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The University of Lethbridge

BIOLOGY 3200 Principles of Microbiology LABORATORY MANUAL

Spring, 2006

Written by: L. A. Pacarynuk Revised: December, 2005



Biology 3200 Laboratory Schedule Grade Distribution Occupational Health and Safety Guidelines Guidelines for Safety Procedures Exercise 1 ­ Introduction to Microscopy Exercise 2 ­ General Laboratory Principles and Biosafety Exercise 3 - Bacterial and Yeast Morphology Exercise 4 ­ Bacterial Reproduction Exercise 5 ­ The Ames Test Exercise 6 ­ Biochemical Tests Exercise 7 ­ Virology Exercise 8 ­ Water Quality Appendix 1 ­ The Compound Light Microscope Appendix 2 ­ Preparation of Scientific Drawings Appendix 3 ­ Aseptic Technique Appendix 4 ­ The Cultivation of Bacteria Appendix 5 ­ Bacterial Observation Appendix 6 ­ Laboratory Reports Appendix 7 ­ Use of the Spectrophotometer Appendix 8 ­ Media, Reagents, pH Indicators Appendix 9 ­ Care and Feeding of the Microscopes


2 3 4 5 7 12 14 20 25 28 30 35 38 41 43 49 54 55 57 59 68



Jan. 10 Jan. 12 Jan. 17 Jan. 19 Jan. 24 Jan. 26 Jan. 31 Feb. 2 Feb. 7 Feb. 9 Feb. 14 Feb. 16 Feb. 21 Feb. 23 Feb. 28 Mar. 2 Mar. 7 Mar. 9 Mar. 14 Mar. 16 Mar. 21 Mar. 23 Mar. 28 Mar. 30 Apr. 4 Introduction, Microscopy General Lab Procedures, Biosafety Bacterial Morphology Bacterial Morphology Bacterial Morphology Bacterial Morphology Bacterial Growth Bacterial Growth ­ Complete Ames Test; Biochemical Tests - Selective and Differential Media Hand in Assignment 1 at the BEGINNING of the lab period; Ames Test ­ Complete Biochemical Tests and Selective/Differential Media Biochemical Tests and Selective/Differential Media - Complete Reading Week Reading Week Virology Start Hand in Lab Report at BEGINNING of lab; Virology Virology Virology ­ Complete; In-class Assignment 2 Water Microbiology Start (enumeration, MPN) Water Microbiology (Plating on EMB) Water Microbiology (select unknown) Water Microbiology (Gram stain; test strip inoculation) Water Microbiology (test evaluation) Water Microbiology Complete; In-class Assignment 3 Free Laboratory Period Final Lab Exam (practical and written)

Thursday Apr. 6


Laboratory Grade Distribution: The laboratory component of Biology 3200 is worth 50% of your course mark. It is distributed as follows: · · Assignments Lab Report · · · Ames Test 15% 20% Due Thursday, March 2 at the beginning of lab 15%

Lab Exam

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 where you are to prepare an in-class assignment requires documentation. Upon presentation of this documentation, you will either have to prepare the assignment as soon as possible, prepare an alternate assignment, or the marks may be added to your final exam. As proficiency in microbiological techniques is considered an essential component of the course, students are only permitted two lab period absences (you do not require any documentation). Missing more than two 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 prior to the 30 minute mark -25% (eg. if the assignment is out of 50 points, you will lose 12.5 marks); after the 30 minute mark, but prior to 4:30 pm ­50%; by 9:00 am the next morning -75%, and after 9:00 am the following day, no marks will be given. Extensions for lab reports and Assignment 1 will only be granted for situations involving prolonged illness (documentation is required). The lab exam (April 6) is comprehensive, covering all aspects of the laboratory. It may contain a practical as well as a theoretical component.


THE UNIVERSITY OF LETHBRIDGE Policies and Procedures Occupational Health and Safety



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: 331.5201 Occupational Health and Safety: 394.8937 394.8716


EMERGENCY CALL LIST 1600 -0800 2345 SECURITY 331-5201 CHEMICAL RELEASE OFFICER 394-8937 OCCUPATIONAL HEALTH AND 394-8716 SAFETY IF THE CHEMICAL RELEASE OFFICER CANNOT BE LOCATED CALL: 328-4833 DBS If the area must be evacuated all employees will be evacuated to the North Parking Lot. GUIDELINES FOR SAFETY PROCEDURES 4


EMERGENCY NUMBERS City Emergency Campus Emergency Campus Security (Emergency - 2483) 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) 2) 3) 4) 5) 6) 7) Closest emergency exit Closest emergency telephone and emergency phone numbers Closest fire alarm Fire extinguisher and explanation of use Safety showers and explanation of operation Eyewash facilities and explanation of operation. First aid kit 9-911 2345 2603

Student Health Centre 2484

GENERAL SAFETY REGULATIONS 1) 2) 3) 4) 5) 6) Eating, drinking or gum chewing is prohibited in the laboratory. Always wash your hands after entering and prior to leaving the laboratory. Laboratory coats are required for all laboratories and must be stored in the lab when not in use. Report equipment problems to instructor immediately. Report all spills to the instructor immediately. Long hair must be kept restrained to keep from being caught in equipment, Bunsen burners, chemicals, etc.


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.


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, severely-curved 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.




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 0.25. 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 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) 2) 3) 4) 5) 6) Place a micrometer slide on the stage and focus the scale using the 40x objective. Turn the eyepiece until the graduations on the ocular scale are parallel with those on the micrometer slide scale and superimpose the micrometer scale. Move the micrometer slide so that the first graduation on each scale coincides. Look for another graduation on the ocular scale that exactly coincides with a graduation on the micrometer scale. 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. Calibrate the ocular divisions for the 40x and the 100x objective lenses. Note that immersion oil is not necessary for calibration.



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 µ m) is equal to 5 ocular divisions (o.d.) on the ocular micrometer. 1 ocular division equals 60 µ m/5 ocular divisions or 12 µ m. 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.


METHODS: For each student: · · · · · 1) Compound light microscope Various prepared slides of bacteria. Stage micrometer Ocular micrometer Immersion oil 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.


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) 2) 3) 4) Tape bench coat onto the bench to cover your working surface. 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. From the same suspension, inoculate one tube of nutrient broth. For steps 4 - 11, work in pairs. Place a watch glass in the centre of the bench coat.


5) 6) 7) 8)

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. Using a pipette pump, draw up 2 mL of bacteria/fluorescein suspension. Remove lids from agar plates and set aside. 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.


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 16-20 hours at 37oC, and then refrigerated at 4oC. 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.


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. 58-59) 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.


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--hydroxybutyric Acid (PHB) Staining PHB granules are common inclusion bodies in bacteria. Monomers of -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


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.


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 1) 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. 2) 3) Add Gram's iodine solution to the slide for 1 min. Wash gently with tap water and remove as above. 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. 4) 5) Counterstain the slide with safranin for 30 seconds - 1 minute. 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) 2) Flood the dried, heat fixed film with Ziehl's carbol fuchsin and place on the rack over the boiling water bath. Steam gently for 5 minutes. Do not let the slide dry out. Add more carbol fuchsin as required.


3) 4) 5)

Wash with tap water to remove excess stain. Decolourise with 20% sulfuric acid until no more stain comes out. Wash with tap water to remove excess. 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) 2) 3) 4) 5) 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. Pour off excess stain (do not wash) and air dry. Clear slide by dipping in a jar of solvent in the fume hood for 5 sec. Air dry in the fume hood. Counterstain for 1 min. with safranin. 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) 2) Prepare smear and heat fix. Cover the dried fixed film with a small piece of paper towel. Saturate this with 5% malachite green. 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) 4) 5) 6) Allow the slide to cool, then rinse with water. Tap over a paper towel to remove excess water Counterstain with safranin for 30 seconds. Rinse slide with water. 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) 2) 3) Prepare a wet mount of the cells using a drop of Methylene Blue. Carefully place a cover slip on the cell/stain mixture. View the cells noting size and shape. If you look carefully, you should be able to see budding cells.


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. 1997. Principles of Microbiology. Wm. C. Brown Publishers, Toronto. Madigan, M. T., and Martinko, 2006. Brock Biology of Microorganisms Eleventh Edition. Prentice-Hall of Canada, Inc., Toronto. Ross, H. 1992-1993. Microbiology 241 Laboratory Manual. The University of Calgary Press, Calgary.


EXERCISE 4 BACTERIAL REPRODUCTION MEASUREMENT OF BACTERIAL GROWTH (See Madigan, M. T and Martinko, J. M., 2006. 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 (td). 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. 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 an E. coli culture by the viable count and turbidimetric methods. You will determine the number of bacteria (CFU) present in your culture following various time points of incubation. You will establish a growth curve and 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 10-100 µL micropipettor 100-1000 µL micropipettor Sterile tips for micropipettors Container of sterile microfuge tubes Microfuge tube racks Sterile d2H2O Spectrophotometer blank containing TB broth Bacterial waste container Vortex Cuvettes Spectrophotometer Culture flask of E. coli


Please work in groups of four. At 20 minute intervals, monitor the growth of your E. coli culture by determining viable counts as well as optical density following the procedures outlined below.

A. 1)

Culture sampling For laboratory sections 1 and 2, 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.


Everyone in the laboratory will be sampling at the same time. Samples will be collected three times at 20 minute intervals. 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. 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.


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).


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. 1) 2)

Determination of optical density (please read Appendix 7) Zero the spectrophotometer as outlined in Appendix 7. 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 provided. 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.


After reading, dispose of your 5 mL sample of culture in the waste beaker provided. Place the spectrophotometer tube into the bleach tray.


C. 1)

Enumeration of viable bacteria 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.


Set up your serial dilutions according to the information in Table 4.1. Aseptically pipette 900 µL of TB 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 1 2 3 4 5 (for labs 3 and 4 only) Amount of sterile TB (µL) 900 990 990 900 900 Amount of Culture 100 µL from culture flask 10 µL from tube 1 10 µL from tube 2 100 µL from tube 3 100 µL from tube 4 10-7 10-6 10-5 10-3 Final Dilution Factor 10-1

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. 5) 6) Follow the instructions provided in step 4 above to plate out the contents of Tube 4. When the agar has solidified, place the inverted plates on a tray at the side of the lab. The plates will be incubated for 16 ­ 20 hours at 37°C and refrigerated until the next lab session. The next laboratory period: 7) 8) 9) 10) Examine the plates carefully and select the plate where the bacterial count ranges between 30 and 300 colonies. 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. Use class data to determine the average number of bacteria per mL of culture. 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. 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 0.75. Based on the differences in ingredients, what are the differences between growing cells on LB versus TB? Why is TB used for generating growth curves of E. coli rather than LB?


EXERCISE 5 THE AMES TEST MUTATION AND RECOMBINATION (See Madigan and Martinko, 2006. Chapter 10 Pg. 257-267) 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 Histester 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.


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 50oC 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 Sterile filter paper disks Forceps 3x micropipettors (10 ­ 100 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 1 2 + 1535 +1535 + 1538 + 1538 +1535 +1538

Unknown 3

Sodium Azide

Hair Dye

1 2 3 4 5

+1535 +1538 +1535 +1538 +1535 +1538 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. 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 oC. 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. 1973. Carcinogens are mutagens: a simple test combining liver homogenates for activation and bacteria for detection. Proc. Natl. Acad. Sci. U.S.A. 70:2281-2285. Ames, B.N., Lee, F.E., and Durston, W.E. 1973. An improved bacterial test system for the detection and classification of mutagens and carcinogens. Proc. Natl. Acad. Sci. U.S.A. 70:782-786. Ames, B.N., McCann, J., and Yamasaki, E. 1975. Methods for detecting carcinogens and mutagens with the Salmonella-microsome mutagenicity test. Mutational Research 31:347364. Madigan, M. T., and Martinko, 2006. Brock Biology of Microorganisms Eleventh Edition. Prentice-Hall of Canada, Inc., Toronto.


EXERCISE 6 BIOCHEMICAL TESTS (Selective and Differential Media; Enterotubes) 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 nonpathogens 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). See the literature associated with the Enterotubes for test and evaluation details. SELECTIVE AND DIFFERENTIAL MEDIA: 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 nonlactose 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 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. METHODS For each bench: · · · 3 plates each of MacConkey and EMB media 5 unknown broth cultures (1/bench) 1 Enterotube and instructions for all students

Please work in groups of four. 1) 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 summarising growth and properties of all bacteria on the two media. 2) Use the instructions provided with the Enterotubes to inoculate the tube using your unknown. After 37°C for 48 hours, observe, and evaluate these results thoroughly. Use the information provided to determine the probable identity of your unknown.

Thought Questions: · · Identify your unknown. Provide evidence to support your choice of organisms. 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 two 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).


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 oC) 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 oC 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.


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 se wage. Tube # Contents (µ L) K12 Ra w Se wage Filtrate 1 2 3 4 5 500 0 0 500 500 0 500 0 500 0 Treated Se wage Filtrate 0 0 500 0 500

3) 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 16 ­ 20 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 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 oC 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 oC) · Sterile test tubes · Test tube indicating 4 mL mark


· Test tube rack · 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 oC overnight. 3) The next day, score as + or ­ for phage growth on each host. This lab will form the basis of an in-class assignment. 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?



(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 MostProbable Number (MPN) procedure in order to assess water quality. A. 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 100 ­ 1000 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) Enumeration and the MPN Procedure


· · · · · ·

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 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) 3) Prepare duplicate pour plates using 1 mL of the 10-3, 10-4 and 10-5 dilutions. These plates will be incubated at 37 °C for 48 hours.

Most-Probable Number Procedure (to be performed by two groups only) 1) 2) 3) 4) 5) 6) 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. 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. Aseptically, add 1 mL of the diluted sewage sample to be tested to each of 5 tubes of singlestrength MacConkey broth. Aseptically, add 0.1 mL of the sewage sample to be tested to each of 5 tubes of singlestrength MacConkey broth. Tubes will be incubated at 37 °C for 48 hours. At that point, count the number of tubes in each dilution series that show acid AND gas formation. 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 water sample, this would be read as 3-2-0 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)

Work individually: 1) 2) 3) Obtain 1 plate of EMB agar and label completely. Work aseptically to prepare a streak plate for single colonies using 1 of the positive MPN tubes. Invert your plate and incubate at 37 °C for 48 hours.

During the next lab period: 4) 5) 6) Obtain your plate and make observations on the results. Are there coliforms present? Are they all the same organism? How can you tell? Prepare a liquid culture from a single colony. Incubate 48 hours at 37 °C Prepare a Gram stain from your colony. Are you studying a Gram negative rod?

During the next lab period: 7) Use your liquid culture to inoculate an Enterotube. Incubate 48 hours at 35 °C, then evaluate your results. Identify your unknown organism to the level of genus. This evaluation and identification will form the basis of the in-class assignment. Identification may necessitate consultation of other sources prior to coming to lab!



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


nosepiece so that the lowest power objective lens (generally the 10x power lens) is in position. To focus the microscope, locate the coarse and fine adjustment knobs at the base of the microscope, and use the coarse adjustment to move the slide close to, but not touching, the objective lens. Look at the stage from the side as you do this. On most microscopes this involves raising the stage, but on some the lenses are lowered. Also, on most microscopes an automatic stop will prevent you from moving the stage closer than about one centimeter from the lens. Now, look through the ocular lenses, and move the slide away from the objective lens until the specimen becomes clear (is in focus). Finish focusing with the fine adjustment knob. Once you have focused with the low objective power lens, you may switch over to the next higher power lens with only fine focus adjustments (the microscope is said to be parfocal). As you switch from one objective lens to another, you will notice that the working distance, the clearance between lens and stage, decreases with increasing lens power. This is illustrated in Figure 2 below.

Figure 2: The working distance (above) and the field of view (below) change with magnification of objective lens.

It should be obvious to you why, on high power objective lenses (40x or 100x), you must use only the fine focus knob to adjust focus; otherwise the risk of (damaging) contact between lens and slide becomes great. Also illustrated in Figure 2 is the diminishing field of view as objective lens power increases; this is due to a smaller and smaller aperture at the bottom of the lens through which light enters. This means that [a] things are harder to find on a slide when you are using high power since only a small fraction of the slide can be seen, and [b] less light enters your eye and everything in the field appears darker. As a consequence, you will learn to [a] switch back to a lower power objective lens when you want to "scan" around the slide, and [b] manipulate the amount of light coming into the lens so that you can see the objects clearly.


The amount and concentration of light coming through the specimen and hence to your eye can be adjusted in several ways. First, of course, is the on/off light switch, generally located at the base of the microscope, and often associated with a rheostat to control light intensity. A condenser lens is mounted below the stage, and concentrates the light on to the specimen; it generally needs no adjustment of position. An iris diaphragm is located below the condenser lens. Find the lever which controls the diaphragm; it can be very useful in adjusting illumination and contrast. Biology 3200 microscopes are binocular, containing two eyepieces. To correct for the slight difference in the focus of your two eyes, precisely fine focus a specimen using only your one eye which is at the non-focusing ocular (if your microscope contains two focusing oculars, either may be used to begin). Next, open the other eye and bring the image into focus for that eye using only the ocular focus. Since other students use these same microscopes during the semester, this exercise of binocular focusing should be performed at the onset of each microscope session. Finally, some useful hints and cautions: · · · · · · Never drag the microscope across the counter-top. Lift it with both hands by its arm, being careful not to tip it. Use lens paper to clean glass slides and lens surfaces before using your microscope. Water damages objective lenses; if water does contact a lens, wipe it off immediately. Also avoid getting water under the slide as it will stick to the stage. If you have used immersion oil, use lens paper dipped in 60 % ethanol to remove it from the 100x objective lens when you are finished. Always start the focusing procedure with low (10x) power lens. When attempting to locate an object on a slide, remember that the image you see is reversed; that is, as you move the slide toward you on the stage, the slide is apparently moving away from you as you view it through the lens. · Some ocular lenses are equipped with pointers; they appear as a dark black line that will rotate if the lens is rotated in its tube. Electron Microscopy Bacterial size places them at the limits of resolution of the light microscope. Even with the best quality lenses, magnification can only be increased slightly beyond 1,500x. Much higher magnification can be achieved with the electron microscope with the scanning electron microscope (SEM) reaching about 100,000x magnification and the transmission electron microscope (TEM) capable of 1,000,000x magnification.


APPENDIX 2 PREPARATION OF SCIENTIFIC DRAWINGS 1) 2) 3) 4) 5) 6) 7) Use a sharpened pencil; never ink. The lead should be hard. Place drawing to one side, usually the left, leaving room for labels to the right. Try to draw with one continuous line and do not retrace your lines. Do not shade. Place label lines horizontally (use a ruler), with no crossed lines. Objects labelled should be singular unless label line branches to multiple objects. Label only what you see, not what you think should be seen. Below the figure you should add: a) b) 8) The title of the diagram The magnification of the drawing (see below)

The magnification of the diagram gives you the relationship between the size of your diagram and the actual size of the specimen. A diagram of a cell would be much larger than the actual cell, whereas a diagram of an elephant could be much smaller than the actual elephant.

Magnification is defined as: Where: · · ·

size of drawing actual size of specimen

size of the drawing is measured with a ruler actual size of specimen is determined by one of the methods in Exercise 1. the number calculated has as many significant figures as the accuracy of your measurement (usually 2, if you measure in mm)



Example of a drawing:

Figure 1. A chain of Bacillus subtilis cells stained with methylene blue (23 000x) · · · · Notice that in the figure, enough organisms are shown such that the arrangement can be seen. Drawing magnification is calculated based on length or width, not both of only one of the organisms (not the whole chain). Figures are given numbers - Figure 1, Figure 2, etc. As much detail as possible is provided in the title (eg Gram reaction seen, type of stain used, type of organism etc.).



Much microbiological work, and to some extent biochemical work, depends on the maintenance of pure cultures of microorganisms. Therefore, there are various essential precautions that MUST be observed to exclude unwanted organisms. Accidental contamination may ruin your results completely. Aseptic technique is largely a matter of common sense, but it is essential to realise that bacterial and fungal spores are present everywhere, and a high standard of technique must be attained. Correct methods of handling cultures and apparatus will be demonstrated. These methods should be followed. Consider carefully and remember the following points: 1. Clean air contains many bacterial and fungal spores carried on dust particles or in water droplets. Any surface exposed to air quickly becomes contaminated, and if material is to be kept sterile it should be exposed only as much as is absolutely necessary for manipulation. Instruments which can be sterilised by heating in a bunsen flame (e.g. inoculating loops) can be left exposed, but they must be flamed thoroughly before use, and again before being replaced in the holder. Items of equipment that cannot be treated in this way (e.g. pipettes) are sterilised in wrappings or containers from which they must not be removed until actually needed. They must not be allowed to touch unsterile surfaces during use. Plugs and caps of tubes and bottles must not be laid on the bench nor must sterile containers be left open to collect falling dust. 2. Clothes, hair, skin and breath all carry a heavy microbial load and where strict asepsis is essential, sterilised gowns, caps, gloves etc. are worn. Even in normal microbiological work care must be taken to prevent contamination from the above mentioned sources. A clean laboratory overall is advised for all lab work. Microbial contamination in the lab is most often due to currents of unsterile air. The chief merit of inoculation chambers and screens therefore lies in the protection they give from drafts. This protection can be supplemented by keeping all windows and doors shut and by cutting down personal movement within the laboratory. These precautions can be offset by careless use of burners that create convection currents.



Before any operation is started, all necessary materials should be assembled in convenient order with provision for protecting sterile objects until needed, and for disposing of used apparatus (so as not to contaminate other material).


Aseptic Culture Manipulation To prevent the contamination of the environment and people working in the laboratory from the cultures used in the exercises 2) To prevent accidental contamination of cultures of microorganisms and of solutions and equipment used in the laboratory

Purposes: 1)

Correct methods of handling cultures and apparatus will be demonstrated. These methods should be followed. Consider carefully and remember the following points: · Prior to starting any work in the laboratory, wash hands with soap, and wash down bench area using 10% bleach. This procedure should be repeated after the lab is complete. · · Avoid working on your lab book or lab notes. Clean laboratory coats must be worn. If you have long hair, tie it back before working in the laboratory environment. · Eating or drinking are not permitted in the laboratory. Do not place pencils, fingers or anything else in your mouth. · Clean air contains many bacteria and fungal spores carried on dust particles or in water droplets. Any surface exposed to air quickly becomes contaminated. If material is to be kept sterile, it should be exposed only as much as is absolutely necessary for manipulation. Plugs and caps of tubes, tops of Petri dishes and bottles of solutions, (even water!!) must not be laid on the bench nor must sterile containers and cultures be left open and exposed to the air.


Inoculation of Culture Tubes Again, the important thing to remember is that exposure of sterile liquids or bacterial cultures to air must be minimised. -Ensure that you have the tubes, plate of inoculum, inoculating loop and a sterile tube of medium available within easy reach. -Flame the inoculating loop until red hot. When removing inoculum from a tube, remove the cap from the tube by grasping the cap between the last finger and the hand which is also holding the inoculating needle (Figure 1). Do not place the cap on the bench!!

Figure 1: Technique for manipulating test tubes aseptically. -Flame the mouth of the tube by passing it rapidly through the Bunsen burner 2-3 times. This sterilises the air in and immediately around the mouth of the tube. -Cool the loop on the inside of the tube, remove the inoculum. -Reflame the mouth of the tube and replace the cap -Flame the inoculating loop before replacing -Note, when removing inoculum from a plate, cool the loop in the agar before picking up the bacteria


Streaking for Single Colonies -A loop of liquid culture or a small amount of bacterial growth from a plate culture is transferred aseptically to a sterile plate in the area shown by Figure 2A. -Once the first set of streaks has been made, the inoculating loop is reflamed until red hot. DO NOT REINTRODUCE THE LOOP INTO THE ORIGINAL CULTURE!!! -Cool the loop, and make a second set of streaks as shown in Figure 2B, only crossing over the initial set of streaks once. -Flame the loop again, cool, and repeat for three more sets (Figure 2C). Note, try not to gouge the agar while streaking the plate.


Preparation of Spread Plates: Generally, volumes of culture greater than 100 µL are NOT plated as it takes too long for the liquid to dry. · · Use aseptic technique to obtain 100 µL of culture and place in the middle of a plate of medium. Use a sterile glass spreader (this may involve dipping a spreader into a beaker of alcohol and waving it through a Bunsen Burner flame. If this is the case, DO NOT hold the spreader in the flame and avoid tipping the spreader so that flaming alcohol runs over your hand. Once the flame has burnt out, the spreader is ready to use). · · Use the same hand that holds the spreader to lift the lid of the plate and keep it just above the plate the entire time. Gently touch the spreader to the side of the medium (not directly in the culture in case the spreader is still a bit warm). Smooth the culture evenly over the surface of the plate ensuring that you cover the entire plate. · Invert the plates and place in the incubator when dry.

C. Sterilisation Media must be sterilised after distribution into tubes, flasks or bottles. Sterilised media may later be transferred aseptically to previously sterilised containers, but this should only be done when really necessary, e.g. in preparing "plate" cultures, since some risk of contamination is unavoidable. Methods of Sterilisation 1. Most media (including agar) can be sterilised by treatment with steam under pressure in an autoclave, the usual treatment being 15-20 minutes at a pressure of two atmospheres. This raises the steam temperature to 121°C. When using an autoclave, the water should be allowed to boil, and the steam to fill the autoclave before shutting the valve. This allows the material to heat up and ensures that the correct steam pressure is attained. Never overfill an autoclave since this will upset the pressure/volume relationship and the correct temperature will not be attained. Materials that might be adversely affected by this treatment may sometimes be treated for a short time or at a lower temperature, but this will not be effective if the material is heavily contaminated to begin with. Screw caps on bottles must be left slightly open during sterilisation and screwed down on removal from the steriliser. 2. Media that are difficult or impossible to autoclave satisfactorily, e.g. gelatin media and some sugar media, may be sterilised by intermittent steaming. Objects to be sterilised are heated over boiling water in a steamer (steam temperature 85°-95°C) for 15-20 minutes on each of three or more successive days. Time must be allowed for the medium to reach the same temperature as the steam. Between treatments the material


must be kept at a temperature allowing spores to germinate (30°-37°C) and so lose their heat resistance. 3. It is often necessary to sterilise some ingredients of a medium separately and to add them to the rest of the medium before use. Heat-labile ingredients, e.g. urea, serum, etc. must be sterilised by filtration through a bacteria-proof filter, i.e. Seitz filters or membrane filters. 4. Dry glassware, e.g. glass petri dishes, empty flasks, pipettes may be sterilised in the autoclave and then dried or may be sterilised in a hot air oven. Any oil material must also be sterilised in a hot air oven. The minimum effective treatment is 1 hour at 150°C. This should be increased to 160°C or the time of heating prolonged to 2 or 3 hours wherever possible.


APPENDIX 4 THE CULTIVATION OF BACTERIA (Please read Madigan et. al., 2003; Chapter 5) In order to grow, microorganisms require a) water, b) macronutrients eg. ­ C, N, K, P, S, Mg, Ca, Na, and Fe c) micronutrients (trace elements) eg. - Fe, W, Zn and d) growth factors ­ vitamins, amino acids, purines and pyrimidines. In general, wild-type organisms are termed prototrophs. An auxotroph is a nutritional mutant, unable to synthesise an essential component for growth from precursors. Note that this essential component is normally synthesised by the wild-type or prototrophic strains of the same species. Scientists study and manipulate nutritional requirements of bacteria or yeast using minimal media. Minimal or defined media are those in which the exact chemical composition of all ingredients is known. A medium where the exact chemical composition is not known is termed complex. Complex media are preferred as they are generally easier to prepare than minimal media, they result in high levels of growth, and are useful when exact nutritional requirements of an organism are not known. Nutritional Classification: The nutritional classification of organisms is based on three parameters: the energy source, the principal carbon source and the source of reducing power. With respect to energy source, phototrophs are photosynthetic organisms that use light as their energy source and chemotrophs are organisms that depend on a chemical energy source. Organisms able to use CO2 as a principal carbon source are autotrophs. Heterotrophs depend on an organic carbon source. To designate the source of reducing power, the term lithotroph or organotroph is applied. Lithotrophs use inorganic compounds as their source of reducing power, and organotrophs use organic compounds as their source of reducing power. To summarise: source of energy source photoautotroph (photolithotroph) photoheterotroph (photoorganotroph) chemoautotroph (chemolithotroph)* chemical (oxidation of reduced inorganic compounds e.g. NH3 , NO2- and H2) chemoheterotroph (chemoorganotroph) chemical organic organic CO2 inorganic light organic light carbon source CO2 reducing power inorganic oxidizable substrate organic


*All chemoautotrophs are chemolithotrophs, but not all lithotrophs are autotrophic. For example, the methylotrophic bacteria can use organic carbon as their carbon source. Common Media Constituents (see Table 5.4, Madigan et. al (2003) for examples): Energy or Carbon sources: · · · · Sugars, alcohols, carbohydrates and amino acids Found in infusions ­ for instance ­ beef infusion Found in extracts ­ for instance ­ yeast extracts Also found in peptones (see below)

Nitrogen sources: · · · · · Inorganic sources such as ammonia or nitrate Nitrogen fixing organisms use atmospheric N2 Extracts, infusions Peptone ­ hydrolysis of proteins produces mixtures of short-chains of amino acids (peptides). Sources of peptones may include meat, fish, blood, or soybeans Tryptone ­ pancreatic digestion of casein

Other Macronutrient Source Examples: · · · MgSO4 CaCl2 Potassium salts

Micronutrient Sources: · May not be necessary to add as these are required in such small concentrations.

Growth Factors: · · Some organisms are able to synthesise all growth factors from precursors. Other organisms require these compounds already synthesised For example ­ thiamine, biotin

Buffering Components Buffers, which prevent large changes in pH, are often required to facilitate growth. This is particularly true of media composed of simple compounds or in which acid-producing bacteria are cultivated. Mixtures of sodium and potassium phosphates are often employed. In complex media, buffering is provided by the peptides and amino acids.


Gelling Agents For a solid medium, agar, a water soluble polysaccharide, is added to the medium. First discovered in 1658 in Japan, agar was first used for microbiological purposes by R. Koch in 1882. It is extracted from members of Class Rhodophyceae (a group of red-purple marine algae). Agar is particularly suited to microbial propagation because: · · · It lacks metabolically useful chemicals such as peptides and fermentable carbohydrates (it cannot be broken down by bacterial enzymes) It melts at a high enough temperature (85 oC) to support growth of different temperature requiring microbes It lacks bacterial inhibitors

Below are two examples of media used for cultivation of microbes. TY is an example of a complex medium whereas VMM is an example of a minimal or defined medium: TY Agar (used for the cultivation of organisms such as Rhizobium leguminosarum, Pseudomonas fluorescens) As with most complex media, ingredients for TY are weighed out, 1 L of water is added, and the mixture autoclaved. After cooling slightly to approximately 60 oC, TY medium is poured into Petri dishes. Ingredient Tryptone Amount (/L) 5.0 g Source of? Macronutrients (primarily nitrogen, also carbon and growth factors in the form of amino acids) Yeast Extract 3.0 g Macronutrients (primarily carbon, also nitrogen and growth factors) CaCl2 MgSO4 Agar 0.5 g 0.1 g 20 g Macronutrients Macronutrients Gelling agent

For the next example ­ VMM ­ three different mixtures (Solutions A, B and C) of ingredients are made up separately, autoclaved separately, and then combined. Finally, a carbon source is added just prior to pouring.


VMM (Vincent's Minimal Medium - Vincent, 1970) (used for the study of nutritional requirements of Rhizobium leguminosarum) Solution A: Compound K2HPO4 KH2PO4 KNO3 For Solid Medium: Agar Solution B (10x): Compound FeCl3 MgSO4 CaCl2 Amount (/L) 0.1 g 2.5 g 1.0 g Source of? Macro/Micronutrients Macronutrients Macronutrients Amount (/L) 1.0 g 1.0 g 0.6 g 12.5 g Source of? Buffering agent/ Macronutrients Buffering agent/Macronutrients Macronutrients (nitrogen in particular) Gelling agent

Autoclave and add to a final concentration of 1x Solution C (100x) Compound Biotin Thiamine Calcium Pantothenate Amount for: 1 L 0.01 g 0.01 g 0.01 g Source of? Growth factors Growth factors Growth factors

Autoclave and add to a final concentration of 1x. Carbon sources: Depending on the organism studied, a variety of carbon sources may be added. For instance, when studying genes required for catabolism of a certain carbon source, a scientist will often first create a mutant or auxotroph unable to catabolise that carbon source. To confirm presence of the mutation, it is necessary to plate the putative auxotroph on medium containing the carbon source of interest, and plating on a medium containing a carbon source that the organism is able to utilise. In Rhizobium leguminosarum, some examples of carbon sources that are useful for these types of experiments are mannitol, sorbitol (both are sugar alcohols), or rhamnose. Each carbon source is prepared as a stock solution, filter sterilised, and added to a final concentration of 0.4% (w/v). Oxygen Requirements of Microorganisms. Many species of bacteria are facultative aerobes, i.e. they can grow under aerobic or anaerobic conditions, the latter ability being dependent upon the presence of some substance that can be utilised as an electron acceptor by the species concerned. Some bacteria are obligate aerobes, unable to use anything but oxygen as a final electron acceptor. Others are obligate anaerobes that cannot use oxygen as an electron acceptor. A few bacteria are somewhat intermediate,


growing best in low oxygen tensions. These are called microaerophilic bacteria. During growth in liquid culture, microorganisms tend to utilise all available oxygen and so reduce the medium. Thus, the oxidation-reduction potential (Eo) of the medium may become low enough to allow anaerobic growth to occur. One example of this is found in the fermentation of sugar to produce alcohol by yeast (Exercise 8 part C). Unless the mixture is stirred frequently, the little oxygen available in the grape juice solution is utilised rapidly by the growing culture. Organisms then switch to anaerobic growth. In order to sample material containing anaerobes, specimens must be obtained and immediately placed into an environment containing an oxygen-free gas and an indicator that changes colour when oxidised to indicate when oxygen has contaminated the sample. Organisms may then be cultured in sealed jars containing gas mixtures of N2 and CO2 or even by cultivation in an anaerobic chamber. Temperature Requirements of Microorganisms Cultures should be incubated at the temperature most favourable to growth or the specific activity being studied. Human pathogens and commensal species grow best at body temperature, i.e. 37°C. Soil organisms and plant pathogens are normally incubated at 20-30°C. The optimum temperature is that temperature at which the growth rate is maximal for a particular organism. Note that for every organism, there is also a minimum temperature below which no growth occurs, and a maximum temperature, above which no growth occurs. The terms used to describe microorganisms according to their temperature requirements are as follows: · · · · thermophiles require temperatures of 45°C-65°C extreme thermophiles (which are usually archaebacteria) will grow at temperatures above 65°C. mesophiles grow best at temperatures of 20°C-45°C. psychrophiles require low temperatures - below 15°C.

References: Difco Manual. 1998. Difco Laboratories, Division of Becton Dickinson and Company, Maryland. Madigan, M. T., Martinko, J. M., and Parker, J. 2003. Brock Biology of Microorganisms 10th Edition. Prentice-Hall Canada Inc., Toronto. Ross, H. 1992/3. Microbiology 241 Lab Manual. University of Calgary Press, Calgary.


APPENDIX 5 BACTERIAL OBSERVATION Bacterial genera may be differentiated in two ways: 1) 2) by the cellular morphology which is observed microscopically by the colony morphology which is observed on a plate culture

Cellular Morphology includes: 1) 2) 3) 4) Shape: rods, cocci, spirilli Size (in µm): diameter (cocci); lengthxwidth (rods) Typical arrangement of the cells: chains, clusters, pairs, random Gram reaction

A diagram drawn to scale accompanies the cellular morphology. Colony Morphology is that of a single isolated colony on the plate, not the morphology of the entire bacterial growth on the plate. Colony morphology is influenced by medium composition; type of medium organism is grown on (defined, complex, specific type) should be noted in conjunction with the description of colony morphology. The following characteristics are those most commonly used to describe colony morphology: 1) Shape or form

Circular 2) 3)


Rhizoid Filamentous

Punctiform (1mm or less in diameter)

Surface: smooth/rough; mucoid/moist/dry/powdery Elevation:

Flat 4) 5)





Size: measure a single colony with a ruler Pigment: cream, white or beige coloured organisms are usually considered to be nonpigmented. Pigments may be purple, red, pink, yellow, brown, blue, grey, etc. Water soluble pigments diffuse into the medium.


Opacity: Transparent (can see through) or opaque.


APPENDIX 6 LABORATORY REPORTS Lab reports shall be in the style of scientific papers published in refereed journals. This scientific style is relatively similar across journals although specific formats vary, including the form of literature citations. The journals Microbiology or Canadian Journal of Microbiology will be used as models for the specific format of Biology 3200 reports. Please do not use formats from journals such as Nature or Proceedings of the National Academy of Sciences as this will result in loss of marks. For detailed information on preparation of scientific reports, please refer to the Biology 3200 web site. The text should be in prose form and standard rules of grammar apply. Check spelling, including technical terms and names of bacterial species which are italicised or underlined; for instance, Escherichia coli or Escherichia coli. The reports shall be double-spaced, single-sided and typed. Staple the report together and do not submit it in a cover. The reports shall contain the normal components of a scientific paper including: Title - the title should identify the experimental topic as completely as possible. Abstract - the abstract is an abbreviated version of the complete report. Typically containing no more than 250 words, the abstract picks out the highlights of the introduction, methods, results and discussion. The abstract should be complete enough that it can be removed from the report and will still provide a meaningful description of the study. Introduction - The introduction serves to (i) provide background information and a description of what is known prior to the study, and (ii) offer a justification for the study. This justification describes why the experiment was performed - how does it fit into science and are there any applied aspects of the knowledge (i.e. is it relevant to medicine, agriculture or other disciplines). Relevant literature is used and cited. Methods - The methods or 'Materials and Methods' describes the materials involved in the study, including biological materials (bacteria, etc.), and outlines the procedures used in the study. Reference must be made to this laboratory manual (Pacarynuk and Danyk, 2004). Other references, the text by Madigan et. al., (2003), or other published materials may be cited. Global referencing ("All of the following methods are taken from...") should be avoided. The methods section should be adequate for the reader to completely understand what was done and also to be able to repeat fully the study. Results - The results describe the observations or experimental outcomes, providing figures, tables or other data as suitable. This section answers the question "What Happened?" The author should decide what is the most suitable format for experimental information and draft the report accordingly. Figures may include drawings that should be in pencil. Graphs or other figures may also be included as appropriate. Experimental results should be presented


only once. If information is presented in a figure then it should not be repeated in a table. Each figure and table must have a caption which is complete enough that the figure and caption can be removed from the report and still be understandable. Figures and tables must be referred to in the text and described so that if the reader did not have the figure or table, trends or highlights of the results would still be evident. Never include a figure or table without referring to it and describing it; to do so will result in loss of marks. Avoid evaluating or interpreting your results in this section. Discussion - The discussion should refer to concepts or questions posed in the introduction and relate these concepts from the literature to the results. Do not restate the results in this section. Your discussion will be graded based on your evaluation of the results with respect to the literature. Any time you use information from another source, it must be immediately cited within the text. Failure to do this constitutes plagiarism and may result in a mark of zero being assigned for the entire document. For examples of how to cite properly, refer to peerreviewed journal articles in Microbiology or Canadian Journal of Microbiology. Never include quotations, such as phrases from the course text or this lab manual. Direct quotes are inappropriate in scientific writing. Always introduce relevant concepts using your own wording and then cite using the format found in Microbiology or in Canadian Journal of Microbiology. Literature Cited ­ This section only includes references cited within the body of the text. Again, use the format found in Microbiology or the Canadian Journal of Microbiology. References will include journal papers, books and most likely, Holt (1989) or Holt (1994) (Bergey's Manual of Systematic Bacteriology). It is important to note that Bergey did not write Bergey's Manual of Systematic Bacteriology; the proper formats for referencing are as follows: Holt, J. G. (editor-in-chief). Bergey's Manual of Systematic Bacteriology, Vol. I, 1984; vol. II, 1986; vols, III and IV, 1989. Williams and Wilkins, Baltimore. Holt, J. G. (editor-in-chief) (1994). Bergey's Manual of Determinative Bacteriology, 9 th edition. Williams and Wilkins, Baltimore.


APPENDIX 7 USE OF THE SPECTROPHOTOMETER Many procedures for the quantitative analysis of compounds in biological fluids are based on the fact that such compounds will selectively absorb specific wavelengths of light. For example, a solution that appears red to us (such as blood) absorbs the blue or green colours of light, while the red is reflected to our eyes. The eye, however, is a poor quantitative instrument, and what appears bright red-orange to one person may appear dull red-purple to another. A spectrophotometer is one instrument that will objectively quantify the amount and kinds of light that are absorbed by molecules in solution. A source of white light is focused on a prism to separate it into its individual bands of radiant energy (Figure 1). One particular wavelength is selected to pass through a narrow slit and then through the sample being measured. The sample, usually dissolved in a solvent, is contained in an optically selected tube or cuvette, which is standardized for wall thickness and has a light path exactly one centimeter across (these tubes are therefore expensive!).

Figure 1. A photoelectric spectrophotometer. After passing through the sample, the selected wavelength of light strikes a photoelectric tube. If the substance in the cuvette has absorbed any of the light, the light transmitted out the far side will then be reduced in total energy content. When it hits the photoelectric tube, it generates an electric current proportional to the intensity of the light energy striking it. By connecting the photoelectric tube to a device that measures electric current (a galvanometer), a means of directly measuring the intensity of the light is achieved. The galvanometer has two scales: one indicates the % transmittance, and the other, a logarithmic scale with unequal divisions graduated from 0.0 to 2.0, indicates the absorbance.


Zeroing the Spectrophotometer Because most biological molecules are dissolved in a solvent before measurement, a source of error can be the absorption of light by the solvent. To assure that the spectrophotometric measurement will reflect only the light absorption of the molecules being studied, a mechanism of "subtracting" the absorbance of the solvent is necessary: 1) Align the needle to 0 on the transmittance scale using the knob on the left hand side of the machine (as you face the machine). Note, this step should be performed prior to placing any tubes into the machine. 2) Insert the reagent "blank" (the solvent) into the instrument, and align the needle to 0 on the absorbance scale using the knob on the right hand side of the machine (as you face the machine). 4) The sample, containing solute plus solvent, is then inserted. Any reading on the scale that is less than 100% transmittance (or greater than 0.0 absorbance) is considered to be due to absorbance by the solute only. Units of measurement: The transmittance scale is a % number; a ratio of the light exiting the sample tube to the light entering the tube. However, this number is not a linear reflection of the concentration of the solute molecules (Figure 2). The absorbance scale, on the other hand, does reflect a linear relationship. Although you do not necessarily know the exact concentration of the solute molecules in your sample, you do know that if the absorbance value doubles, the concentration of solute in your sample has doubled. Absorbance has no units, but the wavelength of the light is usually indicated by a subscript.

Figure 2. The relationship between % transmittance and solute concentration (on the left), and absorbance and solute concentration (on the right).


APPENDIX 8 Media, Reagents and pH Indicators MEDIA: Tryptic Soy Broth: A general purpose medium used to cultivate a variety of microorganisms. Composition (g/L): Bacto tryptone Bacto soytone Dextrose NaCl Dipotassium phosphate 17.0 g 3.0 g 2.5 g 5.0 g 2.5 g

Dissolve in distilled water to a final volume of 1 L, dispense into test tubes, and autoclave for 15 min at 121oC. Tryptic Soy Agar: Used for cultivation of a variety of microorganisms. Composition (g/L): Bacto tryptone Bacto soytone NaCl Agar 15.0 g 5.0 g 5.0 g 15.0 g

Dissolve in distilled water to a final volume of 1 L, autoclave for 15 min at 121oC, and pour into sterile Petri dishes.


LB Medium (Luria-Bertani Medium): Used for cultivation of Enterobactereaceae family members, Sinorhizobium and Agrobacterium Composition (g/L): Tryptone Yeast extract NaCl 10.0 g 5.0g 10.0 g

Dissolve in distilled deionised H2 O to a final volume of 1 L, autoclave for 20 minutes at 15 psi (1.05 kg/cm2) on liquid cycle, and pour into sterile Petri dishes. Terrific Broth (TB) Used for the cultivation of E. coli Composition (g/L) Tryptone Yeast Extract Glycerol 12.0 g 24.0 g 4.0 mL

Dissolve in distilled deionised H2 O to a final volume of 900 mL, autoclave for 20 minutes at 15 psi (1.05 kg/cm2) on liquid cycle. Allow the solution to cool to 60 oC or less, and then add 100 mL of a sterile solution of 0.17M KH2PO4 , 0.72M K2HPO4 (this is the solution resulting from dissolving 2.31 g of KH2PO4 and 12.54g of K2HPO4 in 90 mL of deionised H2 O. After the salts have dissolved, adjust the volume of the solution to 100 mL with deionised H2 O and sterilise by autoclaving for 20 minutes at 15 psi on liquid cycle).


Nutrient Agar: Used for the cultivation of a wide variety of microorganisms. Composition (g/L) Peptone NaCl Yeast extract Beef extract Agar 5.0 g 5.0 g 2.0 g 1.0 g 15.0 g

Dissolve in distilled water to a final volume of 1 L, autoclave for 15 min at 121oC, and pour into sterile Petri dishes. TY Agar Used for the cultivation of Pseudomonas and Rhizobium. Composition (g/L): Tryptone Yeast Extract CaCl2 MgSO4 Agar 5.0 g 3.0 g 0.5 g 0.1 g 13.0 g

Add distilled water to a final volume of 1 L, autoclave for 15 min. at 121 oC, and pour into sterile Petri dishes. Luria Methylene Blue Agar Used for the observation of coliphage plaques. Composition (g/L): Tryptone Yeast Extract NaCl Glucose Methylene Blue Agar 10.0 g 5.0 g 5.0 g 1.0 g 0.02 g 15.0 g


Dissolve in distilled deionised H2 O to a final volume of 1 L, autoclave for 20 minutes at 15 psi (1.05 kg/cm2) on liquid cycle, and pour into sterile Petri dishes. Luria Agar Overlay Used for the propagation of coliphage. Composition (g/L): Tryptone NaCl Glucose CaCl2 Agar 10.0 g 5.0 g 1.0 g 0.11 g 6.0 g

Add 3 mL of NaOH per L and check for a pH of 7.2. Add agar, dissolve, then autoclave for 20 minutes at 15 psi (1.05 kg/cm2) on liquid cycle.

Eosin Methylene Blue Agar: Used for selection of Gram negative bacteria, and differentiation of lactose fermenting organisms. Composition (g/L): Peptones Di-potassium hydrogen phosphate Lactose Sucrose Eosin Y, yellowish Methylene blue Agar 10.0 g 2.0 g 5.0 g 5.0 g 0.4 g 0.07 g 15 g

Dissolve in distilled water to a final volume of 1 L, autoclave 15 min at 121oC, and pour plates. MacConkey Agar: Used for selection of Gram negative bacteria, and differentiation of lactose fermenting organisms.


Composition (g/L): Peptone NaCl Lactose Bile salts Neutral red Agar 20.0 g 5.0 g 10.0 g 5.0 g 0.075 g 12.0 g

Dissolve in distilled water to a final volume of 1 L, autoclave 15 min at 121oC, and pour plates.


References: Atlas, R.M., and Parks, L.C. 1993. Handbook of Microbiological Media. CRC Press, Inc. Boca Raton, Florida. Difco Manual: Dehydrated Culture Media and Reagents for Microbiology. 10th Ed. (1984). Difco Laboratories, Detroit, Michigan. Merck Microbiology Manual 1994. Merck, Darmstadt, Germany. Ross, H. 1992/3. Microbiology 241 Lab Manual. University of Calgary Press, Calgary. Sambrook, J. and Russell, D. W. 2001. Molecular Cloning ­ A Laboratory Manual. 3rd edition. Cold Spring Harbor Laboratory Press, New York.


REAGENTS: Ethanol, 70%: 95% Ethanol Distilled Water Barritt's Reagents: Solution A: Dissolve 6 g alpha naphthol in 100 mL 95% ethanol Solution B: Dissolve 16 g potassium hydroxide in 100 mL distilled water. Crystal Violet Stain: Solution A: Dissolve 2.0 g of crystal violet in 20 mL of 95% ethanol. Solution B: Dissolve 0.8 g of ammonium oxalate in 80 mL of distilled water. Mix solutions A and B. Gram's Iodine: Dissolve 2 g of potassium iodide in 300 mL of distilled water; then add 1 g of iodine crystals. Kovac's Reagent: Mix the following: n-Amyl alcohol Hydrochloric acid p-dimethylamine-benzaldehyde Malachite Green Stain: Dissolve 5 g of malachite green oxalate in 100 mL of distilled water. Nigrosin Solution: Add 10 g of nigrosin (water soluble) to 100 mL of distilled water. Boil for 30 min, and add 0.5 mL of formaldehyde (40%). Filter twice through double filter paper. Store under aseptic conditions. 75 mL 25 mL 5.0 g 36.8 mL 13.2 mL


Oxidase Test Reagent: Dissolve 1 g of dimethyl-p-phenylenediamine hydrochloride in 100 mL of distilled water. Make fresh. Phloxine B: Dissolve 1 g of phloxine in 100 mL of distilled water. Safranin: Dissolve 0.25g safranin in 10 mL of 95% ethanol. Add to 100 mL of distilled water. Sudan Black Stain: Dissolve 0.3 g of Sudan Black in 100 mL of 70% ethanol. Shake before each use.

References: Clark, G. (1984) Staining Procedures. 4th Ed. Williams and Wilkins, Baltimore, Maryland. Benson, H.J. (1985). Microbiological Applications: A Laboratory Manual in General Microbiology, 4th Ed. Wm. C. Brown Publishers, Dubuque, Iowa.


pH INDICATORS: Table 1: Indicators of Hydrogen Ion Concentration.

pH Indicator Cresol Red Meta Cresol Purple (acid range) Thymol Blue Brom Phenol Blue Brom Cresol Green Chlor Cresol Green Methyl Red Chlor Phenol Red Brom Cresol Purple Bromothymol Blue Neutral Red Phenol Red Cresol Red Meta Cresol Purple (alkaline range) Thymol Blue (alkaline range) Cresolphthalein Phenolphthalein

pH Range 0.2 - 0.8 1.2 - 2.8 1.2 - 2.8 3.0 - 4.6 3.8 - 5.4 4.0 - 5.6 4.4 - 6.4 4.8 - 6.4 5.2 - 6.8 6.0 - 7.6 6.8 - 8.0 6.8 - 8.4 7.2 - 8.8 7.4 - 9.0 8.0 - 9.6 8.2 - 9.8 8.3 - 10.0

Full Acid Colour Red Red Red Yellow Yellow Yellow Red Yellow Yellow Yellow Red Yellow Yellow Yellow Yellow Colourless Colourless

Full Alkaline Colour Yellow Yellow Yellow Blue Blue Blue Yellow Red Purple Blue Amber Red Red Purple Blue Red Red

Adapted from: Benson, H.J. (1985). Microbiological Applications: A Laboratory Manual in General Microbiology, 4th Ed. Wm. C. Brown Publishers, Dubuque, Iowa.


APPENDIX 9 Care and Feeding of the Microscopes Checklist For Compound Microscopes Name:________________________________________ Class and section: ______________________________ Date:_________________________________________ Microscope #: _________________________________ Did you find the microscope in proper working order? Y or N If not, what was the problem? _____________________________________________ _____________________________________________ _____________________________________________ _____________________________________________ __ Slide removed from stage __ Slide, stage, and objectives are free of oil __ Mechanical stage is centered __ Stage placed at its lowest position __ 4x objective placed into working position __ Ocular micrometer replaced with regular ocular __ Binocular head secured in "start" position __ Rheostat turned to 0 and lamp is turned off __ Cord is wrapped tightly around arm and lamp __ Cord is secured with cord clip __ Dust cover is placed over scope Checklist For Dissecting Scopes Name:________________________________________ Class and section: ______________________________ Date:_________________________________________ Microscope #: _________________________________ Did you find the microscope in proper working order? Y or N If not, what was the problem? _____________________________________________ _____________________________________________ _____________________________________________ _____________________________________________ __ Turn off transformer __ Unplug transformer and lamp __ Wrap cord tightly around transformer __ Place transformer on stage with binocular head well above the transformer __ Replace dust cover



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