Download Biol 3400 Lab Manual Spring 2016 Final

The
University of
Lethbridge
BIOLOGY 3400
Principles of Microbiology
LABORATORY MANUAL
Spring, 2016
Written by: L. A. Pacarynuk and H.C. Danyk
Revised: December, 2015
TABLE OF CONTENTS
Exercise:
Page
Biology 3400 Laboratory Schedule.................................................................................2
Biology 3400 Laboratory Expectations...........................................................................3
Grade Distribution..........................................................................................................4
Lab Book Preparation ………………………………………………………………………………………………..5
Guidelines for Safety Procedures...................................................................................7
Exercise 1 – Microscopy and Gram Staining: A Review ...............................................10
Exercise 2 – General Laboratory Principles and Biosafety...........................................13
Exercise 3 – Free-Living Nitrogen Fixation....................................................................15
Exercise 4 – Soil and Compost Microbial Ecology ........................................................23
Exercise 5 – Bacterial Reproduction.............................................................................33
Exercise 6 – Fermentation............................................................................................36
Exercise 7 – Virology.....................................................................................................41
Appendix 1 – The Compound Light Microscope...........................................................47
Appendix 2 – Preparation of Scientific Drawings.........................................................54
Appendix 3 – Staining……………………………..……………………………………………………………….56
Appendix 4 – Aseptic Technique..................................................................................60
Appendix 5 – The Cultivation of Bacteria.....................................................................65
Appendix 6 – Bacterial Observation.............................................................................71
Appendix 7 – Laboratory Reports.................................................................................72
Appendix 8 – Use of the Spectrophotometer...............................................................74
Appendix 9 – Media, Reagents, pH Indicators.............................................................76
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BIOLOGY 3400 LAB SCHEDULE
SPRING, 2016
Jan.7
No lab
Jan. 12
Jan. 14
Introduction, Microscopy and Gram Staining: A Review
General Lab Procedures, Biosafety, N-Fixation (inoculate broths)
Jan. 19
General Lab Procedures, Biosafety – Complete; Soil/Compost Bacteria
Enumeration – Begin
Soil Bacteria Enumeration – Complete; Prepare Streak Plates; N-Fixation
Jan. 21
Jan. 26
Jan. 28
Morphological Characterization of Unknown (Gram Stain); Subculture
Unknown; N-Fixation
Endospore/Capsule Stains; Unknown Identification – Develop Outline for
Identification
Feb. 2
Feb. 4
Unknown Identification; Catabolic Activity
Unknown Identification; Catabolic Activity
Feb. 9
Feb. 11
Unknown Identification: N-Fixation PCR
Unknown Identification; N-Fixation Electrophoresis
Feb. 16
Feb. 18
Reading Week
Reading Week
Feb. 23
Feb. 25
Bacterial Growth
N-Fixation: Sequence Analysis
Mar. 1
Mar. 3
Fermentation Part A: Yogurt
Fermentation Part A: Yogurt - Complete
Mar. 8
Mar. 10
Fermentation Part B: Yeast Fermentation
Yeast Fermentation – sample and enumerate
Mar. 15
Mar. 17
Virology (phage isolation); Yeast Fermentation - enumerate
Virology (phage elution); Fermentation Part B: Yeast Fermentation Complete
Mar. 22
Mar. 24
Virology (amplification)
Virology (titre/host range)
Mar. 29
Mar 31
Virology – Complete
No lab
Apr. 5
Apr. 7
Apr. 12
No lab
No lab
Lab Exam
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Biology 3400 Laboratory Expectations:
Biology 2000 is the prerequisite for Biology 3400. In turn, Biology 1010 and Biology
1020 serve as prerequisites for Biology 2000. Biology 3400 laboratories will build upon
concepts introduced in these junior level courses. We will not spend laboratory time
reviewing material covered in lectures and laboratories associated with these courses.
If material/lab techniques are unfamiliar to you, it will be up to you to become
proficient. The following list provides some guidance with respect to concepts and skills
in which you should be proficient to be successful in microbiology labs this semester:
Concepts:
 Prokaryotic cell structure (versus structure of eukaryotic cells)
 Prokaryotic cell division
 Bacterial DNA replication
 Polymerase chain reaction and gel electrophoresis
 Respiration and fermentation
 Catabolism
 Viruses; especially bacteriophage (lytic cycle)
Skills:
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Aseptic technique
Gram staining
Culture inoculation (broths and plates)
Appropriate handling of biologicals and chemicals
Use of a dichotomous key
Microscopy – dissecting and compound light microscopes, oil immersion
Serial dilutions
Keeping a lab book
Scientific drawings
Scientific method principles
Critical thinking
Data analysis and drawing appropriate conclusions
Writing lab reports
Searches of peer-reviewed literature
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Laboratory Grade Distribution:
The laboratory component of Biology 3400 is worth 50% of your course mark. It is
distributed as follows:
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Skills Tests
Assignments and/or Lab Reports
Lab Books
Lab Exam
8%
24%
8% (to be handed in multiple times)
10%
Performance:
Up to 10% of laboratory grade (5 marks out of 50) will be subtracted for poor laboratory
performance. This includes (but is not limited to) failure to be prepared for the
laboratory, missing lab notebook or lab manual, poor time management skills, improper
handling and care of equipment such as microscopes and micropipettors, and unsafe
practices such as not tying hair back, chewing gum, applying lipstick, eating, drinking, or
chewing on pencils, and sloppy technique leading to poor results. As we are working
with potential pathogens, students displaying improper or careless techniques will be
asked to leave the lab and will have at least 5% of their laboratory grade deducted
immediately.
Missing a lab for which there is a skills test or assignment requires documentation.
Upon presentation of this documentation, you will either have to complete the
assignment or skills test as soon as possible or, if this is not possible, your lab grade will
be recalculated.
The lab books will be collected and graded several times during the semester. Although
most exercises are completed as groups, the lab books are to be completed
individually, and must represent individual effort. The following page provides you
with tips on how to construct your books.
Unannounced skills tests will be given during the semester. Students are expected to
work independently on some technical aspect of microbiology and will be graded based
on their techniques and their results.
As proficiency in microbiological techniques is considered an essential component of the
course, students are only permitted three lab period absences (you do not require any
documentation). Missing more than three labs will result in a grade of 0 being assigned
for the lab (at this point, it is recommended that students consult with Arts and Science
Advising for the option of completing the laboratory the following year). Students are
still responsible for the material missed (and their assignments, lab reports etc. will be
graded as such). There are no make-up laboratories.
Late Reports will be penalized as follows: less than 24 hours late, –10%; between 24-48
hours late, -25%, between 48-72 hours late, -50%, after 72 hours, no marks will be
given.
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Extensions for the lab reports and assignments will only be granted for situations
involving prolonged illness (documentation is required).
Preparation of a Lab Book:
Your lab book provides you with a detailed record of your experiments performed. This
record proves invaluable when preparing manuscripts for publication, or, more
immediately, when preparing lab reports. This lab book, as with all of the reports and
assignments is an individual effort.
Choice of Lab Book
Standard black lab books can be purchased from the bookstore but these are not
required for this course. The only required features are:
 Pages are non-removable (no spiral bindings)
 All pages must be numbered in the top outer corner
In General
 all entries must be made in blue or black ink (except drawings)
 date EVERY entry
 never remove a page or use white-out
 if an entry needs to be deleted, strike out the entry with a single straight
line (the deleted entry must be readable)
 keep up to date, a lab book is meant to be filled out as the experiments are
carried out and NOT after the fact
 record anything that may be useful to you when preparing your lab reports
 leave plenty of space throughout the lab book to add comments after the fact.
Many exercises run over multiple periods – it is easier to organize your entries if
you leave several blank pages for each exercise.
Table of Contents
Designate the first 2 pages as the Table of Contents
 record information and page numbers as you go
Lab Entries
For each lab be sure to include the following;
1. Objective
2. Method Summary
 do not rewrite the protocol from the lab manual
 highlight any specific changes to the lab protocol
 include times and dates for when work was performed
 record product and bacterial names and manufacturers used
- enzymes, chemicals, equipment (micropipettors, baths)
 include incubation conditions for cultures and reaction
3. Observations & Results
 record any & all observations, this goes beyond number results
 include diagrams and any other form of raw data
 include calculations as appropriate
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4. Conclusions
 did you achieve your objective? Why or why not?
 use your results to support your conclusions
5. Answer the thought questions at the end of the lab (as applicable)
 use reference citations as needed; these may be graded.
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GUIDELINES FOR SAFETY PROCEDURES
Students enrolled in laboratories in the Biological Sciences should be aware that there
are risks of personal injury through accidents (fire, explosion, exposure to biohazardous
materials, corrosive chemicals, fumes, cuts, etc). The guidelines outlined below are
designed to:
a) minimize the risk of injury by emphasizing safety precautions and
b) clarify emergency procedures should an accident occur.
EMERGENCY NUMBERS
City Emergency
Campus Emergency
Campus Security
Student Health Centre
911
(403) 329-2345
(403) 329-2603
(403) 329-2484
THE LABORATORY INSTRUCTOR MUST BE NOTIFIED AS SOON AS POSSIBLE AFTER
THE INCIDENT OCCURS.
EMERGENCY EQUIPMENT:
Your lab instructor will indicate the location of the following items to you at the
beginning of the first lab period.
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Closest emergency exit
Closest emergency telephone and emergency phone numbers
Closest fire alarm
Fire extinguisher and explanation of use
Safety showers and explanation of operation
Eyewash facilities and explanation of operation
First aid kit
GENERAL SAFETY REGULATIONS:
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Eating and drinking is prohibited in the laboratory. Keep pencils, fingers and
other objects away from your mouth. These measures are to ensure your safety
and prevent accidental ingestion of chemicals or microorganisms.
Personal protective wear is mandatory. Lab coats, safety glasses and closedtoed shoes must be worn at all times during lab exercises which involve potential
for chemical or biological spills.
Coats, knapsacks, briefcases, etc. are to be hung on the hooks provided, stowed
in the cupboards beneath the countertops, or placed along a side designated by
your instructor. Take only the absolute essentials needed to complete the
exercise* with you to your laboratory bench. (* e.g. manual, pen or pencil)
Mouth pipetting is NOT permitted; pipet pumps are provided and must be used.
Always wash your hands prior to leaving the laboratory.
Students are not allowed access to the central Biology Stores area for any
reason. Consult your instructor if you require additional supplies.
Report any equipment problems to instructor immediately. Do NOT attempt to
fix any of the equipment that malfunctions during the course of the lab.
Use caution when handling chemical solutions. Consult the lab instructor for
instruction regarding the clean-up of corrosive or toxic chemicals.
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 Contain and wipe up any spills immediately and notify your lab instructor (see
SPILLS below). Heed any special instructions outlined in the lab manual, those
given by the instructor or those written on reagent bottles.
 Long hair must be restrained to prevent it from being caught in equipment,
Bunsen burners, chemicals, etc.
 Dispose of broken glass, microscope slides, coverslips and pipets in the specially
marked white and blue boxes. There will be NO disposal of glassware in the
wastepaper baskets.
 You are responsible for leaving your lab bench clean and tidy. Glassware must
be thoroughly rinsed and placed on paper toweling to dry.
SPILLS:
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Spill of SOLUTION/CHEMICAL: While wearing gloves, wipe up the spill using
paper towels and a sponge as indicated by the lab instructor.
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Spill of ACID/BASE/TOXIN: Contact instructor immediately. DO NOT TOUCH.
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BACTERIA SPILLS: If necessary, remove any contaminated clothing. Prevent
anyone from going near the spill. Cover the spill with 10% bleach and leave for
10 minutes before wiping up. Discard paper towels in biohazard bag. Discard
contaminated broken glass in designated biohazard sharps container.
DISPOSAL:
•
Broken glass, microscope slides, coverslips and Pasteur pipets are placed in the
upright white ‘broken glass’ cardboard boxes. NO PAPER, CHEMICAL,
BIOLOGICAL OR BACTERIAL WASTE MATERIALS should be placed in this container
•
Petri plates, microfuge tubes, pipet tips should be placed in the orange
biohazard bags. The material in this bag will be autoclaved prior to disposal.
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Bacterial cultures in tubes or flasks should be placed in marked trays for
autoclaving.
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Liquid chemicals should be disposed of as indicated by the instructor. DO NOT
dispose of residual solution in the regent bottles. In case of any uncertainty in
disposal please consult the lab instructor.
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Slides of bacteria should be placed in the trays filled with 10% bleach that are
located at the ends of the laboratory benches.
HEALTH CONCERNS:
Students who have allergies, are pregnant, or who may have other health concerns
should inform their lab instructor so that appropriate precautions may be taken where
necessary.
More information on Laboratory Safety is available on the Risk and Safety Services
webpage: http://www.uleth.ca/risk-and-safety-services/content/lab-safety
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STUDENT SAFETY ACKNOWLEDGEMENT
This form must be completed, signed and submitted to the laboratory instructor
before any laboratory work begins.
I have read carefully and understand all of the safety requirements and procedures for
working safely in the lab. I also agree to read all rules for specific exercises contained in
the laboratory manual or laboratory handouts required for this course. I recognize that
it is my responsibility to strictly follow them.
I understand that I am required to wear personal protective equipment, such as safety
glasses, lab coat, gloves etc., at all times when directed to do so in the laboratory. I
further understand that I am permitted to work in the laboratory only under the
supervision of a laboratory instructor.
If I am unsure of the potential hazards related to lab procedures, I will discuss this
with my instructor prior to undertaking the procedure in question.
Name: ___________________________________________________ (please print)
Student ID No.: _____________________________________________
Signature: ________________________________________________
Course: __________________________________________________
Date: ___________________________________________________
FOIP Notification: The personal information on this form is collected under the authority of the
Post-secondary Learning Act (Alberta) and the Freedom of Information and Protection of Privacy Act
(Alberta) for the purpose of administering the University’s laboratory safety program. For questions
on the collection, use, and disclosure of this information, please contact the University’s FOIP
Coordinator at 4401 University Drive West, Lethbridge, AB T1K 3M4; 403-332-4620; [email protected].
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EXERCISE 1
MICROSCOPY AND GRAM STAINING – A REVIEW
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 m to
1.0 m in width and from 1.0 m to 5.0 m in length, although there are 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.
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6. Calibrate the ocular divisions for the 40x and the 100x objective lenses. Note that
immersion oil is not necessary for calibration.
Figure 1. Calibration of an ocular micrometer using a stage micrometer. The mark on
the stage micrometer corresponding to 0.06 mm (60 m) is equal to 5 ocular divisions
(o.d.) on the ocular micrometer.  1 ocular division equals 60 m/5 ocular divisions or
12 m.
Once an ocular micrometer has been calibrated, objects may be measured in ocular
divisions and this number converted to m 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.
METHODS:
1. Use the diagram in Figure 1 to calibrate the 40x and the 100x objectives on your
compound microscopes. Consult Appendix 1 if you need to review how to operate
a compound light microscope. 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.
2. 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.
3. Prepare Gram stains of the cultures provided (consult Appendix 3 to review the
procedure, if needed).
4. Prepare scientific drawings of organisms stained (consult Appendix 2 to review
how to make scientific drawings).
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THOUGHT QUESTIONS:
1. Why is it necessary to stain microorganisms before viewing them with the
compound light microscope?
2. Explain how the Gram stain differentiates between Gram positive and Gram
negative cells (your answer should address bacterial structure AND purpose of
each of the components of the Gram stain).
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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 non-pathogenic or pathogenic, the microorganisms are treated with the same respect to
ensure 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/artsci/biological-sciences/bio-safety
METHODS
Part 1: General Laboratory Procedures
A. Work individually to prepare a streak plate and a broth culture using the E. coli cultures
provided. Refer to Appendix 3 as necessary.
B. Work individually to prepare serial dilutions of the E. coli culture provided.
1. Prepare two spread plates on LB using 100 l of your 10-6 dilution for each. Place your
plates on the appropriately labelled tray for incubation at 37 oC for 18 h. Record how
you set up your serial dilutions in your lab book.
2. In the next lab period, count the number of colonies on your spread plates, and
calculate the mean CFU/mL of original culture. Record this number on the board and
in your lab book.
3. Calculate the mean CFU/mL (+/- SD) for your class, and compare your value to these.
How successful were you at preparing serial dilutions?
Part 2: Biosafety
You will be provided with the following:
 Sterile swabs
 Sterile water
 Potato Dextrose Agar (PDA) plates and Luria Bertani (LB) plates
Work in pairs to complete the following exercise:
1. Draw a line on the back of each plate to divide the plates in half. Label one half of
the plate with the name of the surface to be tested. Label the other half of the plate
as “after disinfection”.
2. Moisten the swabs provided with a small amount of sterile water. Brush the surface
to be tested with the swab, and then use the swab to inoculate one-half of each of
your two plates.
3. Disinfect the surface, moisten another swab, and repeat using the other half of both
plates. Wrap the plates with parafilm.
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4. Your plates will be incubated for 16-20 hours at 30oC, and then refrigerated at 4oC.
During the next laboratory period, evaluate your plate results and record the number
of colonies present on each half of both plates. Make observations of colony
morphology.
Thought Questions: (Use the Biosafety Web Site as a reference)
1. Were differences in colony morphology and number observed on the two types of
media? Why?
2. Does disinfection of work surfaces completely eliminate all microbial organisms?
What evidence do you have?
3. What is an MSDS and where can you find one?
4. 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?
5. What criteria would characterize an organism classified in Risk Group 1, 2 3 or 4?
6. There are many “Golden Rules” for Biosafety. Identify 4 common sense practices that
will protect you in your microbiology labs.
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EXERCISE 3
FREE-LIVING NITROGEN FIXATION
PART A: ISOLATION OF FREE-LIVING NITROGEN FIXING MICROORGANISMS
Nitrogen is an important component of amino acids, cell walls and other cofactors
present in all cells. Nitrogen gas comprises greater than 75% of our atmosphere, but it is
one of the most stable bonds in nature and is unavailable for use in this form. At one
time early in the evolutionary history of life on earth, all cells may have had the ability to
fix N2 gas into a more usable form (nitrate, nitrite or ammonia). Today however, only a
few species of bacteria and archaea are capable of converting N2; all other organisms rely
on N2-fixing prokaryotes for their fixed nitrogen requirements.
M.W. Beijerinck, a Dutch microbiologist, successfully isolated free living nitrogen fixing
bacteria in 1901. He inoculated soil samples into enrichment media containing glucose
and mineral salts, but lacking any source of nitrogen other than atmospheric nitrogen. He
observed cells that are today identified as members of the genus Azotobacter.
Subsequently, other aerobic, free-living nitrogen fixing genera of bacteria have been
isolated and identified, including Azomonas, Azospirillum and Beijerinckia.
Nitrogen fixation occurs only when an enzyme called nitrogenase is present. The enzyme
consists of two distinct proteins (i) dinitrogenase, which reacts with N2, and (ii)
dinitrogenase reductase, which reduces nitrogen gas to ammonia. The dinitrogenase
reductase component is irreversibly inactivated by the presence of oxygen. Several
strategies have evolved to enable free-living, aerobic organisms like Azotobacter to fix
nitrogen. Azotobacter has a very high respiratory rate, which is thought to prevent any
stray oxygen from coming into contact with the nitrogenase enzyme. Additionally, freeliving nitrogen fixers often secrete copious amounts of slime which may prevent extra
oxygen from entering the cells. There is also evidence suggesting that in the presence of
oxygen nitrogenase can combine with a specific protein inside the cell that shields the
oxygen sensitive site and prevents it from interacting from oxygen. When oxygen levels
drop, nitrogenase can resume its activity.
Over the course of the semester we will isolate free living nitrogen fixing bacteria from
prairie soil, establish pure cultures and attempt to identify cultures using modern day
molecular techniques.
METHODS:
For each lab:
 5, 250 mL flasks containing N-free medium
 10 plates N-free medium
 Balance, weigh boats and spatula
 N-free soil sample
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1. Work in groups of 4 to inoculate your flasks.
2. Weigh out 1 g of the soil sample provided, and add it to an Erlenmeyer flask
containing 100 mL of N-free medium.
3. Swirl gently to mix. Label the flask with your lab, bench number, and date. Make
sure that the cap or foil is loosened sufficiently to allow air to enter the culture.
Incubate for one week at 30oC.
4. After 7 days, remove the flask and look for the presence of a thin film of growth on
the surface of the medium. Use a sterile inoculating loop to remove some of this
film and prepare a streak plate on N-free agar. The streak plate will be incubated
for a further 7 days at 30oC.
5. Examine wet mounts from your first streak plate culture using the phase contrast
microscope. Prepare Gram stains of the film and look for large Gram negative
cells that may be bacillus or coccoid in shape. They may occur singly, or in
arrowhead-shaped pairs. Record observations in your lab book.
6. After the incubation period is complete, examine your streak plate. Look for large,
translucent, mucoid colonies. Prepare a wet mount from an isolated colony and
view it using a phase-contrast microscope. Prepare another streak plate using
cells from the same colony. This plate will be incubated again, and observations
will be made later in the term. Additionally, this culture will be used in Part B of
this exercise.
Thought Questions:
1. Define enrichment. What aspect(s) of the medium used in this exercise made it an
enrichment medium? Why did we use the same medium for plating after freeliving nitrogen fixers were isolated? What term would we use to describe the
medium in this case?
2. Why did we sample the film on top of the culture, rather than the sediment on the
bottom of the flask?
3. When you viewed your Gram stains, you may have observed cells on your slides
that didn’t appear to be Azotobacter. Why might these other genera be present?
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PART B: IDENTIFICATION OF MICROORGANISMS USING PCR OF 16s rDNA
The DNA from microbes can be isolated and may be studied via construction of BAC
(Bacterial Artificial Chromosome) libraries (for an example, see Rondon, et al., 2000).
More simply, an appreciation of diversity may be obtained by using universal primers for
PCR amplification of rDNA genes from the Bacterial domain on a preparation of total DNA
from an environmental sample. The resulting pool of nucleotide fragments may then be
cloned, unique clones sequenced, and the resulting sequences analyzed in order to
characterize and potentially identify the microbes present.
In Part B of this exercise you will perform PCR using primers specific for prokaryotic 16s
rDNA to isolate ribosomal DNA from putative Azotobacter cultures and then visualise this
DNA using Agarose Gel Electrophoresis. In addition, DNA from successful PCRs will be
sent for sequencing and you will then be using online tools to perform sequence analysis
to confirm the identity of your cultures.
PCR of Soil Bacteria
Two different primer sets will be employed. Each group will only be using one set on their
particular culture. Note that the primer designations refer to location of primer binding
site on the 16s rDNA molecule.
For preparation of your reaction mixtures:
Benches 1, 3, and 5:
Working with the people at your bench, each group will be setting up 3x reactions as
outlined below:
Primer Set #1
Template
Source
FP1/1492R
Unknown
Culture
FP1/1492R
E. coli K12
FP1/1492R
No template
Benches 2/4:
Working with the people at your bench, each group will be setting up 3x reactions as
outlined below:
Primer Set #2
Template
Source
27F/805R
Unknown
Culture
27F/805R
E. coli K12
27F/805R
No template
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METHODS:
Reagents:
 Taq (Invitrogen)
 10x PCR buffer
 50 mM MgCl2
Primers (*Y = C or T)
 FP1 (AGAGTTYGATYCTGGCT)*1 (10 pmol/L)
 RP1492 (TACGGYTACCTTGTTACGACT)*1 (10 pmol/L)
 27F (AGAGTTTGATCCTGGCTCAG)2 (10 pmol/L)
 805R (GACTACCAGGGTATCTAATCC)2 (10 pmol/L)
 dNTP mix (8 mM)
 Optima Water (Fisher Scientific)
Cultures:
 Pure culture of organism isolated from soil
 Plate culture of E. coli
Equipment:
 Thermocyclers (BioRad)
 Micropipettors and sterile tips
 Parafilm
 Ice buckets and ice
 Sterile 0.5 mL tubes
 Sterile 0.2 mL PCR tubes
 Biohazard bags
 Permanent markers
Note: Use aseptic technique throughout. Keep your tubes on ice at all times!
1. Obtain three 0.2 mL PCR tubes from the sterile container at the side of the lab.
Decide on appropriate codes for labeling the tubes (keeping in mind that other
groups are carrying out the same reactions). Label the tubes on the tops and on
the sides using permanent marker. Place the tubes on ice.
2. Obtain a 0.5 mL tube for your Master Mix. Keep this tube on ice. Use the
information outlined in Table 1 to set up your Master Mix. This mix contains
everything required in order for DNA replication to occur. Generally, Master
Mixes contain enough volume to set up the number of reactions + 1. In your case,
you will be preparing enough mix for 4 reactions. Work carefully.
18
Table 1. Components, starting concentrations and volumes for set-up of PCRs.
Component and Starting Final
Master Mix vol.
Concentration
Concentration
(enough for 4
reactions) (L)
Optima-Water
10x PCR buffer
50 mM MgCl2
dNTP mix (8 mM of all 4)
Primer 1 (from set assigned to
your bench)
Primer 2 (from set assigned to
your bench)
Taq DNA polymerase (5 U/L)
Template DNA
1x
1.5 mM
40 nmoles (0.8
mM of all 4)
0.4 M
128
20
8
20
8
0.4 M
8
5 U (units)
4
Leave Template
out of Master
Mix!
Note: One primer set per reaction mixture!
3. While the Master Mix is being set up, other group members should be setting up
template preparations. Obtain a small square of parafilm. For each bacterial
culture (soil bacteria and E. coli – what is the role of E. coli?), use a micropipettor
with a sterile tip to pipette 20 L of sterile Optima-water (Fisher Scientific) onto
the parafilm.
4. Take a 10 – 100 L micropipettor and put on a sterile tip. Touch the tip to a single
colony from your soil bacterial culture plate. Pipette up and down into the
Optima water on the parafilm. 1 L of this mixture will be used as your template
source.
5. Mix E. coli in the same fashion with your second drop of Optima water on the
parafilm. Again, 1 L of this mixture will be used as template in your second
reaction.
6. For your third reaction, you will be leaving out template and replacing it with an
equal volume of sterile Optima water. What is the purpose of this reaction?
7. After preparation of Master Mix, add the appropriate volume of template or
sterile water (1 L) to each tube, then check with the Instructor to see where
everyone else is at. When all of the groups are at the same stage, add the
appropriate volume of Master Mix (49 L ) to each tube. Keep your tubes on ice
until in the PCR machine.
GENTLY tap tubes to mix. When everyone is ready, the instructor will then show you how
to operate the thermocycler.
19
The parameters you are using for the PCR are:
12 minutes at 95oC (used not only in initial DNA denaturation, but also to lyse the
bacterial cells)
30 cycles of:
 1 minute at 94oC
 45 seconds at 55oC
 90 seconds at 72oC
A final elongation of:
 20 minutes at 72oC
The samples will be stored at -20oC upon completion.
Thought Questions:
1. What are the purposes of the primers in PCR?
2. What happens at each temperature?
3. How is annealing temperature determined?
4. If you left out the forward primer, would you expect to see a band resulting on the
gel? If you did, explain what this would mean.
Suggested Background Reading:
Amann et. al., 1995. Phylogenetic identification and in situ detection of individual
microbial cells without cultivation. Microbiol. Rev. 59 (1): 143-169.
Cole, J. R., Chai, B., Farris, R. J., Wang, Q., Kulam, S. A., McGarrell, D. M., Bandela, A. M.,
Cardenas, E., Garrity, G. M., and Tiedje, J. M. 2007. The ribosomal database project
(RDPII): introducing myRDP space and quality controlled public data. Nuc. A. Res. 35:
D169-D172.
DeLong and Pace, 2001. Environmental diversity of bacteria and archaea. Syst. Biol. 50(4):
470-478.
Gabor, E. M., deVries, E. J., and Janssen, D. B. 2003. Efficient recovery of environmental
DNA for expression cloning by indirect extraction methods. FEMS. 44(2): 153-163.
Pace, 1997. A molecular view of microbial diversity and the biosphere. Science. 276: 734740.
Whitford, M. F., Forster, R. J., Beard, C. E., Gong, J., and Teather, R. M. 1998. Phylogenetic
analysis of rumen bacteria by comparative sequence analysis of cloned 16S rRNA genes.
Anaerobe. 4: 153-163.
Woese, C. R., Kandler, O., and Wheelis, M. L., 1990. Towards a natural system of
organisms: Proposal for the domains Archaea, Bacteria, and Eucarya. Proc. Natl. Acad. Sci.
USA. 87: 4576-4579.
20
Agarose Gel Electrophoresis
METHODS
Reagents:
 1x TBE buffer
 0.8% agarose gels (1 per 2 benches)
 10x loading dye
 2-log NEB ladder premixed with loading dye
 Ethidium bromide bath
 PCR samples from last lab
Equipment
 Power supplies (1 per 2 benches)
 Micropipettors
 Sterile tips
 Parafilm
 Transilluminator/camera
 Biohazard bags
 Gloves
Note: Two groups will load their samples (6 tubes total) onto one gel.
We will be using 0.8% agarose prepared in 1x TBE.
1.
2.
Obtain and completely thaw your PCR samples.
Using a micropipettor, 'dot' out 1 L aliquots of 10x loading dye in a line on a thin
strip of parafilm. Remove a 7.5 L aliquot of your first sample, mix gently with the
loading dye on the parafilm, and proceed with loading. Aim for approximately 1-2x
final concentration of loading dye per sample loaded (and recognise that this is NOT
exact).
Loading Dye – 1) increases the density of the sample ensuring that it drops evenly into
the well; 2) adds colour to the sample to simplify loading; and 3) contains dyes that in
an electric field move toward the anode at predictable rates. In this laboratory, we
are making use of mixtures containing xylene cyanol FF. This dye migrates in 0.5x TBE
at approximately the same rate as linear DNA of 4000 bp in size. Often, bromophenol
blue is used in conjunction with xylene cyanol, or separately. Bromophenol blue
migrates at approximately the same rate as linear DNA of 300 bp in size in 0.5x TBE
(2.2 fold faster than xylene cyanol FF, independent of agarose concentration).
3.
4.
Load the remainder of the samples in the same manner, leaving at least one well
empty (to be used for a DNA ladder). Be sure to RECORD the order in which the
samples were loaded.
Load 10 L of the ladder.
21
One type of size standard is produced by ligating a monomer DNA fragment of known
size into a ladder of polymeric forms. The 2-log DNA ladder from New England
Biolabs consists of a mixture of a number of proprietary plasmids digested to
completion with different restriction enzymes. Ladders tend to be purchased as
commercial preparations. For an example please see:
http://www.neb.com/nebecomm/products/productn3200.asp
5.
6.
Turn on the power supply and set the voltage to 100 V. Place the lid on the gel and
start the run. The gel will run for 30 minutes, then shut off automatically.
After the run is complete, turn off the power. Designate one group member to put
on gloves, scoop up the gel, and gently slide the gel into the ethidium bromide bath.
Caution: Ethidium bromide is a mutagen and a suspected carcinogen. At very
dilute concentrations and with responsible handling, this risk is minimised.
7.
Stain the gel with gentle shaking for approximately 10 minutes. One group member
again should put on gloves, and transfer the gel to the gel documentation system.
View using the UV transilluminator. Photographs will be taken. Please ensure that
you bring a USB memory stick so that you can obtain the photograph of your gel
(these will NOT be emailed out).
Caution: Ultraviolet light is damaging to naked eyes and exposed skin. Always
view through filter or safety glasses that absorb harmful wavelengths.
8.
Based on gel results and quantification of your DNA, a selection of samples will be
sent off for sequencing. In order to facilitate this, use a piece of tape to completely
label your PCR products ensuring that the label corresponds with that from the gel.
Thought Questions
1. Evaluate your gel results with respect to: expected fragment sizes and reasoning,
and control results. Do we have evidence to suggest that we were successful in
amplifying 16s rDNA? Explain your reasoning.
2. What are some of the advantages and disadvantages of molecular techniques for
identification of bacteria? Compare and contrast with conventional culturing
techniques.
3. After examining your sequence data, can you conclude that you isolated N-fixing
organisms? Explain your reasoning.
4. Can you identify your unknown organisms to genus? To species? Why or why
not?
5. What was the purpose of the control? How does it help you to make conclusions
about your unknown isolates?
22
EXERCISE 4
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 utilize
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 utilized. Bacteria are able to utilize the end products of fungal metabolism.
Nematodes and protozoa feed on the bacteria and mites and 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 characterized 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.
Out of your group of four, one pair will prepare enumeration plates for soil, while the
other pair will prepare enumeration plates for compost.
23
A. ENUMERATION OF BACTERIA IN SOIL AND COMPOST
Period 1:
1. Weigh 1 g of soil or 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, labelled Petri dishes and 1 mL
of the 10 -5 dilution to two labelled Petri dishes.
4. Obtain a 250 mL bottle of molten peptone yeast extract agar containing actidione
from the water bath. (Actidione or cyclohexamide is an antibiotic that inhibits
fungal growth).
5. Add approximately 20 mL of medium to the plates prepared in Step 3. Swirl
carefully to mix the inoculum evenly with the medium. Label the bottle of molten
agar with your group name and replace immediately in the waterbath.
6. 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 10-4, 10-5, and 10-6 dilutions of your compost.
7. 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.
8. Obtain your labelled 250 mL bottle of molten peptone yeast extract agar
containing actidione from the water bath. (Actidione or cyclohexamide is an
antibiotic that inhibits fungal growth).
9. 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 10:
10. The plates will be incubated for 24 hours at 30oC and refrigerated until the next
lab session
Period 2:
Results
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
number of bacteria per g of soil and of compost.
24
B. ISOLATION AND CHARACTERIZATION OF BACTERIA FROM SOIL OR COMPOST
Each class will identify organisms from either soil or compost bacteria plates. Your
instructor will indicate what plates to use.
Work individually to complete Part B of this exercise.
Period 2:
1. Choose a morphologically distinct colony from one of your plates (but approved by
your instructor), and prepare a streak plate for single colonies on peptone yeast
extract agar (Appendix 4).
2. The plates will be incubated for 24 hours at 30oC and then stored at 4oC.
Period 3:
1. During your next lab period, prepare a Gram stain of the pure culture and record the
cell morphology. Record the colony morphology of the culture (Appendix 6).
2. In addition, prepare new streak plates on PYE and NA and a broth culture using TSB
from the colony you used to prepare your Gram stain. These cultures will be
incubated at 30oC; PYE plates for 24 hours, and NA and TSB for 48 hours.
Period 4:
1. Use the NA plate to prepare an endospore stain (Appendix 3). Use the broth culture
and the phase contrast microscopes available at the back of the lab to look for the
presence of capsules.
2. Subculture an isolated colony from your PYE plate into TSB – this culture will be
incubated overnight at 30oC. You will use this culture as your inoculant for
biochemical tests in subsequent periods.
3. Use the dichotomous key provided to develop a detailed outline in your lab book of
the series of steps you plan to take to identify your unknown (this may require some
out of lab time to complete). The outline should include tests to carry out as well as
dates when you intend to do these tests. Your outline must be handed in and
approved before you will be allowed to proceed.
25
The following media and reagents will be available for you to utilize as you attempt to
identify your unknown:















nutrient agar
endospore stain reagents
hydrogen peroxide (catalase test)
oxidase reagent (oxidase test)
IMViC reagents
indole broths
MRVP broths
citrate slants
urea slants
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)
26
1.
Gram stain
Gram positive ....................................................................................................... 2
Gram negative......................................................................................................16
2.
Cell morphology
bacillus or spirillum……......................................................................................... 3
coccus….............................................................................................................. 12
ovoid…................................................................................................. Azotobacter
3.
Endospore stain
positive………………………..................................................................................................4
negative…………………………............................................................................................. 6
4.
1True
endospores……………………………............................................................................ 5
spore types……………………………....................................................................... 6
2 Special
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
1
2
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.
27
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
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 water-soluble 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
28
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
30. Motile……………………………............................................................................................ 31
Non-motile……………………………........................................................................... Shigella
31. Indole (+), swarming growth on BHI agar…………………………................................ Proteus
Indole (-), no swarming growth…………………………....................................................... 32
32. Litmus milk acid……………………………............................................................... Salmonella
Litmus milk acid and peptonised…………………………....................................... Aeromonas
33. Citrate positive………………………….................................................................... Rhizobium
Citrate negative……………………................................................................ Agrobacterium
29
Periods 5-8:
Once your outline has been approved, carry out your tests using materials available in
your kits or on the side bench. You will have 4 lab periods and possibly some open lab
time to complete this part of the exercise.
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).
C.
INVESTIGATION OF CATABOLIC ABILITY OF SOIL AND 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)
Period 5:
Each group of 4 is responsible for replica-plating 20 random colonies from plates of soil or
compost bacteria onto plates containing one of the 4 substrates of interest
(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 labeled
plates (CMC, starch or xylan). For plates containing casein as the substrate, you will
need to estimate placement of the colonies as you will 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.
30
Work individually to determine the catabolic ability of your unknown:
4.
5.
Obtain 4 plates, each containing a different substrate. Label with your name and
with the name of the substrate.
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 30 oC for 48 hours.
Period 6:
Evaluation of Catabolic Ability:
1.
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 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.
2. 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.
3. For those plates containing casein as a substrate, examine the plate closely.
Caseolytic positive isolates should be surrounded by zones of clearing.
Thought Questions:
1. For enumeration, why do you only count plates having between 30 and 300
colonies?
2. Why do you incubate soil and compost bacteria at 28-30oC?
3. Provide one specific example of a differential medium used in the current exercise.
4. What is the differential component in the medium in your answer in (a)?
5. How is the medium differential?
6. Is this medium type selective also? Why or why not? How would you make this
medium selective for the carbon source in question?
7. In addition to manipulating nutrients, how else could you make culture conditions
selective? Provide a specific example.
31
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.
32
EXERCISE 5
BACTERIAL REPRODUCTION
Most bacteria reproduce by an asexual process called binary fission. In this process a
single mother cell produces two identical daughter cells. Cell growth is often equated
with increase in cell number due to the difficulty in measuring changes in cell size. Under
ideal conditions populations of bacterial cells grow exponentially as cell number doubles
at a regular interval or generation time (td).
In the laboratory, pure cultures are routinely grown as batch cultures in test tubes and
Erlenmeyer flasks. A batch culture is prepared by inoculating a fixed amount of liquid
medium with the bacteria. The resulting culture is then 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., PetroffHauser chamber) to count the number of cells in a known volume of culture. Automated
systems such as Coulter counters may also be used to determine cell number.
In contrast, indirect count methods require the growth of cells in culture in order to
enumerate cell numbers. The most common method for enumerating living cells is the
viable plate count. Serial dilutions of a cell suspension are prepared and spread on to the
surface of a solid agar medium (spread plate) or incorporated into molten agar that is
then poured into sterile petri dishes (pour plate). Following a suitable incubation time,
the number of colonies growing on and in the inoculated agar are counted and used to
33
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.
Prelab preparation: Turn on the spectrophotometer and set to 600 nm at least 15
minutes prior to taking readings.
METHODS
 Ethanol/dettol in bottles
 Test tube racks
 Spectrophotometer blank containing LB broth
 Spectrophotometer
 Overnight culture of E. coli K12 strain

 6 Culture flasks of LB (100 mL volume); pre-warmed
 2 Culture flasks of LB with 2% NaCl; pre-warmed
 2 Culture flasks of LB at pH 5; pre-warmed
Please work in pairs. At 20 minute intervals, monitor the growth of your E. coli culture by
optical density following the procedures outlined below.
A. Culture inoculation:
1. Each group of two will be assigned one culture flask and one treatment (see Table
below). Please mark the flask with your treatment, bench number and lab number,
and replicate (A or B; to be assigned by your instructor). Groups in laboratory
section 3 will continue to sample from the flask corresponding to your bench. Data
from all three lab sections will be pooled and posted on the Biology 3400 web site.
Table 1: Growth parameters for bacterial reproduction exercise.
E. coli strain
K12
K12
DH5
K12
K12
Medium
LB
LB
LB
LB with 2.5% NaCl
LB at pH 5
Growth Temperature
37oC
30oC
37oC
37oC
37oC
34
2. Aseptically remove 1 mL of culture from the overnight E. coli culture you were
assigned, and add it to your culture flask. Swirl to mix, and record an initial optical
density value as described in Part B below.
B. Culture sampling:
3. Everyone in the laboratory will be sampling at the same time. Samples will be
collected four times during your regularly scheduled lab period at 20 minute intervals.
For labs 1 and 2, these correspond to: 9:35 am (“time zero”), 9:55am, 10:15 am,
10:35 am, and for labs 3 and 4: 11:00 am, 11:20 am, 11:40 am, and 12:00 pm. Your
laboratory instructor will set a timer so that everyone is coordinated. Just prior to
your readings, zero the spectrophotometer as outlined in Appendix 7.
4. At the times indicated, carefully remove your flask from the shaking incubator,
remove 5 mL of your culture, and place it into a spectrophotometer tube. Replace
your culture flask back into the incubator, and then read the absorbance of the
sample you collected. Record the optical density (Absorbance) reading in your lab
book and in the spreadsheet.
5. Your lab instructors will collect and distribute class data. Prepare graphs for each of
the parameters tested, and use them to address the thought questions below.
Thought Questions:
1. Use your graphs to calculate generation time of E. coli in each of the cultures.
2. What impact does increasing the concentration of NaCl have on growth of E. coli?
3. Do changes in the pH or temperature cause a change in the rate of growth of E.
coli? Provide reasons for any changes observed.
4. Are there differences in the rate of growth of the two strains of E. coli examined?
Why might this be?
5. Compare your values to that from the literature. Do the values differ? Why might
this be?
35
EXERCISE 6
FERMENTATION
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. We are going to
examine the latter two processes in this exercise.
Part A: Yogurt Production
MATERIALS
 Milk
 Skim milk powder
 Balance
 Erlenmeyer flasks – 500 mL
 Hot plates
 Thermometers
 Beakers – 250 mL
 Bacterial inoculant
 Probiotic capsules
 Water baths
 Micropipettors and tips
 pH meter
 phenolphthalein
 0.1N NaOH
Procedure – Period 1
1. Work as a bench group to heat approximately 250 mL of milk to 85 oC using the
Erlenmeyer flask and hot plate provided. Hold the temperature at 85 oC for 2
minutes, and then carefully remove the flask from the hot plate, and let cool
slowly on the bench to about 42-45oC.
2. Transfer 125 mL of heated milk into each of two beakers. Each pair at the bench
will then be responsible for preparing one of the treatments listed in Table 8.1
below. Inoculant “A” is live yogurt culture. Add one rounded teaspoonful to 125
mL milk samples. Inoculant “B” is a commercial yogurt bacterial started – add
36
0.63 g to milk samples. Inoculant “C” is a probiotic capsule, which contains some
of the same bacterial species as those found in the commercial starter. Empty the
contents of one capsule into the cooled milk sample for treatment 11.
3. For treatments where skim milk is added, mix 6 g into the cooled milk prior to
adding the bacterial inoculant. Add the inoculant, and gently stir to mix. Label
your beakers and place them in the water bath appropriate for your treatments.
Your instructor will remove beakers after approximately six hours of incubation,
and cultures will be stored at 4oC until the next lab period.
Table 6.1: Experimental set-up for yogurt cultures; A = yogurt, B = inoculant, C =
probiotic.
Treatment
1
2
3
4
5
6
7
8
9
10
11
12
Inoculant
A
A
A
B
B
B
A
B
A
B
C
none
Skim Milk Powder
no
no
no
no
no
no
yes
yes
no
no
no
no
Incubation
Temperature
32oC
42oC
52oC
32oC
42oC
52oC
42oC
42oC
42oC-with shaking
42oC-with shaking
42oC
42oC
Procedure – Period 2
1. Examine all yogurt cultures (not just the one you set up), and make observations
regarding texture, consistency, smell and general appearance in your lab book.
2. Determine the pH of your cultures using the pH meter provided.
3. The amount of acid produced during the fermentation process can be calculated
by titrating your culture with 0.1 N NaOH in the presence of phenylphthalein, a pH
indicator. Measure 9 mL of culture and add to a small beaker, along with 4 drops
of indicator. Mix well. Slowly add NaOH until a sustainable colour change is
observed. Record the volume of NaOH required, and multiply by 0.1 to convert to
% lactic acid.
4. Record your data in the table on the board and in your lab book.
5. Prepare a Gram stain of your culture and record observations in your lab book.
How many different types of organisms do you see?
37
Part B: Alcohol Production from Yeast
Prior to your lab period, grape juice, water and yeast cells were added to a sterile
container. Over the next ten days, 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 5 - 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 will require some out of lab participation.
Please sign up for two time slots 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
 30oC incubator
 YPD plates
 Spreaders and alcohol
 Spreadsheet for recording results
38
At each time point the following data must be collected by each group:
 Temperature
 OD600
 pH
 Specific gravity
 Viable counts
Procedure
1. Sterilize (by immersion in metabisulfite), the spoon, the wine thief, the sampling
graduated cylinder, the hydrometer and the thermometer, and rinse each in water
prior to touching the wine. Use the sterilized spoon to stir the mixture in the primary
fermentor.
2. Use the sterilized wine thief to remove mixture from the primary fermentor into the
plastic graduated cylinder. Add enough sample to bring the level up to the line
marked.
3. Measure and record the sample temperature in the cylinder.
4. Measure and record specific gravity. Use the diagram posted to assist you.
5. Aliquot some mixture into the 125 mL Erlenmeyer flask provided – label with
sampling time, and place in the tray in the deli cooler. An instructor will read and
record the pH for you.
6. Use a sterile 5 mL pipette to remove 5 mL of sample from the cylinder and place into
a sterile spec tube.
7. Remove the 50 mL Falcon tube of sterile wine must labeled with your time point from
the deli fridge, and use it to make appropriate dilutions.
8. Use the blank provided to zero the machine. Read the optical density of your 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 OD 600 of the diluted sample, then
multiply by the dilution factor to obtain your corrected reading. Record the
corrected reading on the sheet provided.
9. Viable counts:
 Prepare in duplicate. Refer to the posted table to determine the number of
dilutions required.
 Prepare required dilution series – check dilution calculations with your lab
instructor first.
 Label plates with time, name and dilutions. Spread plate (in duplicate) 100 µL of
suggested dilutions on YPD agar.
 Invert plates and incubate at 30ºC – incubator is labeled.
Plates will be removed and placed into the fridge. During lab, you will count and record
values from plates having between 30 and 300 colonies.
Tidy up work area!!– put spec tubes into labeled rack, discard tips, pipettes and
microfuge tubes into Biohazard bag, put everything back where you found it. Pour out
wine sample and resterilize cylinder for the next group.
39
Thought Questions (to be answered in your notebooks):
1.
Describe the metabolic reactions that result in the production of yogurt. What
microorganism(s) is/are required? How does each play a role?
2. Explain the effects of the following on the fermentation process that occurs during
yogurt production: addition of skim milk powder, incubation temperature, shaking
and type of inoculant used.
3. What is the purpose of the heat pre-treatment in the yogurt making procedure?
4. What was the purpose of incubating the treatment containing milk only?
5. 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.
6. Calculate the generation time of the yeast cells in the culture.
7. Name two factors that control the final ethanol concentration in a culture.
8. Although we stirred our culture each time before sampling, winemakers do not. Why
would winemakers not stir the culture?
9. Why did the pH of the culture change as fermentation proceeded?
10. Why did the specific gravity of the culture change over time?
40
EXERCISE 7
VIROLOGY
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.
(Please review the material on sewage treatment posted on the Biology 3400 web page).
Prior to the laboratory, sewage samples were collected at the areas indicated on the
schematic posted on the web page. Both samples were stored at 4 oC prior to filtering, for
up to 1 week. On the morning of the lab, samples were filtered twice using 0.45 m
filters.
PART A - ISOLATION
METHODS:
For each bench:
 Luria Methylene Blue agar plates
 Overnight culture of Escherichia coli LE 392
 Bottle of molten Luria agar overlay (at 60 oC)
 Sterile test tubes
 Test tube rack
 Micropipettor (100 L – 1000 L)
 Sterile tips
 Microbiology kits
For the lab:
 Vortex mixer
 Water bath set to 60 oC
 Raw and treated sewage filtrate
 Test tube showing 4 mL mark
Work in groups of 4.
Note that sewage filtrate contains human pathogens. Work very carefully. Students
who are clearly unprepared or are sloppy will be asked to leave the lab.
41
Period 1 - Procedure
1. Obtain a tube of culture of E. coli LE 392.
2. Obtain 5 Luria Methylene Blue agar plates, and 5 sterile test tubes. Label your 5
tubes according to Table 7.1.
Table 7.1 Experimental set-up for isolation of coliphage from sewage.
Tube #
Contents (L)
LE 392
Raw
Sewage
Filtrate
Treated
Sewage Filtrate
1
500
0
0
2
0
500
0
3
0
0
4
500
500
0
5
500
0
500
500
3. Pipette the appropriate amount of filtrate and/or cells into each of your labeled
test tubes. Leave the tubes at room temperature on your bench to incubate for 20
minutes to allow the phage to adsorb to the cells.
4. While your cultures are incubating, label your Luria Methylene Blue plates
according to Table 7.1. Mark the level of 4 mL on each of your tubes using the
marked test tube on the side bench as a guide.
5. Starting with Tube 1, aseptically pour molten agar into the tube up to the level of 4
mL. Vortex to mix, then immediately pour the contents over the surface of the
appropriately labeled plate. Swirl the plate gently to ensure that the entire surface
is covered with the agar.
6. Repeat step 5 for the remaining tubes and plates.
7. After 10 minutes, the overlay should be set. Invert your plates and place them on
a tray on the side bench to be incubated. Plates will be incubated at 37 °C for 16 –
20 hours, then stored at 4 °C until the next laboratory period.
The next laboratory:
Work in groups of four.
MATERIALS
 Pasteur pipettes
 Bulbs
 Gloves
 Chloroform (in the fume hood)
 Vortex mixer
 Phage dilution buffer
42






Plates from last lab
1 dissecting microscope per bench
Microfuge tubes (sterile)
1 mL pipettes and propipettors
Microfuge racks
Labeled microfuge rack on the side bench for class tubes
Period 2 - Procedure
1. Obtain your plates. Examine them carefully. Record the number of plaques
present for both raw and treated filtrate. Is there any difference?
2. 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.
3. After making observations, obtain a microfuge tube and aseptically add 1 mL of
phage dilution buffer to your tube. Label with your group designation.
4. 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). Release plaque into the prepared tube of
phage dilution buffer.
5. Vortex vigorously to disperse the agar.
6. 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?
7.
Place your tubes in the rack on the side bench. The tubes will be stored at 4 oC
allowing the phage to elute from the agar into the buffer.
PART B – AMPLIFICATION OF PHAGE BY PLATE LYSIS
METHODS
Cultures/phage:
 Overnight culture of E. coli strain LE 392
 Tube containing plaque isolated previous day
Other supplies:
 Waterbaths set at 37 oC, 60 oC
 Test tube racks in the 37 oC waterbath
 15 mL Falcon tubes
 Top agar (molten in 50 oC waterbath)
 Freshly prepared methylene blue LB plates at room temperature
 vortex mixers
 Biohazard disposal bags
43
Period 3 – Procedure
For Plating your Phage:
1. Obtain two freshly poured methylene blue LB plates. Label appropriately.
2. Obtain two test tubes per bench. One is for phage plus cells while the other is for a
control. Label appropriately. Aseptically, transfer 0.1 mL of E. coli LE 392 culture to
one tube.
3. To your cells in the tube, add 1/10 of the volume of your buffer (0.01 mL) containing
the plaque.
4. Set up a negative control containing E. coli only in the same fashion.
5. Place the tubes in the 37 oC waterbath and incubate here for 20 minutes.
What is the purpose of this incubation step?
6. Ensure you have all supplies for plating close at hand. Aseptically, add 3 mL of top
agar to each tube.
7. Working quickly, vortex briefly to mix phage, cells and top agar, then pour over the
surface of the labelled plate corresponding to the mixture. Swirl the plate to evenly
distribute the mixture over the entire surface. Do not swirl for more than a few
seconds!
8. Leave the plates face-up to dry.
9. Transfer the plates also face-up to the labelled tray on the side bench. Plates will be
incubated without inversion for 16-24 hours at 37 oC.
For Harvesting Phage:
Note: One member of each group will be required to come in make observations and add
buffer to plates tomorrow (out of regular lab time)
Additional Supplies
 Chloroform (in the fume hood) -* Caution: Chloroform is toxic*
 Falcon tubes (polypropylene)
 Shaker at 4 oC
 5 mL pipettes and propipettors
 Phage dilution buffer
 Pasteur pipettes and bulbs
 100 L and 1 mL micropipettors and sterile tips
 Biohazard bags
 Centrifuge at 4 oC
 Vortex mixer in fume hood
 Balance and beaker
The day prior to lab:
1. View your plates and record observations for both the control and phage treatments.
2. Aseptically add 5 mL of phage dilution buffer to the surface of each plate containing
phage and cells.
3. Place these plates on the shaker at 4 oC to shake gently overnight.
44
Period 4 - Procedure
1. Using a Pasteur pipette, transfer as much of the liquid as possible into a sterile Falcon
tube.
2. Move to the fume hood and add 0.1 mL of chloroform to each tube.
3. Vortex briefly, then find another group at the same stage for balancing your tubes.
4. Balance your tubes to within 0.1 g as follows:
 place 1 tube in a beaker on the balance pan and zero the balance
 remove the first tube and place the second in the beaker. Note the mass.
 adjust the volume in the tubes with sterile phage dilution buffer such that the
balance reads the same for both tubes.
5.
6.
Place your tubes into the centrifuge in a balanced configuration; ie, the 2 tubes
balanced against each other should be across from each other.
Centrifuge at 4000 g for 10 minutes at 4 oC.
PART C – HOST RANGE AND PHAGE TITRE
METHODS
Overnight cultures of:
 E. coli strains CSH121 and CSH125 and LE 392
 Enterobacter
Other supplies:
 Phage dilution buffer
 Micropipettors and sterile tips
 Autoclave waste disposal
 Luria Methylene Blue agar plates
 LB plates
 Bottle of molten Luria agar overlay (at 60 oC)
 Sterile test tubes
 Test tube indicating 4 mL mark
 Test tube rack
 Micropipettor (100 L – 1000 L)
 Sterile tips
 Microbiology kits
For determining phage titre:
1. Prepare serial dilutions of your phage in dilution buffer (10 -2, 10-4, 10-6, 10-8) in
microfuge tubes. Vortex each tube as you create each dilution. Ensure that you
use fresh tips for each transfer.
2. In separate, labeled sterile test tubes, mix 500 L of each dilution (10o, 10-2, 10-4,
10-6, 10-8) with 500 L of host strain E. coli LE 392. 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.
45
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 L of liquid culture to create a uniform lawn.
2. When lawns are dry, divide plates into four quadrants. Spot 20 L of undiluted
phage, or 20 L of your 10-2, 10-4 or 10-6 dilution in each quadrant. Do not invert.
Plates will be incubated at 37 oC overnight.
Period 5 - Procedure
1. The next day, score as + or – for phage growth on each host.
Thought Questions:
1. 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?
2. Have you isolated more than one type of phage? How might you be able to tell?
3. To what components of the bacterial cell do phage typically adhere?
46
APPENDIX 1
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. Please
review Appendices 1 and 9.
Magnification is achieved by the refraction of light travelling though lenses,
transparent devices with curved surfaces. In general, the degree of refraction, and
hence, magnification, is determined by the degree of curvature. However, rather than
using a single, severely-curved biconvex lens such as that of Leeuwenhoek's simple
microscopes, Hooke determined that image clarity was improved through the use of a
compound microscope, involving two (or more) separate lenses.
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.
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,
47
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.
How to use a compound light microscope:
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.
48
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 forwardto-back. Note also the two micrometer scales on the mechanical stage, which allow you
to note the coordinates of a particular object on the slide you are viewing.
Place a slide on the stage and center it over the hole in the stage. Adjust the distance
between the oculars to match your interpupillary distance (distance between your
pupils). Revolve the nosepiece so that the lowest power objective lens (generally the 10x
power lens) is in position. To focus the microscope, locate the coarse and fine
adjustment knobs at the base of the microscope, and use the coarse adjustment to move
the slide close to, but not touching, the objective lens. Look at the stage from the side as
you do this. On most microscopes this involves raising the stage, but on some the lenses
are lowered. Also, on most microscopes an automatic stop will prevent you from moving
the stage closer than about one centimeter from the lens. Now, look through the ocular
lenses, and move the slide away from the objective lens until the specimen becomes clear
(is in focus). Finish focusing with the fine adjustment knob. Once you have focused with
the low objective power lens, you may switch over to the next higher power lens with only
fine focus adjustments (the microscope is said to be parfocal).
49
As you switch from one objective lens to another, you will notice that the working
distance, the clearance between lens and stage, decreases with increasing lens power.
This is illustrated in Figure 2 below.
Figure 2: The working distance (above) and the field of view (below) change with
magnification of objective lens.
It should be obvious to you why, on high power objective lenses (40x or 100x), you must
use only the fine focus knob to adjust focus; otherwise the risk of (damaging) contact
between lens and slide becomes great. Also illustrated in Figure 2 is the diminishing field
of view as objective lens power increases; this is due to a smaller and smaller aperture at
the bottom of the lens through which light enters. This means that [a] things are harder
to find on a slide when you are using high power since only a small fraction of the slide
can be seen, and [b] less light enters your eye and everything in the field appears darker.
As a consequence, you will learn to [a] switch back to a lower power objective lens when
you want to "scan" around the slide, and [b] manipulate the amount of light coming into
the lens so that you can see the objects clearly.
The amount and concentration of light coming through the specimen and hence to your
eye can be adjusted in several ways. First, of course, is the on/off light switch, generally
located at the base of the microscope, and often associated with a rheostat to control
light intensity. A condenser lens is mounted below the stage, and concentrates the light
on to the specimen; it generally needs no adjustment of position. An iris diaphragm is
located below the condenser lens. Find the lever which controls the diaphragm; it can be
very useful in adjusting illumination and contrast.
50
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.
Phase Contrast Microscope
The phase contrast microscope was developed by Frits Zernike, a Dutch mathematical
physicist, in 1936. His discovery, for which he was awarded a Nobel Prize in 1953, led
to the development of other types of microscopes (confocal and fluorescence
microscopy).
Phase contrast microscopes are based on the principle that cells differ in refractive
index from their surroundings, meaning that light that passes through a cell differs in
phase (the light is slowed as it passes through a cell) from light passing through its
surroundings. The difference in phase is subtle, but can be enhanced, or amplified by
a device called a phase ring. The result is a dark image on a light background. Phase
contrast microscopy is a very powerful tool for observing living cells and their
activities.
Tips for use:
 Prepare a wet mount slide, place it on the stage, and find and focus on the cells using
brightfield optics at a low-power magnification.
 Once the image is in focus, move to the next highest magnification. Switch to phase
optics that match your objective lens (coding is present on the objective lenses) by
rotating the disc underneath the stage.
51


The use of phase contrast requires much more illumination as much of the light is lost
as it passes through the phase ring. Increase the amount of light hitting the specimen
once you switch to phase.
Switch to higher magnifications in the same way that you would for brightfield optics,
but remember to rotate the annular stop corresponding to the objective lens into
position each time.
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
52
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
53
APPENDIX 2
PREPARATION OF SCIENTIFIC DRAWINGS








Use a sharpened pencil; never ink. The lead should be hard.
Place drawing to one side, usually the left, leaving room for labels to the right.
Try to draw with one continuous line and do not retrace your lines. Do not shade.
Place label lines horizontally (use a ruler), with no crossed lines.
Objects labelled should be singular unless label line branches to multiple objects.
Label only what you see, not what you think should be seen.
Below the figure you should add:
 The title of the diagram
 The magnification of the drawing (see below)
The magnification of the diagram gives you the relationship between the size of your
diagram and the actual size of the specimen. A diagram of a cell would be much larger
than the actual cell, whereas a diagram of an elephant could be much smaller than the
actual elephant.

Magnification is defined as: 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

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
STAINING
Bacteria cells are very difficult to observe using compound light microscopes because
the cells appear transparent in the aqueous medium in which they are suspended.
Staining the cells prior to observation increases the contrast between the cell and the
medium, which allows for the visualization of cell structures. However, the application
of stains usually leads to cell death. Phase contrast microscopes enhance the contrast
between cells and their environment without the use of stains, meaning that living
cells and their activities can be observed. We will use both approaches to study the
morphology of microorganisms in this exercise.
Staining
In general, prior to any staining procedure, fixation occurs. Fixation performs two
functions: (i) immobilizes (kills) the bacteria; and (ii) affixes them to the slide.
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.
Gram staining behaviour 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 decolourization, 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.
56
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.
Negative staining is used to characterize external structures, like capsules, that are
associated with living bacterial cells. Negative stains make use of acidic dyes where
acidic means that they owe their coloured properties to an anion (negatively charged
molecule), so they are repelled by the negatively charged cell wall. Hence, the cell is
transparent and its surroundings are coloured.
Negative staining is useful for
determining cell dimensions and visualizing capsules, as heat fixation shrinks both cells
and capsules.
Poly--hydroxybutyric Acid (PHB) Staining
PHB granules are common inclusion bodies in bacteria.
Monomers of hydroxybutyric acid are connected by ester linkages forming long polymers which
aggregate into granules. As these granules have an affinity for fat-soluble dyes such as
Sudan black, they can be stained and then identified with the light microscope. These
granules are storage depots for carbon and energy.
Endospore Staining
Certain bacteria may produce endospores under unfavourable environmental
conditions. Endospores are mainly found in Gram-positive organisms, including the
Gram-positive Clostridium and Bacillus, in the Gram-positive cocci Sporosarcina, and in
some of the filamentous Gram-positive Monosporaceae family. It has also been
discovered that Coxiella burnetii, a small rod found in raw milk that has a variable
Gram stain reaction, but a typical Gram-negative cell wall has a sporogenic cycle.
When conditions become more favourable, the endospores will germinate and the
bacteria will return to the actively growing and dividing form.
Endospores are highly resistant to heat, chemical disinfectants and to desiccation and
therefore allow the bacterial endospore to survive much more rigorous conditions
than the vegetative cells. Endospore resistance is due to several factors, including:
 A decrease in the amount of water compared to vegetative cells
 An increase in the amount of dipicolinic acid and calcium ions
 Enzymes which are more resistant to heat
 A spore coat which is impermeable to many substances
Endospores may be formed in a central, terminal, or sub-terminal position in the cell
and their shape varies from ellipsoidal to spherical. The location of the endospore in
the cell is usually characteristic of the species. For example, the location and shape of
the Bacillus subtilis endospore is different from the location and shape of the
57
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.
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
 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.
 Add Gram’s iodine solution to the slide for 1 min. Wash gently with tap water and
remove as above.
 Decolourise with 95% ethanol by dripping ethanol on surface of slide until no more
colour is removed. Rinse gently with water. If too much alcohol is added, the
Gram-positive organisms may become Gram-negative. Remove the water after
the last wash.
 Counterstain the slide with safranin for 30 seconds - 1 minute.
 Wash the slides with tap water, air dry on paper towels, and examine under oil
immersion.
Gram positive organisms stain purple; Gram negative organisms, red (pink).
Negative Staining – Procedure
 Add a small drop of India ink to a wet mount of bacterial cells on a microscope
slide. Mix with a toothpick.
58


Add a coverslip, place the slide between two sheets of blotting paper, and apply
pressure with a cork until most of the India ink is removed (slide should appear a
light gray in colour)
Observe using phase contrast microscopy.
PHB Staining - Procedure
 Prepare smears of the organism, air dry and heat fix. Flood entire slide with Sudan
Black B and add more stain as the dye solvent evaporates. Stain for at least 10
minutes.
 Pour off excess stain (do not wash) and air dry.
 Clear slide by dipping in a jar of solvent in the fume hood for 5 sec. Air dry in the
fume hood.
 Counterstain for 1 min. with safranin.
 Wash with water, drain, blot and air dry. Examine with oil immersion objective.
Cell is pink, lipids are dark grey or black.
Endospore Staining - Procedure
 Prepare smear and heat fix. Cover the dried fixed film with a small piece of paper
towel. Saturate this with 5% malachite green.
 Place the slide on a rack over a boiling water bath. Steam slide for 5-10 minutes in
this manner. Add additional stain as needed - do not allow the slide to dry out
during this procedure.
Caution: the water bath is at 100oC – the steam will burn your skin. Take
appropriate precautions.




Allow the slide to cool, then rinse with water. Tap over a paper towel to remove
excess water
Counterstain with safranin for 30 seconds.
Rinse slide with water.
Allow to air dry, and view.
Endospores will stain green and the rest of the cell pink.
References:
Atlas, R. M. 1997. Principles of Microbiology. Wm. C. Brown Publishers, Toronto.
Madigan, M. T., and Martinko, 2006. Brock Biology of Microorganisms Eleventh
Edition. Prentice-Hall of Canada, Inc., Toronto.
Ross, H. 1992-1993. Microbiology 241 Laboratory Manual. The University of Calgary
Press, Calgary.
59
APPENDIX 4
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.
methods must be followed.
These
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, sterilized 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.
60
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 must 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 is not permitted in the laboratory. Do not place pencils, fingers
or anything else in your mouth.

Clean air contains many bacteria and fungal spores carried on dust particles or in
water droplets. Any surface exposed to air quickly becomes contaminated. If
material is to be kept sterile, it should be exposed only as much as is absolutely
necessary for manipulation.
Plugs and caps of tubes, tops of Petri dishes and bottles of solutions, (even water!!)
must not be laid on the bench nor must sterile containers and cultures be left open and
exposed to the air.
Inoculation of Culture Tubes
Again, the important thing to remember is that exposure of sterile liquids or bacterial
cultures to air must be minimised.
-Ensure that you have the tubes, plate of inoculum, inoculating loop and a sterile tube of
medium available within easy reach.
61
-Flame the inoculating loop until red hot. When removing inoculum from a tube, remove
the cap from the tube by grasping the cap between the last finger and the hand which is
also holding the inoculating needle (Figure 1). Do not place the cap on the bench!!
Figure 1: Technique for manipulating test tubes aseptically.
-Flame the mouth of the tube by passing it rapidly through the Bunsen burner 2-3 times.
This sterilises the air in and immediately around the mouth of the tube.
-Cool the loop on the inside of the tube, remove the inoculum.
-Reflame the mouth of the tube and replace the cap
-Flame the inoculating loop before replacing
-Note, when removing inoculum from a plate, cool the loop in the agar before picking up
the bacteria
Streaking for Single Colonies
-A loop of liquid culture or a small amount of bacterial growth from a plate culture is
transferred aseptically to a sterile plate in the area shown by Figure 2A.
-Once the first set of streaks has been made, the inoculating loop is reflamed until red hot.
DO NOT REINTRODUCE THE LOOP INTO THE ORIGINAL CULTURE!!!
-Cool the loop, and make a second set of streaks as shown in Figure 2B, only crossing over
the initial set of streaks once.
-Flame the loop again, cool, and repeat for three more sets (Figure 2C). Note, try not to
gouge the agar while streaking the plate.
62
Figure 2: How to prepare a streak plate.
Preparation of Spread Plates:
Generally, volumes of culture greater than 100 L are NOT plated as it takes too long for
the liquid to dry.



Use aseptic technique to obtain 100 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.
63

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. Sterilization
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 Sterilization
1.
Most media (including agar) can be sterilised by treatment with steam under
pressure in an autoclave, the usual treatment being 15-20 minutes at a pressure of
two atmospheres. This raises the steam temperature to 121°C. When using an
autoclave, the water should be allowed to boil, and the steam to fill the autoclave
before shutting the valve. This allows the material to heat up and ensures that the
correct steam pressure is attained. Never overfill an autoclave since this will upset
the pressure/volume relationship and the correct temperature will not be
attained. Materials that might be adversely affected by this treatment may
sometimes be treated for a short time or at a lower temperature, but this will not
be effective if the material is heavily contaminated to begin with. Screw caps on
bottles must be left slightly open during sterilisation and screwed down on
removal from the steriliser.
2.
Media that are difficult or impossible to autoclave satisfactorily, e.g. gelatin media
and some sugar media, may be sterilised by intermittent steaming. Objects to be
sterilised are heated over boiling water in a steamer (steam temperature 85°-95°C)
for 15-20 minutes on each of three or more successive days. Time must be
allowed for the medium to reach the same temperature as the steam. Between
treatments the material must be kept at a temperature allowing spores to
germinate (30°-37°C) and so lose their heat resistance.
3.
It is often necessary to sterilise some ingredients of a medium separately and to
add them to the rest of the medium before use. Heat-labile ingredients, e.g. urea,
serum, etc. must be sterilised by filtration through a bacteria-proof filter, i.e. Seitz
filters or membrane filters.
4.
Dry glassware, e.g. glass petri dishes, empty flasks, pipettes may be sterilised in
the autoclave and then dried or may be sterilised in a hot air oven. Any oil
material must also be sterilised in a hot air oven. The minimum effective
treatment is 1 hour at 150°C. This should be increased to 160°C or the time of
heating prolonged to 2 or 3 hours wherever possible.
64
APPENDIX 5
THE CULTIVATION OF BACTERIA
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:
photoautotroph
(photolithotroph)
photoheterotroph
(photoorganotroph)
chemoautotroph
(chemolithotroph)*
chemoheterotroph
(chemoorganotroph)
energy source
light
carbon source
CO2
light
organic
chemical (oxidation of CO2
reduced inorganic
compounds e.g. NH3,
NO2- and H2)
chemical
organic
65
source of
reducing power
inorganic
oxidizable
substrate
organic
inorganic
organic
*All chemoautotrophs are chemolithotrophs, but not all lithotrophs are autotrophic. For
example, the methylotrophic bacteria can use organic carbon as their carbon source.
Common Media Constituents:
Energy or Carbon sources:
 Sugars, alcohols, carbohydrates and amino acids
 Found in infusions – for instance – beef infusion
 Found in extracts – for instance – yeast extracts
 Also found in peptones (see below)
Nitrogen sources:
 Inorganic sources such as ammonia or nitrate
 Nitrogen fixing organisms use atmospheric N2
 Extracts, infusions
 Peptone – hydrolysis of proteins produces mixtures of short-chains of amino acids
(peptides). Sources of peptones may include meat, fish, blood, or soybeans
 Tryptone – pancreatic digestion of casein
Other Macronutrient Source Examples:
 MgSO4
 CaCl2
 Potassium salts
Micronutrient Sources:
 May not be necessary to add as these are required in such small concentrations.
Growth Factors:
 Some organisms are able to synthesise all growth factors from precursors. Other
organisms require these compounds already synthesised
 For example – thiamine, biotin
Buffering Components
Buffers, which prevent large changes in pH, are often required to facilitate growth. This is
particularly true of media composed of simple compounds or in which acid-producing
bacteria are cultivated. Mixtures of sodium and potassium phosphates are often
employed. In complex media, buffering is provided by the peptides and amino acids.
Gelling Agents
For a solid medium, agar, a water soluble polysaccharide, is added to the medium. First
discovered in 1658 in Japan, agar was first used for microbiological purposes by R. Koch in
66
1882. It is extracted from members of Class Rhodophyceae (a group of red-purple marine
algae). Agar is particularly suited to microbial propagation because:
 It lacks metabolically useful chemicals such as peptides and fermentable
carbohydrates (it cannot be broken down by bacterial enzymes)
 It melts at a high enough temperature (85 oC) to support growth of different
temperature requiring microbes
 It lacks bacterial inhibitors
Below are two examples of media used for cultivation of microbes. TY is an example of a
complex medium whereas VMM is an example of a minimal or defined medium:
TY Agar (used for the cultivation of organisms such as Rhizobium leguminosarum,
Pseudomonas fluorescens)
As with most complex media, ingredients for TY are weighed out, 1 L of water is
added, and the mixture autoclaved. After cooling slightly to approximately 60 oC,
TY medium is poured into Petri dishes.
Ingredient
Tryptone
Amount (/L)
5.0 g
Yeast Extract
3.0 g
CaCl2
MgSO4
Agar
0.5 g
0.1 g
20 g
Source of?
Macronutrients (primarily
nitrogen, also carbon and
growth factors in the
form of amino acids)
Macronutrients (primarily
carbon, also nitrogen and
growth factors)
Macronutrients
Macronutrients
Gelling agent
For the next example – VMM – three different mixtures (Solutions A, B and C) of
ingredients are made up separately, autoclaved separately, and then combined. Finally,
a carbon source is added just prior to pouring.
67
VMM (Vincent’s Minimal Medium - Vincent, 1970) (used for the study of nutritional
requirements of Rhizobium leguminosarum)
Solution A:
Source of?
Compound
Amount (/L)
Buffering
agent/
K2HPO4
1.0 g
Macronutrients
Buffering
KH2PO4
1.0 g
agent/Macronutrients
Macronutrients (nitrogen
KNO3
0.6 g
in particular)
Gelling agent
For Solid Medium: 12.5 g
Agar
Solution B (10x):
Compound
FeCl3
Amount (/L)
0.1 g
Source of?
Macro/Micronutrient
s
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
0.01 g
Growth factors
Pantothenate
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).
68
Oxygen Requirements of Microorganisms
Many species of bacteria are facultative aerobes, i.e. they can grow under aerobic or
anaerobic conditions, the latter ability being dependent upon the presence of some
substance that can be utilised as an electron acceptor by the species concerned. Some
bacteria are obligate aerobes, unable to use anything but oxygen as a final electron
acceptor. Others are obligate anaerobes that cannot use oxygen as an electron acceptor.
A few bacteria are somewhat intermediate, growing best in low oxygen tensions. These
are called microaerophilic bacteria. During growth in liquid culture, microorganisms tend
to utilise all available oxygen and so reduce the medium. Thus, the oxidation-reduction
potential (Eo) of the medium may become low enough to allow anaerobic growth to occur.
One example of this is found in the fermentation of sugar to produce alcohol by yeast
(Exercise 8 part C). Unless the mixture is stirred frequently, the little oxygen available in
the grape juice solution is utilised rapidly by the growing culture. Organisms then switch
to anaerobic growth.
In order to sample material containing anaerobes, specimens must be obtained and
immediately placed into an environment containing an oxygen-free gas and an indicator
that changes colour when oxidised to indicate when oxygen has contaminated the sample.
Organisms may then be cultured in sealed jars containing gas mixtures of N 2 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.
69
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.
70
APPENDIX 6
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 m): 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)
Shape or form
Circular
Irregular
RhizoidFilamentous Punctiform (1mm or less
in diameter)
2)
Surface: smooth/rough; mucoid/moist/dry/powdery
3)
Elevation:
Flat
4)
5)
6)
Raised
Convex
Umbonate
Umbilicate
Size: measure a single colony with a ruler
Pigment: cream, white or beige coloured organisms are usually considered to be
non-pigmented. Pigments may be purple, red, pink, yellow, brown, blue, grey, etc.
Water soluble pigments diffuse into the medium.
Opacity: Transparent (can see through) or opaque.
71
APPENDIX 7
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
72
information and draft the report accordingly. Figures may include drawings that should
be in pencil. Graphs or other figures may also be included as appropriate. Experimental
results should be presented only once. If information is presented in a figure then it
should not be repeated in a table. Each figure and table must have a caption which is
complete enough that the figure and caption can be removed from the report and still be
understandable. Figures and tables must be referred to in the text and described so that
if the reader did not have the figure or table, trends or highlights of the results would still
be evident. Never include a figure or table without referring to it and describing it; to do
so will result in loss of marks. Avoid evaluating or interpreting your results in this section.
Discussion - The discussion should refer to concepts or questions posed in the
introduction and relate these concepts from the literature to the results. Do not restate
the results in this section. Your discussion will be graded based on your evaluation of the
results with respect to the literature. Any time you use information from another source,
it must be immediately cited within the text. Failure to do this constitutes plagiarism and
may result in a mark of zero being assigned for the entire document. For examples of
how to cite properly, refer to 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.
73
APPENDIX 8
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.
74
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).
1)
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).
75
APPENDIX 9
Media, Reagents and pH Indicators
MEDIA:
Tryptic Soy Broth:
A general purpose medium used to cultivate a variety of microorganisms.
Composition (g/L):
Bacto tryptone
Bacto soytone
Dextrose
NaCl
Dipotassium phosphate
17.0 g
3.0 g
2.5 g
5.0 g
2.5 g
Dissolve in distilled water to a final volume of 1 L, dispense into test tubes, and
autoclave for 15 min at 121oC.
Tryptic Soy Agar:
Used for cultivation of a variety of microorganisms.
Composition (g/L):
Bacto tryptone
Bacto soytone
NaCl
Agar
15.0 g
5.0 g
5.0 g
15.0 g
Dissolve in distilled water to a final volume of 1 L, autoclave for 15 min at 121 oC,
and pour into sterile Petri dishes.
LB Medium (Luria-Bertani Medium):
Used for cultivation of Enterobactereaceae family members, Sinorhizobium and
Agrobacterium
Composition (g/L):
Tryptone
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.
76
Terrific Broth (TB)
Used for the cultivation of E. coli
Composition (g/L)
Tryptone
Yeast Extract
Glycerol
12.0 g
24.0 g
4.0 mL
Dissolve in distilled deionised 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 KH PO , 0.72M
2
4
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).
Nutrient Agar:
Used for the cultivation of a wide variety of microorganisms.
Composition (g/L)
Peptone
NaCl
Yeast extract
Beef extract
Agar
5.0 g
5.0 g
2.0 g
1.0 g
15.0 g
Dissolve in distilled water to a final volume of 1 L, autoclave for 15 min at 121 oC,
and pour into sterile Petri dishes.
TY Agar
Used for the cultivation of Pseudomonas and Rhizobium.
Composition (g/L):
Tryptone
Yeast Extract
CaCl2
MgSO4
Agar
5.0 g
3.0 g
0.5 g
0.1 g
13.0 g
77
Add distilled water to a final volume of 1 L, autoclave for 15 min. at 121 oC, and
pour into sterile Petri dishes.
Luria Methylene Blue Agar
Used for the observation of coliphage plaques.
Composition (g/L):
Tryptone
Yeast Extract
NaCl
Glucose
Methylene Blue
Agar
10.0 g
5.0 g
5.0 g
1.0 g
0.02 g
15.0 g
Dissolve in distilled deionised 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
NaCl
Glucose
CaCl2
Agar
10.0 g
5.0 g
1.0 g
0.11 g
6.0 g
Add 3 mL of NaOH per L and check for a pH of 7.2. Add agar, dissolve, then
autoclave for 20 minutes at 15 psi (1.05 kg/cm2) on liquid cycle.
78
Eosin Methylene Blue Agar:
Used for selection of Gram negative bacteria, and differentiation of lactose
fermenting organisms.
Composition (g/L):
Peptones
Di-potassium hydrogen phosphate
Lactose
Sucrose
Eosin Y, yellowish
Methylene blue
Agar
10.0 g
2.0 g
5.0 g
5.0 g
0.4 g
0.07 g
15 g
Dissolve in distilled water to a final volume of 1 L, autoclave 15 min at 121oC, and
pour plates.
MacConkey Agar:
Used for selection of Gram negative bacteria, and differentiation of lactose
fermenting organisms.
Composition (g/L):
Peptone
NaCl
Lactose
Bile salts
Neutral red
Agar
20.0 g
5.0 g
10.0 g
5.0 g
0.075 g
12.0 g
Dissolve in distilled water to a final volume of 1 L, autoclave 15 min at 121 oC, and
pour plates.
79
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.
80
REAGENTS:
Ethanol, 70%:
95% Ethanol
Distilled Water
36.8 mL
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
Hydrochloric acid
p-dimethylamine-benzaldehyde
75 mL
25 mL
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.
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.
81
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.
82
pH INDICATORS:
Table 1: Indicators of Hydrogen Ion Concentration.
pH Indicator
pH Range
Full Acid Colour
Full Alkaline Colour
Cresol Red
0.2 - 0.8
Red
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)
7.4 - 9.0
Yellow
Purple
Thymol
(alkaline range)
Blue 8.0 - 9.6
Yellow
Blue
Cresolphthalein
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
83