The University of Lethbridge
BIOLOGY 3200 Principles of Microbiology LABORATORY MANUAL Spring, 2005 Written by: L. A. Pacarynuk and H. C. Danyk Revised: January, 2005
TABLE OF CONTENTS
Exercise
Page
Biology 3200 Laboratory Schedule
2
Grade Distribution
3
Occupational Health and Safety Guidelines
4
Guidelines for Safety Procedures
5
Exercise 1 – Introduction to Microscopy
7
Exercise 2 – General Laboratory Principles and Biosafety
12
Exercise 3 - Bacterial and Yeast Morphology
14
Exercise 4 – Bacterial Reproduction
20
Exercise 5 – The Ames Test
25
Exercise 6 – Biochemical Tests
28
Exercise 7 – Virology
34
Exercise 8 - Soil and Compost Microbial Ecology
38
Exercise 9 - Applications of Microbiology*
47
Appendix 1 – The Compound Light Microscope
51
Appendix 2 – Preparation of Scientific Drawings
54
Appendix 3 – Aseptic Technique
56
Appendix 4 – The Cultivation of Bacteria
61
Appendix 5 – Bacterial Observation
66
Appendix 6 – Laboratory Reports
67
Appendix 7 – Use of the Spectrophotometer
69
Appendix 8 – Media, Reagents, pH Indicators
71
Appendix 9 – Care and Feeding of the Microscopes
84
*Will require some out of laboratory time for sampling.
1
BIOLOGY 3200 LAB SCHEDULE SPRING, 2005 Jan. 11 Jan. 13
Introduction, Microscopy General Lab Procedures, Biosafety
Jan. 18 Jan. 20
Bacterial Morphology Bacterial Morphology
Jan. 25 Jan. 27
Bacterial Morphology Bacterial Morphology; Hand in Assignment 1
Feb. 1 Feb. 3
Bacterial Growth Bacterial Growth - Complete
Feb. 8 Feb. 10
Ames Test; Biochemical Tests - Selective and Differential Media Ames Test – Complete; Biochemical Tests - Selective and Differential Media – Complete; Hand in Assignment 2
Feb. 15 Feb. 17
Biochemical Tests - IMViC Tests Biochemical Tests - IMViC Tests – Complete
Feb. 22 Feb. 24
Reading Week Reading Week
Mar. 1 Mar. 3
Virology Virology
Mar. 8 Mar. 10
Virology; Enumeration of Soil and Compost Bacteria Virology – Complete; Enumeration – Complete; Selection of Unknown and Streak Plate
Mar. 15
Microscopic and Macroscopic Observations of Unknown, Wine Fermentation Microbial Products (using soil/compost samples and unknown), Wine Fermentation
Mar. 17 Mar. 22 Mar. 24
Evaluation of Microbial Products; Experimental Design Experimental Design; Wine Fermentation – Complete
Mar. 29 Mar. 31
Identification of Unknown Identification of Unknown
Apr. 5 Apr. 7
Identification of Unknown Identification of Unknown – Complete; Hand in Lab Report
Thursday Apr. 14
Final Lab Exam (practical)
2
Laboratory Grade Distribution: The laboratory component of Biology 3200 is worth 50% of your course mark. It is distributed as follows: •
Assignments
•
Lab Report •
Wine Fermentation
•
Due Thursday, Apri.l 7 by 4:30 PM
7.5% 15%
•
In-Lab Skills Tests
7.5%
•
Lab Exam
20%
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. Unannounced skills tests will be given during the semester. Students are expected to work independently on some technical aspect of microbiology and will be graded based on their techniques and their results. As proficiency in microbiological techniques is considered an essential component of the course, students are only permitted 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: After 4:30 pm but prior to 9:00 am the next day - -25% (eg. if the assignment is out of 50 points, you will lose 12.5 marks); between 9:00 am and 4:30 pm –50%; etc. Extensions will only be considered upon application to your lab instructor no less than two days prior to the due date of the assignment. This application should include documentation and the portion of the assignment completed at that point. Failure to include any evidence of work completed will result in no extension being granted. The lab exam (April 14) is comprehensive, covering all aspects of the laboratory. It may contain a practical as well as a theoretical component.
3
THE UNIVERSITY OF LETHBRIDGE Policies and Procedures Occupational Health and Safety
SUBJECT:
CHEMICAL RELEASE PROCEDURE
Precaution must be taken when approaching any chemical release. 1. Unknown/Known Release • • • • •
Clear the area Call Security 2345 Do not let anyone enter the area Call Utilities at 2600 and request the air be turned off at the release site Security will immediately notify: Chemical Release Officer: 331.5201 Occupational Health and Safety:
394.8937 394.8716
EMERGENCY CALL LIST 0800 – 1600 2345 331-5201 2301 394.8937 394.8716
SECURITY CHEMICAL RELEASE OFFICER ADMIN. ASSISTANT OCCUPATIONAL HEALTH AND SAFETY
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.
4
•
GUIDELINES FOR SAFETY PROCEDURES
EMERGENCY NUMBERS City Emergency
9-911
Campus Emergency
2345
Campus Security
2603
Student Health Centre
2484
(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)
Closest emergency exit
2)
Closest emergency telephone and emergency phone numbers
3)
Closest fire alarm
4)
Fire extinguisher and explanation of use
5)
Safety showers and explanation of operation
6)
Eyewash facilities and explanation of operation.
7)
First aid kit
GENERAL SAFETY REGULATIONS 1)
Eating, drinking or gum chewing is prohibited in the laboratory.
2)
Always wash your hands after entering and prior to leaving the laboratory.
3)
Laboratory coats are required for all laboratories and must be stored in the lab when not in use.
4)
Report equipment problems to instructor immediately.
5)
Report all spills to the instructor immediately.
6)
Long hair must be kept restrained to keep from being caught in equipment, Bunsen burners, chemicals, etc.
5
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.
6
EXERCISE 1 INTRODUCTION TO MICROSCOPY MICROSCOPY To view microscopic organisms, their magnification is essential. The microscope is the instrument used to magnify microscopic images. Its function and some aspects of design are similar to those of telescopes although the microscope is designed to visualize very small close objects while telescopes magnify distant objects. Magnification is achieved by the refraction of light travelling though lenses, transparent devices with curved surfaces. In general, the degree of refraction, and hence, magnification, is determined by the degree of curvature. However, rather than using a single, severelycurved biconvex lens such as that of Leeuwenhoek's simple microscopes, Hooke determined that image clarity was improved through the use of a compound microscope, involving two (or more) separate lenses. Operation of the Compound Microscope Students should be familiar with all names and functions of the components of their compound light microscopes as demonstrated in Appendix 1. Properties of the Objective Lenses 1.
Magnification
Magnification is a measure of how big an object looks to your eye. The number of times that an object is magnified by the microscope is the product of the magnification of both the objective and ocular lenses. The magnification of the individual lenses is engraved on them. Your microscope is equipped with ocular lenses that magnify the specimen ten times (10X), and four objectives which magnify the specimen 4X, 10X, 40X, and 100X. Each lens system magnifies the object being viewed the same number of times in each dimension as the number engraved on the lens. When using a 10X objective, for instance, the specimen is magnified ten times in each dimension to give a primary or "aerial" image inside the body tube of the microscope. This image is then magnified an additional ten times by the ocular to give a virtual image that is 100 times larger than the object being viewed.
7
2.
Resolution
Resolution is a measure of how clearly details can be seen and is distinct from magnification. The resolving power of a lens system is its capacity for separating to the eye two points that are very close together. It is dependent upon the quality of the lens system and the wavelength of light employed in illumination. The white light (a combination of different wavelengths of visible light) used as the light source in the lab limits the resolving power of the 100X objective lens to about 0.25 µm. Objects smaller than 0.25 µm cannot be resolved even if magnification is increased. Spherical aberration (distortion caused by differential bending of light passing through different thicknesses of the lens center versus the margin) results from the air gap between the specimen and the objective lens. This problem can be eliminated by filling the air gap with immersion oil, formulated to have a refractive index similar to the glass used for cover slips and the microscope's objective lens. Use of immersion oil with a 100X special oil immersion objective lens can increase resolution to about 0.18 µm. Resolving power can be increased further to 0.17 µm if only the shorter (violet) wavelengths of visible light are used as the light source. This is the limit of resolution of the light microscope. The resolving power of each objective lens is described by a number engraved on the objective called the numerical aperture. Numerical aperture (NA) is calculated from physical properties of the lens and the angles from which light enters and leaves. Examine the three objective lenses. The NA of the 10X objective lens is 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.
8
Observing Bacteria Three fundamental properties of bacteria are size, shape and association. Bacteria generally occur in three shapes: coccus (round), bacillus (rod-shaped), and spirillum (spiral-shaped). Size of bacterial cells used in these labs varies from 0.5 µm to 1.0 µm in width and from 1.0 µm to 5.0 µm in length, although there is a range of sizes which bacteria demonstrate. Association refers to the organization of the numerous bacterial cells within a culture. Cells may occur singly with cells separating after division; showing random association. Cells may remain together after division for some interval resulting in the presence of pairs of cells. When cells remain together after more than a single division, clusters result. Cell divisions in a single plane result in chains of cells. If the plane of cell division of bacilli is longitudinal, a palisade results, resembling a picket fence. Both bacterial cell shape and association are usually constant for bacteria and hence, can be used for taxonomic identification. However, both properties may be influenced by culture condition and age. Further, some bacteria are quite variable in shape and association and this may also be diagnostic. Micrometry When studying bacteria or other microorganisms, it is often essential to evaluate the size of the organism. By tradition, the longest dimension (length) is generally stressed, although width is sometimes useful for identification or other study. Use of an Ocular Micrometer (Figure 1) An ocular micrometer can be used to measure the size of objects within the field of view. Unfortunately, the distance between the graduations of the ocular micrometer is an arbitrary measurement that only has meaning if the ocular micrometer is calibrated for the objective being used. 1) Place a micrometer slide on the stage and focus the scale using the 40x objective. 2) Turn the eyepiece until the graduations on the ocular scale are parallel with those on the micrometer slide scale and superimpose the micrometer scale. 3) Move the micrometer slide so that the first graduation on each scale coincides. 4) Look for another graduation on the ocular scale that exactly coincides with a graduation on the micrometer scale. 5) Count the number of graduations on the ocular scale and the number of graduations on the micrometer slide scale between and including the graduations that coincide. 6) Calibrate the ocular divisions for the 40x and the 100x objective lenses. Note that immersion oil is not necessary for calibration.
9
Stage Micrometer (each division = 0.01 mm)
0
Ocular Micrometer 0
5
10
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 µm/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.
10
METHODS: For each student: •
Compound light microscope
•
Various prepared slides of bacteria.
•
Stage micrometer
•
Ocular micrometer
•
Immersion oil
1) Use the diagram in Figure 1 to calibrate the 40x and the 100x objectives on your compound microscopes. Record these values in your lab book as you will then use these values when measuring cells and structures for the rest of the lab. Note: Do NOT use immersion oil when calibrating the 100x objective. This is the ONLY time during the term that you will not use immersion oil with this objective. 2) Use the compound microscope to observe the prepared slides of bacteria using the 10x and 40x objective lenses. Observe the same slides under the 100x objective using immersion oil. 3) Diagram two of the organisms viewed following the instructions found in Appendix 2.
11
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: http://www.uleth.ca/fas/bio/safety/biosafety.html EXPERIMENTAL OBJECTIVES Students will use fluorescein dye-labelled E. coli cultures to perform a series of exercises designed to illustrate the potential for contamination that is always present when working with microorganisms. As well, students will become familiar with using aseptic techniques to handle microorganisms. METHODS Benches will be provided with the following: •
Fluorescein-labelled broth culture of E. coli (ATCC strain)(2/bench)
•
Nutrient agar plates (8/bench)
•
Nutrient broth (4 tubes/bench)
•
Bench coat
•
Tape
•
Gloves
•
Hand-held UV lamp
•
Watch glasses (2/bench)
•
Sterile pipettes
•
Pipette pump
•
Tray containing bleach disinfectant
Wear gloves for the entire exercise. 1) Tape bench coat onto the bench to cover your working surface. 2) Work individually over the bench coat and prepare a streak plate for single colonies. Label and place in the tray on the side to be incubated. 3) From the same suspension, inoculate one tube of nutrient broth. For steps 4 - 11, work in pairs. 4) Place a watch glass in the centre of the bench coat. 5) Obtain and label 2 NA plates (name, date, organism, distance). Place agar plates on either side of the glass plate, one 5 cm and the other 10 cm from the watch glass. 6) Using a pipette pump, draw up 2 mL of bacteria/fluorescein suspension.
12
7) Remove lids from agar plates and set aside. 8) Hold pipette tip 30 cm from glass plate and allow 10 drops to fall (one drop at a time) onto the glass plate. Put any remaining bacterial culture back into the original culture tube. 9) Remove glass plate to disinfectant tray and cover agar plates. Place on a tray on the side bench. 10) Use the hand-held UV lamp in C741 to inspect your bench coat, gloves, and lab coat. What do you observe? 11) Your plates will be incubated for 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.
13
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 et al, 2003, pp. 56-63) or dark-field microscopy, which is also referred to as negative staining (Negative staining will be utilised later on this laboratory). 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 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.
14
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. β-hydroxybutyric Acid (PHB) Staining Poly-β PHB granules are common inclusion bodies in bacteria. Monomers of β-hydroxybutyric acid are connected by ester linkages forming long polymers which aggregate into granules. As these granules have an affinity for fat-soluble dyes such as Sudan black, they can be stained and then identified with the light microscope. These granules are storage depots for carbon and energy. Endospore Staining Certain bacteria may produce endospores under unfavourable environmental conditions. Endospores are mainly found in Gram-positive organisms, including the Gram-positive Clostridium and Bacillus, in the Gram-positive cocci Sporosarcina, and in some of the filamentous Gram-positive Monosporaceae family. It has also been discovered that Coxiella burnetii, a small rod found in raw milk that has a variable Gram stain reaction, but a typical Gram-negative cell wall has a sporogenic cycle. When conditions become more favourable, the endospores will germinate and the bacteria will return to the actively growing and dividing form. Endospores are highly resistant to heat, chemical disinfectants and to desiccation and therefore allow the bacterial endospore to survive much more rigorous conditions than the vegetative cells. Endospore resistance is due to several factors, including: •
A decrease in the amount of water compared to vegetative cells
•
An increase in the amount of dipicolinic acid and calcium ions
•
Enzymes which are more resistant to heat
•
A spore coat which is impermeable to many substances
Endospores may be formed in a central, terminal, or sub-terminal position in the cell and their shape varies from ellipsoidal to spherical. The location of the endospore in the cell is usually characteristic of the species. For example, the location and shape of the Bacillus
15
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.
16
Prepare scientific diagrams (Appendix 2) showing results from each stain on separate pieces of paper. These will be collected and graded. If the stain is for a specific structure, ensure this structure is diagrammed and labelled. 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) Add Gram’s iodine solution to the slide for 1 min. Wash gently with tap water and remove as above. 3) 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) Counterstain the slide with safranin for 30 seconds - 1 minute. 5) Wash the slides with tap water, air dry on paper towels, and examine under oil immersion. Gram positive organisms stain purple; Gram negative organisms, red (pink). Acid-fast Staining - Procedure Perform on Mycobacterium smegmatis and on Escherichia coli 1) Flood the dried, heat fixed film with Ziehl’s carbol fuchsin and place on the rack over the boiling water bath. 2) Steam gently for 5 minutes. Do not let the slide dry out. Add more carbol fuchsin as required. 3) Wash with tap water to remove excess stain.
17
4) Decolourise with 20% sulfuric acid until no more stain comes out. Wash with tap water to remove excess. 5) Counterstain with methylene blue for 1 minute. Acid fast organisms retain the red stain while others are stained blue. PHB Staining - Procedure Perform on Bacillus thuringiensis. 1) Prepare smears of the organism, air dry and heat fix. Flood entire slide with Sudan Black B and add more stain as the dye solvent evaporates. Stain for at least 10 minutes. 2) Pour off excess stain (do not wash) and air dry. 3) Clear slide by dipping in a jar of solvent in the fume hood for 5 sec. Air dry in the fume hood. 4) Counterstain for 1 min. with safranin. 5) Wash with water, drain, blot and air dry. Examine with oil immersion objective. Cytoplasm is pink, lipids are dark grey or black. Endospore Staining - Procedure Perform on Bacillus thuringiensis. 1)
Prepare smear and heat fix. Cover the dried fixed film with a small piece of paper towel. Saturate this with 5% malachite green.
2)
Place the slide on a rack over a boiling water bath. Steam slide for 5-10 minutes in this manner. Add additional stain as needed - do not allow the slide to dry out during this procedure.
3)
Allow the slide to cool, then rinse with water. Tap over a paper towel to remove excess water
4)
Counterstain with safranin for 30 seconds.
5)
Rinse slide with water.
6)
Allow to air dry, and view.
Endospores will stain green and the rest of the cell pink. Yeast Staining – Procedure Perform on Saccharomyces bayanus 1) Prepare a wet mount of the cells using a drop of Methylene Blue. 2) Carefully place a cover slip on the cell/stain mixture. 3) View the cells noting size and shape. If you look carefully, you should be able to see budding cells.
18
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., Martinko, J. M., and Parker, J. 2000. Brock Biology of Microorganisms Ninth Edition. Prentice-Hall of Canada, Inc., Toronto. Ross, H. 1992-1993. Microbiology 241 Laboratory Manual. The University of Calgary Press, Calgary.
19
EXERCISE 4 BACTERIAL REPRODUCTION MEASUREMENT OF BACTERIAL GROWTH (See Madigan, et. al., 2003. Chapter 6 Pg.137151) 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). For example Escherichia coli has a generation time of 20 minutes under optimal conditions (e.g., 37°C, vigorous aeration and a rich growth medium). 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.
20
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 bottles of molten Luria-Bertani (LB) agar
•
10% bleach
•
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
•
65 oC water bath
•
Sterile d2H2O
•
Spectrophotometer blank containing TB broth
•
Bacterial waste container
•
Vortex
•
Cuvettes
•
Spectrophotometer
•
Culture flask of E. coli
21
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.
Culture sampling
1) 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. 2) 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. 3) At 20 minute intervals aseptically obtain one 5 mL sample of culture and immediately place it in a spectrophotometer tube. This material will be used to measure optical density (OD) (Section B). After reading, dispose of your 5 mL sample of culture in the waste beaker provided. Rinse the spectrophotometer tube using the squirt bottle of bleach provided and then dispense the solution into the waste beaker. 4) Remove another 100 µL of the culture and place it into a sterile microfuge tube. Label this tube Tube 1. Use this culture for Section C.
B.
Determination of optical density (please read Appendix 7)
1)
Zero the spectrophotometer as outlined in Appendix 7.
2)
Place the spectrophotometer tube containing your culture into the spectrophotometer and record the optical density (Absorbance) reading in your lab book and in the table on the blackboard. If the reading is greater than 0.7, you must dilute your sample and remeasure the optical density. It is suggested that you begin by diluting your sample 1:1 with the TB 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.
22
C.
Enumeration of viable bacteria
1)
Remove four sterile microfuge tubes from the container on the side bench. In order that you don’t contaminate all of the tubes, gently tap out four tubes from the container rather than using your hand to grab tubes.
2)
Set up your serial dilutions according to the information in Table 4.1. Aseptically pipette 900 µL of TB 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
Amount of
Amount of
Final Dilution
Number
µL) sterile TB (µ
Culture
Factor
1
900
100 µL from
10-1
culture flask 2
990
10 µL from tube
10-3
1 3
990
10 µL from tube
10-5
2 4
900
5 (for labs
900
100 µL from
10-6
tube 3 3 and 4
100 µL from
10-7
tube 4
only)
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). Labs 3 and 4 will be plating the contents of Tube 4 and Tube 5 (10-6 and 10-7). 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)
Follow the instructions provided in step 4 above to plate out the contents of Tube 4.
23
6)
When the agar has solidified, place the inverted plates on a tray at the side of the lab. The plates will be incubated for 16 – 20 hours at 37°C and refrigerated until the next lab session.
The next laboratory period: 7)
Examine the plates carefully and select the plate where the bacterial count ranges between 30 and 300 colonies.
8)
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.
9)
Use class data to determine the average number of bacteria per mL of culture.
10)
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, et. al, 2003)
Prepare a Results and Discussion section upon conclusion of this laboratory according to the information found in Appendix 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?
24
EXERCISE 5 THE AMES TEST MUTATION AND RECOMBINATION (See Madigan, et. al., 2003. Chapter 106 Pg. 265-276) You have learned about some of the advantages of using a model system in your study of the effect of UV light on DNA in Biology 2000 (Introduction to Genetics). The Ames test also makes use of a model system in order to measure the mutagenic potential of compounds. This test is a reversion mutagenesis assay and uses strains of the bacterium Salmonella that have point mutations in various genes in the histidine operon. These His- mutants are unable to synthesise histidine and therefore unable to grow on minimal media lacking histidine. When the His- tester cells are cultured on a minimal agar medium containing trace amounts of histidine, a small and relatively constant number of cells per plate spontaneously revert to His+ and subsequently reproduce and form colonies. Incorporation of a mutagen into the agar increases the number of revertant colonies per plate, usually in a dose dependent manner. EXPERIMENTAL OBJECTIVE You will make use of the Ames test in order to evaluate the mutagenicity of a selection of compounds. PRE-LAB PREPARATION Each class should bring in a total of three household compounds they would like to test. These will be decided in advance. Note that these compounds must be known (ie “mystery liquid” from the garage is not acceptable) and they must be taken home again once Period 1 of the lab is finished.
METHODS: For each lab: •
100 mg/mL Sodium Azide (CAUTION: MUTAGEN!)
•
Ethidium bromide (10 mg/mL)
•
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
25
•
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 #
1 2
Compound to be Tested Water Unknown Unknown 1 2 + 1535 +1535 + 1538 + 1538 +1535 +1538
Unknown 3
3
Sodium Azide
+1535 +1538
4
+1535 +1538
5
1)
Ethidium bromide
+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 redhot! 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. For the cigarette extract, you will need to go to the fume hood to do this.
26
7) 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. 8) 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. Thought Questions: •
What specific mutations in the His operon do each of the Salmonella strains used contain?
•
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)
•
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., Martinko, J. M., and Parker, J. 2003. Brock Biology of Microorganisms Tenth Edition. Prentice-Hall of Canada, Inc., Toronto.
27
EXERCISE 6 BIOCHEMICAL TESTS (Selective and Differential Media; IMViC Tests) Normally, the coliform group of bacteria is used to indicate the pollution of water with fecal wastes of humans and animals, and thus, the suitability of a particular water supply for domestic use. The term coliform is used to describe aerobic and facultatively anaerobic Gram negative rods that ferment lactose with gas formation. Most, but not all organisms within this group are intestinal in origin; for instance, Escherichia coli. Consequently, presence of lactose fermentors in a sample of water provides circumstantial evidence of pollution by fecal wastes, and may suggest the presence of pathogenic bacteria such as members of the genera Salmonella and Shigella. These pathogens, in addition to non-pathogens such as E. coli are members of the Enterobacteriaceae family. In order to identify the organisms present in the water, several biochemical tests that rely on differences in the chemical composition of media used may be performed (see Appendix 4 and Appendix 8 for more details). SELECTIVE AND DIFFERENTIAL MEDIA: I. Media for Isolation of Enterobacteriaceae A strategy for bacterial isolation involves the use of selective media, media with specific components that promote the growth of some bacteria and inhibit the growth of others. Selectivity may be achieved in three ways: •
by adding something to the medium to discourage the growth of species not required
•
by altering the pH of the medium
•
by omission of some ingredient required by most bacteria, but not by the organism to be isolated
Differential media contain specific biochemical indicators that demonstrate the presence of certain substances characteristic of certain bacteria. Thus, differential media are useful for bacterial identification. Eosin Methylene Blue Agar (EMB Agar) EMB is both a differential and selective plating medium recommended for use in the isolation of Gram-negative bacilli and the differentiation of lactose fermentors from non-lactose fermentors. EMB agar contains the two indicators, eosin Y and methylene blue as well as the carbohydrate lactose. Eosin (an acidic dye) reacts with methylene blue (a basic stain) to form a compound of either acidic or neutral nature. The acid produced by lactose fermentors is sufficient to cause this dye compound to be taken up by the cells. Non-lactose fermentors are colourless because the eosin and methylene blue compound cannot be taken up by the cells. The basic stain methylene blue inhibits bacterial growth, particularly that of Gram positive
28
bacteria (due to their cell wall composition). Eosin methylene blue (EMB) agar is thus selective for Gram negative bacteria. MacConkey Agar MacConkey agar is a differential and selective plating medium recommended for use in the isolation of Gram-negative bacilli and the differentiation of lactose fermentors from nonlactose fermentors. The differential action of the MacConkey agar is indicated by the colonies of coliform bacteria becoming “brick red” in colour. This occurs when the coliforms utilise the lactose producing acids. The decrease in pH results in the uptake of the indicator neutral red by the cells. Non-lactose fermentors are colourless and transparent. Production of acid may also result in a zone of precipitated bile surrounding the colony. Bile salts and crystal violet present in the medium inhibit Gram-positive bacteria from growing. II. Acid Production from Carbohydrates As demonstrated with MacConkey Agar, bacteria vary in their ability to ferment various sugars. Products of fermentation are often acids and hence, pH changes can demonstrate successful fermentation. In addition, gas (usually but not always CO2) is often produced during fermentation, offering another indicator. Hugh and Leifson’s method for demonstrating the presence of the products of fermentation consists of a semi-solid medium containing peptone (short chains of amino acids), the carbohydrate of interest (usually glucose or lactose), and a pH indicator, Bromothymol blue. Tubes are stab-inoculated all the way to the bottom of the tube, so as not to introduce oxygen into the medium. Several reactions may be observed. Facultative organisms will produce an acid reaction (the indicator changes to yellow) throughout the entire tube of medium. The acid reaction produced by oxidative organisms is apparent first at the surface, extending gradually downwards into the medium. Note that organisms that oxidise glucose are generally unable to ferment any carbohydrate. Strict fermentors will produce an acid reaction at the bottom of the tube. Organisms unable to use the carbohydrate may be able to grow using the peptone in the medium. Production of alkaline products result in the formation of a blue colour at the top of the tube (although this does not indicate that the organism is aerobic). III. Motility Medium This medium contains triphenyl tetrazolium chloride and a small concentration of agar in order to make the medium semi-solid. TTC is reduced when broken down by the organism, and the TTC turns red where this has occurred. If the organism is facultative and motile, it moves throughout the entire tube of medium and the whole tube becomes red. If the organism is aerobic and motile, the top of the tube becomes red.
29
METHODS For each bench: •
3 plates each of MacConkey and EMB media
•
5 known broth cultures
•
1 ‘unknown’ broth culture
•
6 tubes of Hugh and Leifson’s (H & L) lactose medium
•
6 tubes each of motility medium
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 in your lab book. Generate a table of results summarising growth and properties of all bacteria on the two media. 2) Determine the lactose fermentation ability of all of the bacteria provided. These tubes are inoculated using a stab technique. Use the probe to aseptically remove a small amount of bacterial culture, and then stab the probe to the bottom of the tube of medium without mixing the medium around. Inoculate each tube with one of the bacterial species and label appropriately. Tubes will be incubated for 48h at 37°C. After incubation, observe tubes and record results in your lab book. 3) Work collectively to inoculate your motility medium tubes. Again, as this medium is semi-solid (the stab technique is used for all semi-solid media in this course), use a probe and stab the culture down to the bottom of the tube and remove the probe. Do not mix the probe around in the tube. Tubes will be incubated for 48 h at 37oC. After incubation, observe tubes and record the results in your lab book. IMViC TESTS Only preliminary taxonomic assessment of bacteria can be made on the basis of microscopic size, shape, association, and Gram staining. Information regarding natural occurrence is also valuable since bacteria generally occur in specific habitats. This is particularly the case for fastidious bacteria, those with very specific nutritional and environmental requirements. However, even when supplemented with habitat information, bacterial identification based on microscopic assessment is generally incomplete. Confident bacterial identification can be made based on biochemical tests, and for certain pathogens, or for examining microbial presence in specific environments, series of diagnostic tests have been developed. For example, the IMViC tests are used routinely to confirm the presence of coliform organisms in water. “IMViC” is an acronym for ‘Indole, Methyl Red, Voges-Proskauer, and Citrate utilisation’ tests (the “i” is inserted for ease of pronunciation). I. Indole Formation - Utilisation of Tryptophan
30
When cultured on peptone water, a liquid medium containing tryptophan, certain bacteria will produce indole. The presence of this indole is readily revealed through addition of Kovak's reagent, producing a pink colour. This reagent contains the organic solvent amyl alcohol that extracts the coloured (pink) substance. II. The Methyl Red (MR) Test - Mixed Acid Fermentation Pathway Fermentation of glucose via the mixed acid fermentation pathway results in the formation of a number of organic acids such as lactic and acetic acid. If this is a primary fermentation pathway of a bacterium, a noticeable drop in pH will occur with incubation on MRVP media. This decrease in pH can be revealed by a methyl red solution which is yellow under neutral conditions and red at a pH less than 5. III. The Voges-Proskauer (VP) Test - The Butanediol Fermentation Pathway An alternate fermentation pathway performed by some other bacteria results in the formation of a non-acidic product, butanediol and hence, is named for this product. The occurrence of the pathway may be determined by a biochemical test for an intermediate compound in the pathway, acetoin (acetyl methyl carbinol), which is detected by the VogesProskauer test.
IV. Citrate Utilisation - Growth Using A Single Carbon Source The nutritional requirements of different bacteria vary considerably and these can provide useful information contributing to biochemical identification. In Simmon's citrate agar, citrate, in the form of sodium citrate, is the sole carbon source. Organisms able to utilise the citrate grow on the surface of the medium and due to oxidative formation of sodium carbonate, raises the pH of the medium changing it from green to blue (bromothymol blue is the indicator). V. Urea Hydrolysis Some bacteria can produce urease, an enzyme which hydrolyses urea into ammonium and carbon dioxide. The presence of this enzyme is detected by growing the bacteria in a medium containing urea and a pH indicator, phenol red. If ammonium is produced as a result of urea hydrolysis, the increase in pH will turn the medium to a violet-red colour.
31
METHODS: For each bench: •
6 broth cultures, one of which is an ‘Unknown’
•
6 MRVP broth tubes
•
6 indole broth tubes
•
6 Simmons citrate agar slants,
•
6 Urea broth tubes
Please work in groups of four. 1) Inoculate 6 Indole broth tubes separately with the 6 bacteria. After 48h of incubation at 37 oC, add 20 drops (1 mL) of Kovak's reagent (work in the fume hood and leave your tubes in the fume hood to develop and observe). Shake and look for the formation of a pink colour in the top (organic) phase; it may take 20 minutes to develop. The pink colour is a positive result, indicating the ability to use tryptophan. Note, please place tubes containing amyl alcohol in a separate rack in the fume hood as this material needs to be disposed of separately. 2) A single culture solution (peptone, glucose, potassium phosphate) will be used for both the methyl red and Voges-Proskauer tests. Inoculate 6 MRVP tubes with the 6 bacteria provided, one culture into each tube. •
After 48h of incubation at 37°C, remove about 1/4 of the broth (=2 mL = 40 drops) from the MRVP tube and transfer that to another test tube. Add 3-5 drops of methyl red solution. An immediate red reaction provides a positive response to the test, indicating the presence of mixed acid fermentation. A yellow or orange colour represents a negative response.
•
As the same solutions are used for the MR and VP, remove an additional 2 ml of culture solution and add 1 ml α-napthol (Barritt's reagent A - 1 ml is about 20 drops) and 1 ml 40% KOH (Barritt’s reagent B; caution - this is caustic). Shake vigorously for 30 seconds.
•
Shake the tubes frequently and observe for up to 30 minutes for the formation of a red colour that represents a positive VP test. A yellow or brown colour is a negative result.
3) Inoculate 6 Simmon's citrate agar slants separately with the 6 bacteria. For these inoculations, use your loop to smear cells along the surface of the slant. Incubate tubes for 48 h at 37°C.
32
•
After incubation, observe colours on the surface and down through the tubes. A dark blue colour is a positive result while green indicates a negative test for citrate utilisation.
4) Inoculate 6 urea slants separately with the 6 bacteria. After incubation for 48h at 37°C, observe for the development of a violet-red colour. 5) After completing the Indole, MR, VP, Citrate and Urea tests, collaborate with the other students at your bench to generate in your lab book tables of results for all bacteria in all tests.
Thought Questions: •
Identify your unknown. Can it be any of the knowns? Why or why not? 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).
33
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 o
on the web page. Both samples were stored at 4 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 C)
•
Sterile test tubes
• •
Test tube rack Micropipettor (100 µL – 1000 µL)
•
Sterile tips
•
Microbiology kits
o
For the lab: •
Vortex mixer
•
Water bath set to 60 C
•
Raw and treated sewage filtrate
•
Test tube showing 4 mL mark
o
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.
34
2) Obtain 5 Luria Methylene Blue agar plates, and 5 sterile test tubes. Label your 5 tubes according to Table 7.1. Table 7.1 Experimental set-up for isolation of coliphage from sewage. µ L) Contents (µ Tube # K12
Raw Sewage
Treated Sewage
Filtrate
Filtrate
1
500
0
0
2
0
500
0
3
0
0
500
4
500
500
0
5
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 o
o
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
35
•
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? o
Place your tubes in the rack on the side bench. The tubes will be stored at 4 C allowing the phage to elute from the agar into the buffer. PART B – HOST RANGE METHODS Overnight cultures of: •
Salmonella typhimurium strain 1535
•
E. coli strains CSH121 and CSH125 and K12
•
Proteus vulgaris
•
Enterobacter
Other supplies: • Phage dilution buffer • Micropipettors and sterile tips • Autoclave waste disposal • Luria Methylene Blue agar plates • LB plates • Bottle of molten Luria agar overlay (at 60 C) o
• Sterile test tubes • Test tube indicating 4 mL mark • Test tube rack • Micropipettor (100 µL – 1000 µL) • Sterile tips
36
•
Microbiology kits
For determining phage titre: -2
-4
-6
-8
1) Prepare serial dilutions of your phage in dilution buffer (10 , 10 , 10 , 10 ) 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 o
dilution. Do not invert. Plates will be incubated at 37 C overnight. 3) The next day, score as + or – for phage growth on each host. Thought Questions: •
Based on the schematic found on Dr. Brent Selinger’s web site, what step(s) is/are most likely responsible for the difference in coliphage numbers between raw and treated sewage?
•
Have you isolated more than one type of phage? How might you be able to tell?
•
To what components of the bacterial cell to phage typically adhere?
37
EXERCISE 8 SOIL AND COMPOST MICROBIAL ECOLOGY Soil Bacteria The microflora of the soil exist as a complex food chain that brings about the release of nutrients from dead plant material on the surface. The surface layer of newly fallen plant material is called the litter and chemically it is composed of insoluble materials such as cellulose, hemicellulose and lignin. Only a few organisms, usually fungi, are able to utilise these high molecular weight compounds since carbohydrates, amino acids, vitamins and other growth factors are lacking. However, once microorganisms do begin to decompose the litter, the chemical structure of the litter is modified and the organisms produce end-products that are released into the environment and become available for use by others. Death of these organisms also provides new small molecular weight compounds that may then be utilised. Bacteria are able to utilise the end-products of fungal metabolism. Nematodes and protozoa feed on the bacteria and mites and other animals live on the nematodes and protozoa. In this way nutrients are recycled. Compost Bacteria Composting is a microbial process whereby plant matter including lignin is partially converted to humus, therefore supplementing the organic content of soil. The process is initiated by mesophilic heterotrophs and initially is characterised by a temperature increase up to 55 – 60oC for a few days where thermophiles such as Bacillus stearothermophilus and Thermomonospora are active. The temperature then decreases, followed by several months of curing at mesophilic temperatures, where again, mesophiles predominate. Composting is not exclusively carried out by bacteria; fungi such as Aspergillus fumigatus, and Geotrichum candidum, are also involved. EXPERIMENTAL OBJECTIVES In this experiment, you will prepare serial dilutions of compost and of soil samples and plate out the appropriate dilutions. After incubation, you will determine the number of bacteria isolated in your two samples, and assess the microorganisms for their ability to utilise carboxymethylcellulose, casein, starch and xylan. You will choose one organism from either soil or compost, use biochemical tests to identify the microorganism you have chosen, and use the class results to compare and contrast microbial diversity in soil and in compost.
38
METHODS: •
Microbiology kits
•
Soil
•
Compost
•
Balance
•
Bottles containing 100 mL sterile water
•
Vortex
•
Petri dishes
•
250 mL bottles of molten peptone yeast extract agar in 60oC water bath
•
9 mL water blanks
•
Sterile pipettes, propipettors
•
Crystal violet
•
Safranin
•
Carbol fuchsin
•
Gram’s iodine
•
PYE broths
•
Sudan black
•
95% ethanol
Out of your group of four, one pair will prepare enumeration plates for soil, while the other pair will prepare enumeration plates for compost. A. Enumeration of bacteria in soil and in compost 1)
Weigh 1 g of soil or of compost provided and add to 100 mL of sterile distilled water. This is dilution #1 (1:100). Shake the suspension for 5 minutes.
For those pairs working with soil, please follow the instructions outlined in steps 2-5; those pairs working with compost, please follow steps 6-9.
2)
Use the sterile 9 mL distilled water blanks provided to create serial dilutions of your soil. Please ensure that you vortex your samples well prior to making each new dilution. You will require 10-4 and 10-5 dilutions of soil.
3)
Add 1 mL of the 10 -4 dilution to each of two sterile, labeled Petri dishes and 1 mL of the 10 -5 dilution to two labeled Petri dishes.
4)
Obtain a 250 mL bottle of molten peptone yeast extract agar – label with tape and leave in waterbath when not in use.
5)
Use the sterile 9 mL distilled water blanks to create serial dilutions of your compost. Please ensure that you vortex your samples well prior to making each new dilution. You will require -4
-5
-6
10 , 10 , and 10 dilutions of your compost.
39
6)
Add 1 mL of the 10 -4 dilution to each of two sterile Petri dishes, 1 mL of the 10 -5 dilution to two sterile Petri dishes, and 1 mL of the 10-6 dilution to two sterile Petri dishes.
7)
Obtain your labeled 250 mL bottle of molten peptone yeast extract agar from the water bath.
8)
Add approximately 20 mL of medium to the plates prepared in Step 7. Swirl carefully to mix the inoculum evenly with the medium.
Both pairs should make note of step 9: 9)
The plates will be incubated for 24 hours at 30oC and refrigerated until the next lab session.
Results (please work as a group of 4 to enumerate the organisms) Examine both sets of plates carefully and select the plates where the bacterial count ranges between 30 and 300 colonies. Record the number of colonies on the plates in your notebooks and on the board and determine the mean (± standard deviation) number of bacteria per g of soil and of compost. B. Isolation and characterization of bacteria from soil or compost Two classes will identify organisms from compost bacteria plates while the remaining classes will identify organisms from soil bacteria plates. Your instructor will indicate what plates to remove your unknown from. Work individually to complete Part B of this exercise. Choose a morphologically distinct colony from the plate provided by your instructor and prepare a streak plate for single colonies on peptone yeast extract agar (Appendix 3). The plates will be incubated for 24 hours at 30oC and then stored at 4oC. Prepare a Gram stain of the pure culture and record the cell morphology. Record the colony morphology of the culture (see Appendix 6). Prepare a liquid culture of a single colony using PYE broth. Use this culture to inoculate all of the biochemical test media you use. Use the dichotomous key provided to develop a detailed outline of the series of steps you plan to take to identify your unknown. This outline should include tests to carry out as well as dates when you intend to do these tests. This outline must be handed in and approved before you will be allowed to proceed. The following media and reagents will be available for you to utilise as you attempt to identify your unknown:
40
•
nutrient agar
•
endospore stain reagents
•
capsule stain reagents
•
hydrogen peroxide (catalase test)
•
oxidase reagent (oxidase test)
•
IMViC reagents
•
indole broths
•
MRVP broths
•
citrate slants
•
urea broths
•
litmus milk broths
•
mannitol broth (with phenol red indicator)
•
H & L medium containing glucose
•
H & L medium containing lactose
•
sucrose agar
•
motility medium (with TTC)
41
1.
Gram stain Gram positive ......................................................................................…………………...... 2 Gram negative...........................................................................................………………….16
2.
Cell morphology bacillus or spirillus……...................................................................………….........………...3 coccus….................................................................................................……………………..12 ovoid….....................................................................................……………………Azotobacter
3.
Endospore stain positive.......................................................................................................……..……………..4 negative ...................................................................................…………………................... 6
4.
1
5.
Aerobe or facultative anaerobe .................................................................………………..Bacillus Obligate anaerobe..............................................................................…………………..Clostridium
6.
Bacillus cells may be branched, no true mycelium................….....……………........................ 7 Bacilli form true mycelium...............................................................................………………….10
7.
Cells pleomorphic depending on age of culture...................................................……………...8 Cells bacillus-shaped only. Club-shaped swelling may be present in young cultures....................................…................................................…………………..9
8.
Cells pleomorphic becoming coccoid with age. Gram reaction of bacilli and cocci usually positive....................................................………………Nocardia Gram-positive coccoid cells in older cultures. Coccoid cells germinate to produce bacillus-shaped cells. Bacilli may be Gram negative with Gram positive granules..............................……………..Arthrobacter
9.
Catalase positive ........................................................................………………….Corynebacterium Catalase negative .............................................................................…………………..Lactobacillus
10.
Conidia or sporangia formed within one week ..................................................……………...11 No conidia formed, anaerobic or microaerophilic ...........................………………Actinomyces
11.
Chains of conidia formed; colony may produce brown water soluble pigment and have an “earthy” smell ............................……………..Streptomyces Single conidia only.............................……………………Micromonospora or Thermoactinomyces
12.
Cells arranged singly, or in chains or clusters................................................…………………13 Cells in cubical packets..............................................................................……………………….15
13.
Catalase negative...........................................................................................……………………..14 Catalase positive ............................................................................................……………………15
1
Endospores are seen within the vegetative cells on an endospore stain after growing on sporulation agar for 48 hours. Rod shaped cells break up into coccoid shapes or conidia after growing on an agar plate for several days
2
2
True endospores......................................................................................…………………........... 5 Special spore types..........................................................................................……………………6
42
14.
Large, mucoid colonies on sucrose agar; microaerophilic or facultative anaerobic; capsule present.......................................………………..Leuconostoc Small, round (1-2 mm) colonies on sucrose agar, no capsule.........………………Streptococcus
15.
Glucose fermented .......................................................................…………………..Staphylococcus Glucose not fermented ..........….........................................................……………….. Micrococcus
16.
bacillus shaped...............................................................................................…………………….17 coccus shaped .......................................................................................…………………...Neisseria ovoid .................................................................................................……………………Azotobacter
17.
Red or purple pigmented colonies on agar plate.............................................………………. 18 No red or purple pigmented colonies; not associated with root formations in plants.................................................................................…………………..19 No red or purple pigmented colonies; associated with root formations in plants. .................................................................................………………... 33
18.
Acid from mannitol, pigment soluble in acetone:alcohol ................…………...…....... Serratia No acid from mannitol.............................................................………………….Chromobacterium or Rhodopseudomonas or Rhodospirillum
19.
Organisms produce a green, blue, brown or yellow watersoluble pigment which diffuses into the medium. Glucose respired; oxidase positive; aerobic; motile............................………………...Pseudomonas No water-soluble pigment produced.................................................................………………..20
20.
Curved or bent bacilli on Gram stain...............................................................…………………21 Straight bacilli................................................................................................……………………..22
21.
Bent bacilli, methyl red (-), Voges Praskauer (+), catalase (+)......................……………..Vibrio Spiral bacilli, no growth in peptone water (indole broth) without cellulose strip....................................................................…………………Spirillum
22.
Glucose not utilised...........................................................…………………................................ 23 Glucose utilised facultatively.......................................................................………………….... 25 Glucose utilised aerobically..............................................................................………………….28
23.
Yellow pigmented colony.............................................................…………………Flavobacterium Non-pigmented colony......................................................................................…………………24
24.
Litmus milk alkaline, oxidase positive, aerobic, motile….…………….................... Alcaligenes Litmus milk alkaline, oxidase negative, non-motile…......................……………..Acinetobacter
25.
Lactose fermentation produces acid.................................................................…………………26 No acid from lactose........................................................................................…………………...29
26.
Methyl red (+), no growth on citrate, fecal odor on BHI......................……………..Escherichia Methyl red (-), growth on citrate......................................................................………………….27
27.
Non-motile...........................................................................................…………………….Klebsiella Motile..............................................................................................……………………..Enterobacter
28.
Yellow pigmented colonies..........................................................……………….....Xanothomonas Non-pigmented colonies....................................................................………………….Acetobacter
29.
Urease (+)........................................................................................................…………………….30 Urease (-).........................................................................................................…………………….31
43
30.
Motile................................................................................……………………............................... 31 Non-motile.............................................................................................…………………….Shigella
31.
Indole (+), swarming growth on BHI agar..................................................………………Proteus Indole (-), no swarming growth.........................................................................………………...32
32.
Limus milk acid.................................................................................……………………Salmonella Litmus milk acid and peptonised........................................................………………...Aeromonas
33.
Citrate positive.....................................................................................…………………..Rhizobium Citrate negative..............................................................................………………….Agrobacterium
Once your outline has been approved, carry out your tests using materials available in your kits or on the side bench. You will have four lab periods. Record all of your results in your lab book. Include diagrams of all staining results, as well as descriptions of cell and colony morphology and tables of biochemical test results. Include the results from Part C below. Use reference material to identify your organism to species (note, identify all possible species). Note: you will be pooling your results with those from the other classes and examining class results as well. C
Investigation of Catabolic Ability of Soil and of Compost Bacteria
This exercise will be completed concurrently with exercise B, above. Methods Each bench will require: •
2 casein agar plates
•
2 NA plates containing carboxymethylcellulose
•
2 NA plates containing starch
•
2 NA plates containing xylan
•
Each individual will require:
•
1 casein agar plate
•
1 NA plate containing carboxymethylcellulose
•
1 NA plate containing starch
•
1 NA plate containing xylan
•
4 replica-plating templates per bench
•
sterile toothpicks – 1 beaker per bench
•
waste beakers for used toothpicks
•
Enumeration plates (of soil and of compost bacteria)
Each group of 4 is responsible for replica-plating 40 random colonies from plates of soil or compost bacteria onto plates containing one of the 4 substrates of interest
44
(carboxymethylcellulose, starch, casein or xylan). Note that plates containing CMC, starch or xylan all contain these substrates added to a nutrient agar base whereas plates containing casein are composed of skim milk and agar only. 1)
Using a sterile toothpick for each new colony, carefully scrape a well-defined colony from one of your soil or compost bacteria enumeration plates.
2)
Stroke the toothpick across square number 1 on each of the three different labelled plates (CMC, starch or xylan). For plates containing casein as the substrate, you may need to estimate placement of the colonies as you may not be able to see the template through the plate. Place the used toothpick into the beaker provided.
3)
Select a fresh toothpick and repeat steps 1-2 for 40 different colonies of bacteria.
Work individually to determine the catabolic ability of your unknown:
4)
Obtain 4 plates, each containing a different substrate. Label with your name and with the name of the substrate.
5)
Use an inoculating loop to streak out your unknown (from your PYE plate of pure culture) onto each of the 4 plates (you want to streak for single colonies).
Invert all of the plates. These plates will be incubated at 30oC for 48 hours. The Next Laboratory Period - Evaluation of Catabolic Ability:
6)
For those plates containing xylan or carboxymethylcellulose as substrates, flood the plate with a 0.1% (w/v) aqueous solution of Congo Red. Incubate for at least 5-10 minutes, then pour off the excess solution into a waste beaker (not down the drain!), and flood the plate with 1M NaCl to destain. Swirl the plate and let stand. Over the next 30 minutes, perform this destaining step 2 more times. Be generous with the NaCl. Cellulase- or xylanase-producing colonies will be surrounded by yellow haloes visible against the red or orange background. If there are no obvious haloes, then score the results as negative.
7)
For those plates containing starch as a substrate, flood the plate with a 0.13% iodine/0.3% potassium iodine solution. Swirl to cover the surface of the plate, then discard immediately. Destain by flooding the plate with 1 M NaCl and allowing the plates to stand. Amylase positive colonies should be surrounded by a zone of clearing.
8)
For those plates containing casein as a substrate, examine the plate closely. Caseolytic positive isolates should be surrounded by zones of clearing.
Thought Questions:
45
•
For enumeration, why do you only count plates having between 30 and 300 colonies?
•
Why do you incubate soil and compost bacteria at 28-30oC?
•
Provide one specific example of a differential medium used in the current exercise.
•
What is the differential component in the medium in your answer in (a)?
•
How is the medium differential?
•
Is this medium type selective also? Why or why not? How would you make this medium selective for the carbon source in question?
•
In addition to manipulating nutrients, how else could you make culture conditions selective? Provide a specific example.
References: Atlas, R.M. and Richard Bartha. 1998 Microbial Ecology: Fundamentals and Applications, Fourth Edition. Benjamin/Cummings Publishing Company, Inc. 640 pp. Poulsen, O. M. and Petersen, L. W. 1989. Electrophoretic and enzymatic studies on the crude extracellular enzyme system of the cellulolytic bacterium Cellulomonas sp. ATCC21399. Biotechnol. Bioeng. 34: 59-64. Ross, H. 1993. Cellular, Molecular and Microbial Biology 343 Laboratory Manual. The University of Calgary press, Calgary AB. Teather, R. M. and Wood, P. J. 1982. Use of Congo red polysaccharide interactions in the enumeration and characterization of cellulolytic bacteria from the bovine rumen. Appl. Environ. Microbiol. 43: 777-780.
46
EXERCISE 9 APPLICATIONS OF MICROBIOLOGY A number of industrial processes make use of the end products of bacterial and fungal fermentations. For instance, in the presence of acid producing bacteria, and often the enzyme renninase, milk will form curds (solid) and whey (liquid). Once the solids are compressed, salted, and aged, the resulting product is cheese. Different cheeses are produced by varying the bacterial inoculum, varying the milk used, or even by introducing fungi such as certain species of Penicillium into the curds. Some Streptococcus species and some Lactobacillus species produce only lactic acid as a result of reduction of pyruvic acid. These organisms are responsible for the production of yogurt. Yogurt can be made from milk simply by inoculating with a starter culture of yogurt that contains live bacterial culture. Conversely, yeasts produce alcohol and CO2 rather than lactic acid as a result of the reduction of pyruvic acid. EXPERIMENTAL OBJECTIVE This experiment will illustrate fermentation pathways and organisms involved in the production of alcohol. A
Alcohol Production
Prior to your lab period, grape juice, water and yeast cells were added to a sterile container. Over the next two weeks, you will be responsible for sampling the fermenting juice at various time intervals. The primary fermentor is inoculated with a high cell density (~106 yeast cells/mL). The bulk of the must (grape juice medium) is rapidly depleted of oxygen by the yeast and remains anaerobic, despite the primary fermentor remaining open to the atmosphere. Yeast cells continue to reproduce by acquiring the needed energy and carbon through fermentation. The fermentation is an ethanolic fermentation because ethanol and CO2 are the fermentation endproducts. Growth of the yeast culture can be monitored by measuring optical density and enumerating CFU/mL (Recall Exercise 4 - Bacterial Reproduction). Ethanol concentration can be estimated indirectly by measuring the specific gravity of the wine must with a hydrometer. The specific gravity of the must decreases as the grape juice sugars are converted to ethanol and CO2. A "specific gravity to percent ethanol" conversion chart supplied with the hydrometer is then used to determine ethanol content of the must. Note: this exercise requires some out of lab participation. Failure to sign up will result in a deduction of 5% from your lab grade.
47
Please sign up for a time slot when at least half of your group members can attend. MATERIALS (in C741) •
Wine thief
•
Spectrophotometer (warm up 15 minutes prior to reading OD values)
•
Primary fermentor
•
pH paper
•
Micropipettors and sterile tips
•
Sterile microfuge tubes
•
Rack for microfuge tubes
•
Filter sterile wine must for diluting samples
•
Bunsen burner
•
Hydrometer
•
Sterilising solution
•
Graduated cylinder
•
Thermometer
•
28 oC incubator
•
YPD plates
•
Spreaders and alcohol
•
Spreadsheet for recording results
At each time point the following data must be collected by each group: •
Temperature
•
OD600
•
pH
•
Specific gravity
•
Viable counts
Procedure – Work very carefully. These results will form the basis of your major lab report. 1)
Mix the culture well, then remove a sample from the primary fermentor.
2)
From the sample (not the primary fermentor), measure and record the sample temperature.
3)
Measure and record the specific gravity.
4)
Measure and record the pH of the sample.
5)
Measure and record OD600. Use the blank provided to zero the machine. Read the optical density of at least 5 mL of the sample. If the OD600 exceeds 0.7, you will have to dilute the sample with the sterile must provided (start by creating a 1:1 dilution). Read the OD600 of the diluted sample, then multiply by the dilution factor to obtain your corrected reading. Record the corrected reading on the sheet provided.
48
Please save the blank for the next group. 5)
For viable counts, Table 8.1 provides you with dilutions to create depending upon your sample time, as well as guidelines for what dilutions to plate out. Ensure that you plate out duplicates of each dilution.
6)
For your dilutions, prepare the required number of 900 µL dilution blanks = 900µL of filter sterile wine must in 1.5 mL microfuge tubes.
7)
Clearly label plates with time, name and dilutions. Spread plate (in duplicate) 100 µL of suggested dilutions on YPD agar.
8)
Invert plates and incubate at 28ºC for 48 hours.
9)
Tidy up work area.
Thought Questions: •
Numerous data relating to alcohol fermentation were collected by the class over the sampling period, including measurements of pH, temperature, specific gravity, optical density, and CFU’s/mL of culture. Design and construct a series of figures to graphically represent the data that were collected.
•
Calculate the generation time and the specific growth rate of the yeast cells in the culture.
•
Name two factors that control the final ethanol concentration in a culture.
•
Although we stirred our culture each time before sampling, winemakers do not. Why would winemakers not stir the culture?
•
Why did the pH of the culture change as fermentation proceeded?
•
Why did the specific gravity of the culture change over time?
49
Table 8.1 Dilutions of yeast mixture to create and to plate out for all time points. Time point
Dilutions to Create
Dilutions to Plate
(hours) Spread plate 100 µL of filter 2
None
sterile wine must in duplicate on YPD Spread plate 100 µL of each of
3
10-3; 10-4; 10-5
the 3 dilutions in duplicate on YPD Spread plate 100 µL of each of
7
-3
-4
10 ; 10 ; 10
-5
the 3 dilutions in duplicate on YPD Spread plate 100 µL of each of
12
-3
-4
10 ; 10 ; 10
-5
the 3 dilutions in duplicate on YPD Spread plate 100 µL of each of
24
-4
-5
10 ; 10 ; 10
-6
the 3 dilutions in duplicate on YPD Spread plate 100 µL of each of
28
-4
-5
10 ; 10 ; 10
-6
the 3 dilutions in duplicate on YPD Spread plate 100 µL of each of
32
-4
-5
10 ; 10 ; 10
-6
the 3 dilutions in duplicate on YPD Spread plate 100 µL of each of
48
-4
-5
10 ; 10 ; 10
-6
the 3 dilutions in duplicate on YPD Spread plate 100 µL of each of
53
-4
-5
10 ; 10 ; 10
-6
the 3 dilutions in duplicate on YPD Spread plate 100 µL of each of
79
10-4; 10-5; 10-6
the 3 dilutions in duplicate on YPD Spread plate 100 µL of each of
217
-4
-5
10 ; 10 ; 10
-6
the 3 dilutions in duplicate on YPD
50
APPENDIX 1 THE COMPOUND LIGHT MICROSCOPE As you label Figure 1, your Instructor will review the use of this microscope with you. Locate the ocular lens (eyepiece); there will be one if the microscope is monocular, or two if it is binocular. Then locate the objective lenses, the ones nearest the object to be studied. These two lenses (ocular and objective) are connected by the body tube of the microscope. The objective lenses (there will be two or more, the smallest being that with the least magnifying power, and the largest being that with the greatest magnifying power) are mounted on a revolving nosepiece above a flat stage on which the study specimen (slide) is placed.
Figure 1: The Compound Microscope Your microscope is equipped with a mechanical stage. This consists of a clip to hold the slide in place (the clip is spring-loaded; the Instructor will demonstrate how it works) and two knobs at the side of the microscope body to move the slide side-to-side, or forward-to-back. Note also the two micrometer scales on the mechanical stage, which allow you to note the coordinates of a particular object on the slide you are viewing.
51
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
52
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.
53
APPENDIX 2 PREPARATION OF SCIENTIFIC DRAWINGS 1)
Use a sharpened pencil; never ink. The lead should be hard.
2)
Place drawing to one side, usually the left, leaving room for labels to the right.
3)
Try to draw with one continuous line and do not retrace your lines. Do not shade.
4)
Place label lines horizontally (use a ruler), with no crossed lines.
5)
Objects labelled should be singular unless label line branches to multiple objects.
6)
Label only what you see, not what you think should be seen.
7)
Below the figure you should add:
8)
a)
The title of the diagram
b)
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:
size of drawing actual size of specimen
Where: •
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)
54
9)
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.).
55
APPENDIX 3 ASEPTIC TECHNIQUE A.
Aseptic Technique
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.
56
3.
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).
B.
Aseptic Culture Manipulation
Purposes: 1)
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 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.
57
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
58
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.
59
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 µ L 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.
60
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.
61
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 photoautotroph
energy source
carbon source
light
CO2
(photolithotroph)
reducing power inorganic oxidizable substrate
photoheterotroph
light
organic
organic
chemoautotroph
chemical (oxidation of
CO2
inorganic
(chemolithotroph)*
reduced inorganic
organic
organic
(photoorganotroph)
compounds e.g. NH3, NO2- and H2) chemoheterotroph
chemical
(chemoorganotroph)
62
*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.
63
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
Amount (/L)
Source of?
Tryptone
5.0 g
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
0.5 g
Macronutrients
MgSO4
0.1 g
Macronutrients
Agar
20 g
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.
64
VMM (Vincent’s Minimal Medium - Vincent, 1970) (used for the study of nutritional requirements of Rhizobium leguminosarum) Solution A: Compound
Amount (/L)
Source of?
K2HPO4
1.0 g
KH2PO4
1.0 g
KNO3
0.6 g
For Solid Medium:
12.5 g
Buffering agent/ Macronutrients Buffering agent/Macronutrients Macronutrients (nitrogen in particular) Gelling agent
Agar Solution B (10x): Compound
Amount (/L)
Source of?
FeCl3
0.1 g
Macro/Micronutrients
MgSO4
2.5 g
Macronutrients
CaCl2
1.0 g
Macronutrients
Autoclave and add to a final concentration of 1x Solution C (100x) Compound
Amount for: 1 L
Source of?
Biotin
0.01 g
Growth factors
Thiamine
0.01 g
Growth factors
Calcium Pantothenate
0.01 g
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
65
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.
66
APPENDIX 5 BACTERIAL OBSERVATION Bacterial genera may be differentiated in two ways: 1)
by the cellular morphology which is observed microscopically
2)
by the colony morphology which is observed on a plate culture
Cellular Morphology includes: 1)
Shape: rods, cocci, spirilli
2)
Size (in µm): diameter (cocci); lengthxwidth (rods)
3)
Typical arrangement of the cells: chains, clusters, pairs, random
4)
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)
Circular
Shape or form
Irregular
Rhizoid Filamentous
Punctiform (1mm or less in diameter)
2)
Surface: smooth/rough; mucoid/moist/dry/powdery
3)
Elevation:
Flat
Raised
Convex
Umbonate
Umbilicate
4)
Size: measure a single colony with a ruler
5)
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.
6)
Opacity: Transparent (can see through) or opaque.
67
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
68
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 peer-reviewed 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, 9th edition. Williams and Wilkins, Baltimore.
69
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.
70
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).
3)
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).
71
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
17.0 g
Bacto soytone
3.0 g
Dextrose
2.5 g
NaCl
5.0 g
Dipotassium phosphate
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
15.0 g
Bacto soytone
5.0 g
NaCl
5.0 g
Agar
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.
72
LB Medium (Luria-Bertani Medium): Used for cultivation of Enterobactereaceae family members, Sinorhizobium and Agrobacterium Composition (g/L): Tryptone
10.0 g
Yeast extract
5.0g
NaCl
10.0 g
Dissolve in distilled deionised H2O 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
12.0 g
Yeast Extract
24.0 g
Glycerol
4.0 mL
Dissolve in distilled deionised H2O 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 H2O. After the salts have dissolved, adjust the volume of the solution to 100 mL with deionised H2O and sterilise by autoclaving for 20 minutes at 15 psi on liquid cycle).
73
Nutrient Agar: Used for the cultivation of a wide variety of microorganisms. Composition (g/L) Peptone
5.0 g
NaCl
5.0 g
Yeast extract
2.0 g
Beef extract
1.0 g
Agar
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
5.0 g
Yeast Extract
3.0 g
CaCl2
0.5 g
MgSO4
0.1 g
Agar
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
10.0 g
Yeast Extract
5.0 g
NaCl
5.0 g
Glucose
1.0 g
Methylene Blue
0.02 g
Agar
15.0 g
74
Dissolve in distilled deionised H2O 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
10.0 g
NaCl
5.0 g
Glucose
1.0 g
CaCl2
0.11 g
Agar
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. Peptone Yeast Extract Medium (PYE) Used for the propagation of soil and compost microorganisms. Composition (g/L) Peptone
10.0 g
NaCl
5.0 g
Yeast Extract
5.0 g
For solid medium: Agar
15.0 g
Dissolve in distilled deionised H2O 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.
75
Yeast Peptone Dextrose Medium (YPD; YEPD) YPD is a complex medium used for yeast. Composition (g/L): Yeast extract
10.0 g
Peptone
20.0 g
Glucose
20.0 g
For solid medium: Agar
15.0 g
Dissolve in distilled deionised H2O 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. Hugh and Leifson’s Medium Used for detection of glucose or lactose degradation by microorganisms. Composition (g/L): Peptone from meat
2.0 g
NaCl
5.0 g
K2HPO4
0.3 g
Carbohydrate
10.0 g
Agar
5.0 g
Bromothymol blue
3.0 mL of a 1% solution
Bromothymol blue is dissolved in water. Alcoholic solutions of indicators should not be used as acid may be produced from the alcohol. For critical studies, the carbohydrate should be sterilised completely and added to the otherwise complete sterile medium. Autoclave 20 minutes on liquid cycle (121 oC; 15 psi). Dispense into sterile test tubes to a final volume of 5 mL per tube. Note, could be dissolved by heating first, dispensed, then the tubes autoclave to sterilise.
76
Eosin Methylene Blue Agar: Used for selection of Gram negative bacteria, and differentiation of lactose fermenting organisms. Composition (g/L): Peptones
10.0 g
Di-potassium hydrogen phosphate
2.0 g
Lactose
5.0 g
Sucrose
5.0 g
Eosin Y, yellowish
0.4 g
Methylene blue
0.07 g
Agar
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
20.0 g
NaCl
5.0 g
Lactose
10.0 g
Bile salts
5.0 g
Neutral red
0.075 g
Agar
12.0 g
Dissolve in distilled water to a final volume of 1 L, autoclave 15 min at 121oC, and pour plates.
77
Casein Agar: Used for the determination of caseolytic activity of microorganisms. Composition (g/L): Agar
10.0 g
Skim milk
100.0 g
Add distilled water to a final volume of 1 L, autoclave for 15 min at 121 oC and pour into sterile Petri dishes. Indole Broth: Used for the differentiation of organisms that can metabolise tryptophan to produce indole. Composition (g/L): K2HPO4
15.65g
L-Tryptophan
5.0 g
NaCl
5.0 g
KH2PO4
1.35g
Dissolve in distilled water to a final volume of 1 L, dispense into test tubes, and autoclave 15 min at 121oC. Methyl Red Voges Proskauer (MRVP) Broth: Used for the differentiation of bacteria based on acid production (methyl red test) or acetoin production (Voges Proskauer reaction). Composition (g/L): Glucose:
5.0 g
KH2PO4
5.0 g
Pancreatic digest of casein
3.5 g
Peptic digest of animal tissue
3.5 g
Dissolve in distilled water to a final volume of 1 L, dispense into test tubes, and autoclave 15 min at 121oC.
78
Simmons Citrate Agar: Used for the detection of citrate degradation by microbes. Organisms able to utilise citrate exhibit growth and the medium changes from green to blue. Citrate negative organisms do not grow and the medium remains green. Composition (g/L): Ammonium dihydrogen phosphate
1.0 g
Potassium dihydrogen phosphate
1.0 g
NaCl
5.0 g
Sodium citrate
2.0 g
Magnesium sulphate
0.2 g
Bromthymol blue
0.08 g
Agar
15.0 g
Dissolve in distilled water to a final volume of 1 L, dispense into test tubes, autoclave 15 min at 121oC, and cool in the slant position. Urea Broth: Urea broth supports the growth of microorganisms that can utilise urea as their sole carbon source. Organisms that are able to metabolise urea change the incorporated indicator to red. Composition (g/L): Yeast extract
0.1 g
KH2PO4
9.1 g
Disodium hydrogen phosphate
9.5 g
Urea
20.0 g
Phenol red
0.01g
Dissolve in distilled water to a final volume of 1 L, heating to 60oC if neccessary. Sterilise by filtration and dispense into sterile test-tubes. Do not autoclave.
79
Sucrose Agar: Used for differentiation of bacteria based on their ability to produce dextran – a polymer of sucrose. Composition (g/L): Beef Heart (solids from infusion)
500.0 g
Sucrose
50.0 g
Agar
20.0 g
Tryptose
10.0 g
NaCl
5.0 g
Dissolve in distilled water to a final volume of 1 L, autoclave for 15 min at 121oC, and pour into sterile Petri dishes. Litmus Milk: Used for the maintenance of lactic acid bacteria and the differentiation of bacteria based on their action in milk. Composition (g/L): Skim milk
100.0 g
Azolitmin
0.5 g
Na2SO3
0.5 g
Dissolve in distilled water to a final volume of 1 L, heat to boiling and dispense into test tubes. Autoclave for 20 min at 115oC.
80
Motility Medium S: Used for the determination of bacterial motility. Composition (g/L): Beef heart solids from infusion
500.0 g
Gelatin
30.0 g
Enzymatic hydrolyzate of protein
10.0g
NaCl
5.0 g
K2HPO4
2.0 g
KNO3
2.0 g
Agar
1.0 g
2,3,5-triphenyltetrazolium chloride solution*
10 mL
Add all components except triphenyltetrazolium chloride solution to distilled water; bring volume to 990 mL. Mix thoroughly, and heat to boiling. Autoclave for 15 min. at 121oC, let cool to 60oC and add 10 mL of triphenyltetrazolium chloride. Dispense into sterile test tubes. * To prepare 2,3,5-triphenyltetrazolium chloride solution, dissolve 0.1 g in 10 ml of distilled water. Mix thoroughly and filter sterilise.
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.
81
REAGENTS: Ethanol, 70%: 95% Ethanol
36.8 mL
Distilled Water
13.2 mL
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
75 mL
Hydrochloric acid
25 mL
p-dimethylamine-benzaldehyde
5.0 g
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.
82
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.
83
pH INDICATORS: Table 1: Indicators of Hydrogen Ion Concentration. pH Indicator Cresol Red
pH Range 0.2 - 0.8
Full Acid Colour Red
Full Alkaline C l Yellow
Meta Cresol Purple (acid range) Thymol Blue
1.2 - 2.8
Red
Yellow
1.2 - 2.8
Red
Yellow
Brom Phenol Blue
3.0 - 4.6
Yellow
Blue
Brom Cresol Green
3.8 - 5.4
Yellow
Blue
Chlor Cresol Green
4.0 - 5.6
Yellow
Blue
Methyl Red
4.4 - 6.4
Red
Yellow
Chlor Phenol Red
4.8 - 6.4
Yellow
Red
Brom Cresol Purple
5.2 - 6.8
Yellow
Purple
Bromothymol Blue
6.0 - 7.6
Yellow
Blue
Neutral Red
6.8 - 8.0
Red
Amber
Phenol Red
6.8 - 8.4
Yellow
Red
Cresol Red
7.2 - 8.8
Yellow
Red
Meta Cresol Purple (alkaline range) Thymol Blue (alkaline range) Cresolphthalein
7.4 - 9.0
Yellow
Purple
8.0 - 9.6
Yellow
Blue
8.2 - 9.8
Colourless
Red
Phenolphthalein
8.3 - 10.0
Colourless
Red
Adapted from: Benson, H.J. (1985). Microbiological Applications: A Laboratory Manual in General Microbiology, 4th Ed. Wm. C. Brown Publishers, Dubuque, Iowa.
84
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
85
