its use is less common since many bacteria can digest it. Liquid culture media are usually referred to as broths. Solid media can be in the form of plates or slants. All media must include the necessary nutrients required for microbial growth. Other conditions such as temperature and the presence or absence of oxygen are controlled by other pieces of equipment such as an incubator. These conditions allow the microbiologist to grow and maintain large numbers of bacteria, which are necessary for experimental studies. 17

19 INOCULATING SOLID MEDIA: SPREADING In contrast to streaking, spreading is only used with liquids. It allows spreading the bacteria evenly over the surface of the plate. However, in contrast to streaking it does not allow any dilutions to be performed on the plate itself. Consequently, if dilutions are required, these must be prepared independently before. The instrument used is a glass spreader commonly referred to as a hockey stick. As with the loop, the spreader must be sterilized before each use. o STERILIZING THE SPREADER In order to sterilize the spreader, it is dipped into ethanol, ignited, and the ethanol is allowed to burn off. Do not hold the spreader in the flame as it will get to hot! The spreader is then allowed to cool and used to spread the sample of bacteria onto the surface of the plate. Ethanol Dip spreader in ethanol Ignite ethanol Allow ethanol to burn off Spread bacteria 18

20 EXERCISE 1.2: VIABLE COUNTS OF A SOIL SAMPLE (Groups of 2) Materials 1g of soil Flask with 100 ml of sterile water 100 ml sterile water 3 sterile test tubes 3 TSA plates 3 plates of Sabouraud Dextrose Agar containing 100 µg/ml chloramphenicol 3 glycerol agar plates with yeast extract Method 1. Transfer 1g of soil to the flask containing 100 ml of sterile water. (this represents a 10-2 dilution) Shake on shaking platform for 10 minutes. 2. Allow the soil to settle for 10 minutes. 3. Prepare 10-3, 10-4 and 10-5 dilutions of the soil suspension in a final volume of 10 ml. (see figure below) 4. Plate 0.1 ml from each of the three highest dilutions on 3 appropriately labelled TSA plates. 5. Plate 0.1 ml from each of the three highest dilutions on 3 appropriately labelled Sabouraud Dextrose Agar plates. 6. Plate 0.1 ml from each of the three highest dilutions on 3 appropriately labelled glycerol agar plates with yeast extract. 7. Incubate the inverted plates at room temperature until next week. 1g of soil ml Water Dilution 0.1ml 0.1ml 0.1ml 19

21 LAB N O 2 Schedule 1 2 pm Exercise 2.0: Preparation of PCR reactions 2 3 pm Lecture 3 4 pm Exercise :30 pm Exercise 2.0: Preparation of agarose gel 4 :30 5 :30 pm Exercise :30 6 pm Exercise 2.0: Loading and migration of agarose gels 6 6:30 pm Exercise 2.5 COLIFORMS Total coliforms are small rods of the Enterobacteriacae family that ferment lactose at 37 o C with formation of acid and gas in 48 hours. These include both bacteria native to the mammalian intestine or the environment such as Escherichia coli, Klebsiella sp., Enterobacter sp., Citrobacter sp., Serratia sp., Shigella sp., and Proteus sp. Fecal coliforms (Escherichia coli): This is the only coliform that is not normally found outside of the intestinal environment. Its presence is therefore an excellent indication of fecal contamination. However, because of its relatively low viability outside of its natural environment, a negative test is not necessarily indicative of the absence of fecal contamination. MOLECULAR TECHNIQUE TO EVALUATE THE PRESENCE OF MICROBES An alternative to viable counts consists of using molecular techniques to verify for the presence of microorganisms. One of these techniques is PCR. EXERCISE 2.0: DETECTION OF COLIFORMS BY PCR (Groups of 2) Materials: PCR Cocktail: Taq buffer (2X), 5.0mM MgSO 4, 400µM dntp, 4µM of each primer against LacZ (Keep on ice) 10-5 dilution in saline of overnight culture of E.coli Thin walled PCR tubes Microcentrifuge tubes Saline Sterile water Primers : LacZfor : GTTGGCAATTTAACCGCCAGTCAGG LacZrev : CTGTAGCGGCTGATGTTGAACTGGA 20

22 Method (PCR) 1. Prepare using sterile saline as a diluent 10-1 and 10-3 dilutions of the E.coli suspension provided. 2. Transfer 1 ml from each of your samples (undiluted, 10-1 and 10-3 samples) to a labelled microcentrifuge tube. Make sure to keep the remainder of your samples for future experiments. 3. Centrifuge the tubes in the microcentrifuge at maximum speed for 5 minutes. 4. Pour out the supernatant and then add 50µL of sterile water. 5. Mix vigorously on the vortex for 20 sec.. 6. Boil for 5 minutes and then place on ice for 1 minute. 7. Centrifuge a maximum speed for 5 minutes. You will use the supernatant for your PCR analysis. 8. Transfer 25µL from each of your samples to labelled PCR tubes. 9. Add 25µL of the PCR cocktail to each PCR tube. 10. Bring your PCR tubes to the front. The teaching assistants will add 0.5 µl of Taq polymerase to each tube and then perform the PCR. 11. Your reactions will be returned to you once the PCR is finished. PCR amplification conditions: 1 cycle of 5min, 94 o C for denaturation. 35 cycles of 30sec, 94 o C denaturation; 1 minute at 68 o C annealing and extension. 1 cycle of 5min, 72 o C. Cool down at 4 o C, indefinitely. AGAROSE GEL MIGRATION (Groups of 4) Agarose gel electrophoresis involves the separation of DNA molecules based on their size and conformation. The electrophoretic migration of a DNA fragment through agarose is inversely proportional to the logarithm of its molecular weight. After the electrophoresis the gels will be viewed under ultraviolet light and a digital picture will be taken. The intensity of fluorescence is proportional to the amount (and length) of linear DNA. Ethidium bromide, which is used in staining agarose gels to visualize DNA under ultraviolet light, is a potential carcinogen; so always wear gloves when handling anything containing it. The UV light source is also extremely hazardous to skin and particularly your eyes, so be sure to use proper protection (gloves, lab coat) when viewing your agarose gels. Materials Electrophoresis apparatus Agarose 5X loading buffer 10X TBE buffer PCR reactions 21

23 Method (Electrophoresis) 1. Prepare 200mL TBE buffer at a final concentration of 1X in your graduated cylinder. Mix well. 2. Mix the appropriate amount of agarose to obtain a final concentration of 1.0% m/v) in 25 ml of 1X TBE. 3. To dissolve the agarose, microwave (25-45 sec.). Loosely cover the mouth of the flask with plastic to prevent evaporation. Once the agarose is totally dissolved, allow the flask to cool to o C (5 minutes or so). 4. Add 50 L of a stock solution of ethidium bromide at 1mg/mL (CAUTION! CARCINOGEN!) and mix well. 5. Pour into the gel tray. After pouring the gel, place the 8 well comb (Well capacity approx. 14 L) and remove any air bubbles (e.g. with a small tip). Allow to solidify at least minutes. 6. Once the gel has solidified, remove the dams. Pour a sufficient amount of 1X TBE to cover the gel by approx. 0.5cm. 7. Carefully remove the comb. 8. Obtain your PCR reactions. Add 10 µl of the DNA loading dye to each reaction. 9. Load 10μL of each reaction. 10. Carry out the electrophoresis at 100V. 11. After the electrophoresis, examine your gel under the UV light and have a picture taken for your analysis. MOST PROBABLE COUNTS A variation of viable counts is based on probabilities to determine the number of bacteria in a sample. As with viable counts, this method requires the growth in an appropriate medium. However, in contrast to viable counts, detection is based on the presence or absence of growth or on the production of a by-product. To understand the theory behind the most probable counts (MPN), think about 10 fold serial dilutions with 1mL samples from each dilution inoculated in different tubes containing a given growth medium. Following the incubation, the broths are examined for the presence or the absence of growth. In theory, if a least one organism was present in any of the inoculums visible growth should be observed for that tube. If the broth inoculated from the 10-3 dilution shows growth, but the broth inoculated from the 10-4 dilution does not, it is thus possible to affirm that there were more than 1X10 3 organisms per ml of sample, but less than 1 X 10 4 per ml. Bacteria are only rarely, if ever, evenly distributed within a sample. For example, if a 10 ml sample contains a total of 300 organisms, not all 1 ml aliquots will contain 30 organisms; some will have more or less than 30 organisms, but on average all ten aliquots in the whole 10 ml sample will be 30. This holds true for any of the dilutions from which an inoculum is taken. To increase the statistical accuracy of this type of test, more than one broth is inoculated for each dilution. The standard MPN makes use of a minimum of three dilutions and 3, 5 or 10 tubes per dilution. Following the incubation, the pattern of positive and negative tubes is recorded after which a table of standard MPN is consulted in order to determine the most probable number of organisms (which cause the positive results) Per unit volume of the original sample. In the following example, sets of three tubes of broth are inoculated with 1 ml from each of the 10 fold dilutions of a soil suspension at 1 g/100 ml. 22

24 1g of soil ml water Dilution 10-2 Inoculum: 1mL of mL of mL of mL of mL of 10-6 Results: Following the incubation, the number of tubes which show growth is recorded and expressed as the number of positive tubes over the total number of tubes for that dilution. For example, for the 10-3 dilution the result would be expressed as 2/3. At a certain point the dilution will be so high that no organism is found within the inoculums in any of the tubes for that dilution. In this case the result would be expressed as 0/3. MPN determination When more than three dilutions are used in a decimal series of dilutions, use the results from only three of these to determine the MPN. To select the three dilutions to be used in determining the MPN index, determine the highest dilution (most dilute sample) that gives positive results in all three samples tested (so 3/3) and for which there are no lower dilution (less dilute sample) giving any negative results. Use the results for this dilution set and the two next succeeding higher dilutions to determine the MPN index from the MPN table. (See examples a and b below) If none of the dilutions yield all positive tubes, then select the three lowest dilutions for which the middle dilution contains the most positive results, as shown in example c and d. If after following these rules, there is a series showing positive results in higher dilutions than the chosen three, add the result to the highest dilution as in example e. Example Combination of positives MPN index/ml a 3/3 3/3 2/3 0/ b 3/3 2/3 1/3 0/ c 0/3 1/3 0/3 0/ d 1/3 1/3 2/3 0/ e 1/3 2/3 0/3 1/ Once you ve obtained the MPN index, multiply it by the dilution factor of the middle set of dilutions. For instance in example a you would get 9.3/10-2 = 9.3 X

25 Pos. tubes MPN/g Pos. tubes MPN/g (ml) (ml) < >

26 EXERCISE 2.1: MPN OF COLIFORMS IN YOUR ENVIRONMENT (Groups of 2) Principle: The presumptive test is specific for detection of coliform bacteria. Measured aliquots of the water to be tested are added to a lactose fermentation broth containing an inverted gas vial (Durham tubes). Because these bacteria are capable of using lactose as a carbon source (other enteric organisms are not), their detection is facilitated by use of this medium. Tubes of this lactose medium are inoculated with 10-mL, 1-mL, and 0.1-mL aliquots of the water sample. The series consists of at least three groups, each composed of three tubes of the specified medium. The tubes in each group are then inoculated with the designated volume of the water sample. Development of gas in any of the tubes is presumptive evidence of the presence of coliform bacteria in the sample. The presumptive test also enables the microbiologist to obtain some idea of the number of organisms present by means of the most probable number test (MPN). The MPN is estimated by determining the number of tubes in each group that show gas following the incubation period. Lauryl-trypotose broth. This medium contains peptones as a carbon and nitrogen source as well as other nutrients. In addition this medium contains lactose as a fermentable carbohydrate and sodium lauryl sulfate which inhibits the growth of organisms other than coliforms. An inverted vial is included to detect the accumulation of gas. A positive test, gas accumulation following a 48 hour incubation period at 37 o C indicates the presumptive presence of coliforms. Materials Undiluted and diluted E.coli suspensions 3 sterile tubes 9 X 5 ml Lauryl-tryptose broths Sterile water Method 1. Perform an MPN count as indicated below in 3 sets of 3 tubes containing 5 ml of Lauryltryptose broth. 2. Incubate at 37 degrees. These will be returned to you next week. Sample Volume of inoculum Vol. of medium 3 tubes Undiluted (provided) 1.0 ml 5 ml ml 5 ml ml 5 ml 25

27 EXERCISE 2.2: BACTERIAL COUNTS OF A SOIL SAMPLE (Groups of 2) Last week you performed viable counts of a soil sample. Among the various microorganisms in the soil, bacteria are the most numerous organisms that can be cultured (viruses are more numerous, but are difficult to grow, because they require a suitable host). The predominant genera are Arthrobacter, Bacillus, Pseudomonas, and Streptomyces. Arthrobacter and Streptomyces are actinomycetes that produce cells similar to molds. Note that the results you get will be dependent on the medium used. The media for the isolation of bacteria are usually not ideal for mold growth, while those used for molds often have antibiotics to inhibit bacterial growth. Materials Viable counts of soil on TSA plates from last week Method 1. Obtain your TSA plates on which you spread the different dilutions of the soil sample. 2. Count the number of CFUs observed for each of the dilutions. Record these counts in your lab note book. EXERCISE 2.3: COUNTS OF ACTINOMYCETES IN SOIL (Groups of 2) Materials Viable counts of soil on glycerol with yeast extract agar from last week Method 1. Obtain your glycerol with yeast extract agar plates on which you spread the different dilutions of the soil sample. 2. Count the number of CFUs observed for each of the dilutions. Record these counts in your lab note book. 26

28 FUNGI Fungi are eukaryotic organism and they are classified into two main groups; yeasts and molds. These groups can easily be discriminated based on the macroscopic appearance of the colonies formed. The yeasts produce moist, creamy, opaque or pasty colonies, while molds produce fluffy, cottony, woolly or powdery colonies. These microorganisms are useful as well as harmful to human beings. Useful because they produce many antibiotics, natural products and are used in industrial fermentation processes. They may be harmful since they may cause human diseases, produce toxic substances as well as harm important crops. It is therefore very important to study fungal species. The branch of science that deals with study of fungal species is called Mycology. In contrast to bacterial counts, viable counts of fungi are estimated from plates containing between CFUs. EXERCISE 2.4: COUNTS OF FUNGI IN SOIL (Group of 2) Materials Viable counts on Sabouraud dextrose agar from last week Method 1. Obtain your Sabouraud dextrose agar plates on which you spread the different dilutions of the soil sample. 2. Count the number of CFUs observed for each of the dilutions. Record these counts in your lab note book. 27

29 INOCULATING SOLID MEDIA: STREAKING FOR SINGLE COLONIES Streaking for single colonies is simply a more elaborate method of streaking which is used to generate pure cultures in which only one organism can be found. Several different methods exist to generate pure cultures. In general, all of these methods rely on the principal of dilutions to isolate the desired organism from all others. Let us take for example a population of millions of individuals from which you want to identify and pick out a specific individual. If one could dilute the population such that in any given area there were only a couple of individuals, it would then be easy to pick out the individual you are interested in. As mentioned above, diluting a heterogeneous starting culture to such a point usually generates pure cultures. Streaking for single colonies achieves this. Streaking can be done from a solid or a liquid medium, whereas spreading is always performed from a liquid medium. The instrument used in this case is an inoculation loop, which is basically a metal wire used to pick up and deliver the bacteria. Do not confuse this instrument with the inoculation needle, which has a straight end rather than a loop. Before using the loop, it must be sterilized. To do this, place the loop in the flame of the Bunsen burner until it turns red. Allow the loop to cool for a minute or so before using it. DO NOT PUT IT ON YOUR BENCH. Once it has cooled down, grab some bacteria from the plate culture supplied. Be careful not to pick up too much. 28

30 The procedure is essentially as follows. Initially bacteria are picked up from a broth or solid culture with a sterile loop. The bacteria are then streaked on an area of the plate, essentially diluting it. The loop is then sterilized once again and used to streak a new area of the plate by picking up bacteria from the initial streak, thus diluting it even more. (See figure below) Note: You must flame the loop between each streaking and you must not go back to the source. 29

31 EXERCISE 2.5: STREAKING FOR SINGLE COLONIES (Individually) Materials Plates of soil viable counts 2 TSA plates Method 1. Each person will perform single colony isolations from a bacterial colony and an actinomycetes colony. 2. Using a sterile inoculation loop, pick up a small sample of a bacterial colony from your soil viable counts and streak a TSA plate as illustrated in the first panel of the figure on the previous page. STERILIZE YOUR LOOP after this initial streaking. 3. Make a second set of streaks with the sterile loop as shown. 4. Continue as many times as possible, sterilizing the loop each time between streaks to isolate single colonies. 5. On a new plate, repeat the procedure for single colony isolation from one of the colonies obtained on the actinomycetes plates. 6. Place both single colony isolations in the designated area so that they can be incubated at room temperature. 30

32 1 2 pm Lecture 2 2:30 pm Exercise :30 3 pm Exercise :30 pm Exercise :30 5 pm Exercise 3.3 LAB N O 3 Schedule EXERCISE 3.0: MPN OF COLIFORMS IN YOUR ENVIRONMENT (Groups of 2) Materials MPN broths from last week Method 1. Examine your broths for the presence or absence of growth and gas. 2. Fill out the following table to indicate your results. Dilution Growth : Tube

33 DIRECT COUNTS (hemocytometer slide) As the name implies, direct counts involves taking a direct measurement of the actual number of microorganisms present within a sample without a priori growing them. This can be achieved by different visualization techniques two of which you will examine in the following exercises. Note that a direct count does not distinguish between whether a microorganism is alive or dead. One method involves using a special slide, a hemocytometer slide, which possesses a counting chamber of a fixed and known volume. A standard hemocytometer slide has two identical counting areas consisting of nine 1mm 2 etched squares (1mm X 1mm; see figure). When the counting chamber is overlaid with a coverslip, the free space available is 0.1mm deep. The volume of each square is thus 0.1mm 3 or 10-4 cm 3. After applying an aliquot of the sample to be counted, the cells are visualized and enumerated. The cells within each of three squares are counted. The average is then calculated and the total number of cells within the original sample is interpolated. Example of calculation: If 10, 14, and 6 bacterial cells were counted in each of three independent squares; the average number of bacteria per square is 10. This number is equivalent to having 10 bacteria/ 0.1 mm 3, or 10 bacteria/ cm 3 or 10 bacteria/ ml. Therefore, the concentration of bacteria in the sample being examined is 1 X 10 5 /ml. Hemocytometer Y Y 32

34 EXERCISE 3.1: DIRECT COUNT OF A YEAST SUSPENSION (Groups of 2) Materials Yeast suspension Hemocytometer slide Sterile water Tubes Pasteur pipettes/micropipette Method 1. Prepare the following dilutions of the yeast suspension: 10-1, 10-2, and Make sure to thoroughly mix the yeast suspension before sampling. 2. Fill the hemocytometer counting chamber with a sample from the highest dilution (see image below or ask a teaching assistant to show you). Make sure to thoroughly mix the yeast suspension before sampling. 3. Count the number of cells observed in each of three squares of the size indicated by a Y in the above image. 4. If the number of cells is too low, start over with the previous dilution. If the number of cells is too high, prepare a 10 fold higher dilution and start over. 5. Record the following information in your lab note books: Number of cells counted per square, dimensions of the chosen squares, dilution used. 33

35 VIEWING MICROORGANISMS Microbiology is the study of very small organisms, microorganisms that cannot be seen with the naked eye. In order to observe and study their morphological characteristics, it is necessary to examine clusters of cells - macroscopic viewing of colonies or visualizing individual cells using a microscope. There are several groups of organisms that fall into this category, including bacteria, algae, fungi and protists. In this group, there are several species of human interest because of their ability to cause disease or their use in the food industry. The emphasis during the semester will be on bacteria. MACROSCOPIC VISUALIZATION COLONY MORPHOLOGIES Preliminary identification of microorganisms can be based on colonial morphology. Each single colony represents a population of bacterial cells originating from a single cell that, after multiple rounds of cell division, generated a colony of cells stacked on top of each other in a characteristic shape according to the bacterial type (just as a cluster of bananas has a different appearance of a heap of potatoes). The morphology of a bacterial colony is a function of several factors such as the shape of the cell, its size and physiology. It is also important to note that the colonies of a particular bacterial type often have colors, textures and distinct odors. Refer to the figure below to determine the morphologies of the observed colonies. EXERCISE 3.2: COLONY MORPHOLOGIES (Groups of 2) Materials Viable counts on TSA and glycerol + yeast extract agar plates from last week Method 1. Obtain your TSA and glycerol + yeast extract agar plates from last week on which you spread different dilutions of a soil. 2. Examine the different colony morphologies observed. 3. Record in your lab note book the different numbers of bacteria with distinct morphologies observed. 34

36 MICROSCOPIC VISUALIZATION The light microscopes used in this lab are binocular and have ocular lenses with a magnification of 10X. In addition to this magnification there are also four different objective lenses to choose from 4X, 10X, 40X, and 100X. Magnification of the object being viewed is the product of the ocular objective multiplied by the lens objective currently in use. For instance when viewing an object on the 4X objective lens, the object is magnified a total of 40X. The highest objective for our microscopes is 100X, which has a magnification of 1000X. With this objective, it is necessary to place a drop of oil on top of the slide and immerse the lens. The oil immersion lens (100X) is specially sealed and IS THE ONLY LENS THAT SHOULD BE PLACED IN OIL! The importance of proper handling and use of the microscope is vital. It is critical that you clean the microscopes before and after use. Use a new KimWipe to gently wipe the ocular lenses and then wipe the 10x, 40x, and 100x objective lenses. If there is any excess oil on the microscope be sure and remove that. If you have trouble removing oil, use the microscope cleaner provided. Viewing a specimen 1. Use the coarse adjustment knob (1) to move the stage to its lowest level. 2. Clean all objective lenses. 3. Before use, clean all slides, top and bottom, with KimWipes. 4. Adjust the condenser (2) to its highest level. Turn on the lamp. 5. Rotate the objectives until the 10X objective clicks into place. 6. Place the slide on the stage so that it is held within the slide holder clamping device. The slide must lie flat on the stage. Using the mechanical stage knobs (3), position the slide so that the specimen is in the exact center of the light coming through the condenser. 7. While looking through the eyepieces (NOT AT THE COMPUTER SCREEN), adjust the width between the eyepieces until a single, circular field is seen simultaneously with both eyes. 8. Light intensity is an essential aspect of microscopy. For optimal viewing, the light must be adjusted at each magnification. Always adjust the light while looking through the eyepieces. a. Initially adjust the light intensity (4) at a low/medium setting. The entire viewing area (field) must be filled with light. The lighted area may become smaller while focusing or changing magnifications. b. Locate the thin, black iris diaphragm (5) lever under the stage. Adjust this lever to a medium/low light level. The iris diaphragm will need to be adjusted as magnifications increase. 9. Under low power (10X), SLOWLY focus with the coarse adjustment knob until the specimen comes into view. Adjust the light as required. 10. Focus the image with the fine adjustment knob and by adjusting the light. 11. Before switching to the next objective, move the slide so that the desired specimen is located in the center of the field. 35

37 Using immersion oil at 100X Immersion oil is used with the 100X objective because it increases the resolution. The oil should come in contact with both the lens of the 100X objective and the slide. 1. Be sure that the specimen is in the EXACT CENTER of the viewing field under the 40X objective. 2. Rotate the 40X objective away from the slide but do not yet click the 100X objective into place. 3. Put a small drop of immersion oil on the slide directly over the light. 4. Rotate the nosepiece until the 100X oil immersion objective is clicked into place. 5. DO NOT USE THE COARSE ADJUSTMENT KNOB WHEN FOCUSING UNDER THE 100X OBJECTIVE! ONLY USE THE FINE ADJUSTMENT KNOB! 6. Adjust the light for optimal viewing Taken from:

38 Common Problems The field is dark. Is the light on? Is the objective securely clicked into place? Is the diaphragm open? Is the slide lying flat on the stage? You are not sure if you are looking at dirt on the objective lens or the specimen. Use the mechanical stage knobs to move the slide slightly while looking through the eyepieces. If what you are looking at does not move, it is probably dust or dirt on the objective. If it does move, it is on the slide. Rotate the ocular gently between your fingers. If what you are looking at rotates, it is probably dirt on the ocular. You cannot find the specimen. Is the specimen directly over the light? Is the slide secure and flat in the mechanical stage? Did you start with a low power objective and focus on the lower objectives first? Have you adjusted the light? Are you moving the adjustment knobs too quickly? Work slowly so you do not miss the specimen. Remember, bacteria look like specks at low magnifications. You are having trouble focusing. Always start on a low power objective, and focus here first. Focus SLOWLY! It is very easy to move past the specimen if the adjustment knobs are moved too quickly. Be sure to look in the ocular (NOT THE COMPUTER SCREEN) while you are focusing with the adjustment knobs or changing the light intensity. Adjust the light. You lose the specimen when switching from the 40X objective to the oil immersion objective. Was the specimen in the exact center of the field before switching to the 100X objective? Is the 100X objective lens clean? Have you adjusted the light? Have you refined the image with the fine adjustment? 37

39 EXERCISE 3.3: MICROSCOPIC VISUALIZATION OF BACTERIA SIMPLE STAINS (Groups of 2) Microbial cells are colorless and thus very difficult to visualize even with a microscope. Consequently, various staining procedures have been developed to both visualize and classify microbes. Staining procedures are generally classified into two categories; either simple staining or differential staining. Simple staining involves the use of a single stain such as methylene blue which stains all cells the same color. This type of stain is useful to look at the different shapes and aggregations of bacterial cells. (See figure on page 42) Materials Streaking for single colonies from last week Microscope slides Stains Method 1. Prepare smears from the bacterial and actinomycetes colonies. 2. Use a sterile loop to transfer a drop of water onto a microscope slide. Then use the sterile loop once again to transfer some bacteria from a chosen colony (VERY LITTLE) to the drop of water on the microscope slide. 3. Suspend the bacteria in the drop of water and spread over a small area of the slide. 4. Allow your smears to completely air dry on your work bench. Too little OK Too thick 5. Heat fix your bacterial smears by rapidly passing the slides through the flame of the Bunsen burner. Do not over heat!! Burnt bacteria do not stain well. 38

40 6. Setup your staining station as follows: Absorbent paper mat (Plastic coating under) Staining Container (Gladware) 7. Deposit the slides over the slits of the Gladware. Add one drop of methylene blue. 8. Leave the stain on the smears for approximately 30 seconds. 9. Wash off the excess dye with distilled water. 10. Blot the slides dry by pressing them between the layers of absorbent paper. 11. Examine under the microscope and obtain digital pictures. Save these. Dispose of the staining solutions in the drums provided to that effect. DO NOT THROW STAINS DOWN THE DRAINS 39

41 BACTERIAL CELL MORPHOLOGIES Neisseria Micrococci Micrococci 40

42 LAB N O pm Lecture 2 3 pm Exercise pm Exercise :15 pm Exercise 4.2 Schedule MICROSCOPIC VISUALIZATION - GRAM STAINING Previously, you performed a simple staining procedure, which allowed you to compare microbial cell sizes and shape. Several other staining procedures have been developed which are said to be differential. These procedures stain microbes differently as a function of different cellular properties. Dr. Christian Gram developed a differential staining technique, the Gram stain. Gram staining allows not only to observe cell shape and size, but also to classify bacteria in one of two groups: Gram positive or Gram negative. The Gram stain technique makes use of four chemical components: Crystal violet, a primary dye that stains all cells blue indiscriminately, Gram s iodine, which acts as a mordant that interacts with the primary stain, Ethanol, whose which dehydrates the cell wall and Safranin, a counter stain that stains all cells indiscriminately red. The sequential addition of these 4 components will cause some bacterial species to be stained red (Gram negative), whereas others will be stained blue (Gram positive), according to their species. The differential property of Gram staining is a function of their different cell wall composition. Typically, Gram-positive bacteria possess a very thick cell wall composed of peptidoglycan layers. In Contrast, Gram-negative organisms have relatively thin cell walls consisting of one-three peptidoglycan layers covered by a lipid layer. The critical step in the Gram staining technique, which confers its discriminatory properties, is the ethanol wash. In the case of Gram-positive organisms, the cell walls are very rapidly dehydrated due to the lack of lipids. Consequently, these cells tend to retain the crystal violet-gram s iodine complex. In contrast, the cell wall of Gram-negative organisms is not easily dehydrated by the ethanol wash due to its high lipid content and thus retains its high permeability. Consequently, the ethanol wash effectively leaches any of the crystal violet-gram s iodine complexes, and the cells remain susceptible to safranin staining. These cells will thus appear red. 41

43 EXERCISE 4.0: GRAM STAINING (Groups of 2) Materials Microscope slides Broth cultures of S. aureus and E. coli Stains Method 1. Prepare heat fixed bacterial smears of the broth cultures supplied. 2. Stain with crystal violet for one minute. 3. Carefully wash of the stain with distilled water. 4. Apply Gram s iodine for one minute 5. Carefully wash of the Gram s iodine with distilled water. 6. Carefully wash with 95% ethanol by adding it one drop at a time until the alcohol runs clear. 7. Wash the excess ethanol with distilled water 8. Counterstain with safranin for 45 seconds. 9. Wash off with distilled water 10. Blot dry with bilbous paper. 11. Examine under the microscope and take digital pictures. Save these. MICROSCOPIC VISUALIZATION - ACID-FAST STAINING Another specialized staining procedure is the Acid-Fast stain. This staining technique is particularly useful to colorize bacteria, which possess wax-like constituents in their cell wall. These include many human pathogens such as the Mycobacteriacae, some members of which cause tuberculosis and leprosy. The high content of mycolic acid, a wax like compound, of the cell wall of bacteria within this family makes them literally impermeable to stains. However, once a stain has penetrated, it cannot be easily removed by decolorizing agents such as ethanol. The principal of the acid-fast technique is thus to render the bacteria permeable to the stain and to then verify resistance to harsh decolorizing conditions. These objectives are reached by using carbol fuchsin as the primary stain. Carbol fuchsin is a highly lipid soluble compound and thus easily permeates the waxy layer of Mycobacteriacae. These bacteria then resist decolonization by the harsh acid wash treatment done with acid alcohol. 42

44 EXERCISE 4.1: ACID FAST STAINING (Groups of 2) Materials Microscope slides Slant cultures of B. subtilis and M. smegmantis Carbol Fuschin of Kinyoun Acid alcohol Method 1. Prepare heat fixed bacterial smears of each of the cultures. 2. Flood each of the smears with carbol fuchsine for 5 minutes. 3. Rinse with distilled water. 4. Destain with acid alcohol until no more stain runs off. 5. Rinse with distilled water. 6. Counter stain with methylene blue for 30 seconds. 7. Destain with distilled water. 8. Blot dry the slide with blotting paper. 9. Examine under the microscope and take digital pictures. Save these. Exercise 4.2: MICROSCOPIC VISUALIZATION SPORE STAINING Bacterial cells belonging to both the Bacillus and the Clostridium genera can assume one of two states: A vegetative cell, which is metabolically active, or that of a spore, which is metabolically inactive. The spore represents a dormant state of the cell that is highly resistant to several different harsh conditions such as heat and dehydration. When the environmental conditions become unfavorable for continued growth, vegetative cells initiate sporogenesis, which gives rise to a novel intracellular structure, the endospore. Just like the seeds of plants, the endospore is composed of several layers, which confer to it a high level of resistance. Eventually, the endospore is released as a spore that is independent of the vegetative cell. When the environmental conditions become favorable for growth, the spores germinate and return to their vegetative state. The resilient nature of both the spore and the endospore make them resistant to standard staining techniques and thus require specialized staining procedures to stain them. Briefly, spore staining is accomplished as follows. A bacterial smear is initially exposed to a primary stain, malachite green. Due the high level of impermeability of the spores, the stain is simultaneously exposed to heat, to increase the permeability to the stain. At this stage, both the spores and the vegetative cells are stained green. The smear is subsequently thoroughly washed with water to remove the excess of the stain. Since malachite green has relatively little affinity for structures of the cell, the vegetative cell is decolorized by this treatment whereas the spores retain the stain. Finally, a counterstain is applied, safranin, which colors the cells red whereas it does not stain the spore. See the demonstration slide 43

45 PowerPoint Presentation Each group must prepare a PowerPoint presentation of their images. The first slide must include a title, the names of the group members, the group number and the date. The presentation must include the following images (one per slide): A Gram stain of each of the broth culture supplied including the following information. (2 images) o Bacterial genus and species o Type of staining used o Cell morphology o Cell aggregation o Magnification An acid fast stain of Mycobacterium smegmantis and Bacillus subtilis including the following information (2 images) o Bacterial genus and species o Type of staining used o Cell morphology o Cell aggregation o Magnification 44

46 LAB N O Lecture 2 6 Exercise :30 5 :30 Exercise :30 4 Exercise 5.2 Schedule GROWTH OF BACTERIA GROWTH CURVE Microbial growth is generally defined as the increase in cell number. A growth curve represents growth as a function of time. Growth can be divided into four different stages: the lag phase, exponential or log phase, the stationary phase and the death or the phase of decline. When an inoculum is transferred to a new environment, the lag phase occurs first. This is the time required by the microbial cells to adapt to new conditions. If they come out of a prolonged stationary phase, they may have to adjust metabolically. The lag phase may be either very short or very long. The latency phase is followed by the logarithmic phase. In this phase, the cells divide and grow most actively. This is the period where the population is growing exponentially. After the exponential phase, the population enters the stationary phase. The limitation of nutrients or the accumulation of metabolic waste slows population growth. A prolonged period in the stationary phase eventually leads to cell death where the population is in decline. This phase can be caused by the accumulation of toxic waste or by a very low nutrient availability, or damage to cells. Microbial growth can be measured in different ways. One way is to measure the optical density (OD) of a culture broth as a function of time. The change in the apparent absorbance (actually the loss of light due to light scattering by the particles) can be used to obtain an estimate of the generation time. EXERCISE 5.0: E.COLI GROWTH CURVE (Groups of 2) Materials Each group of 2 will be assigned a 25 ml culture of E. coli in a 250 ml flask. The cultures will be initiated before the lab period at a time which represents approximately 10h, 11h, 12h and 13h. The time when the inoculation was performed shall be indicated on the bottle. Two growth conditions will be available LB medium at 37 o C with shaking LB medium at room temperature with shaking Spectrophotometer with disposable cuvettes 20 ml LB broth One test tube 45

47 Method 1. Obtain the culture flask which you were assigned. Record in your note book the following information: Growth condition and the time at which the inoculation was done. 2. Transfer 2 ml of the culture to a test tube and place on ice. Record the exact time at which the sampling was done. Immediately return the flask to the shaker at the appropriate temperature. 3. Blank the spectrophotometer at a wavelength of 600nm with 1 ml of LB. 4. Empty the cuvette in the waste beaker. Transfer 1 ml of the sample you collected to the cuvette. Obtain the optical density and record it in your note book. 5. Repeat steps 1-4 every 30 minutes until you ve obtained 8 optical density readings. YEAST FERMENTATION Yeasts are facultative anaerobes; that is to say they can grow in the presence or absence of oxygen by metabolisms involving either respiration or fermentation, respectively. One of the main differences between respiration and fermentation is the way that the NADH produced during glycolysis is recycled to NAD +. During aerobic respiration, the hydrogen electrons from NADH are transferred to oxygen in the electron transport chain generating 3 ATP per NADH while during fermentation the hydrogen electrons are transferred to acetaldehyde producing ethanol without ATP generation: Yeast s growth rate can thus be assessed by examining the efficiency with which this biochemical pathway takes place. In the following exercise, you will examine the efficiency of fermentation as a function of different carbon sources. 46

48 EXERCISE 5.1: YEAST FERMENTATION BIOASSAY (Groups of 2) Materials Dry yeast Sterile water 20% (m/v) Glucose 20% (v/v) Coke (assigned) 20% (v/v) Orange juice (assigned) Phosphate buffer ph 7 2 Fermentation assemblies Method 1. Prepare a yeast suspension by adding 10g of dry yeast to 50 ml of water. Rehydrate at room temperature for 5 minutes. 2. Prepare the following fermentation media in Falcon tubes : 20 % Glucose Buffer ph 7.0 Water 1.0 ml 5.0 ml 3.0 ml 0.5 ml 5.0 ml 3.0 ml 0.1 ml 5.0 ml 3.0 ml 0.05 ml 5.0 ml 3.0 ml 3. Once the media have been prepared, add 10 ml of the yeast suspension to the first two tubes. 4. Transfer all the contents to 50cc syringes. 5. Assemble the apparatus as illustrated on the next page to follow the production of CO Record the volume of water at time 0 in the collection tube. 7. Record the volume of water every 10 minutes for a total period of 1 hour. 8. Repeat the experiment with the other two glucose based media. 9. Repeat the experiment as indicated above with the drink you were assigned. Prepare the following fermentation media in falcon tubes: 20 % Coke or Buffer ph 7.0 Water Orange juice 1.0 ml 5.0 ml 3.0 ml 0.5 ml 5.0 ml 3.0 ml 0.1 ml 5.0 ml 3.0 ml 0.05 ml 5.0 ml 3.0 ml 47

49 50 ml Falcon tube Water CO 2 released 20 ml Yeast suspension Lid with holes 50 cc syringe Water 48

50 DEATH KINETICS Different microorganisms have different susceptibilities to treatments. For example, spores are particularly resistant. As with bacterial growth, mortality occurs exponentially. Consequently, mathematical functions that describe the profile of cell death under a given condition were made. Such functions are useful for determining the minimum time required to reduce a microbial population below a harmful level. The decimal reduction time, called the D value, represents the period of time under given circumstances necessary to reduce a population of microorganisms by a value of one log or 90%. In other words, if the value of 1 D of E.coli is 1 minute, this indicates that an exposure of one minute is required to reduce the bacterial population by 90%. So to reduce a population of 1 x 10 6 to 1 x 10 4 cells of E. coli would take 2D which is equivalent to 2 minutes. The D values are influenced by the bacterial species, form, and the conditions in which they find themselves. For example, the D value of spores is usually much higher than that of vegetative cells. The D value is an important parameter used in several areas to evaluate and compare the effectiveness of different treatments. EXERCISE 5.2: DIFFERENTIAL STAINS AND STERILIZATION (Groups of 2) As we have seen, dyes can be used as an indicator of metabolism. Since metabolism is an indication of viability, it is possible to use such dyes to assess the viability of microorganisms in a sample. An example where this would be desired is to assess the effectiveness of different methods of sterilization. Sterilization is a term referring to any process that eliminates (removes) or kills all forms of life (such as fungi, bacteria, viruses, spores, etc.) in a given region, on a surface, a volume of fluid, drugs or compounds such as culture media. Sterilization may be performed by one or more of the following processes: heat, chemicals, irradiation, high pressure, and filtration. Sterilization is distinct from disinfection and pasteurization in that the sterilization kills or inactivates all forms of life. In the following exercise, you will evaluate the effectiveness of different sterilization methods on different yeast preparations. To obtain a measure of the effectiveness, a metabolic dye, methylene blue, will be used. During respiration and fermentation, hydrogen electrons are removed from glucose molecules by enzymes called dehydrogenases and given to various chemical compounds such as NAD +. In this way, substances such as glucose provide energy for the vital reactions in living organisms. In this exercise we will be using methylene blue as an artificial electron acceptor (oxidative agent). When methylene blue is reduced, it becomes colorless. Thus, after staining with methylene blue, yeast cells which are metabolically active are colorless, while dead cells are blue. 49

51 Materials Dry yeast preparation (1g) 0.01% methylene blue, 2% (w/v) sodium citrate dihydrate in PBS 1% Glucose 4 petri dishes Hemocytometer slide Method 1. Suspend approximately 0.1g of dry yeast in 10 ml of the glucose solution. 2. Mix well for about 2 minutes. 3. Transfer 0.1 ml to a microcentrifuge tube labelled time Transfer the suspension to a petri dish. 5. Treat the suspension in the petri dish in the microwave at a power setting of 20 for 20 seconds. 6. Transfer 0.1 ml to a labelled microcentrifuge tube. 7. Swirl the yeast suspension in the plate and repeat steps 4 and 5 for additional 20 seconds intervals for a total exposure time of 2 minutes. You should therefore have collected samples representing exposure times of 0, 20, 40, 60, 80, 100 and 120 seconds. 8. Add 0.5 ml methylene blue to each of the tubes. Wait 1 minute. 9. Examine with the microscope on a hemocytometer slide. Count the number of colorless cells and the number of blue cells per square. Obtain counts from a number of squares which represents approximately a total of 50 cells. 10. Determine the percentage of live cells in each sample as follows. Number of colorless cells X 100 Total number of cells total de cellules 11. Determine the percentage reduction of viability for each treatment. % of viable cells before treatment - % of viable cells after treatment % of viable cells before treatment X

52 LAB N O 6 Schedule 1 2 pm Lecture 2 2:30 pm Exercise 6.0 2:30 2:45 pm Exercise 6.1 2:45 4 pm Exercise 6.2 CONTROL OF MICROBIAL GROWTH - ANTIBIOTICS Antibiotics can be synthetic, semi-synthetic or natural compounds, which inhibit the growth or kill bacteria. Antibiotics can be generally classified according to their mode of action as being bacteriostatic, bacteriolytic, or bactericidal. Bacteriostatic antibiotics inhibit growth but do not kill bacteria. Bacteriolytic antibiotics kill bacteria by causing their lysis. Bactericidal antibiotics kill bacteria without lysis. KIRBY-BAUER DISC DIFFUSION METHOD When faced with a newly discovered bacterial pathogen or a new derivative of a known pathogen, it is essential to assess the antibiotic susceptibility of the isolate. Initially, one must test the effect of a wide variety of different antibiotics to determine which may be potentially used. This is usually assessed by a semi-quantitative assay referred to as the Kirby-Bauer disc sensitivity method. In this test, the sensitivity of a bacterium to a variety of different antibiotics is tested. This test is performed on a Mueller-Hinton agar plate of which the surface is covered with an inoculum of the bacterium to be tested. Filter discs containing known amounts of different antibiotics are then placed on the surface of the plate. Following a suitable incubation period for optimal growth of the bacterium being tested, the plate is examined for the absence of bacterial growth in proximity to the antibiotic discs. These zones of inhibition are observed as halos around the antibiotic discs. The diameter of the halo is used as a measure of the relative sensitivity of the bacterium to the antibiotic (see figure). Zone size recommendations for the interpretation of the Kirby-Bauer (resistant, intermediate, sensitive) are established by disease control organizations. 14mm 51

53 EXERCISE 6.0: KIRBY-BAUER ASSAY (Groups of 2) Materials 5 ml broth culture of S. faecalis 5 ml broth culture of E. coli 4 chocolate agar plates Sterile swabs Filter discs 24 wells plate 10 ml of the assigned antibiotic Approx. 50 ml TSB Forceps Antibiotic Ampicillin Kanamycin Nalidixic acid Erythromycin Class Beta-lactam Aminoglycoside Quinolone Macrolide Method 1. Label two chocolate agar plates Streptococcus and the other two E. coli. 2. Immerse a cotton swab into the broth culture until it is thoroughly wet. Remove surplus suspension from the swab by rotating against the inside of the culture tube. 3. Spread the entire surface of each plate with the appropriate culture. Even distribution is essential; spread evenly in three directions so that even, confluent growth will result. 4. With plates covered, allow the inoculums to dry for 15 min. 5. Divide each plate into six sections and label as follows: A1 A2 A3 B1 B2 B3 A6 A5 A4 B6 B5 B4 Preparation of the dilutions of the assigned antibiotic : 6. Add 0.5 ml TSB to each of the wells of row A and B of a 24 well plate. 7. Add 0.5 ml of the assigned antibiotic to the first well of the first row (A1). 8. Perform 2 fold serial dilutions by transferring 0.5 ml from well A1 to well A2, from well A2 to well A3 and so on to generate 12 dilutions from A1 to B6. 52

54 Preparing the antibiotic discs: 9. Using forceps, dip one of the filter discs in the antibiotic well A1. Deposit the disc in the section A1 of the inoculated agar plates. Apply a light pressure on the disc to ensure a good contact with the agar 10. Repeat the previous step with a new disc dipped in well A2 and deposit on section A2 of the inoculated plates. 11. Repeat these steps for each antibiotic dilution. 12. Incubate the inverted plates at 37 C until next week. E-TEST This test combines the principals of the Kirby Bauer assay and the dilution method described in the next experiment. The E-test is a well-established method for antimicrobial resistance testing in microbiology laboratories around the world. E-test consists of a predefined gradient of antibiotic concentrations on a plastic strip and is used to determine the Minimum Inhibitory Concentration (MIC) of antibiotics. EXERCISE 6.1: SENSITIVITY OF S. FAECALIS TO VANCOMYCIN (Groups of 2) Materials 5 ml of S. faecalis broth 1 Chocolate agar plate Sterile swab Vancomycin E-test Forceps Method 1. Use the same approach as that in exercise 6.1 to inoculate a chocolate agar plate with S. faecalis. 2. Using forceps, deposit the E-test strip in the center of the plate. Gently tap the strip to ensure full contact with the agar surface. 3. Incubate the inverted plate at 37 C. 53

55 DETERMINING THE THERAPEUTIC DOSE When an antibiotic is to be used for therapeutic purposes, it is essential to determine the minimal concentration of the antibiotic, which is going to be effective. Lack of this information can result in the prescription of too high or too low a dose. Why would either of these scenarios represent a problem? The most common way of determining both the minimal inhibitory concentration (MIC) and the minimal bactericidal concentration (MBC) of an antibiotic is the broth dilution method. A fixed amount of bacteria is inoculated into broth containing varying amounts of the antibiotic being tested. The lowest antibiotic concentration at which no bacterial growth is observed after a suitable incubation period is referred to as the MIC. EXERCISE 6.2: DETERMINING MICs (Groups of 2) Materials 5 ml broth culture of S. faecalis 5 ml broth culture of E. coli 10 ml of assigned antibiotic (1mg/mL in TSB) Approx. 50 ml TSB 24 well plate Method 1. Follow the approach described in exercise 6.0 to generate 12 2 fold serial dilutions of the assigned antibiotic in rows A and B. 2. Repeat step 1 for rows C and D. 3. Dilute in TSB each of the provided cultures in order to obtain 5 ml of broth representing a 10 5 X dilution factor. 4. Inoculate each of the wells from rows A and B with 0.5 ml of the diluted S. faecalis culture. 5. Inoculate each of the wells from rows C and D with 0.5 ml of the diluted E. coli culture. 6. Seal the plate with parafilm and incubate at 37 o C until next week. S. faecalis E. coli 54

56 LAB N O 7 CONTROL OF MICROBIAL GROWTH (Cont d) EXERCISE 7.0: KIRBY BAUER DIFFUSION ASSAY Materials Kirby Bauer assay from last week Method Obtain the diameters of the zones of inhibition for each of the antibiotics with each of the bacteria tested. EXERCISE 7.1: SENSITIVITY OF S. FAECALIS TO VANCOMYCIN - E-TEST (Groups of 2) Materials E-Test plate from last week. Method 1. Read the MIC from the strip, which is represented by the concentration indicated on the strip which gives rise to the smallest diameter for the zone of inhibition. CMI 55

57 BACTERIAL METABOLISM AND DIFFERENTIAL TESTS Bacteria are amongst the most diverse organisms on earth; indeed they can be found everywhere. It is therefore obvious that they must have very different metabolisms that are adapted to the environments they inhabit. This great diversity is the biochemical basis of the diagnostic approach that is used by microbiologists to identify microorganisms. The tests used for identification and diagnosis are called differential or biochemical tests. This week you will use these tests to examine different biochemical characteristics dependent on specific enzymatic reactions in certain metabolic pathways. A summary of the different tests that you will perform is presented below: Starch agar Milk agar Spirit Blue agar DNA agar Phenol red broths MR-VP Bacteria B. cereus B. subtilis E. coli E. aerogenes P. aeruginosa P. mirabilis TSI slant Simmon s citrate slant Urea slant Ornithine decarboxylase broth Phenylalanine agar slant Lysine agar slant Sim tube Nitrate broth 56

58 UTILIZATION OF COMPLEX CARBON SOURCES: EXOCELLULAR ENZYMES Simple carbon sources such as monosaccharides, disaccharides, and amino acids enter the cell either by simple diffusion or by making use of specific transporter systems. In contrast, more complex carbon sources such as polysaccharides and proteins are too large to use either of these transport mechanisms. Consequently, they must first be broken down into smaller manageable units before they can be utilized. This is achieved by the secretion of specialized exocellular enzymes, which operate outside of the cell to break down polymers into smaller monomeric compounds. Examples of complex polymeric carbon sources utilized by some bacteria are starch, a polysaccharide, casein, a polypeptide or protein, tributyrin, a lipid or fatty acid polymer and DNA, a nucleotide polymer. Each of these complex carbon sources require the secretion of a specific enzyme, which degrades them into monomers, which can be easily transported into the cell to be metabolized. -amylase, is an enzyme that can cleave the -1, 4 linkage joining the glucose monomers in starch. Caseinase is a protease, which can cleave the peptide linkages joining the amino acid monomers in the protein casein. Lipase can degrade into individual fatty acids. Finally, DNase cleaves the phosphodiester linkages between the nucleotides within a polynucleotide chain. EXERCISE 7.2: DEGRADATION OF COMPLEX CARBON SOURCES (Groups of 2) Materials B. cereus B. subtilis E. coli 3 starch plates 3 milk plates 3 Spirit Blue plates 3 DNA plates Method 1. Appropriately label the different media plates with the bacteria to be tested. Four media/bacteria. 2. Streak for single colonies each of the bacteria indicated above on one of each the agar media. 3. Incubate at 28 o C. 57

59 SUGAR METABOLISM PHENOL RED BROTH The phenol red broth is a differential medium to which a sugar is added. The base medium contains a source of protein, peptone, and the ph indicator, phenol red. Phenol red turns yellow in acid medium and turns red in alkaline medium. In addition, an inverted vial is used to detect the accumulation of gas. Generally, the presence of acid with or without a gas build-up indicates a fermentative metabolism. EXERCISE 7.3: METABOLISM IN PHENOL RED BROTH (Groups of 2) Materials P. miribalis P. aeruginosa E. coli E. aerogenes 3 phenol red broths of each of the following sugars: glucose, lactose, and sucrose Method 1. Label each set of 3 phenol red broths (glucose, lactose and sucrose) with one of the bacteria indicated above. 2. Inoculate each bacterium in the appropriate broths. 3. Incubate at 37 o C. GROWTH IN TSI MEDIUM (TRIPLE SUGAR IRON) This growth medium is commonly used to obtain a preliminary identification of bacterial members of the Enterobacteriaceae family. It contains four different potential carbon sources, glucose, lactose, sucrose, and proteins, as well as a ph indicator that allows one to discriminate between the use of proteins or sugars. The fermentation of the sugars as carbon source gives rise to an acidic reaction, whereas the oxidation of proteins results in an alkaline reaction. The medium is supplied as a slant, which allows the simultaneous observation of growth under both aerobic and anaerobic conditions. The surface of the slant provides good aerobic conditions, whereas the butt is essentially devoid of oxygen thus favoring fermentation or anaerobic respiration. Amongst the three sugars, glucose is limiting (0.1%) whereas sucrose and lactose are available in excess (1.0%). Since all the Enterobacteriaceae can metabolize glucose, this metabolism initially makes the medium acidic (yellow). For continued growth, after the glucose has been exhausted, one of the other carbon sources must be used. If neither sucrose nor lactose can be used, the carbon source which will be metabolized will then be proteins, generating alkaline by-products. The resulting increase in ph will thus change the medium color from yellow to a neutral or an alkaline (orange or red respectively. However, if sucrose and/or lactose can be used anaerobically the acids produced will cause the medium to remain acid (yellow). In addition to allowing the distinction between the fermentation of different sugars, TSI medium allows one to determine whether a bacterium can degrade amino acids with a sulfur group, such as methionine and cysteine. Degradation of these amino acids generates as a by-product hydrogen sulfide, which reacts with ferrous sulfate in the medium, giving rise to a black precipitate. 58

60 EXERCISE 7.4: GROWTH IN TSI MEDIUM (Groups of 2) Materials P. miribalis P. aeruginosa E. coli 3 TSI Slants Method 1. Inoculate the surface and the butt of TSI slants with your two bacteria. (See image) 2. Incubate at 37 o C. Stab the inoculation loop down into the bottom of the butt. As you withdraw the loop, streak the surface of the slant. Slant Butt 59

61 USE OF CITRATE AS A CARBON SOURCE This test is designed to determine whether a microorganism can use citrate, an intermediate of the Krebs cycle, as sole source of carbon and inorganic ammonium salts as sole nitrogen source. Bacteria which possess the enzyme citrate permease can transport the citrate within the cell to convert it to pyruvate. The use of citrate generates alkali by-products which are identified by the inclusion of a ph indicator, bromthymol blue, which is green at a ph of 6.8 and blue at a ph of 7.6 and above. EXERCISE 7.5 GROWTH ON SIMMON S CITRATE SLANT (Groups of 2) Materials E. coli E. aerogenes 2 Simmon s citrate slants Method 1. Streak each of the above indicated bacteria on a Simon s citrate slant 2. Incubate at 37 o C. UREA METABOLISM Urea is a product resulting from the decarboxylation of specific amino acids and thus the degradation of proteins in vertebrates. Some microorganisms can use urea as a nitrogen source. Urea catabolism requires urease, an enzyme hydrolyzing the urea in ammonia, carbon dioxide and water. Ammonia released in the culture medium makes it alkaline. A ph indicator in the culture medium, phenol red, can detect the presence of alkali products, changing the color of the medium to pink. EXERCISE 7.6: GROWTH ON UREA SLANT (Groups of 2) Materials E. coli P. mirabilis 2 Urea slants Method 1. Streak each of the above indicated bacteria on a urea slant 2. Incubate at 37 o C. 60

62 DECARBOXYLASES AND DEAMINASES Deamination and decarboxylation tests are used for differentiating bacteria of the Enterobacteriaceae family. The majority of these bacteria produce several enzymes necessary for the degradation of amino acids. The enzymes that remove a COOH group are decarboxylases while those that remove NH2 groups are deaminases. The media used to verify the presence of different amino acid decarboxylase contain glucose, a fermentable carbon source, and the desired amino acid. The acid produced from glucose fermentation reduces the ph of the medium and changes the color of the ph indicator from purple to yellow. These acidic conditions stimulate the activity of decarboxylases. The decarboxylation of amino acids causes a rise in ph, which changes the color of the indicator, Bromcresol purple, from yellow to purple. If the organism does not produce the appropriate enzyme the medium remains acid (yellow). One of the deaminases produced by many bacteria allows for the deamination of phenylalanine generating phenylpyruvic acid. This reaction can be detected in the phenylalanine agar medium that provides a high source of phenylalanine. After incubation in this medium, phenylpyruvic acid reacts with a reagent in this medium, ferric chloride (FeCl3) generating a green color. Deamination and decarboxylation of lysine can be detected in lysine iron agar medium. This medium contains, amongst other things, glucose, lysine, sodium thiosulfate as a sulfur source that can be reduced and the ph indicator, bromcresol purple. If an organism that has lysine decarboxylase is inoculated into this medium, glucose fermentation creates an acidic environment that induces the production of decarboxylase. The fermentation of glucose initially causes the medium to turn yellow, but after decarboxylation the medium becomes alkaline and turns purple. In contrast, if the organism produces lysine deaminase, the generated products reacts with ammonium ferric citrate and produces a red color. EXERCISE 7.7: DECARBOXYLASE AND DEAMINASE ASSAYS (Groups of 2) Materials E. coli P. mirabilis E. aerogenes 3 decarboxylase broths without any amino acid 3 decarboxylase broths with ornithine 3 phenylalanine agar slants 3 Lysine with iron agar slants Method 1. Inoculate the decarboxylase broths lacking any amino acids with each of the bacteria. 2. Inoculate the decarboxylase broths with ornithine with each of the bacteria. 3. Overlay all the broths with mineral oil to create anaerobic conditions. 4. Inoculate the phenylalanine agar slants with each of the above indicated bacteria. 5. Inoculate the lysine agar slants with each of the above indicated bacteria. 6. Incubate at 37 o C. 61

63 SIM: PRODUCTION OF HYDROGEN SULFIDE, INDOLE AND MOTILITY H 2 S PRODUCTION As with the TSI medium, the degradation of sulfur containing amino acids can be determined by the production of a black precipitate. MOTILITY Some microorganisms have the ability to move with the help of flagella. This characteristic can easily be observed in semi-solid media in which non-motile bacterial growth is restricted to the site of inoculation whereas the growth of motile bacteria can be observed to spread beyond the site of inoculation. INDOLE PRODUCTION: DEGRADATION OF TRYPTOPHAN Another amino acid that some microorganisms can use as carbon source is the aromatic amino acid tryptophan. The degradation of tryptophan generates the by-products pyruvic acid and indole. Pyruvic acid can then be metabolized as a carbon source whereas indole is secreted within the growth medium as a waste product. Indole production can be detected by its reaction with the reagent dimethylaminobenaldehyde (Kovac's reagent). EXERCISE 7.8: SIM TEST (Groups of 2) Materials E. coli P. mirabilis E. aerogenes 3 SIM tubes Method 1. Use an inoculation needle to inoculate each of the above indicated bacteria down to the bottom of the tube along a straight line. 2. Incubate at 37 o C. 62

64 NITRATE AND NITRITE REDUCTION Some bacterial species can reduce nitrates to nitrite and subsequently the nitrite to ammonia, which can be used for the synthesis of amino acids. Enzymes called nitrate reductases, which are necessary for the assimilation of nitrates, catalyze these reactions. Other bacterial species can use nitrates instead of oxygen as a final electron acceptor for the generation of energy. This nitrate reduction pathway is said to be dissimilatory. The use of nitrates as a final electron acceptor is an example of anaerobic respiration. NO 3 - NO 2 - NH 4 + Nitrate reductase Nitrite reductase other enzymes N 2 EXERCISE 7.9: NITRATE REDUCTION ASSAY (Groups of 2) Materials E. coli P. aeruginosa P. mirabilis 3 nitrate broths Method Inoculate each of the above indicated bacteria in nitrate broths and incubate at 37 o C. 63

65 ENTEROPLURI TEST: ENTEROBACTERIACEAE SYSTEM This test is a 12-sector system containing special culture media that permits identification of the Enterobacteriaceae and other gram negative, oxidase negative bacteria. The system allows the simultaneous inoculation of all media present in the sectors and the execution of 15 biochemical reactions. The microorganism is identified by evaluating the color change of the different culture media after hours of incubation at 36 ± 1 C and by a code number obtained from the biochemical reactions. The combination of reactions obtained permits, with the help of the codebook to identify significant Enterobacteriaceae from a clinical point of view. EXERCISE 7.10: ENTEROPLURI TEST (Groups of 2) Materials Gram negative unknown Enterotube Method 1. Remove both caps. The tip of the inoculating wire is under the white cap. Do not flame the wire. 2. Pick a well isolated colony directly with the tip of the Enteropluri inoculating wire (Figure 1 on next page). A visible amount of inoculum should be seen at the tip and the side of the wire. 3. Inoculate the Enteropluri test by first twisting wire, then withdrawing wire through all twelve compartments applying a turning motion (Figure 2 on next page). 4. Reinsert wire (without sterilizing) into Enteropluri tube, using a turning motion through all compartments, until the notch on the wire is aligned with the opening of the tube (Figure 3 on next page). 5. Break wire at notch by bending. The portion of the wire remaining in the tube maintains anaerobic conditions necessary for true fermentation of glucose, production of gas and decarboxylation of lysine and ornithine. 6. With the broken off part of the wire, punch holes through the film covering the air inlets of the compartments Adonitol, Lactose, Arabinose, Sorbitol, VP, Dulcitol/PA, Urea, and Citrate in order to support aerobic growth. (Figure 4 on next page). Replace both caps. 7. Incubate at 37 C until the next lab period with Enteropluri test lying on its flat surface (See figure on next page). 64

66 See the video at the following link: Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 65

67 THE STREPTOCOCCI AND THE STAPHYLOCCOCI As previously mentioned bacteria are ubiquitous and can normally be found on several parts of the human body. Unless there is an infection, the majority of the human body which is not in contact with the external environment is sterile. This includes your blood, all your internal fluids, internal organs, cavities, etc.. In contrast, various microorganisms inhabit every surface of the human body exposed to the environment. The majority of these bacterial species are harmless unless their growth becomes uncontrolled. The human body provides a wide variety of different environments that differ with regards to the availability of oxygen, ph, water content, etc. Accordingly, various microorganisms have established niches in different regions of the body. In the following exercise, you will sample your throat to isolate microorganisms representative of this niche. The upper respiratory tract is populated by many Gram-positive species, many of which are potential pathogens if their growth is unchecked. These include Staphylococci and Streptococci. Amongst the Staphylococci, S. aureus and S. epidermidis, are species of clinical importance. It is estimated that in 10-40% of the population S. aureus is present in the natural flora. However, this species is associated with several medical problems, such as abscesses, bacteremia, and endocarditis. S. epidermidis, one of the resident species of the skin, is an opportunistic pathogen. These species are not usually pathogenic in healthy individuals, but can cause serious infections in weakened individuals, such as immunocompromised individuals. BLOOD HEMOLYSIS Blood agar (BA) contains general nutrients and 5% sheep blood. It is useful for cultivating fastidious organisms and to determine the hemolytic activities of an organism. Some bacteria produce exoenzymes that lyse red blood cells and degrade hemoglobin; called hemolysins. Bacteria can produce different types of hemolysins. Beta-hemolysin breaks down the red blood cells and hemoglobin completely. This leaves a clear zone around the bacterial growth. This is referred to as β-hemolysis (beta hemolysis). Many pathogenic species of Streptococcus and some of Staphylococcus belong to this group. Alpha-hemolysin partially breaks down the red blood cells and leaves a greenish color behind. This is referred to as α-hemolysis (alpha hemolysis). The greenish color is caused by the presence of biliverdin, a by-product of the breakdown of hemoglobin. Many non-pathogenic species of Streptococcus belong to this group. If the organism does not produce hemolysins and does not break down the blood cells, no clearing will occur. This is called γ-hemolysis (gamma hemolysis). Most Streptococci within this group are non-pathogenic. 66

68 EXERCISE 7.11: THROAT SAMPLING ON BLOOD AGAR PLATES (Groups of 2) Materials 6 blood agar plates 20 ml TSB Cotton swabs Method You will have to sample the throat of two different people. 1. Ask someone to use a cotton swab to sample the tonsil area of your throat (See the illustration below) 2. Drop the swab in a tube containing 1 ml of TSB. 3. After having vigorously mixed, prepare 10-1 and 10-2 dilutions in TSB. 4. Spread 0.1mL of the undiluted sample and each of the two dilutions on blood agar plates. 5. Incubate at 37 o C in the candle jar. 67

69 DIFFERENTIAL TESTS CONTINUED LAB N O 8 EXERCISE 8.0: DEGRADATION OF COMPLEX CARBON SOURCES (Groups of 2) 1. Examine your milk, spirit blue and DNA plates for any clearing surrounding the bacterial growth. Such a clearing is indicative of the production of exocellular enzymes. 2. For the starch plate, flood the bacterial growth with Gram's iodine. Gram's iodine reacts with starch to produce a dark blue color. Lack of color development indicates that starch was degraded. Amylase Caseinase Lipase DNAse 68

70 EXERCISE 8.1: METABOLISM IN PHENOL RED BROTH (Groups of 2) Method 1. Obtain your pairs of phenol red broths and make the following observations : Was there any growth What is the color of the broth Is there gas accumulation Acid Alkaline Acid + Gas 2. According to your observations, determine which sugars were metabolized and by what metabolism. Possible results Observation Interpretation Yellow broth with bubble in vial Fermentation with acid by-products and gas Yellow broth without bubble in vial Fermentation with acid by-products without gas Orange broth (Original color) No fermentation Red broth without bubble in vial Degradation of peptones with alkali by-products 69

71 EXERCISE 8.2: GROWTH IN TSI (Groups of 2) Do an analysis of your TSI slants in order to obtain the following information: What is the reaction of the slant: acid (A), alkaline (K) or neutral (NC) What is the reaction in the butt : acid (A), alkaline (K) or neutral (NC) Is there H 2 S production o If so, is it produced aerobically, anaerobically or both Is there any gas accumulation A B C D A. Acid butt and slant + accumulation of gas B. Alkaline slant and acid butt C. Alkaline butt and alkaline slant + H 2 S anaerobically D. Alkaline slant and neutral butt 70

72 Possible results: Results (slant/butt) Symbol Interpretation Red/yellow Yellow/yellow Red/red Red/no color change Yellow/yellow with bubbles Red/yellow with bubbles Red/yellow with bubbles and black precipitate Red/yellow with black precipitate Yellow/yellow with black precipitate K/A A/A K/K K/NC A/A,G K/A,G K/A,G, H 2 S K/A, H 2 S A/A, H 2 S Glucose fermentation only; Peptone catabolized Glucose and lactose and/or sucrose fermentation No fermentation; Peptone catabolized No fermentation; Peptone used aerobically Glucose and lactose and/or sucrose fermentation; Gas produced Glucose fermentation only; Gas produced Glucose fermentation only; Gas produced; H 2 S produced Glucose fermentation only; H 2 S produced Glucose and lactose and/or sucrose fermentation; H 2 S produced No change/no change NC/NC No fermentation A=acid production; K=alkaline reaction; G=gas production; H 2 S =sulfur reduction Typical results Slant Butt Gas H 2 S Escherichia coli A A + - Proteus mirabilis K A - + Pseudomonas aeruginosa K K - - Salmonella typhimurium K A - + Shigella flexneri K A - - A : Acid (yellow) K: Alkaline (Red) 71

73 GLUCOSE FERMENTATION: PRODUCTION OF MIXED ACIDS OR ACETOIN Some bacteria ferment glucose generating large quantities of acids that cause a significant reduction in the ph of their environment. This can be detected by the ph indicator methyl red which is red at ph 4.4 or less. In contrast, other bacterial species generate only small amounts of acids, but large amounts of neutral by-products such as ethanol and butanediol. An intermediate for the production of butanediol is acetoin, which can be detected using two reagents, alpha naphthol and KOH. EXERCISE 8.3: METHYL RED - VOGUES-PROSKAUER TEST (MRVP) (Groups of 2) Materials Cultures of E. aerogenes and E. coli in MRVP broths MRVP reagents 4 test tubes Method 1. To complete the MR and VP tests, transfer 1 ml of the MRVP broth culture indicated above to each of two new test tubes. 2. For the MR test, add 2 drops of methyl red to one of the two tubes. Observe the color at the surface of the broth. A red color indicates the presence of a large quantity of acids whereas a yellow color indicates the absence of any acids. 3. To complete the VP test, add 6 drops of alpha-naphtol to the second tube. 4. Add 3 drops of KOH to the tube. 5. Mix well and allow the reaction to proceed for minutes. 6. Observe whether there is a color change. If a red color develops it indicates the presence of acetoin. Negative Positive Positive Negative 72

74 EXERCISE 8.4 GROWTH ON SIMMON S CITRATE AGAR SLANT (Groups of 2) Method 1. Examine your slants for a color change to blue. This color indicates that alkaline by-products were generated indicating citrate could be used as a carbon source. Negative Positive EXERCISE 8.5: GROWTH ON UREA SLANT (Groups of 2) Method 1. Examine your slants for a color change to pink. This color indicates that alkaline by-products were generated by the action of urease. Negative Positive EXERCISE 8.6: DECARBOXYLASE AND DEAMINASE ASSAYS (Groups of 2) ORNITHINE DECARBOXYLASE Method 1. Obtain your decarboxylase assay broths and compare the color of the broths with and without ornithine O -O +O -O +O -O +O: with ornithine; -O: Without ornithine 1. Negative result : Alkaline reaction in the presence and absence of amino acids 2. Positive result : Alkaline reaction with the amino acid but an acid reaction in its absence 3. Negative result : Acid reaction with or without amino acids 73

75 PHENYLALANINE DEAMINASE Method 1. Obtain your phenylalanine slants. 2. Add a few drops of 12% ferric chloride. 3. Note any change in color. (A : Negative, B : Positive) LYSINE DECARBOXYLASE AND DEAMINASE Method 1. Obtain your lysine agar slants. 2. Note any color change and the corresponding area. Color Purple butt and slant Yellow butt and purple slant Yellow butt and red slant Black precipitate Interpretation Lys deaminase negative Lys decarboxylase positive Lys deaminase negative Lys decarboxylase negative Fermentation of glucose Lys deaminase positive Lys decarboxylase negative Fermentation of glucose Sulfur reduction EXERCISE 8.7: SIM TEST (Groups of 2) Do an analysis of your SIM tubes to obtain the following information: Are your bacteria motile Is there production of H 2 S Is tryptophan used as a carbon source o Is there indole production? To obtain this result, add a few drops of Kovacs reagent. If indole was produced the reagent becomes red A. Motile +H 2 S B. Motile + Indole C. Non-motile D. No growth 74

76 EXERCISE 8.8: NITRATE REDUCTION ASSAY (Groups of 2) Method 1. Add 3 drops of sulfanilic acid and 3 drops of alpha-naphtylamine to your nitrate broth culture. These reagents react with nitrite to give a red color. 2. Wait for 1 minute. If no color change occurs, add a pinch of zinc powder. 3. Observe if a color change occurs. Zinc reduces nitrates to nitrites which then react with the sulfanilic acid and the alpha-naphtylamine to give a red color. Results after the addition of sulfanilic acid and alpha-naphtylamine Results following the addition of zinc 75

77 EXERCISE 8.9: ENTEROPLURI TEST (Groups of 2) Method At the end of incubation: Observe the change in color of culture media in the different sectors and interpret results using the table below. NOTE: if there is no change in color in the sector Glucose/Gas while in some other sectors there are color changes, the microorganism being tested does not belong to the family of Enterobacteriaceae. Record the results obtained on the data chart; except for the Indole (sector H 2 S/Indole) and Voges-Proskauer tests (sector VP). Sector Biochemical reactions Sector colors Positive reaction Negative reaction Glucose/Gas Glucose fermentation Yellow Red Gas production Lifted wax Overlaying wax Lysine Lysine decarboxylation Violet Yellow Ornithine Ornithine decarboxylation Violet Yellow H 2 S/Indole Hydrogen sulfide production Black-brown Beige Indole test Pink-red Colorless Adonitol Adonitol fermentation Yellow Red Lactose Lactose fermentation Yellow Red Arabinose Arabinose fermentation Yellow Red Sorbitol Sorbitol fermentation Yellow Red VP Acetoin production Red Colorless Dulcitol/PA Dulcitol fermentation Yellow Green Phenylalanine deamination Dark brown Green Urea Urea hydrolysis Purple Beige Citrate Citrate utilization Blue Green Perform Indole and Voges-Proskauer tests. o Indole test Lay the EnteroPluri-Test with its flat surface pointing upward and, by punching the plastic film, add 3 or 4 drops of Kovac s Indole Reagent in the sector H 2 S/Indole. The reaction is positive if a pink-red color develops in the added reagent within seconds. o Voges-Proskauer test Lay EnteroPluri-Test with its flat surface pointing upward and, by punching the plastic film, add 3 drops of α-naphtol and 2 drops of potassium hydroxide. The reaction is positive if a red color develops within 20 minutes. 76

78 Generate the 5-digit code as follows: 1. The 15 biochemical tests are divided into 5 groups each containing 3 tests and each one is indicated with a positivity value of 4, 2, or 1. Value 4: first test positive in each group (Glucose, Ornithine, Adonitol, Sorbitol, PA) Value 2: second test positive in each group (Gas, H 2 S, Lactose, VP, Urea) Value 1: third test positive in each group (Lysine, Indole, Arabinose, Dulcitol, Citrate) Value 0: every test negative 2. Adding the number of positive reactions in each group, you will obtain a 5 digit code which, by the use of the Codebook, allows the identification of the microorganism under examination as in the following example: Group 1 Group 2 Group 3 Group 4 Group 5 Test Positivity code Result Code sum Numerical code Microorganism: 77

79 DIFFERENTIAL TESTS FOR THE IDENTIFICATION OF GRAM POSITIVE COCCI Micrococcaceae family The Micrococcaceae family includes pathogenic and non-pathogenic organisms often associated to the natural human flora. This family includes two main genera, the Staphylococcus and the Micrococcus. Both can make use of oxygen and possess a typically respiratory metabolism. Specifically, members of the genus Micrococcus are strict aerobes whereas those from the genus Staphylococcus are facultative aerobes. Indeed, the Micrococci produce acid from glucose only under aerobic conditions whereas the Staphylococci do so under both aerobic and anaerobic conditions. Several species of the genus Micrococcus have pigmented colonies, such as M. luteus (yellow) or M. roseus, (pink). Their cells often have a tetrad arrangement. Residents of the skin, this genus is rarely pathogenic, being rather opportunistic. Contrary to the Micrococci, the Staphylococci are human parasites which very often under certain conditions are the cause of serious illness. The three major species of the genus Staphylococcus are S. aureus, S. saprophyticus and S. epidermidis. S. epidermidis is a non-pigmented non-pathogenic organism usually found on the skin and mucus membranes. S. aureus, a yellow colored species, is commonly associated to acne, pneumonias, meningitis, and toxic shock syndrome. S. saprophyticus, another organism often found on skin is non-pigmented and often implicated in urinary infections. Streptococcaceae family This family includes bacteria of the genus Streptococcus, which includes pathogenic and nonpathogenic species. This genus is divided into three groups of related species; the Lactococcus, streptococci of importance to the dairy industry, the Enterococcus, which includes streptococci of fecal origins, and the Streptococcus, which includes most of the pathogenic species. The latter are classified according to the Lancefield classification system, which divides these bacteria into 8 groups (A-H and K-U). This classification is based on the immunological reaction of polysaccharides within their cell walls. Members of clinical importance of the Streptococcus genus include those from the group A; S. pyogenes. S. pyogenes is the principal cause of strep throats and in rare cases can cause the massive destruction of tissues (flesh eating bacteria). S. agalactiae, only member of the B group, causes septicemias in newborns resulting in death in 75% of cases. The group D Enterococci are implicated in urinary infections, endocarditis, and wound infections. Other Streptococci that are not classified according to the Lancefield classification system include S. pneumoniae, the principal causing agent of Pneumonias as well as S. mutans and S. mitis which are the main causes of cavities. 78

80 EXERCISE 8.10: BLOOD HEMOLYSIS (Groups of 2) Blood agar (BA) contains general nutrients and 5% sheep blood. It is useful for cultivating fastidious organisms and to determine the hemolytic activities of an organism. Some bacteria produce exoenzymes that lyse red blood cells and degrade hemoglobin; called hemolysins. Bacteria can produce different types of hemolysins. Betahemolysin breaks down the red blood cells and hemoglobin completely. This leaves a clear zone around the bacterial growth. This is referred to as β-hemolysis (beta hemolysis). Many pathogenic species of the genus Streptococcus and some of the genus Staphylococcus belong to this group. Alphahemolysin partially breaks down the red blood cells and leaves a greenish color behind. This is referred to as α-hemolysis (alpha hemolysis). The greenish color is caused by the presence of biliverdin, a by-product of the breakdown of hemoglobin. Many non-pathogenic species of the genus Streptococcus belong to this group. If the organism does not produce hemolysins and does not break down the blood cells, no clearing will occur. This is called γ-hemolysis (gamma hemolysis). Most Streptococci genera within this group are non-pathogenic. Materials Sampling of the throat on blood agar Materials 1. Examine your blood agar plates and determine the type of hemolysis observed: alpha, beta or gamma. 2. Perform a Gram stain on a colony representing each type of hemolysis EXERCISE 8.11: CATALASE (Groups of 2) This test represents the first step in the discrimination of bacteria of the Micrococcaceae family (catalase positive) from the Streptococcaceae family (catalase negative). Catalase is an enzyme, which is found in most organisms that live in the presence of oxygen, such as aerobic and facultative microorganisms. Oxygen metabolism generates free radicals, such as hydrogen peroxide, which damages the cell. Catalase reduces peroxide converting it to water and oxygen. 2H 2 O 2 2H 2 O + O 2 The generation of oxygen can easily be detected as fine bubbles. 79

81 Materials Gram positive cocci unknown on TSA plate (Labelled 1 or 2) 3% (v/v) Peroxide Method 1. Add 1-2 drops of 3% hydrogen peroxide to the growth of your bacterial unknowns. 2. Observe if there is the production of air bubbles. If your unknown is catalase positive, proceed with the PPT presentation «A». If your unknown is catalase negative, proceed with the PPT presentation «B». These presentations are available on K: /BIO3126. A description of the different tests follows: BILE-ESCULIN This test is useful for the identification of group D streptococci; the Enterococcus. These bacteria hydrolyze esculin to esculitin and glucose. Esculitin reacts with an iron salt, ferric citrate, generating a dark brown or black complex. Bile is included to inhibit the growth of Gram positive bacteria other than the Enterococci. BACITRACIN, OPTOCHIN AND NOVOBIOCIN SENSITIVITY The bacitracin and optochin sensitivity tests are utilized to identify Streptococcus pyogenes and Streptococcus pneumoniae, respectively. Only these two species of Streptococcus are sensitive to the respective antibiotics when the test is performed. Novobiocin sensitivity allows the discrimination of S. saprophyticus from other bacteria of the Staphylococcus genus; S. saprophyticus being resistant. Bacitracin Optochin Novobiocin S. pneumoniae Resistant Sensible Sensitive S. pyogenes Sensitive Resistant Sensitive S. saprophyticus N.A. N.A. Resistant MANNITOL + SALTS AGAR This medium contains a high salt concentration (7.5%) which allows the enrichment of bacteria of the genus Staphylococcus. As with phenol red broths, this medium provides two carbon sources, either mannitol or proteins. The inclusion of a ph indicator, phenol red, allows the discrimination of Staphylococcus fermenters, such as S. aureus, from the nonfermenters. 80

82 TELLURITE AGAR OR BAIRD PARKER AGAR This is a selective medium for the presumptive identification of coagulase-positive staphylococci. The selectivity of the medium is due to Lithium Chloride and a 1% Potassium Tellurite Solution, suppressing growth of organisms other than staphylococci. The differentiation of coagulase-positive staphylococci is based on Potassium Tellurite and Egg Yolk Emulsion. Staphylococci that contain lecithinase break down the Egg Yolk and cause clear zones around the colonies. Reduction of Potassium Tellurite, a characteristic of coagulase-positive staphylococci, causes blackening of colonies. PYR TEST The PYR test is a qualitative procedure for determining the ability of streptococci to enzymatically hydrolyze L- pyrrolidonyl-β-naphthylamide (PYR). Most group A streptococci and group D enterococci hydrolyze PYR. Whereas most group B streptococci and non-group A, B and D streptococci, yield negative PYR test results. L-pyrrolidonyl-β-naphthylamide (PYR) is hydrolyzed by bacteria that possess the enzyme pyrrolidonyl peptidase. Demonstrating PYR hydrolysis involves two reactions. Pyrrolidonyl peptidase, if present, hydrolyzes PYR to liberate L-pyrrolidone carboxylic acid and β-naphthylamine. Β-naphthylamine, reacts with p-dimethylaminocinnamaldehyde to form a pink/fuchsia precipitate. 81