BIOLOGY Principles of Microbiology LABORATORY MANUAL

Transcription

1 The University of Lethbridge BIOLOGY 3200 Principles of Microbiology LABORATORY MANUAL Spring, 2008 Written by: L. A. Pacarynuk Revised: December, 2006

2 TABLE OF CONTENTS Exercise Page Biology 3200 Laboratory Schedule 2 Grade Distribution 3 Occupational Health and Safety Guidelines 5 Guidelines for Safety Procedures 6 Exercise 1 Introduction to Microscopy 9 Exercise 2 General Laboratory Principles and Biosafety 14 Exercise 3 - Bacterial and Yeast Morphology 16 Exercise 4 The Ames Test 22 Exercise 5 Biochemical Tests 25 Exercise 6 Bacterial Reproduction 31 Exercise 7 Virology 36 Exercise 8 Water Quality 40 Appendix 1 The Compound Light Microscope 43 Appendix 2 Preparation of Scientific Drawings 46 Appendix 3 Aseptic Technique 48 Appendix 4 The Cultivation of Bacteria 54 Appendix 5 Bacterial Observation 59 Appendix 6 Laboratory Reports 60 Appendix 7 Use of the Spectrophotometer 62 Appendix 8 Media, Reagents, ph Indicators 64 Appendix 9 Care and Feeding of the Microscopes 73 1

3 BIOLOGY 3200 LAB SCHEDULE SPRING, 2008 Jan. 8 Jan. 10 Jan. 15 Jan. 17 Jan. 22 Jan. 24 Jan. 29 Jan. 31 Feb. 5 Feb. 7 Feb. 12 Feb. 14 Feb. 19 Feb. 21 Feb. 26 Feb. 28 Mar. 4 Mar. 6 Mar. 11 Mar. 13 Mar. 18 Mar. 20 Mar. 25 Mar. 27 Apr. 1 Apr. 3 Introduction, Microscopy General Lab Procedures, Biosafety Bacterial Morphology Bacterial Morphology Bacterial Morphology Bacterial Morphology Ames Test; Biochemical Tests - Selective and Differential Media Ames Test Complete; Selective and Differential Media - Complete IMViC Tests IMViC Tests - Complete Bacterial Growth Bacterial Growth Complete; Hand in Assignment 1 at the BEGINNING of the lab period Reading Week Reading Week Virology (phage isolation) Virology (phage elution) Virology (titre/host range) Virology Complete Water Microbiology Start (enumeration, MPN) Water Microbiology (Plating on EMB) Water Microbiology (select unknown; Gram stain) Water Microbiology (TSA plate) ; Hand in Virology Lab Report at BEGINNING of the lab period Water Microbiology (enterotube inoculation) Water Microbiology Complete Free Laboratory Period Free Laboratory Period Thursday Apr. 10 Final Lab Exam (practical and written) 2

4 Laboratory Grade Distribution: The laboratory component of Biology 3200 is worth 50% of your course mark. It is distributed as follows: Assignments and Skills Tests 10% Lab Report Virology 15% Due Thursday, March 20 at the beginning of lab Lab Books 10% (to be handed in twice) Lab Exam 15% Performance: Up to 10% of laboratory grade (5 marks out of 50) will be subtracted for poor laboratory performance. This includes (but is not limited to) failure to be prepared for the laboratory, missing lab notebook or lab manual, poor time management skills, improper handling and care of equipment such as microscopes and micropipettors, and unsafe practices such as not tying hair back, chewing gum, applying lipstick, eating, drinking, or chewing on pencils, and sloppy technique leading to poor results. As we are working with potential pathogens, students displaying improper or careless techniques will be asked to leave the lab and will have at least 5% of their laboratory grade deducted immediately. Missing a lab for which there is a skills test or assignment requires documentation. Upon presentation of this documentation, you will either have to complete the assignment or skills test as soon as possible or, if this is not possible, the marks may be added to your final exam. The lab books will be collected and graded twice during the semester. Although most exercises are completed as groups, the lab books are to be completed individually, and must represent individual effort. The following page provides you with tips on how to construct your books. Unannounced skills tests will be given during the semester. Students are expected to work independently on some technical aspect of microbiology and will be graded based on their techniques and their results. As proficiency in microbiological techniques is considered an essential component of the course, students are only permitted three lab period absences (you do not require any documentation). Missing more than three labs will result in a grade of 0 being assigned for the lab (at this point, it is recommended that students consult with Arts and Science Advising for the option of completing the laboratory the following year). Students are still responsible for the material missed (and their assignments, lab reports etc. will be graded as such). There are no make-up laboratories. Late Assignments will be penalised as follows: For Assignment 1 and the Lab Report: after the start of lab, but by 4:30 pm on the due date 25%; by 9:00 am the next morning -50%, and after 9:00 am the following day, no marks will be given. 3

5 Extensions for the lab report and Assignment 1 will only be granted for situations involving prolonged illness (documentation is required). The lab exam (April 10) is comprehensive, covering all aspects of the laboratory. It may contain a practical as well as a theoretical component. PREPARATION OF A LAB BOOK Your lab book provides you with a detailed record of your experiments performed. This record proves invaluable when preparing manuscripts for publication, or, more immediately, when preparing lab reports. This lab book, as with all of the reports and proposals is an individual effort. Choice of Lab Book Standard black lab books can be purchased from the book store but these are not required for this course. The only required features are; Pages are non-removable (no spiral bindings) All pages must be numbered in the top outer corner page numbers may be hand-written on EVERY page in INK In General all entries must be made in blue or black ink (except drawings) date EVERY entry never remove a page or use white-out if an entry needs to be deleted, strike out the entry with a single straight line (the deleted entry must be readable) keep up to date, a lab book is meant to be filled out as the experiments are carried out and NOT after the fact record anything that may be useful to you when preparing your lab reports leave plenty of space throughout the lab book to add comments after the fact Table of Contents Designate the first 2 pages as the Table of Contents record information and pages numbers as you go Lab Entries For each lab be sure to include the following; Objective Method Summary do not rewrite the protocol from the lab manual highlight any specific changes to the lab protocol include times and dates for when work was performed record product names and manufacturers used - enzymes, chemicals, equipment (micropipettors, baths) 11) include incubation conditions for cultures and reactions 4

6 Observations & Results record any & all observations, this goes beyond number results include diagrams and any other form of raw data include calculations as appropriate Conclusions 1) did you achieve your objective? Why or why not? 2) use your results to support your conclusions Answer the thought questions at the end of the lab (as applicable) 1) use reference citations as needed 2) these may be graded 5

7 THE UNIVERSITY OF LETHBRIDGE Policies and Procedures Occupational Health and Safety SUBJECT: CHEMICAL RELEASE PROCEDURE Precaution must be taken when approaching any chemical release. 1. Unknown/Known Release Clear the area Call Security 2345 Do not let anyone enter the area Call Utilities at 2600 and request the air be turned off at the release site Security will immediately notify: Chemical Release Officer: Occupational Health and Safety: EMERGENCY CALL LIST SECURITY CHEMICAL RELEASE OFFICER 2301 ADMIN. ASSISTANT OCCUPATIONAL HEALTH AND SAFETY EMERGENCY CALL LIST SECURITY CHEMICAL RELEASE OFFICER OCCUPATIONAL HEALTH AND SAFETY IF THE CHEMICAL RELEASE OFFICER CANNOT BE LOCATED CALL: DBS If the area must be evacuated all employees will be evacuated to the North Parking Lot. 6

8 GUIDELINES FOR SAFETY PROCEDURES EMERGENCY NUMBERS City Emergency 911 Campus Emergency 2345 Campus Security 2603 Student Health Centre 2484 (Emergency ) THE LABORATORY INSTRUCTOR MUST BE NOTIFIED AS SOON AS POSSIBLE AFTER THE INCIDENT IF NOT PRESENT AT THE TIME IT OCCURRED. EMERGENCY EQUIPMENT: Know the location of the following equipment, which will be indicated to you at the beginning of the first lab: 1) Closest emergency exit 2) Closest emergency telephone and emergency phone numbers 3) Closest fire alarm 4) Fire extinguisher and explanation of use 5) Safety showers and explanation of operation 6) Eyewash facilities and explanation of operation. 7) First aid kit GENERAL SAFETY REGULATIONS 1) Eating, drinking or gum chewing is prohibited in the laboratory. 2) Always wash your hands after entering and prior to leaving the laboratory. 3) Laboratory coats are required for all laboratories and must be stored in the lab when not in use. 4) Report equipment problems to instructor immediately. 5) Report all spills to the instructor immediately. 6) Long hair must be kept restrained to keep from being caught in equipment, Bunsen burners, chemicals, etc. 7

9 SPILLS Spill of ACID/BASE/TOXIN: Contact instructor immediately! BACTERIA SPILLS: If necessary, remove any contaminated clothing. Prevent anyone from going near the spill. Cover the spill with dilute bleach and leave for 10 minutes before wiping up. DISPOSAL Upright Blue Cardboard Boxes: CLEAN LAB GLASSWEAR - broken glass, Pasteur pipettes, etc. NO CHEMICAL, BIOLOGICAL, OR RADIOACTIVE MATERIALS. Orange Biohazard Bags: Petri plates, microfuge tubes, tips, plastic pipettes, etc. All of this material will be autoclaved prior to disposal. Bacterial Cultures: Tubes and flasks containing liquid cultures are placed in marked trays for autoclaving. Bacterial Slides Used microscope slides are placed into the trays of bleach found at the end of each of the laboratory benches. Liquid Chemicals: Place in labelled bottles in fume hood. 8

10 EXERCISE 1 INTRODUCTION TO MICROSCOPY Please review Appendices 1 and 9 MICROSCOPY To view microscopic organisms, their magnification is essential. The microscope is the instrument used to magnify microscopic images. Its function and some aspects of design are similar to those of telescopes although the microscope is designed to visualize very small close objects while telescopes magnify distant objects. Magnification is achieved by the refraction of light travelling though lenses, transparent devices with curved surfaces. In general, the degree of refraction, and hence, magnification, is determined by the degree of curvature. However, rather than using a single, severelycurved biconvex lens such as that of Leeuwenhoek's simple microscopes, Hooke determined that image clarity was improved through the use of a compound microscope, involving two (or more) separate lenses. Operation of the Compound Microscope Students should be familiar with all names and functions of the components of their compound light microscopes as demonstrated in Appendix 1. Properties of the Objective Lenses 1. Magnification Magnification is a measure of how big an object looks to your eye. The number of times that an object is magnified by the microscope is the product of the magnification of both the objective and ocular lenses. The magnification of the individual lenses is engraved on them. Your microscope is equipped with ocular lenses that magnify the specimen ten times (10X), and four objectives which magnify the specimen 4X, 10X, 40X, and 100X. Each lens system magnifies the object being viewed the same number of times in each dimension as the number engraved on the lens. When using a 10X objective, for instance, the specimen is magnified ten times in each dimension to give a primary or "aerial" image inside the body tube of the microscope. This image is then magnified an additional ten times by the ocular to give a virtual image that is 100 times larger than the object being viewed. 9

11 2. Resolution Resolution is a measure of how clearly details can be seen and is distinct from magnification. The resolving power of a lens system is its capacity for separating to the eye two points that are very close together. It is dependent upon the quality of the lens system and the wavelength of light employed in illumination. The white light (a combination of different wavelengths of visible light) used as the light source in the lab limits the resolving power of the 100X objective lens to about 0.25 µm. Objects smaller than 0.25 µm cannot be resolved even if magnification is increased. Spherical aberration (distortion caused by differential bending of light passing through different thicknesses of the lens center versus the margin) results from the air gap between the specimen and the objective lens. This problem can be eliminated by filling the air gap with immersion oil, formulated to have a refractive index similar to the glass used for cover slips and the microscope's objective lens. Use of immersion oil with a 100X special oil immersion objective lens can increase resolution to about 0.18 µm. Resolving power can be increased further to 0.17 µm if only the shorter (violet) wavelengths of visible light are used as the light source. This is the limit of resolution of the light microscope. The resolving power of each objective lens is described by a number engraved on the objective called the numerical aperture. Numerical aperture (NA) is calculated from physical properties of the lens and the angles from which light enters and leaves. Examine the three objective lenses. The NA of the 10X objective lens is Which objective lens is capable of the greatest resolving power? 3. Working Distance The working distance is measured as the distance between the lowest part of the objective lens and the top of the coverslip when the microscope is focused on a thin preparation. This distance is related to the individual properties of each objective. 4. Parfocal Objectives Most microscope objectives when firmly screwed in place are positioned so the microscope requires only fine adjustments for focusing when the magnification is changed. Objectives installed in this manner are said to be parfocal. 5. Depth of Focus The vertical distance of a specimen being viewed that remains in focus at any one time is called the depth of focus or depth of field. It is a different value for each of the objectives. As the microscope is focused up and down on a specimen, only a thin layer of the specimen is in focus at 10

12 one time. To see details in a specimen that is thicker than the depth of focus of a particular objective you must continuously focus up and down. Observing Bacteria Three fundamental properties of bacteria are size, shape and association. Bacteria generally occur in three shapes: coccus (round), bacillus (rod-shaped), and spirillum (spiral-shaped). Size of bacterial cells used in these labs varies from 0.5 µm to 1.0 µm in width and from 1.0 µm to 5.0 µm in length, although there is a range of sizes which bacteria demonstrate. Association refers to the organization of the numerous bacterial cells within a culture. Cells may occur singly with cells separating after division; showing random association. Cells may remain together after division for some interval resulting in the presence of pairs of cells. When cells remain together after more than a single division, clusters result. Cell divisions in a single plane result in chains of cells. If the plane of cell division of bacilli is longitudinal, a palisade results, resembling a picket fence. Both bacterial cell shape and association are usually constant for bacteria and hence, can be used for taxonomic identification. However, both properties may be influenced by culture condition and age. Further, some bacteria are quite variable in shape and association and this may also be diagnostic. Micrometry When studying bacteria or other microorganisms, it is often essential to evaluate the size of the organism. By tradition, the longest dimension (length) is generally stressed, although width is sometimes useful for identification or other study. Use of an Ocular Micrometer (Figure 1) An ocular micrometer can be used to measure the size of objects within the field of view. Unfortunately, the distance between the graduations of the ocular micrometer is an arbitrary measurement that only has meaning if the ocular micrometer is calibrated for the objective being used. 1) Place a micrometer slide on the stage and focus the scale using the 40x objective. 2) Turn the eyepiece until the graduations on the ocular scale are parallel with those on the micrometer slide scale and superimpose the micrometer scale. 3) Move the micrometer slide so that the first graduation on each scale coincides. 4) Look for another graduation on the ocular scale that exactly coincides with a graduation on the micrometer scale. 5) Count the number of graduations on the ocular scale and the number of graduations on the micrometer slide scale between and including the graduations that coincide. 6) Calibrate the ocular divisions for the 40x and the 100x objective lenses. Note that immersion oil is not necessary for calibration. 11

13 0 Stage Micrometer (each division = 0.01 mm) Ocular Micrometer Figure 1. Calibration of an ocular micrometer using a stage micrometer. The mark on the stage micrometer corresponding to 0.06 mm (60 µm) is equal to 5 ocular divisions (o.d.) on the ocular micrometer. 1 ocular division equals 60 µm/5 ocular divisions or 12 µm. Once an ocular micrometer has been calibrated, objects may be measured in ocular divisions and this number converted to µm using the conversion factor determined. Bacterial size is generally a highly heritable trait. Consequently, size is a key factor used in the identification of bacterial taxa. However, for some bacteria, cell size can be modified by nutritional factors such as culture media composition, environmental factors such as temperature, or other factors such as age. EXPERIMENTAL OBJECTIVE In this first exercise, you will calibrate the 40x and 100x objectives of your compound microscope. Then you will use the compound light microscope to assess the shape and associations of bacteria that have already been fixed to slides and stained. You will also use your determined calibration factors to evaluate sizes of organisms viewed. 12

14 METHODS: For each student: Compound light microscope Various prepared slides of bacteria. Stage micrometer Ocular micrometer Immersion oil 1) Use the diagram in Figure 1 to calibrate the 40x and the 100x objectives on your compound microscopes. Record these values in your lab book as you will then use these values when measuring cells and structures for the rest of the lab. Note: Do NOT use immersion oil when calibrating the 100x objective. This is the ONLY time during the term that you will not use immersion oil with this objective. 2) Use the compound microscope to observe the prepared slides of bacteria using the 10x and 40x objective lenses. Observe the same slides under the 100x objective using immersion oil. 3) Diagram two of the organisms viewed following the instructions found in Appendix 2. 13

15 EXERCISE 2 GENERAL LABORATORY PROCEDURES AND BIOSAFETY A primary feature of the microbiology laboratory is that living organisms are employed as part of the experiment. Most of the microorganisms are harmless; however, whether they are nonpathogenic or pathogenic, the microorganisms are treated with the same respect to assure that personal safety in the laboratory is maintained. Careful attention to technique is essential at all times. Care must always be taken to prevent the contamination of the environment from the cultures used in the exercises and to prevent the possibility of the people working in the laboratory from becoming contaminated. Ensure that you have read over the guidelines on Safety, and those on Aseptic technique (Appendix 3). As well, you should be familiar with the contents of the University of Lethbridge Biosafety web site: EXPERIMENTAL OBJECTIVES Students will use fluorescein dye-labelled E. coli cultures to perform a series of exercises designed to illustrate the potential for contamination that is always present when working with microorganisms. As well, students will become familiar with using aseptic techniques to handle microorganisms. METHODS Benches will be provided with the following: Fluorescein-labelled broth culture of E. coli (ATCC strain)(2/bench) Nutrient agar plates (8/bench) Nutrient broth (4 tubes/bench) Bench coat Tape Gloves Hand-held UV lamp Watch glasses (2/bench) Sterile pipettes Pipette pump Tray containing bleach disinfectant Wear gloves for the entire exercise. 1) Tape bench coat onto the bench to cover your working surface. 2) Work individually over the bench coat and prepare a streak plate for single colonies. Label and place in the tray on the side to be incubated. 3) From the same suspension, inoculate one tube of nutrient broth. For steps 4-11, work in pairs. 4) Place a watch glass in the centre of the bench coat. 14

16 5) Obtain and label 2 NA plates (name, date, organism, distance). Place agar plates on either side of the glass plate, one 5 cm and the other 10 cm from the watch glass. 6) Using a pipette pump, draw up 2 ml of bacteria/fluorescein suspension. 7) Remove lids from agar plates and set aside. 8) Hold pipette tip 30 cm from glass plate and allow 10 drops to fall (one drop at a time) onto the glass plate. Put any remaining bacterial culture back into the original culture tube. 9) Remove glass plate to disinfectant tray and cover agar plates. Place on a tray on the side bench. 10) Use the hand-held UV lamp in C741 to inspect your bench coat, gloves, and lab coat. What do you observe? 11) Your plates will be incubated for hours at 37 o C, and then refrigerated at 4 o C. During the next laboratory period, evaluate your plate results and record the number of colonies present. Thought Questions: (Use the Biosafety Web Site as a reference) What is an MSDS and where can you find one? In Canada, the Laboratory Centre for Disease Control has classified infectious agents into 4 Risk Groups using pathogenicity, virulence and mode of transmission (among others) as criteria. What do these terms mean? What criteria would characterise an organism classified in Risk Group 1, 2 3 or 4? Provide an example of an organism found within each group. There are many Golden Rules for Biosafety. Identify 4 common sense practices that will protect you in your microbiology labs. 15

17 EXERCISE 3 BACTERIAL and YEAST MORPHOLOGY The Microscopic Examination of Bacteria Prior to viewing bacteria, two procedures must be performed: 1) fixation and 2) staining. Fixation performs 2 functions: (i) immobilises (kills) the bacteria; and (ii) affixes them to the slide. The most common fixation procedure for bacteria is heat fixation, whereby the slide containing a drop or smear of bacterial culture is passed rapidly once or twice through the heat of a Bunsen flame. Staining Bacteria are almost transparent and hence, unstained bacteria are not readily visible without special techniques such as phase contrast microscopy (see: Madigan and Martinko, 2006, pp ) or dark-field microscopy, which is also referred to as negative staining. Any procedure that results in the staining of whole cells or cell parts is referred to as positive staining. Most positive stains used involve basic dyes where basic means that they owe their coloured properties to a cation (positively charged molecule). When all that is required is a general bacterial stain to show morphology, basic stains such as methylene blue or carbol fuchsin result in the staining of the entire bacterial cell. Differential stains are used to distinguish bacteria based on certain properties such as cell wall structure. Differential stains are useful for bacterial identification, contributing to information based on bacterial size, shape, and association. Differential staining relies on biochemical or structural differences between the groups that result in different affinities by various chromophores (Appendix 4). Gram staining behavior relies on differences in cell wall structure and biochemical composition. Some bacteria when treated with para-rosaniline dyes and iodine retain the stain when subsequently treated with a decolourising agent such as alcohol or acetone. Other bacteria lose the stain. Based on this property, a contemporary of Pasteur, Hans Christian Gram, developed a rapid and extremely useful differential stain, which subsequently bears his name - the Gram stain (see Figure 4-4, page 59 in Madigan and Martinko, 2006) used to distinguish two types of bacteria, Gram positive and Gram negative. Gram negative forms, which are those that lose the stain on decolourisation, can be made visible by using a suitable counterstain. The strength of the Gram stain rests on its relatively unambiguous separation of bacterial types into two groups. However, variables such as culture condition, age or environmental condition, can influence Gram staining of some bacteria. 16

18 The bacterial cell wall is very important for many aspects of bacterial function and hence, the Gram stain also provides valuable information about the physiological, medicinal and even ecological aspects of the bacteria. Acid Fast Staining Members of the genus Mycobacterium contain groups of branched-chain hydroxy lipids called mycolic acids. Robert Koch first described this property; it allowed him to determine the organisms present in lesions resulting from tuberculosis. As a result of the presence of these lipids, these organisms are not readily stained via Gram staining. Instead, cells require heat treatment so that a basic fuchsin and phenol dye penetrate the lipids. Once stained, these lipids resist decolourisation when treated with acid. Poly-β-hydroxybutyric Acid (PHB) Staining PHB granules are common inclusion bodies in bacteria. Monomers of β-hydroxybutyric acid are connected by ester linkages forming long polymers which aggregate into granules. As these granules have an affinity for fat-soluble dyes such as Sudan black, they can be stained and then identified with the light microscope. These granules are storage depots for carbon and energy. Endospore Staining Certain bacteria may produce endospores under unfavourable environmental conditions. Endospores are mainly found in Gram-positive organisms, including the Gram-positive Clostridium and Bacillus, in the Gram-positive cocci Sporosarcina, and in some of the filamentous Gram-positive Monosporaceae family. It has also been discovered that Coxiella burnetii, a small rod found in raw milk that has a variable Gram stain reaction, but a typical Gram-negative cell wall has a sporogenic cycle. When conditions become more favourable, the endospores will germinate and the bacteria will return to the actively growing and dividing form. Endospores are highly resistant to heat, chemical disinfectants and to desiccation and therefore allow the bacterial endospore to survive much more rigorous conditions than the vegetative cells. Endospore resistance is due to several factors, including: A decrease in the amount of water compared to vegetative cells An increase in the amount of dipicolinic acid and calcium ions Enzymes which are more resistant to heat A spore coat which is impermeable to many substances Endospores may be formed in a central, terminal, or sub-terminal position in the cell and their shape varies from ellipsoidal to spherical. The location of the endospore in the cell is usually characteristic of the species. For example, the location and shape of the Bacillus 17

19 subtilis endospore is different from the location and shape of the Clostridium endospore. Therefore, the presence or absence of endospores and the description of the endospore is useful to a microbiologist as an aid in identification. The resistant properties of endospores make them difficult to stain, hence heat is used in conjunction with staining to enable the stain to penetrate into the spore coat. EXPERIMENTAL OBJECTIVE The objective of this series of exercises is to perform specialised staining procedures in order to examine different properties of microorganisms, both bacteria and yeast. These exercises will also reinforce proper techniques for handling of microorganisms. METHODS: For each bench: Stains Crystal violet Safranin 5% Malachite green Carbol fuchsin Methylene blue 20% Sulfuric acid Gram s iodine Sudan black 95% ethanol Hemo-D (in fume hood) Equipment microbiology kits compound microscopes slides Bacteria Mycobacterium smegmatis Bacillus thuringiensis Escherichia coli Staphylococcus epidermidis Yeast Saccharomyces bayanus Follow the guidelines for each stain as described below. Work individually. 18

20 Prepare scientific diagrams (Appendix 2) showing results from each stain. For each set of results, students should plan on looking up the correct reactions and/or morphological features using the resources available (ie your textbook, Dr. Selinger s web page) and using this information to evaluate their techniques. Preparation of Films for Staining Procedure Obtain a clean slide and draw a circle on it approximately 1.5 cm in diameter. Turn the slide over. Flick the tube of culture to mix up the cells, and use a loop to obtain aseptically a drop of culture. Place this loopful of culture within the circle. Alternatively, if using a plate culture, first use your loop to add a drop of water to the circle on the slide. Remove a small quantity of culture and mix with the water to make a smooth suspension. Allow the suspension to air dry. When dry, the film should be only faintly visible; a thick opaque film is useless. The only fixation required is to pass the slide several times (maximum 10) through the bunsen burner flame until the slide is warm but not too hot. If the slide is fixed until too hot to the touch, the bacteria will be misshapen when observed under the microscope. Gram Staining - Procedure Perform on Bacillus thuringiensis, Escherichia coli, and Staphylococcus epidermidis 3) Prepare smear, dry and heat fix. Flood the smear with crystal violet solution for 1 min. Gently wash with tap water for 2-3 seconds and remove the water by tapping the slide gently on paper towel. 4) Add Gram s iodine solution to the slide for 1 min. Wash gently with tap water and remove as above. 5) Decolourise with 95% ethanol by dripping ethanol on surface of slide until no more colour is removed. Rinse gently with water. If too much alcohol is added, the Gram-positive organisms may become Gram-negative. Remove the water after the last wash. 6) Counterstain the slide with safranin for 30 seconds - 1 minute. 7) Wash the slides with tap water, air dry on paper towels, and examine under oil immersion. Gram positive organisms stain purple; Gram negative organisms, red (pink). Acid-fast Staining - Procedure Perform on Mycobacterium smegmatis and on Escherichia coli 1) Flood the dried, heat fixed film with Ziehl s carbol fuchsin and place on the rack over the boiling water bath. 2) Steam gently for 5 minutes. Do not let the slide dry out. Add more carbol fuchsin as required. 19

21 3) Wash with tap water to remove excess stain. 4) Decolourise with 20% sulfuric acid until no more stain comes out. Wash with tap water to remove excess. 5) Counterstain with methylene blue for 1 minute. Note that the term counterstaining implies rinsing the excess stain off with water and blotting dry. Acid fast organisms retain the red stain while others are stained blue. PHB Staining - Procedure Perform on Bacillus thuringiensis. 1) Prepare smears of the organism, air dry and heat fix. Flood entire slide with Sudan Black B and add more stain as the dye solvent evaporates. Stain for at least 10 minutes. 2) Pour off excess stain (do not wash) and air dry. 3) Clear slide by dipping in a jar of solvent in the fume hood for 5 sec. Air dry in the fume hood. 4) Counterstain for 1 min. with safranin. 5) Wash with water, drain, blot and air dry. Examine with oil immersion objective. Cytoplasm is pink, lipids are dark grey or black. Endospore Staining - Procedure Perform on Bacillus thuringiensis. 1) Prepare smear and heat fix. Cover the dried fixed film with a small piece of paper towel. Saturate this with 5% malachite green. 2) Place the slide on a rack over a boiling water bath. Steam slide for 5-10 minutes in this manner. Add additional stain as needed - do not allow the slide to dry out during this procedure. 3) Allow the slide to cool, then rinse with water. Tap over a paper towel to remove excess water 4) Counterstain with safranin for 30 seconds. 5) Rinse slide with water. 6) Allow to air dry, and view. Endospores will stain green and the rest of the cell pink. Yeast Staining Procedure Perform on Saccharomyces bayanus 1) Prepare a wet mount of the cells using a drop of Methylene Blue. 2) Carefully place a cover slip on the cell/stain mixture. 3) View the cells noting size and shape. If you look carefully, you should be able to see budding cells. 20

22 Thought Questions: Why do we stain microorganisms before viewing them with a microscope? What is a differential stain? Give two examples of differential stains used in Biology 3200 labs. Why is immersion oil used to view microscopic organisms? Gram stains separate microorganisms into two major groups: Gram negative bacteria and Gram positive bacteria. Describe the differences in the structure of the cell wall of each type of bacteria that results in the differential stain result. What are endospores? How do they form? Which organisms can produce endospores? What is the mode of transmission of acid fast organisms? Relate the mode of transmission to the cell wall structure. References: Atlas, R. M Principles of Microbiology. Wm. C. Brown Publishers, Toronto. Madigan, M. T., and Martinko, Brock Biology of Microorganisms Eleventh Edition. Prentice-Hall of Canada, Inc., Toronto. Ross, H Microbiology 241 Laboratory Manual. The University of Calgary Press, Calgary. 21

23 EXERCISE 4 THE AMES TEST MUTATION AND RECOMBINATION (See Madigan and Martinko, Chapter 10 Pg ) You have learned about some of the advantages of using a model system in your study of the effect of UV light on DNA in Biology 2000 (Introduction to Genetics). The Ames test also makes use of a model system in order to measure the mutagenic potential of compounds. This test is a reversion mutagenesis assay and uses strains of the bacterium Salmonella that have point mutations in various genes in the histidine operon. These His - mutants are unable to synthesise histidine and therefore unable to grow on minimal media lacking histidine. When the His - tester cells are cultured on a minimal agar medium containing trace amounts of histidine, a small and relatively constant number of cells per plate spontaneously revert to His + and subsequently reproduce and form colonies. Incorporation of a mutagen into the agar increases the number of revertant colonies per plate, usually in a dose dependent manner. EXPERIMENTAL OBJECTIVE You will make use of the Ames test in order to evaluate the mutagenicity of a selection of compounds. PRE-LAB PREPARATION Each class should bring in a total of three household compounds they would like to test. These will be decided in advance. Note that these compounds must be known (ie mystery liquid from the garage is not acceptable) and they must be taken home again once Period 1 of the lab is finished. METHODS: For each lab: 100 mg/ml Sodium Azide (CAUTION: MUTAGEN!) Hair Dye (Miss Clairol) Micro Kits Gloves Sterile water 3x Liquid cultures of Salmonella strains 1535 and 1538 in NB supplemented with NaCl Top agar overlay in 50 o C water bath (2 ml per tube) Test tube with 2 ml mark indicated (at pouring station) Minimal salts plates (15 per lab) Vortex mixer (at pouring station) Bunsen burner (at pouring station) Test tube racks 22

24 Sterile filter paper disks Forceps 3x micropipettors ( µl) Sterile tips 5x beakers with biohazard bags Small vials containing 95% ethanol for flaming Set up your experiment as follows in the Table: Bench # Compound to be Tested Water Unknown Unknown Unknown Sodium Azide Hair Dye ) For each plate, you will be creating an overlay using a single strain mixed with the top agar. The top agar has had a trace amount of histidine and biotin added. Using the Table as a guide, obtain and label the appropriate number of minimal salts plates. Why is it necessary to add a trace amount of histidine to the top agar? 2) Have your plates labelled, and take to the station set up at the back bench. Set a micropipettor to 50 µl. Remove one tube of agar overlay from the waterbath, and aseptically add 50 µl of liquid culture to the tube. Vortex to mix and pour over the surface of your agar plate. Clean up your work surface prior to going back to your bench. Note: you must work very quickly in order to avoid the top agar solidifying. 3) Allow your agar to solidify for 10 minutes. Wear gloves for any handling of the potential mutagens! 4) Flame forceps to sterilise. Note that this does not mean holding forceps in the flame of your Bunsen burner until red hot! Rather, dip the forceps in ethanol, and wave through the flame. Allow the ethanol to burn off. Pick up a sterile filter paper disk and dip in the appropriate mutagen. 23

25 5) Tap the filter paper several times to remove excess liquid. Hold the filter paper for a few moments to ensure that liquid doesn t drip all over your plates. Place the filter paper in the centre of the plate with the solidified overlay. Tap gently to ensure that the filter paper stays in place. 5) Incubate your plates for 48 hours at 37 o C. In the next lab, enumerate the number of colonies on each plate and record the results on the board. Prepare a lab report based on your class results using the information found in Appendix 6. Thought Questions: a. What specific mutations in the His operon do each of the Salmonella strains used contain? b. Evaluate the compounds tested for mutagenicity. What kind of mutations are being caused by the compounds tested? (use the information from the first Thought Question to answer this) c. Typically, mutagens are first mixed with liver extract prior to carrying out the Ames test. What would be the purpose of this step? References: Ames, B.N., Durston, W.E., Yamasaki, E., and Lee, F.E Carcinogens are mutagens: a simple test combining liver homogenates for activation and bacteria for detection. Proc. Natl. Acad. Sci. U.S.A. 70: Ames, B.N., Lee, F.E., and Durston, W.E An improved bacterial test system for the detection and classification of mutagens and carcinogens. Proc. Natl. Acad. Sci. U.S.A. 70: Ames, B.N., McCann, J., and Yamasaki, E Methods for detecting carcinogens and mutagens with the Salmonella-microsome mutagenicity test. Mutational Research 31: Madigan, M. T., and Martinko, Brock Biology of Microorganisms Eleventh Edition. Prentice-Hall of Canada, Inc., Toronto. 24

26 EXERCISE 5 BIOCHEMICAL TESTS (Selective and Differential Media; IMViC Tests) Normally, the coliform group of bacteria is used to indicate the pollution of water with fecal wastes of humans and animals, and thus, the suitability of a particular water supply for domestic use. The term coliform is used to describe aerobic and facultatively anaerobic Gram negative rods that ferment lactose with gas formation. Most, but not all organisms within this group are intestinal in origin; for instance, Escherichia coli. Consequently, presence of lactose fermentors in a sample of water provides circumstantial evidence of pollution by fecal wastes, and may suggest the presence of pathogenic bacteria such as members of the genera Salmonella and Shigella. These pathogens, in addition to non-pathogens such as E. coli are members of the Enterobacteriaceae family. In order to identify the organisms present in the water, several biochemical tests that rely on differences in the chemical composition of media used may be performed (see Appendix 4 and Appendix 8 for more details). SELECTIVE AND DIFFERENTIAL MEDIA: I. Media for Isolation of Enterobacteriaceae A strategy for bacterial isolation involves the use of selective media, media with specific components that promote the growth of some bacteria and inhibit the growth of others. Selectivity may be achieved in three ways: by adding something to the medium to discourage the growth of species not required by altering the ph of the medium by omission of some ingredient required by most bacteria, but not by the organism to be isolated Differential media contain specific biochemical indicators that demonstrate the presence of certain substances characteristic of certain bacteria. Thus, differential media are useful for bacterial identification. Eosin Methylene Blue Agar (EMB Agar) EMB is both a differential and selective plating medium recommended for use in the isolation of Gram-negative bacilli and the differentiation of lactose fermentors from non-lactose fermentors. EMB agar contains the two indicators, eosin Y and methylene blue as well as the carbohydrate lactose. Eosin (an acidic dye) reacts with methylene blue (a basic stain) to form a compound of either acidic or neutral nature. The acid produced by lactose fermentors is sufficient to cause this dye compound to be taken up by the cells. Non-lactose fermentors are colourless because the eosin and methylene blue compound cannot be taken up by the cells. The basic stain methylene blue inhibits bacterial growth, particularly that of Gram positive 25

27 bacteria (due to their cell wall composition). Eosin methylene blue (EMB) agar is thus selective for Gram negative bacteria. MacConkey Agar MacConkey agar is a differential and selective plating medium recommended for use in the isolation of Gram-negative bacilli and the differentiation of lactose fermentors from nonlactose fermentors. The differential action of the MacConkey agar is indicated by the colonies of coliform bacteria becoming brick red in colour. This occurs when the coliforms utilise the lactose producing acids. The decrease in ph results in the uptake of the indicator neutral red by the cells. Non-lactose fermentors are colourless and transparent. Production of acid may also result in a zone of precipitated bile surrounding the colony. Bile salts and crystal violet present in the medium inhibit Gram-positive bacteria from growing. II. Acid Production From Carbohydrates As demonstrated with MacConkey Agar, bacteria vary in their ability to ferment various sugars. Products of fermentation are often acids and hence, ph changes can demonstrate successful fermentation. In addition, gas (usually but not always CO 2 ) is often produced during fermentation, offering another indicator. Hugh and Leifson's method for demonstrating the presence of the products of fermentation consists of a semi-solid medium containing peptone (short chains of amino acids), the carbohydrate of interest (usually glucose or lactose), and a ph indicator, Bromothymol blue. Tubes are stab-inoculated all the way to the bottom of the tube, so as not to introduce oxygen into the medium. Several reactions may be observed. Facultative organisms will produce an acid reaction (the indicator changes to yellow) throughout the entire tube of medium. The acid reaction produced by oxidative organisms is apparent first at the surface, extending gradually downwards into the medium. Note that organisms that oxidize glucose are generally unable to ferment any carbohydrate. Strict fermentors will produce an acid reaction at the bottom of the tube. Organisms unable to use the carbohydrate may be able to grow using the peptone. Production of alkaline products result in the formation of a blue colour at the top of the tube (although this does not indicate that the organism is aerobic). III. Motility Medium This medium contains triphenyl tetrazolium chloride (TTC) and a small concentration of agar in order to make the medium semi-solid. TTC is reduced when broken down by the organism, and the TTC turns red where this has occurred. If the organism is facultative and motile, it moves throughout the entire tube of medium and the whole tube becomes red. If the organism is aerobic and motile, the top of the tube becomes red. 26

28 METHODS For each bench: 3 plates each of MacConkey and EMB media 5 known broth cultures 1 'unknown' broth culture 6 tubes of Hugh and Leifson's (H & L) lactose medium 6 tubes of motility medium Please work in groups of four. 3) Divide your three MacConkey and three EMB plates in half and streak inoculate them with the six bacterial species provided. After incubation at 37 C for 48 hours, observe, and describe the various cultures on the plates. Generate a table of results summarizing growth and properties of all bacteria on the two media. 4) Work collectively to determine the lactose fermentation ability of all of the bacteria provided. These tubes are inoculated using a stab technique. Use the probe to remove aseptically a small amount of bacterial culture, then stab the probe to the bottom of the tube of medium without mixing the medium around. Inoculate each tube with one of the bacterial species and label appropriately. Tubes will be incubated for 48 h at 37 C. After incubation, observe tubes and record the results. 5) Work collectively to inoculate your motility medium tubes. Again, as this medium is semi-solid, use a probe and stab the culture down to the bottom of the tube, and remove the probe carefully. Do not mix the probe around in the tube. Tubes will be incubated for 48 h at 37 C. After incubation, observe tubes and record the results. IMViC TESTS Only preliminary taxonomic assessment of bacteria can be made on the basis of microscopic size, shape, association, and Gram staining. Information regarding natural occurrence is also valuable since bacteria generally occur in specific habitats. This is particularly the case for fastidious bacteria, those with very specific nutritional and environmental requirements. However, even when supplemented with habitat information, bacterial identification based on microscopic assessment is generally incomplete. Confident bacterial identification can be made based on biochemical tests, and for certain pathogens, or for examining microbial presence in specific environments, series of diagnostic tests have been developed. For example, the IMViC tests are used routinely to confirm the presence of coliform organisms in water. IMViC is an acronym for Indole, Methyl Red, Voges-Proskauer, and Citrate utilisation tests (the i is inserted for ease of pronunciation). 27

29 I. Indole Formation - Utilisation of Tryptophan When cultured on peptone water, a liquid medium containing tryptophan, certain bacteria will produce indole. The presence of this indole is readily revealed through addition of Kovak's reagent, producing a pink colour. This reagent contains the organic solvent amyl alcohol that extracts the coloured (pink) substance. II. The Methyl Red (MR) Test - Mixed Acid Fermentation Pathway The mixed acid fermentation pathway results in the formation of a number of organic acids such as lactic and acetic acid. If this is a primary fermentation pathway of a bacterium, a noticeable drop in ph will occur with incubation on MRVP media. This decrease in ph can be revealed by a methyl red solution which is yellow under neutral conditions and red at a ph less than 5. III. The Voges-Proskauer (VP) Test - The Butanediol Fermentation Pathway An alternate fermentation pathway performed by some other bacteria results in the formation of a non-acidic product, butanediol and hence, is named for this product. The occurrence of the pathway may be determined by a biochemical test for an intermediate compound in the pathway, acetoin (acetyl methyl carbinol), which is detected by the Voges-Proskauer test. IV. Citrate Utilisation - Growth Using A Single Carbon Source The nutritional requirements of different bacteria vary considerably and these can provide useful information contributing to biochemical identification. In Simmon's citrate agar, citrate, in the form of sodium citrate, is the sole carbon source. Organisms able to utilise the citrate grow on the surface of the medium and due to oxidative formation of sodium carbonate, raises the ph of the medium changing it from green to blue (bromothymol blue is the indicator). V. Urea Hydrolysis Some bacteria can produce urease, an enzyme which hydrolyses urea into ammonium and carbon dioxide. The presence of this enzyme is detected by growing the bacteria in a medium containing urea and a ph indicator, phenol red. If ammonium is produced as a result of urea hydrolysis, the increase in ph will turn the medium to a violet-red colour. 28

30 METHODS: For each bench: 6 broth cultures, one of which is an Unknown 6 MRVP broth tubes 6 indole broth tubes 6 Simmons citrate agar slants 6 Urea broth tubes Please work in groups of four. 1) Inoculate 6 Indole broth tubes separately with the 6 bacteria. After 48h of incubation at 37 o C, add 20 drops (1 ml) of Kovak's reagent. Shake and look for the formation of a pink colour in the top (organic) phase; it may take 20 minutes to develop. The pink colour is a positive result, indicating the ability to use tryptophan. Note, please place tubes containing amyl alcohol in a separate rack on the side bench as this material needs to be disposed of separately. 2) A single culture solution (peptone, glucose, potassium phosphate) will be used for both the methyl red and Voges-Proskauer tests. Inoculate 6 MRVP tubes with the 6 bacteria provided, one culture into each tube. After 48h of incubation at 37 C, remove about 1/4 of the broth (=2 ml = 40 drops) from the MRVP tube and transfer that to another test tube. Add 3-5 drops of methyl red solution. An immediate red reaction provides a positive response to the test, indicating the presence of mixed acid fermentation. A yellow or orange colour represents a negative response. As the same solutions are used for the MR and VP, remove an additional 2 ml of culture solution and add 1 ml α-napthol (Barritt's reagent A - 1 ml is about 20 drops) and 1 ml 40% KOH (Barritt s reagent B; caution - this is caustic). Shake vigorously for 30 seconds. Shake the tubes frequently and observe for up to 30 minutes for the formation of a red colour that represents a positive VP test. A yellow or brown colour is a negative result. 3) Inoculate 6 Simmon's citrate agar slants separately with the 6 bacteria. For these inoculations, smear cells along the surface of the slant. Incubate tubes for 48 h at 37 C. 29

31 After incubation, observe colours on the surface and down through the tubes. A dark blue colour is a positive result while green indicates a negative test for citrate utilisation. 6) Inoculate 6 urea broth tubes separately with the 6 bacteria. After incubation for 48h at 37 C, observe for the development of a violet-red colour. 7) After completing the Indole, MR, VP, Citrate and Urea tests, collaborate with the other students at your bench to generate in your lab books tables of results for all bacteria in all tests. Thought Questions: Compare and contrast chemically defined and complex media. Provide two examples of complex media used in this exercise and explain why these media are considered complex. Provide 2 examples of compounds responsible for buffering in media. Is agar a nutritionally complete substrate for microbes? Why or why not? Design a defined medium for an organism that can grow aerobically on acetate as a carbon and energy source. In this laboratory, would you classify the organisms used as photoautotrophs, photoheterotrophs, chemoautotrophs, or chemoheterotrophs? Explain your choice(s). Identify your unknown. Provide evidence to support your choice of organisms. 30

32 EXERCISE 6 BACTERIAL REPRODUCTION MEASUREMENT OF BACTERIAL GROWTH (See Madigan, M. T and Martinko, J. M., Chapter 6) Most bacteria reproduce by an asexual process called binary fission. In this process a single mother cell produces two identical daughter cells. Cell growth is often equated with increase in cell number due to the difficulty in measuring changes in cell size. Under ideal conditions populations of bacterial cells grow exponentially as cell number doubles at a regular interval or generation time (t d ). In the laboratory, pure cultures are routinely grown as batch cultures in test tubes and Erlenmeyer flasks. A batch culture is prepared by inoculating a fixed amount of liquid medium with the bacteria then the resulting culture is incubated for an appropriate period of time with no further addition of microorganisms or growth substrates. Cell growth in batch cultures can be divided into four phases. Initially the culture is in a lag phase where cells are preparing to reproduce. During this time cells are adjusting their metabolism to prepare for a new cycle of growth. There is an increase in cell size without increasing numbers. As cells begin to divide and their growth approaches the maximal rate for the particular set of incubation conditions established, the culture enters the exponential growth phase (log phase). One cell gives rise to two, two cells give rise to four, and so on. In this phase, cells are growing and dividing at the maximum growth rate possible for the medium and incubation conditions. Growth rate is determined by a number of factors, including available nutrients, temperature, ph, oxygen and other physical parameters as well as genetic determinants. As nutrients become limiting or waste products accumulate, the growth rate once again slows and the culture enters the stationary phase. During this phase, there is no further net increase in cell number, as growth rate equals the rate of cell death. The final phase of a batch culture is the death phase. During this phase, there is an exponential decline in viable cell numbers. This decline may be reversed if environmental parameters are modified by the addition of nutrients, for example. The rate of growth of bacterial cells is usually monitored by measuring the increase in cell number. Bacterial cell numbers may be enumerated by a number of methods. Direct count methods enumerate all cells whether they are viable or not. The most common direct count method uses a microscope and a specialized counting chamber (e.g., Petroff-Hauser chamber) to count the number of cells in a known volume of culture. Automated systems such as Coulter counters may also be used to determine cell number. In contrast, indirect count methods require the growth of cells in culture in order to enumerate cell numbers. The most common method for enumerating living cells is the viable plate count. 31

33 Serial dilutions of a cell suspension are prepared and spread on to the surface of a solid agar medium (spread plate) or incorporated into molten agar that is then poured into sterile petri dishes (pour plate). Following a suitable incubation time, the number of colonies growing on and in the inoculated agar are counted and used to determine the number of viable cells in the original suspension. This method makes the assumption that each colony arose from a single viable cell or colony forming unit (CFU). Turbidimetric methods can be used to rapidly assess biomass (e.g., cell numbers). The amount of light passing through a cell suspension can be determined with a spectrophotometer. The optical density (OD) is a measure of the amount of light passing through the suspension. A calibration curve can be generated using suspensions of known numbers of bacteria. EXPERIMENTAL OBJECTIVE In this experiment you will monitor the growth of E. coli cultures using the viable count and turbidimetric methods. You will determine the number of bacteria (CFU) present following various time points of incubation. You will establish a growth curve and a calibration curve for OD using the viable count data you collect. Prelab preparation: Turn on the spectrophotometer and set to 600 nm at least 15 minutes prior to taking readings. METHODS 100 ml in 125 ml bottles of molten Luria-Bertani (LB) agar in 65 C waterbath Ethanol/dettol in bottles 5x bleach trays Test tube racks Sterile Petri dishes Sterile 5 ml pipettes Pipette pump µl micropipettor µl micropipettor Sterile tips for micropipettors Container of sterile microfuge tubes Microfuge tube racks Sterile d 2 H 2 O Spectrophotometer blank containing TB broth Bacterial waste container Vortex Cuvettes Spectrophotometer Culture flasks of E. coli (200 ml volume) 32

34 Please work in groups of four. At 20 minute intervals, and again periodically during the afternoon, monitor the growth of your E. coli culture by determining viable counts as well as optical density following the procedures outlined below. A. Culture sampling 1) Each group of four will be assigned a culture flask. Please mark the flask with your bench number and lab number. Groups in laboratory sections 3 and 4 will continue to sample from the flask corresponding to your bench. Data from all four lab sections will be pooled and posted on the Biology 3200 web site. 2) Everyone in the laboratory will be sampling at the same time. Samples will be collected three times during your regularly scheduled lab period at 20 minute intervals, and again during the afternoon. For labs 1 and 2, these correspond to: 9:45 am, 10:05 am, 10:25 am, and for labs 3 and 4: 11:10 am, 11:30 am, and 11:50 am. Afternoon sampling times will be announced during the lab period. Your laboratory instructor will set a timer so that everyone is coordinated. Prior to beginning, designate two individuals in your group to be responsible for obtaining optical density (OD) readings at each time point. The other two individuals will prepare and plate appropriate serial dilutions for viable counts. 3) At 20 minute intervals aseptically obtain one 5 ml sample of culture and immediately place it in a spectrophotometer tube. This material will be used to measure optical density (OD) (Section B). 4) Remove another 100 µl of the culture and place it into a sterile microfuge tube. Label this tube Tube 1. Use this culture for Section C. B. Determination of optical density (please read Appendix 7) 1) Zero the spectrophotometer as outlined in Appendix 7. 2) Place the spectrophotometer tube containing your culture into the spectrophotometer and record the optical density (Absorbance) reading in your lab book and in the table on the blackboard. If the reading is greater than 0.7, you must dilute your sample and remeasure the optical density. It is suggested that you begin by diluting your sample 1:1 with the TB or LB provided (use the medium that corresponds to your bacterial culture). Make note of the dilution that you prepare in order to obtain an accurate absorbance reading. Multiply the absorbance by the dilution factor to obtain the final reading. 3) After reading, dispose of your 5 ml sample of culture in the waste beaker provided. Place the spectrophotometer tube into the bleach tray. 33

35 C. Enumeration of viable bacteria 1) Remove four sterile microfuge tubes from the container on the side bench. In order that you don t contaminate all of the tubes, gently tap out four tubes from the container rather than using your hand to grab tubes. 2) Set up your serial dilutions according to the information in Table 4.1. Aseptically pipette 900 µl of TB (or LB) into Tube 1 that already contains 100 µl of bacterial culture. You have now created a 1:10 dilution. Mix well using the vortex mixer. Create the remaining serial dilutions (tubes 2-4) in the same manner. Use fresh tips for each transfer. Table 4.1. Preparation of serial dilutions from E. coli culture sampled at 20 minute intervals Tube Number Amount of sterile TB or LB (µl) Amount of Culture µl from culture flask µl from tube µl from tube µl from tube 3 5 (for labs µl from 3 and 4 tube 4 only) Final Dilution Factor The dilution sequence will be set up each time you take a sample from your culture flask. 3) Labs 1 and 2 will be plating the contents of Tube 3 and Tube 4 (10-5 and 10-6 dilutions) FOR ALL of the time points at which these labs are sampling. Labs 3 and 4 will be plating the contents of Tube 4 and Tube 5 (10-6 and 10-7 ) FOR ALL of the time points at which these labs are sampling. Obtain 2 sterile Petri dishes. Label the bottom (not the lid) of the plate with the time the sample was taken, your group name, and the dilution. 20 ml corresponds to where the bottom edge of the lid is when the lid is on the Petri dish. 4) Add the contents of Tube 3 to the appropriately labelled sterile Petri dish. Obtain a bottle of molten LB agar from the water bath at the side of the lab, and add approximately 20 ml of molten agar (after flaming the mouth of the bottle) to the diluted culture. Swirl carefully to mix the inoculum evenly with the medium. Label the bottle of molten agar with your group name and replace it immediately in the water bath. 34

36 5) Follow the instructions provided in step 4 above to plate out the contents of Tube 4. 6) When the agar has solidified, place the inverted plates on a tray at the side of the lab. The plates will be incubated for hours at 37 C and refrigerated until the next lab session. 7) Your lab instructor will provide you with details about further sampling times during the remainder of the day. It will be necessary to sample outside of the lab period in order to capture more of the growth curve than could be obtained by in-lab sampling alone. The next laboratory period: 8) Examine the plates carefully and select the plate where the bacterial count ranges between 30 and 300 colonies. 9) Record the number of colonies on the plates in your lab notebook and in the chart on the board. Complete data sets will be available on the Biology 3200 web site. 10) Use class data to determine the average number of bacteria per ml of culture. 11) Prepare graphs from class data comparing i) OD vs time (on semi-log graph paper); 2) CFU/mL vs time (on semi-log graph paper); 3) OD vs CFU/mL (on arithmetic graph paper). The first two graphs are growth curves; the third graph is a standard curve allowing for correlation between OD and CFU/mL (Please see Madigan and Martinko, 2006 Chapter 6) Thought Questions: Use your graph(s) to calculate generation time of E. coli in LB vs TB. Compare your value to that from the literature. Do the values differ? Why might this be? Compare and contrast indirect and direct methods of counting bacteria. Use your standard calibration curve to calculate the CFU/mL of culture for an undiluted sample in which the OD was

37 EXERCISE 7 VIROLOGY (Please review the material on sewage treatment posted on the Biology 3200 web page) EXPERIMENTAL OBJECTIVE The objectives of this series of exercises are first to isolate coliphage from filtered raw and treated sewage obtained from the Lethbridge Wastewater Treatment Plant, to examine the plaque morphologies, and to prepare phage isolate from one particular plaque. Using this phage isolate, the phage titre will be determined, and the host specificity of the phage will be examined using several enteric bacterial strains. These exercises will demonstrate standard techniques in phage isolation and manipulation. Prior to the laboratory, sewage samples were collected at the areas indicated on the schematic posted on the web page. Both samples were stored at 4 o C prior to filtering, for up to 1 week. On the morning of the lab, samples were filtered twice using 0.45 µm filters. PART A - ISOLATION METHODS: For each bench: Luria Methylene Blue agar plates Overnight culture of Escherichia coli K12 Bottle of molten Luria agar overlay (at 60 o C) Sterile test tubes Test tube rack Micropipettor (100 µl 1000 µl) Sterile tips Microbiology kits For the lab: Vortex mixer Water bath set to 60 o C Raw and treated sewage filtrate Test tube showing 4 ml mark Work in groups of 4. Note that sewage filtrate contains human pathogens. Work very carefully. Students who are clearly unprepared or are sloppy will be asked to leave the lab. Procedure 1) Obtain a tube of culture of E. coli K12. 36

38 2) Obtain 5 Luria Methylene Blue agar plates, and 5 sterile test tubes. Label your 5 tubes according to Table 7.1. Table 7.1 Experimental set-up for isolation of coliphage from sewage. Tube Contents (µl) # K12 Ra w Se wage Filtrate Treated Se wage Filtrate ) Pipette the appropriate amount of filtrate and/or cells into each of your labeled test tubes. Leave the tubes at room temperature on your bench to incubate for 20 minutes to allow the phage to adsorb to the cells. 4) While your cultures are incubating, label your Luria Methylene Blue plates according to Table 7.1. Mark the level of 4 ml on each of your tubes using the marked test tube on the side bench as a guide. 5) Starting with Tube 1, aseptically pour molten agar into the tube up to the level of 4 ml. Vortex to mix, then immediately pour the contents over the surface of the appropriately labeled plate. Swirl the plate gently to ensure that the entire surface is covered with the agar. 6) Repeat step 5 for the remaining tubes and plates. 7) After 10 minutes, the overlay should be set. Invert your plates and place them on a tray on the side bench to be incubated. Plates will be incubated at 37 C for hours, then stored at 4 C until the next laboratory period. The next laboratory: Work in groups of four. MATERIALS Pasteur pipettes Bulbs Chloroform (in the fume hood) Vortex mixer Phage dilution buffer 37

39 Plates from last lab 1 dissecting microscope per bench Microfuge tubes (sterile) 1 ml pipettes and propipettors Microfuge racks Labeled microfuge rack on the side bench for class tubes 5) Obtain your plates. Examine them carefully. Record the number of plaques present for both raw and treated filtrate. Is there any difference? 6) Make detailed observations of plaque morphology. Features to look for include size, shape, and turbidity (clear vs cloudy). Use the dissecting microscopes for your observations. 7) After making observations, obtain a microfuge tube and aseptically add 1 ml of phage dilution buffer to your tube. Label with your group designation. 8) Use a Pasteur pipette (with a rubber bulb attached) to remove a plaque (squeeze the bulb, insert pipette into the agar over a plaque, gently release bulb to remove a plug of agar containing the plaque). Note that for each group of 4, two morphologically distinct plaques should be chosen. Release plaque into the prepared tube of phage dilution buffer. 9) Vortex vigourously to disperse the agar. 10) Move to the fume hood and use a Pasteur pipette to add a drop of chloroform to your tube. Vortex the mixture once again. What does the chloroform do? Place your tubes in the rack on the side bench. The tubes will be stored at 4 o C allowing the phage to elute from the agar into the buffer. PART B HOST RANGE METHODS Overnight cultures of: Salmonella typhimurium strain 1535 E. coli strains CSH121 and CSH125 and K12 Proteus vulgaris Enterobacter Other supplies: Phage dilution buffer Micropipettors and sterile tips Autoclave waste disposal Luria Methylene Blue agar plates LB plates Bottle of molten Luria agar overlay (at 60 o C) Sterile test tubes Test tube indicating 4 ml mark Test tube rack 38

40 Micropipettor (100 µl 1000 µl) Sterile tips Microbiology kits For determining phage titre: 1) Prepare serial dilutions of your phage in dilution buffer (10-2, 10-4, 10-6, 10-8 ) in microfuge tubes. Vortex each tube as you create each dilution. Ensure that you use fresh tips for each transfer. 2) In separate, labeled sterile test tubes, mix 500 µl of each dilution with 500 µl of host strain E. coli K12. Sit for 20 minutes of incubation time at room temperature. Mark the 4 ml mark on each test tube while mixtures are incubating. 3) Plate your mixtures as per Part A of this exercise. 4) The next day, count plaques and determine the titre of your phage. For determining host range: 1) Prepare spread plates on LB for each organism to be tested. (use the instructions found in Appendix 3, although this should be a review from previous courses!). Label each plate clearly. Use 100 µl of liquid culture to create a uniform lawn. 2) When lawns are dry, divide plates into four quadrants. In each quadrant, spot 20 µl of each phage dilution. Do not invert. Plates will be incubated at 37 o C overnight. 3) The next day, score as + or for phage growth on each host. Thought Questions: Based on the schematic found on Dr. Brent Selinger s web site, what step(s) is/are most likely responsible for the difference in coliphage numbers between raw and treated sewage? Have you isolated more than one type of phage? How might you be able to tell? To what components of the bacterial cell to phage typically adhere? 39

41 EXERCISE 8 WATER QUALITY (Please see Madigan and Martinko, 2006 Chapter 28) Although the term coliform refers to a group of genera of bacteria that are facultatively anaerobic, Gram negative, non-spore forming rods, the operational definition of coliform is an organism that ferments lactose with gas and acid formation within 48 hours at 35 C. The presence of coliforms is routinely used as part of the regular examination of public water. Coliforms are considered to be useful indicator organisms. Many coliforms are members of the enteric bacterial group and as such, inhabit the intestinal tracts of humans and other animals. Consequently, the presence of coliforms in a water supply suggests fecal contamination of that supply has occurred. In addition, coliforms and pathogens have been demonstrated to behave similarly in water purification. Rather than screening a water supply for individual organisms, large-scale tests have been devised to demonstrate the presence of coliforms and these tests are based on the operational definition of coliforms. It is often convenient to use an indirect method to assess the presence of coliforms in a water supply. The Most Probable Number procedure uses tubes containing liquid medium designed to select for the presence of organisms that ferment lactose producing gas. Replicates of these tubes are set up, each replicate set containing one of three dilutions of water in question. In this case, MacConkey Broth (same as MacConkey plates used previously) is used. It contains an indicator, bromcresol purple that is yellow at ph 5.2 and purple at ph 6.8. These tubes also contain inverted Durham vials designed to show gas production. Positive results from these tubes are scored, and a number or an MPN index is obtained is based on certain probability formulas. This value represents an index of mean density of coliforms per 100 ml of original sample, NOT an actual number of organisms. Coliform density obtained in this fashion provides us with a good initial assessment of water quality. EXPERIMENTAL OBJECTIVES Students will use samples of water from two sources to enumerate total bacteria, then perform standard tests for the coliform group of organisms including the membrane filter (MF) procedure and the Most- Probable Number (MPN) procedure in order to assess water quality. A. Enumeration and the MPN Procedure METHODS: Sample of raw sewage (200 ml of 10-3 dilution per lab) 5x 100 ml sterile nanopure water Sterile test tubes 5 ml pipettes and pipette pumps µl micropipetters Sterile tips Sleeves of Petri dishes (30 per lab) Molten TSA agar held in a 65 C waterbath (5 bottles of 120 ml per lab) 40

42 Microfuge tubes Vortex Biohazard bags Gloves and goggles 10 tubes of double strength MacConkey broth containing Durham vials 20 tubes of single strength MacConkey broth containing Durham vials Each group of four should set up duplicates of enumeration pour plates. For the entire lab, two groups only will set up duplicates of the MPN test. *Everyone should wear gloves and goggles for all parts of this exercise where raw sewage is handled extreme biohazard* Enumeration of bacteria in water samples Period 1: 1) Use aseptic technique to create enough 10-4 and 10-5 dilutions of your water sample (remember, you are starting with a 10-3 dilution of the sewage) to plate duplicates of 1 ml per plate. 2) Prepare duplicate pour plates using 1 ml of the 10-3, 10-4 and 10-5 dilutions. 3) These plates will be incubated at 37 C for 48 hours. Period 2: 4) Record the number of colonies on and in the agar, and calculate the number of colony forming units per ml of water sample. Most-Probable Number Procedure (to be performed by two groups only) Period 1: 1) Prepare 100 ml of a 10-6 dilution of the sewage. This can most easily be achieved by adding 100 µl of the 10-3 dilution into 100 ml of sterile water. Shake vigourously to mix. 2) Aseptically, add 10 ml of this prepared dilution of the sewage sample to be tested to each of 5 tubes containing 10 ml of double-strength MacConkey broth. 3) Aseptically, add 1 ml of the diluted sewage sample to be tested to each of 5 tubes of singlestrength MacConkey broth. 4) Aseptically, add 0.1 ml of the sewage sample to be tested to each of 5 tubes of singlestrength MacConkey broth. 5) Tubes will be incubated at 37 C for 48 hours. Period 2: 6) Following incubation, count the number of tubes in each dilution series that show acid AND gas formation. 7) Find this combination on the chart, and record the MPN index. For instance, if you recorded 2 positive results in the double strength MacConkey broths containing 10 ml of water samples, 3 positive results in the single-strength MacConkey broths containing 1 ml of water samples and 0 positive results in the single-strength MacConkey broths containing 0.1 ml of 41

43 water sample, this would be read as and would correspond to an MPN value of 14/100 ml. B. Confirmation of Presence of Coliforms in Water Samples A more direct method for confirming the presence of coliforms in a water supply is termed the Membrane Filter (MF) procedure. Samples of the water to be tested are passed through a sterile membrane filter of 0.45 m, removing the bacteria. The filter is then plated on EMB medium. This allows determination of exact numbers of coliform bacteria, as well as the potential for identification of the organisms in question. Rather than having you perform this procedure, you will choose one tube from the MPN series that shows a positive result for acid and gas formation and prepare a streak plate for single-colonies using this mixture. This will allow for confirmation of presence of coliforms and allow for selection of a unique colony for further characterisation. METHODS EMB agar (20 plates per lab) Gram stain reagents (in kits for the next lab period) Period 2: Work in groups of 4: 1) Obtain 1 plate of EMB agar and label completely. 2) Work aseptically to prepare a streak plate for single colonies using 1 of the positive MPN tubes. 3) Invert your plate and incubate at 37 C for 48 hours. Period 3: 4) Obtain your plate and make observations on the results. Are there coliforms present? Are they all the same organism? How can you tell? Period 4: 5) Select one of the isolated colonies on your plate and prepare a Gram stain. Are you studying a Gram negative rod? What other non-morphological evidence do you have to support your observation? 6) Inoculate a TSA plate using cells from the same colony. Period 5: 7) Use an isolated colony from your TSA plate to inoculate an Enterotube. Incubate 48 hours at 35 C. Period 6: 8) Evaluate your results. Identify your unknown organism to the level of genus. Identification may necessitate consultation of other sources prior to coming to lab! 42

44 APPENDIX 1 THE COMPOUND LIGHT MICROSCOPE As you label Figure 1, your Instructor will review the use of this microscope with you. Locate the ocular lens (eyepiece); there will be one if the microscope is monocular, or two if it is binocular. Then locate the objective lenses, the ones nearest the object to be studied. These two lenses (ocular and objective) are connected by the body tube of the microscope. The objective lenses (there will be two or more, the smallest being that with the least magnifying power, and the largest being that with the greatest magnifying power) are mounted on a revolving nosepiece above a flat stage on which the study specimen (slide) is placed. Figure 1: The Compound Microscope Your microscope is equipped with a mechanical stage. This consists of a clip to hold the slide in place (the clip is spring-loaded; the Instructor will demonstrate how it works) and two knobs at the side of the microscope body to move the slide side-to-side, or forward-to-back. Note also the two micrometer scales on the mechanical stage, which allow you to note the coordinates of a particular object on the slide you are viewing. Place a slide on the stage and center it over the hole in the stage. Adjust the distance between the oculars to match your interpupillary distance (distance between your pupils). Revolve the 43