JAMES COOK UNIVERSITY
MI2011
MICROBIAL DIVERSITY
PRACTICAL NOTEBOOK – 2022
MICROBIOLOGY & IMMUNOLOGY
CPHMVS: COLLEGE OF PUBLIC HEALTH, MEDICAL AND
VETERINARY SCIENCES
2
MI2011
MICROBIAL DIVERSITY
PRACTICAL NOTEBOOK – 2022
| Subject Coordinator Lecturers Technical Staff |
Dr Jennifer Elliman x16257 Dr Constantin Constantinoiu x16635 Helen Long x14268 |
| Helma Antony x14209 | |
| Elena Constantinoiu x16186 |
AssignmentTutorOnline
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TABLE OF CONTENTS
INTRODUCTION …………………………………………………………………………………. 4
PRACTICAL 1: MICROSCOPY AND ASEPTIC TECHNIQUE ………………….. 6
PART 1: MICROSCOPY …………………………………………………………………………………………………………. 6
PART 2: ASEPTIC TECHNIQUE ………………………………………………………………………………………….. 10
PRACTICAL 2: BACTERIOLOGY – PART 1 ………………………………………… 16
PRACTICAL 3: BACTERIOLOGY – PART 2 ………………………………………… 24
PRACTICAL 4: BASIC MOLECULAR BIOLOGY OF BACTERIA –
MECHANISMS OF GENE TRANSFER ………………………………………………… 28
PRACTICAL 5: MYCOLOGY ………………………………………………………………. 36
APPENDIX 1: MACROSCOPIC EXAMINATION OF BACTERIA …………… 53
Part 2 of this Practical book will be provided later. Some alterations are still
being made to these practicals at the time of writing.
Practicals still to be provided are;
PRACTICAL 6: THE ARTHROPODS
PRACTICAL 7: PARASITOLOGY
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Introduction
Welcome to the practical component of the MI2011: Microbial Diversity.
There are a number of things to note before you begin these classes. Please make
sure you are prepared beforehand.
It is essential that you have read and understood the laboratory safety rules.
You must ensure that you have completed and understood the online component of
the induction into the laboratory prior to commencement of the first practical session.
Failure to do so will result in your not being permitted to attend the practical
classes. The practical classes are not a separate entity to lectures and your
successful completion of the course depends a great deal on these practical
exercises.
Signing into the practical is required. If your signature is not on the sign in
sheet, or on the electronic sign-in register your practical will not be marked
Some of these exercises cannot be completed in a single three hour practical
slot. In some cases you are required to continue with the exercise on subsequent
days. This will usually involve a maximum of an hour and it is your responsibility to
arrange suitable times for these continued practicals with the technician. The
demonstrator will choose a time that suits the majority of students, and this will be
regarded as the timetabled class allocation. Lack of attendance will be noted.
Many of the exercises rely upon you being able to communicate with each
other. In some cases you may be asked to exchange results in order to complete
the write ups. It is entirely your responsibility to obtain these results from each other.
Please endeavour to be punctual for the practical classes and to attend all
practical classes. As mentioned above, these classes are essential to your
understanding of this subject.
It is to your great advantage to read the notes for a practical exercise prior to
the class. Some of the procedures are complex and will require some organisation
of your time. The demonstrators will therefore assume that the notes have been
read and the introduction of the class will rely on this assumption.
Practical reports must be handed in within ONE WEEK of the completion of the
practical unless another time is stated due to the extended nature of the experiment.
Extension of time beyond this may be given in special circumstances if a medical or
other certificate is produced and given to the lecturer in charge. See subject outline
for submission rules and penalties for late submission. No work will be marked
after the return of marked scripts to other course participants.
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Practical Assessment
Practicals may be marked out of different amounts (example 10, 40, 100) depending
on the nature of the questions asked. However each practical session will be worth
a defined amount of your final grade (see table below). LearnJCU will calculate the
percentage and a running total of the percentage you have gained out of the current
assessed work will be available in the LearnJCU Gradebook.
Practicals form 30% of your final grade and the breakdown is as follows. Any
changes to this breakdown will be conveyed to you at the beginning of the relevant
practical.
| Practical | Component | % of grade |
| 1 | microscope set up Report (introduction) Aseptic technique |
0 (hurdle) 3 |
| 2 & 3 | Report – combined (results) Bacteriology |
4 |
| 4 | Report (discussion) Conjugation |
5 |
| 5 | Report (full report 5% and drawn images 5%) fungi |
10 |
| 6 | short answer worksheet Parasitology |
4 |
| 7 | short answer worksheet Parasitology |
4 |
While you will carry out most practicals in pairs, covid safe distancing will be
required and all assessable components will be assessed individually unless
specifically indicated. As such, please write up your reports individually.
Summary for turning up
1. Make sure you have all the gear you’ve been asked to bring and are
dressed appropriately before walking through the door, leaving your
bag outside.
2. Get a lab coat from the hooks and put it on
3. Sign in
4. Ethanol clean the bench and chair
5. Put the minimum required books/pens on the bench away from any
media
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Practical 1: Microscopy and Aseptic Technique
PART 1: MICROSCOPY
Background Information
A microscope is a device for magnifying objects that are too small to be seen with
the naked eye. Microscopes form a fundamental piece of equipment in microbiology,
for this reason it is important that you understand how to set up and use your
microscope optimally. Sub-optimum set-up can result in not being able to see
structures or differentiate different organisms. This can result in incorrect
identification. This practical aims to ensure that you can set up a microscope
correctly for all future practicals.
Resolution is reduced unless the surfaces of all objects in the light path, from the
lamp filament to the eye of the observer, are clean and undamaged. Attention must
therefore be given to the illuminator and condenser lenses, the specimen, and
objective and eyepiece lenses. To maintain the microscope in good working order,
dust, dirt, and fluids, especially corrosive fluids must be avoided.
With a given microscope, correctly adjusted illumination is the most important factor
in obtaining maximum resolution. The following procedure should become second
nature to you. Assume that the person who previously used the microscope left it
dirty and out of adjustment. Therefore, check that the microscope is clean and
properly set up each time you use it. Report any damage or missing parts
immediately to staff in charge.
When you have finished with your microscopes for the day, check that they are
clean, and replace them, with all attachments, where you got them from.. If the
microscope must be left on the bench, cover with a plastic dust cover.
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Principle Parts of a Microscope
The microscope consists of a number of optical and mechanical components
arranged in exact relation to each other and are illustrated in Figure 1.1.
The eyepiece is a removable lens system and may be x5 or x10 magnification.
The microscope is fitted with three objectives, two low power (LP)
magnifications x4 and x10, and two high power (HP) magnifications x40 and x100.
The stage, on which specimens are placed for examination.
Attached to the main body of the microscope and situated on either side of the
limb are the focussing knobs. These consist of a large coarse adjustment knob
which moves the stage rapidly and is generally used with low power objectives, and
a smaller fine adjustment knob which moves the stage slowly and is used with high
power objectives.
The condenser is a lens system which serves to focus light on the specimen
on the stage. Focussing is effected by the condenser adjustment knob.
The aperture iris diaphragm is situated at the base of the condenser and
serves to relate the amount of light passing through the condenser. It is opened and
closed by means of a lever.
The Illuminator is the light source for your microscope
The field iris diaphragm, is located above the illuminator, and serves to
reduce stray light, thereby increasing image definition and contrast.
Figure 1.1 Labeled image of a microscope.
Eyepieces
Objectives
Course
adjustment
knob
Fine
adjustment
knob
Stage
Aperture iris
diaphragm
Condenser
Illuminator
Field iris diaphragm
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Care of the Microscope
Do not dismantle ANY part of the microscope.
The microscope should always be carried using two hands supporting from
beneath.
Take particular care when racking (moving the stage up and down) with coarse
adjustment or swinging objectives across to a higher power objective – make sure
there is sufficient clearance.
DO NOT TILT THE STAGE WHEN EXAMINING WET PREPARATIONS.
After use, the low power objective must be in position and the stage clear of
slides.
Special lens tissue (KimWipes) must be used for cleaning eyepieces. All
microscopes must be returned, cleaned at the end of each use.
INSTRUCTIONS FOR PART 1: Use of a Microscope (Work Individually)
You are required to set up a microscope using the stained slides provided. This
activity will be marked and must be successfully completed by the end of the
Practical 3: Bacteriology session.
Setting up the Microscope
1. Look at the slide macroscopically and take note of the shape, colour and
position of material.
2. Turn in the 4X objective and place the slide on the stage with the coverslip
uppermost, on the front of the stage. Gently holding back the spring arm of the
mechanical stage, push the slide back into the slide holder and release the arm
slowly. The specimen will be held firmly.
3. Turn the brightness control to setting number 5 or halfway.
4. Turn on the electricity at the power point and on the base of the microscope.
Adjust the light intensity until it can be seen shining through the condenser.
5. Adjust your seat so that you can look comfortably into the eyepieces of the
microscope without stretching upwards or bending forwards.
6. Look down the microscope and adjust the distance between the eyepieces to
merge the left and right view fields into one. This can be achieved by pulling
the two bodies of the binocular upwards or towards, or sliding the eyepieces
nearer or further apart; depending on which microscope model you are
operating.
7. Focus individual eyepieces:
a) Look into the right eyepiece (with your right eye) and focus on the slide
using the course and/or fine focus of the microscope.
b) Look into the left eyepiece (with your left eye) and focus on the slide
using the knurled adjusting ring on the eyepiece tube.
8. Turn in the 10X objective and refocus the microscope if necessary.
9. Close the field iris diaphragm.
10. Rack the condenser up and down until an image of the edge of the field iris
diaphragm is as sharply in focus as possible (Figure 1.2).
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Figure 1.2
11. If the image of the field iris diaphragm is not centred, inform a technician who
will show you how this can be fixed. Do not play with the adjustment screws
yourself at this stage.
12. Open the field iris diaphragm until its image moves just outside the field of
view. Close the aperture iris diaphragm (lever on condenser). The image of
the specimen becomes darker and more refractile. Now open the aperture
diaphragm until the field of view just reaches its maximum brightness. The
microscope is now set for Koehler illumination for the 10X objective
Using the 40X Objective
1. After focussing the microscope using the 10X objective carefully revolve the
nosepiece to bring the 40X objective into place.
2. Only slight focusing with the fine focusing control should be necessary to bring
the specimen into sharp focus. Open the aperture iris (condenser) more, and
increase the brightness control to obtain a bright, clear image.
If the 40X objective has been damaged, it may not be possible to focus. If
foxus is not possible after wiping across the lens with a kimwipe, please check this
with a technician. If damage is confirmed, skip to 100X instructions.
Using the 100X oil immersion objective
1. After focussing the microscope using the 40X objective carefully revolve the
nosepiece anti-clockwise to move to the 4X objective, place a drop of
immersion oil on the specimen.
2. Turn the nosepiece anti-clockwise to locate the 100X objective. Do not move
the other objectives through the immersion oil.
3. Only use the fine focusing control to focus the specimen. Open the aperture
iris (on condenser) more to obtain a bright, clear image.
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Removing the Specimen and Cleaning the Microscope
1. Revolve the nosepiece clockwise, so that the objective moves to one side.
2. Remove the slide from the slide holder. At this point it is important to clean the
eyepieces, condenser lens, stage, illuminator lens and lastly the objectives,
with the 100X objective to be cleaned last.
3. To clean immersion oil off the 100X objective, a KimWipe lens tissue
moistened with xylene should be used and should be dried immediately with a
dry KimWipe.
4. All microscopes must be returned, cleaned at the end of each use.
PART 2: ASEPTIC TECHNIQUE
Background
Aseptic technique refers to all procedures performed in a sterile manner. These are
procedures carried out in such a way that contamination of media does not occur.
Contamination of media can result in the media being discarded or incorrect
identification of organisms. Aseptic techniques are defined as methods used to
prevent unwanted organisms from contaminating media and needs to be developed
and practiced to develop good skills. This practical will refresh skills from previous
subjects as well as develop an understanding of the microbial ecologies found on
everyday surfaces.
General Rules of Aseptic Techniques
| | Remove all unnecessary items from the work area; keep the work area as clean as possible. Avoid spluttering of attached material during the heating process so as not to create aerosols. Liquid held in the loop can be dried relatively safely by |
| |
holding the loop in the blue zone of flame and then moving the loop upward
through the flame.
| | Keep cultures closed as much as possible, working quickly and accurately is essential. Carry out all manipulations within the zone of sterility, which is within 15 |
| |
centimeters of the Bunsen burner flame. This ensures an upward air draft that
will take any aerosols into the flame thus keeping your work area sterile.
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INSTRUCTIONS FOR PART 2: Aseptic Techniques (Work Individually)
Use of Wire Loop
Subculture Serratia marcescens provided on nutrient agar (NA) plates onto:
a) 1 X nutrient agar plate using the streak plate method
1. Label the base of the agar plate with your name, the date and the organism
that you are subculturing.
2. Place the agar plate for subculturing and the previously cultured agar plate
upside down on the bench so that the lids are on the bench.
3. Sterilise your wire loop as follows:
Adjust the flame of the Bunsen burner to produce a flame with a short, blue,
central zone. Hold the metal handle of the loop at 45 degrees and place in the
hottest part of the flame, which is just above the blue cone, until the whole
length of the wire is glowing red. Allow to cool before use.
4. Using the loop pick a single colony of bacteria from a previously cultured plate.
5. Replace the lid of the previously cultured agar plate.
6. Hold the agar plate for subculturing within the zone of sterility leaving the lid on
the bench.
7. Gently rub the inoculated loop across the plate spreading the bacteria over part
of the plate as illustrated below (Figure 1.3) and continue the streaking as
depicted, sterilizing the loop in between.
8. Place your agar plate in the container provided to be incubated at 37˚C for 24
hours.
Figure 1.3 Steak plate method to isolate single colonies of bacteria
Sterilise
loop
Sterilis
e loop
Sterilise
loop
1
st
streak
2
nd
streak
3
rd
streak final streak
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b) 1X slope of nutrient agar (solid medium)
1. Label the slope medium lid with your name, the date and the organism that
you are subculturing.
2. Place the previously cultured agar plate upside down on the bench so that the
lids are on the bench.
3. Hold the previously cultured agar plate within the zone of sterility leaving the
lid on the bench.
4. Using a sterile loop pick a single colony of bacteria from a previously cultured
plate.
5. Replace the agar plate lid.
6. Remove the lid of the slope medium bottle and retain in the hand avoiding
contamination (Figure 1.4).
7. Sterilise the top of the bottle; by waving it through the hottest part of the
flame.
8. Gently rub the inoculated loop back and forth gently over the agar slope; from
bottom to top in a zigzag pattern.
9. Sterilise the top of the bottle again.
10. Replace the lid of the sample bottle.
11. Re-sterilise the loop.
12. Place your slope in the container provided for incubation at 37˚C for 24 hours.
Figure 1.4 Illustration demonstrating one way in which
bottles can be manipulated to maintain sterility and
minimise contaminated. This method can be employed to
transfer bacteria from agar plates to slopes and broths or
to transfer materials between bottles.
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c) 1 X nutrient broth
1. Label the liquid medium lid with your name, the date and the organism that
you are subculturing.
2. Place the previously cultured agar plate upside down on the bench so that the
lid is on the bench.
3. Hold the agar plate within the zone of leaving the lid on the bench.
4. Using a sterile loop pick a single colony of bacteria from the plate.
5. Replace the agar plate lid.
6. Remove the lid of the liquid medium bottle and retain in the hand avoiding
contamination (Figure 1.4).
7. Sterilise the top of the bottle by waving it through the hottest part of the flame.
8. Move the inoculated loop back and forth within the liquid medium, fairly
vigorously to remove all bacterial cells from the wire loop.
9. Sterilise the top of the bottle.
10. Replace the lid of the sample bottle.
11. Re-sterilise the loop.
12. Place your slope in the container provided for incubation at 37˚C for 24
hours.
Use of Disposable Pipette
1. Label the liquid medium lid with your name, the date and the organism that
you are subculturing.
2. Remove the lid of the liquid medium bottle and retain in the hand avoiding
contamination.
3. Sterilise the top of the bottle; by waving it through the hottest part of the
flame.
4. Transfer one drop of Escherichia coli suspension into the medium, using a
plastic disposable pipette. Microbial growth should occur after incubation.
Keeping Media Sterile
1. Label 2 bijoux of uninoculated nutrient broth with your name and date.
2. Using a plastic disposable pipette, transfer a small amount of medium back
and forth between the 2 bijoux 6 times.
3. Place your media in the container provided for incubation at 37˚C for 24
hours. No microbial growth should occur after incubation.
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Part 3: Microorganisms on Everyday Objects (Work in Pairs)
1. Using a sterile swab, moistened with sterile water, swab an object within the
laboratory. For example; pens, books, jewellery, door handles or bench
surfaces.
2. Label the base of a nutrient agar plate with your name, date and the location
that the swab was taken from.
3. Place the agar plate upside down on the bench so that the lid is on the bench.
4. Hold the agar base within the zone of sterility leaving the lid on the bench.
5. Move the swab over the entire surface of the agar plate. This can be achieved
by streaking the swab back and forth across the plate working up and down
several times. Turn the plate 90 degrees and repeat rubbing the swab back
and forth and up and down across the entire plate. Turn the plate 45 degrees
and streak a third time (Figure 1.5).
6. Place your media in the container provided for incubation at 37˚C for 24 hours.
No microbial growth should occur after incubation.
Figure 1.5 Illustration demonstrating how to inoculate an agar plate evenly for the
lawn plating method. If the microbes are plentiful this method provides the possibility
of confluent growth over the entire area of the agar plate. Based on the diameter of
the plate, it is possible to calculate the surface area sampled and report the results
as a count/cm3.
15
Preparation for Practical 2: Bacteriology
Water Survey (Working in Pairs)
Take a sterile specimen container and collect a water sample which will be
used in Practical 2: Bacteriology. Preferably obtain the sample on the day of the
practical session, alternatively refrigerate following collection (refrigeration can lower
the number of environmental bacteria found).
Examination of Results (to be done during practical 2)
• Examine your cultures from the microscopy and sterile technique practical. All
plates should have well separated colonies. Please see Appendix 2.2: Macroscopic
Examination of Bacteria if you want to describe your colonies. The slopes should
have confluent growth and the liquid cultures should be turbid, indicating the growth
of microorganisms.
• Examine your sterile bijouxs; which should appear clear and transparent;
turbidity and cloudiness indicates contamination and poor aseptic technique.
Total Assessment (3%)
Microscope setup (Hurdle only) – assessed by the end of practical 3 (about ½
of students should be assessed today)
Report (3%) – Introduction (aseptic technique only – don’t introduce
microscopy) will be graded as per rubric – due Tuesday week 3
BEFORE LEAVING THE LABORATORY!
PLACE ALL CULTURES TO BE INCUBATED IN THE CONTAINERS
PROVIDED.
ENSURE ALL BOTTLES AND PLATES ARE LABELLED CORRECTLY
AND SEALED APPROPRIATELY.
REMOVE LABELS FROM GLASSWARE AND PLASTICWARE
ALL USED MATERIALS SHOULD BE PLACED IN THE CORRECT
RECEPTACLES, AS DETAILED IN, BIOSAFETY PRECAUTIONS FOR
PRACTICAL CLASSES IN MICROBIOLOGY AND IMMUNOLOGY; “DISPOSAL
OF WASTE MATERIAL”.
RETURN YOUR MICROSCOPE TO ITS PLACE OF ORIGIN.
WIPE DOWN THE BENCH AND CHAIR WITH 70% ETHANOL AND
PAPER TOWEL. DO NOT USE KIMWIPES.
REMOVE YOUR LAB COAT AND PLACE IN BAG FOR CLEANING.
LASTLY WASH YOUR HANDS, PRIOR TO LEAVING THE LABORATORY.
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Practical 2: Bacteriology – Part 1
All sections of this practical session will be completed in pairs.
Background Information
This practical provides the opportunity to practice commonly used bacterial
identification techniques such as Gram stains, motility tests and production of media.
Both laboratory strains of bacteria and swabs from the environment will be used.
Pouring agar plates is a necessary skill for the microbiologist, both for producing
sterile plates and for producing pour plates of bacteria. You must be able to carry
out this technique without contaminating your agar or producing lumps in the plate.
Part 1 and 6 of this practical provides you with the opportunity to practice these
techniques.
Selective and differential media are used extensively in microbiology laboratories to
identify particular bacteria in mixed cultures and confirm suspected identifications. In
this practical you will become familiar with three selective agars; Mannitol Salt agar,
Cetrimide agar and MacConkey agar (which is also differential). You should be
familiar with the circumstances under which you might use each agar.
Before beginning the inoculation component of this practical, examine the plates etc
you handled last week. Ensure they look the way they should and adjust your
technique today if necessary.
Part 1: Agar Plate Preparation
Please ensure this procedure is completed with the supervision of a staff member.
1. Label the base of 3 petri dishes provided with your name and the date.
2. Obtain 3 x 20ml bottles of molten nutrient agar from the water bath provided.
The agar medium is maintained in a molten state at 45° C.
3. Pour the molten agar into the 3 petri dishes within the C2 cabinet.
4. Immediately mix the molten agar to ensure a continuous distribution within
the petri dish by sliding the agar plate on the bench in a figure of 8 motion.
5. Replace the agar plate lid, slightly ajar, and allow for the agar to solidify.
These agar plates will be inoculated in Part 3: Microbes in the Environment section
of the practical session.
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Part 2: Selective and Differential Media
Specialised media have been devised to facilitate the isolation and identification of
bacteria. Selective media contains components which selectively inhibit the growth
of certain microorganisms. Differential media typically have a pH indicator which
allows the differentiation between various chemical reactions during growth.
Nutrient agar supports the growth of all conventional bacterial organisms.
MacConkey agar is a selective and differential medium that inhibits Gram positive
organisms and allows for the differentiation of Pseudomonas (appears translucent)
from E. coli (appears pink in colour). Cetrimide agar is a selective medium that
contains cetrimide to inhibit the growth of most organisms except Pseudomonas
which produces a green pigment. Mannitol salt agar (MSA) is used to selectively
isolate Staphylococcus. Pathogenic Staphylococcus (coagulase positive
Staphylococcus) will form small yellow colonies on the MSA plates, as the organism
ferments mannitol. However, non-pathogenic Staphylococcus (coagulase negative
Staphylococcus) will form small colourless colonies on the MSA plates as it does not
ferment mannitol.
A broth containing a mixture of 3 bacteria (Staphylococcus aureus, E. coli and
Pseudomonas aeruginosa) is provided.
| 1. | Streak the mixed broth onto the following media: |
| |
Nutrient agar (NA) MacConkey agar (MAC) Cetrimide agar (CET) |
NOTE: Remember to label your plates with your name and the date
Table 2.1 Observations mixed cultures streaked on different agar plates
| Agar | Colony morphologies noted | Presumptive ID |
| nutrient | ||
| MacConkey | ||
| Cetrimide |
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Part 3: Microbes in the Environment:
Airborne
1. Expose 1 plate (prepared in Part 1: Agar Plate Preparation) to the air for 20
minutes. Select various positions inside and outside of the laboratory building.
Body Surface (External):
1. Use a sterile swab moistened with sterile water to swab the interdigital
spaces of the left hand of one individual in your pair.
2. Inoculate a nutrient agar plate (prepared in Part 1: Agar Plate Preparation),
using the lawn plating method (refer to Practical 1, Part 3 for method).
3. Wash hands of the subject with soap and water OR disinfectant and water (as
directed by the instructing staff members). Lightly dry hands with a paper
towel.
4. Use a sterile swab moistened with sterile water to swab the interdigital
spaces of the opposite hand as completed previously.
5. Inoculate a nutrient agar plate (prepared in Part 1: Agar Plate Preparation),
using the lawn plating method.
Body Surface (Internal)
1. Using a sterile swab, which has been moistened with sterile water, swab the
nasal cavity of one individual in your pair.
2. Inoculate a mannitol salt agar (MSA) plate using the lawn plating method.
Table 2.2 Observations of environmental contamination
| Location | Colony morphology/number of colonies |
Diversity (high/low) and likely ID if possible |
| Airborne: | ||
| External pre clean | ||
| External post clean | ||
| Internal |
19
Part 4: Motility Test:
Observe live unstained P. aeruginosa and S.aureus by completing the wet
preparation method. Ensure that you observe a positive control initially, and then
progress onto the sample.
1. Place a drop of liquid culture onto the centre of a glass slide.
2. Cover with a coverslip.
3. Observe under the microscope, setting it up as previously instructed.
NOTE: An effective method to use is to focus the view at the edge of the coverslip,
and then move to the centre once you have achieved this initial focal point.
Commence with the 10x objective and then once in focus progress to the 40x
objective. It is much easier to see the bacteria when the iris diaphragm is almost
completely to the right (this increases the contrast). Attempt to observe the individual
bacteria moving to avoid mistaking motility with Brownian movement. Brownian
movement is where the organisms may appear quite active, but they typically remain
in the same relative position to other organisms or debris in the visible field.
Which one of these organisms is motile and which is non-motile? Fill in your
observations in table 2.3
Part 5: Gram Stain
Gram staining is the most widely used staining procedure in bacteriology. It is a
differential stain differentiating between Gram-positive and Gram-negative bacteria.
Bacteria that stain purple are termed Gram-positive; those that stain pink are Gramnegative bacteria.
Gram-positive and gram-negative bacteria stain differently because of differences in
the structure of their cell walls; bacterial cell walls contain peptidoglycan. Grampositive bacterial cell walls appear thick and consist of numerous interconnecting
layers of peptidoglycan. Typically, 60% to 90% of the cell wall is peptidoglycan.
Gram-negative bacterial cell walls contain a much thinner, single layer of
peptidoglycan only 2 or 3 layers thick which forms only 10% to 20% of the cell wall.
Prepare a Gram stain of S.aureus and E.coli from the agar plate cultures provided
using the instructions below. Fill in observations in table 2.3
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Preparation of bacterial film
1. Label the slide, with your name and the organisms that you are preparing
(Figure 2.1).
2. Place the slide as close to the Bunsen burner as possible
3. Remove the lid of a sterile bottle of saline and retain in the hand avoiding
contamination.
4. Sterilise the top of the bottle; by waving it through the hottest part of the
flame.
5. Sterilise a loop in the Bunsen burner and transfer a loop of the water on the
slide for each sample. Re-sterilise the wire loop.
6. Place the previously cultured agar plate upside down on the bench so that the
lid is on the bench.
7. Pick up the plate leaving the lid on the bench.
8. Sterilise a loop in the Bunsen burner and pick a single colony of bacteria.
9. Replace the agar plate lid.
10. Smear the loop, containing bacteria, in the saline that you have placed on the
slide and spread over an area of approximately 1cm2.
11. Re-sterilise the loop.
12. Allow the film to air dry, or if you are pressed for time, pass the slide over the
Bunsen burner flame. However ensure the slide does not become too hot,
you can assess the temperature of the slide by touching it to the back of your
hand. If it is too hot to touch, allow the slide to cool down in temperature.
Figure 2.1 Example of how to position your samples for Gram staining to include
both a negative and positive control.
21
Gram Staining
Staining kits are on the side benches under the windows at the stainless steel
sinks.
1. Cover the entire bacterial film with crystal violet.
2. Leave the stain on for 1 minute. Drain off the crystal violet, do not rinse with
water.
3. Cover the entire bacterial film with iodine solution
4. Leave the stain on for 1 minute. Rinse the slide with tap water
5. Cover the entire bacterial film with decolouriser for 5-10 seconds. Please
ensure that you do not over-decolourise, by leaving the decolouriser on the
slide for an extended period of time.
6. Immediately rinse with tap water. Cover the entire bacterial film with safranin.
7. Leave the stain on for 1 minute then wash gently with water and pat dry with
paper towel or tissue.
Microscopic Examination of Bacteria
Bacterial cell morphologies/arrangements are described by their shapes, groupings,
and organisation (refer to Figure 2.2)
Figure 2.2 Microscopic bacterial cell morphologies
22
Table 2.3 Microscopy observations
| Slide | Cell morphology/movement |
| Motility: P. aeruginosa | |
| Motility: S.aureus | |
| Gram stain: S. aureus | |
| Gram stain: E. coli |
Part 6: Viable count of water samples: This is the only result that will be used
in your practical report along with week 3 practical results
You will need to use the water that you have collected prior to arriving to the practical
session.
1. Gently shake the water sample container to ensure the sample is well
distributed through the container.
2. Pipette 1ml of the water sample aseptically into 9ml of saline.
3. Mix well. This is a 10-1 dilution.
4. Transfer 1ml of the 10-1 dilution to 9ml of saline.
5. Mix well. This is a 10-2 dilution.
6. Label the base of the 3 empty petri dishes provided, with your name, the date
and dilution.
7. Plate 1ml of each of the 10-1 dilution and 10-2 dilutions onto 2 of the petri
dishes.
8. Within close proximity to the Bunsen burner, pour the molten agar into these
petri, avoiding direct contact with the sample.
9. Additionally, pour molten agar into the third, empty petri dish, within close
proximity to the Bunsen burner. This will be your negative control plate; which
will determine the sterility of the plate formation.
10. Immediately mix the molten agar by sliding the agar plate on the bench in a
figure of 8 motion.
11. Replace the agar plate lid, slightly ajar, and allow for the agar to solidify.
Table 2.4 Viable count of water sample
| Water source: | ||
| Dilution | Raw counts | Cfu/ml |
| 10-1 | ||
| 10-2 | ||
| Negative control plate |
23
NOTE:
Remember to label all your plates with your name, date and organism or
sample name.
All samples from this practical session are incubated in a 37°C
incubator for 24 hours and are then placed in a refrigerator to prohibit
further bacterial growth.
Total Assessment practical 2&3 combined (4%) (due Tuesday week 5)
Results (of section 6 only) will be graded as per rubric
BEFORE LEAVING THE LABORATORY!
PLACE ALL CULTURES TO BE INCUBATED IN THE CONTAINERS
PROVIDED.
ENSURE ALL BOTTLES AND PLATES ARE LABELLED CORRECTLY
AND SEALED APPROPRIATELY.
REMOVE LABELS FROM GLASSWARE AND PLASTICWARE
ALL USED MATERIALS SHOULD BE PLACED IN THE CORRECT
RECEPTACLES, AS DETAILED IN, BIOSAFETY PRECAUTIONS FOR
PRACTICAL CLASSES IN MICROBIOLOGY AND IMMUNOLOGY; “DISPOSAL
OF WASTE MATERIAL”.
RETURN YOUR MICROSCOPE TO ITS PLACE OF ORIGIN.
WIPE DOWN THE BENCH AND CHAIR WITH 70% ETHANOL AND
PAPER TOWEL. DO NOT USE KIMWIPES.
REMOVE YOUR LAB COAT AND PLACE IN BAG FOR CLEANING.
LASTLY WASH YOUR HANDS, PRIOR TO LEAVING THE LABORATORY.
24
Practical 3: Bacteriology – Part 2
All sections of this practical session will be completed individually.
Background information
When identifying unknown bacteria, there are several common classical biochemical
tests that are normally used prior to a wide variety of genus specific tests. Numerous
bacteria produce hydrogen peroxide; some bacteria are susceptible to hydrogen
peroxide and produce catalase which provides protection from the harmful effects of
hydrogen peroxide. Organisms containing catalase release gaseous oxygen from
hydrogen peroxide. This principle forms the basis for the catalase production test. The
oxidase reagent (tetra methyl para-phenylene diamine dihydrochloride) used in the
oxidase test is colourless when freshly prepared but is rapidly oxidised by the enzyme,
cytochrome c oxidase. Bacteria that are unable to oxidise the dye are lacking this
enzyme.
This practical will provide the opportunity to utilise these tests to differentiate
between three possible identifications for an unknown bacterial isolate. Alternative
biochemical systems such as API’s are also examined as are some specialised
bacterial stains that are used for specific bacteria.
You will also need check the results of last weeks practical to confirm your technique
is sufficient before repeating those protocols today. Ensure you gather all of these
results today and fill in the tables in Practical 2. The Practical report for practical 2
and three will be based on today’s practical and the results of tables 2.2 and 2.4 from
practical 2. Use these and the results from today as a basis for your results section
report.
Primary Biochemical Tests
There are 3 unknown bacterial samples that need to be identified. You will be
provided with 1 of the 3 unknowns and will need to identify this using the methods
outlined in Table 3.1. Compare your results with Table 3.2 to identify your unknown
bacterial sample. To ensure results are relevant, reliable and interpreted correctly the
use of positive and negative control organisms is essential. To achieve this all
control procedures need to be conducted at the same time and in the same manner
as the sample being tested.
25
Table 3.1
| Gram stain +/- and shape |
Catalase Test |
Oxidase Test | Motility Test |
| Unknown organism |
|||
| Positive control |
|||
| Negative control |
X | ||
| Identified organism as: |
Gram Stain
Refer to Practical 2, Part 5 for method.
Catalase Production
1. Place a glass slide, your cultured plate and your tootpick container within the
zone of sterility.
2. Select a colony from a previously cultured plate using a toothpick. Take care to
avoid agar. Smear this onto the surface of a glass slide
3. Place a drop of catalase reagent (hydrogen peroxide) onto the deposited bacteria
on the glass slide.
4. Observe the bacteria for the production of bubbles
Note: If the organism is catalase positive, bubbles will develop around the
inoculum almost immediately.
Oxidase Production
1. Place a filter paper onto the surface of a glass slide.
2. Add 2 to 3 drops of the oxidase reagent to the filter paper
3. Place the previously cultured agar plate upside down on the bench so that the
lid is on the bench.
4. Hold the plate within the zone of sterility leaving the lid on the bench.
5. Using a sterile toothpick pick a single colony of bacteria from a previously
cultured plate. Do not use a metal loop.
6. Replace the agar plate lid
7. Smear the bacterial colony onto the filter paper on the surface of a glass slide
Note: If the organism is oxidase positive, a deep blue/purple colour will develop
on the filter paper within approximately 10 seconds.
26
Motility Test
Refer to Practical 2, Part 4 for methods. Use the broth cultures provided to complete
this test. Bacteria grown on solid media often demonstrate reduced motility.
Table 3.2 Expected results of biochemical tests for various bacteria
| Gram Reaction | Catalase Test |
Oxidase Test | Motility Test | |
| Staphylococcus aureus |
positive cocci in clumps |
positive | negative | negative |
| Escherichia coli | negative rods | positive | negative | positive |
| Pseudomonas aeruginosa |
negative rods | positive | positive | positive |
| Lactococcus lactis |
positive cocci in pairs or chains |
negative | negative | negative |
Appareils et Procedes d’Identification System (API)
The API 20E demonstrated in this practical session, was inoculated with an unknown
bacteria. Interpret the results of the API and compare the results you interpreted to
the results illustrated in Table 3.3, and identify the unknown bacteria.
Table 3.3
| ONPG | ADH | LDC | ODC | CIT | H2S | URE | TDA | IND | VP | GEL | GLU | MAN | INO | SOR | RHA | SAC | MEL | AMY | ARA | OX | |
| E.coli | + | – | + | + | – | – | – | – | + | – | – | + | + | – | + | + | – | + | – | + | – |
| Proteus mirabilis |
– | – | – | + | + | + | + | + | – | – | + | + | – | – | – | – | – | – | – | – | – |
| Klebsiella sp. |
+ | – | + | – | + | – | + | – | – | + | – | + | + | + | + | + | + | + | + | + | – |
| Unknown |
Acid-Fast Stain (Ziehl-Neelsen)
Observe the slide or laminate of Mycobacterium provided as a demonstration and
and illustrate, label and describe your observations.
Spore Stain
Endospores may be observed during the interpretation of a Gram stain, however a
special spore stain will demonstrate bacterial spores much more clearly. Cells will
stain red while the spores will stain green.
Observe the spore stain preparation or laminate provided as a demonstration and
illustrate, label and describe your observations.
27
Complete Tasks from the Previous Bacteriology Practical
Examine the plates exposed to the airborne and external body surfaces
bacteria. Count the number of colonies formed by the different microorganisms.
Describe the bacterial colonies, as outlined in Appendix 1 Macroscopic Examination
of Bacteria. Consider What effects do the different hand washing treatments
have on the microbial flora?
Examine the plate inoculated with the nasal swab collection and external
body surface swab. Describe the bacterial colonies, as outlined in Appendix 1
Macroscopic Examination of Bacteria. Does your plate indicate the person
sampled is carrying a pathogen?
Examine the plates prepared for the water quality assessment. Count the
number of colonies for each dilution. Note: colony morphology is not required for
pour plates. How much bacteria was in the original water sample? – include this
section only in Practical 3 results
Total Assessment practical 2 and 3 (4%) (due Tuesday week 5)
RESULTS will be graded as per rubric
You must process your results individually, even though you worked in a team to
produce them.
Please include results from 2.4 of practical 2. Please check through practical 3 for
ALL instances where you have been asked to describe observations and ensure
they are included and tables and figures referenced in text . Please check that you
have noted any unusual or incorrect results.
BEFORE LEAVING THE LABORATORY!
PLACE ALL CULTURES TO BE INCUBATED IN THE CONTAINERS
PROVIDED.
ENSURE ALL BOTTLES AND PLATES ARE LABELLED CORRECTLY
AND SEALED APPROPRIATELY.
REMOVE LABELS FROM GLASSWARE AND PLASTICWARE
ALL USED MATERIALS SHOULD BE PLACED IN THE CORRECT
RECEPTACLES, AS DETAILED IN, BIOSAFETY PRECAUTIONS FOR
PRACTICAL CLASSES IN MICROBIOLOGY AND IMMUNOLOGY; “DISPOSAL
OF WASTE MATERIAL”.
RETURN YOUR MICROSCOPE TO ITS PLACE OF ORIGIN.
WIPE DOWN THE BENCH AND CHAIR WITH 70% ETHANOL AND
PAPER TOWEL. DO NOT USE KIMWIPES.
REMOVE YOUR LAB COAT AND PLACE IN BAG FOR CLEANING.
LASTLY WASH YOUR HANDS, PRIOR TO LEAVING THE LABORATORY.
28
Practical 4: Basic Molecular Biology of Bacteria –
Mechanisms of Gene Transfer
Background Information
From lectures, you will be aware that there are 3 main mechanisms of gene transfer
between bacterial cells – conjugation, transformation and transduction. Gene transfer
can occur between cells of the same species or between cells of different species or
even different genera. If the transferred DNA becomes stable within the recipient
cell, this cell acquires the characteristics carried by that DNA. Hence, gene transfer is
a means of providing variation within a species.
In the practical exercise you will demonstrate conjugation within the controlled
environment of the laboratory. You should be aware however, that this
mechanism can and does occur naturally.
You should take note that these exercises are extensions of the lectures and the 2
should not be treated separately. You will find that your lecture notes will help you
with, and are necessary to, complete your reports. Likewise, you will find that the
practical exercises will assist in understanding of the lectures.
You will note that the exercises require some input on the days following the
practical class. This cannot be avoided. Whilst in other classes, your plates can be
stored until the following week; this cannot be applied here as pigment, plasmids
and other transient factors which are crucial to the exercises may be lost upon
prolonged storage. For the follow-up sessions, at least 1 person per group should
arrange to attend although it is obviously better for you all to observe the results.
Conjugation
Conjugation is a means of plasmid-mediated transfer of DNA between bacteria.
Some plasmids (F+) contain specific transfer genes (tra) which allow the plasmid to
be copied and transferred to another cell. These donor cells are able to produce a
sex pilus which is a rigid protein tube which extends out from the cell wall. The pilus
contains recognition molecules to locate a sensitive cell (plasmid minus, F), and will
attach to the receptor on this cell. The pilus retracts, pulling the cells together and
forming a short conjugation bridge. The F+ plasmid replicates itself within the donor
cell using a rolling circle model and a linear single stranded copy of the plasmid
crosses the bridge into the recipient cell. Once within the cell, the DNA circularises
and replicates itself into a more stable double stranded plasmid. The recipient cell
has now become transconjugant (F+) and is able to produce pili and become a donor
29
cell. It should be remembered that the plasmid will also undergo vertical transfer and
will be copied into all daughter cells as F+ bacteria divide.
In this practical exercise, you will attempt the conjugative transfer of violacein genes
(which confer a violet pigment to the host cell) and kanamycin resistance. You will
be provided with two strains of E. coli, one of which will be the donor and the other
the recipient. Based on colony morphology and growth of the species before and
after conjugation, you should be able to determine which is the donor and which is
the recipient and what order the donated genes sit on the conjugative plasmid.
You will be supplied with:
1 x LB agar plate of E. coli Q358/R751 (24 hour culture)
1 x LB agar plate of E. coli 1B609 (24 hour culture)
LB agar plates containing kanamycin (4)
LB agar plates containing streptomycin (4)
LB agar plates containing kanamycin and streptomycin (4)
LB broth
Microfuge tubes
A flow chart is provided after the instructions to aid in following the instructions.
1. Note the morphology of each E. coli strain, with particular regard to any
colour pigments, size and smoothness/roughness of the colonies. Use the
space below to make these observations.
2. Add 500µl of LB broth to a microfuge tube. Using a sterile loop, suspend a
0.5 McFarland suspension (only just turbid) of E. coli IB609 in this 500µl LB.
Flick/shake/agitate tube to break any clumps.
a) Prepare a suspension by obtaining a fresh, pure culture of the organism
and inoculating a suitable broth.
b) Visually compare the turbidity of the created suspension with that of the
McFarland standard demonstration provided.
c) If the created suspension is too light, inoculate with additional organisms
until turbidity matches that of the standard. If dilution is necessary, use a
sterile pipette and add sufficient broth to obtain a turbidity that matches that of
the standard.
30
3. Repeat step 2 with E. coli Q358/R751 in a separate microfuge tube.
4. Streak both E. coli cultures onto each of the LB agar plates containing the
single antibiotics and spread plate onto the LB agar plates containing both
antibiotics – as per step 9 below (you should now have inoculated 6 plates):
a) Kanamycin (LB kan) (streak plate)
b) Streptomycin (LB strep) (streak plate)
c) Kanamycin & Streptomycin (LB k&s) (spread plate)
5. Set these plates aside for later incubation.
6. Aseptically transfer 1 culture into the other, label top of tube clearly with your
name and mix by tube inversion.
7. Incubate your E. coli mixture in the rack provided in the 37°C hotblock for the
allotted time assigned to you by the instructors (10 or 30 min).
8. Streak the mixture on each of the single antibiotic LB agar plates (LBkan,
LBstrep).
9. Using the spread plate method, pipette 100µl of the mixture onto the mixed
antibiotic (LBk&s) plates. Details of this method are below;
a) Add the bacteria to the plate.
b) Remove the lid from the ethanol container and dip the glass
spreader in.
c) Replace the lid onto the ethanol container.
d) Rapidly pass the glass spreader through the Bunsen burner once
and then hold the glass spreader in the zone of sterility to allow
the alcohol to burn off. Prolonged exposure to heat will cause the
glass rod to become deformed or shatter.
e) Allow the glass spreader to cool and then spread the sample
evenly over the entire surface of the agar plate by moving the
spreader backward and forward across the plate while turning the
plate with your other hand. Do not lift the plate from the table
during this process.
f) Re-sterilise the glass spreader.
g) Replace the lid onto the agar plate and allow the sample to dry
onto the agar before inverting the plate for incubation.
10. Return the liquid bacterial mixture to the incubator as soon as possible to
incubate overnight.
11. Place a rubber band around all inoculated agar plates and place in the box
for incubation at 37oC for 48 hours
31
Return to the laboratory the next day (Wednesday) and repeat the streak/spread
plating (steps 8, 9 and 11) of your mixture onto fresh plates (all 3 types – this is your
24 hour conjugation). Remember to clearly differentiate between the two spreading
times. You should have 4 sets of each plate type to observe any changes. Your
mixed liquid culture can be discarded at this point into the yellow waste bags.
Return to the laboratory two days after each streak plate time to count the colonies
on the mixed antibiotic plate (Thursday and Friday), and take note of the colony
morphology, including colour and surface characteristics, on all plates (see Appendix
1 for suggested characteristics to include). Table 4.1 will help you to document the
results:
Figure 4.1 Flowchart for conjugation experiment
| Day 1 Note morphology of the 2 different strains |
| Making broth Suspend 1cfu of IB609 in 1ml LB Suspend 1cfu of Q318/R751 in 1ml LB 0.5ml LB 0.5ml LB |
Streaking/spreadi
ng and combining
Short incubation
Streaking/spreading 10
or 30 minute
conjugation
Day 2: Streak/spread
24 hour incubation
Day 3: Check initial and
10 and 30 minute
conjugation
Day 4: Check 24 hour
conjugation
Streak/spread onto all
3 agars
Combine remaining
broth with remaining
Q318/R751 broth
Streak/spread onto all
3 agars
Incubate for 10 min or
30 min
Streak/spread onto all
3 agars
Return broth to
incubator overnight
Incubate
Count colonies/colony
morphology
Streak/spread onto all
3 agars
Incubate
Count colonies/colony
morphology
33
Table 4.1 Results of your streak and spread plating
| KANAMYCIN PLATES* | STREPTOMYCIN PLATES* | KANAMYCIN + STREPTOMYCIN PLATES |
| Purple colonies# | White Colonies# | |
| IB609 | ||
| Q358/R751 | ||
| 10 MINUTE CONJUGATION |
||
| 30 MINUTE CONJUGATION |
||
| 24 HOUR CONJUGATION |
*Record colony morphology of isolates colonies
# count and record cfu as well as colony morphology
34
Practical 4 assessment
There are two parts to this assessment.
The first is to fill in your table with data and answer the following 5 questions (2.5%).
You can work with your partner on the table but the questions must be answered
independently. Please submit this on learnJCU in quiz 1 -questions. You should
work out your answers first and then copy them in. Once this is submitted you will
have access to part 2 of your assessment.
Part 2 involves you analyzing an example of a discussion for this practical (2.5%). In
your analysis you will identify which parts of the discussion relate to which parts of a
rubric for marking the discussion. This will assist you in learning how to structure and
write a discussion when you have a rubric to work with. Once you have carried out
this analysis on paper and are happy with it, you can submit your results to the
conjugation part 2 submission quiz. This is just an online format of what you will
have done on a hard copy.
Questions.
1. Define the terms donor strain and recipient strain and Identify which of the cultures
you used were the donor and recipient strains, using justifications and reasoning
based on expected results (provided on LJCU). Did conjugation occur in your case
(explain why/why not)?
2. Describe the process by which the vio gene was transferred from the donor to
the recipient.
3. Explain why you streaked both cultures onto three antibiotic-containing agars
at the start of the experiment. Explain how both antibiotics being added together to
LB agar will select for transconjugants.
4. Explain why you think you were asked to spread your mixture after two
different time periods? How could this timed procedure (if you did more time
intervals) help you in the study of the plasmids?
5. From the literature, do you consider conjugation to be the primary method for
transfer of antibiotic resistance between bacteria in nature? If not, which method do
you consider to be more likely, use scientific literature to justify your choice?
Total Assessment (5%) Questions due 29/3 (Tuesday week 6), Analysis due
05/4 (Tuesday week 7)
35
BEFORE LEAVING THE LABORATORY!
PLACE ALL CULTURES TO BE INCUBATED IN THE CONTAINERS
PROVIDED.
ENSURE ALL BOTTLES AND PLATES ARE LABELLED CORRECTLY
AND SEALED APPROPRIATELY.
REMOVE LABELS FROM GLASSWARE AND PLASTICWARE
ALL USED MATERIALS SHOULD BE PLACED IN THE CORRECT
RECEPTACLES, AS DETAILED IN, BIOSAFETY PRECAUTIONS FOR
PRACTICAL CLASSES IN MICROBIOLOGY AND IMMUNOLOGY; “DISPOSAL
OF WASTE MATERIAL”.
RETURN YOUR MICROSCOPE TO ITS PLACE OF ORIGIN.
WIPE DOWN THE BENCH AND CHAIR WITH 70% ETHANOL AND PAPER
TOWEL. DO NOT USE KIMWIPES.
REMOVE YOUR LAB COAT AND PLACE IN BAG FOR CLEANING.
LASTLY WASH YOUR HANDS, PRIOR TO LEAVING THE LABORATORY.
36
Practical 5: Mycology
All culturing sections of this practical session will be completed in pairs,
however all other procedures and observations need to be conducted
individually.
Background Information
Fungi are a large, varied and diverse group of organisms that are distributed
worldwide. They are found in diverse habitats and commonly survive on
organic debris of plant, animal or microorganism origin. Fungi function, with
bacteria and other small organisms, to break down organic debris into simpler
forms consequently recycling essential nutrients. All fungi require preformed
organic substrates as an energy source. Fungal activity is particularly strong
in acid soils where bacterial activity is minimal.
Fungi are capable pathogens. This action is a particularly significant activity
in the plant kingdom where fungi are the most important group of microbial
parasites; and in the animal kingdom, the significance of mycotic diseases is
increasing. Additionally fungi have created issues of poisoning, with fungal
toxins found in items of food for human and animal consumption.
The economic importance of fungi overall is as great as that of bacteria.
Fungal use in industrial applications is substantial, ranging from use within the
food industry to the production of industrial products, alcohol, organic acids,
vitamins and antibiotics.
37
Objective
Part A
To examine three divisions of the kingdom fungi including:
Zygomycota
Ascomycota
Basidiomycota
With the intent of developing skills to identify taxonomic characteristics
Identification information for Part A of this practical can be found at the end of
the practical after the information on writing up your practical report. Where
the instructions say to confirm with figure X, please look up the identification
information.
Part B
To compare two different fungi with respect to growth characteristics in
selected environmental conditions. This is the information that will be written
up as a full practical report.
38
Part A
For the recording of your observations please include in your illustrations: the
Phyla, Genus, the viewing magnification; and where possible label the fungal
structures observed. All observations of demonstration slides will be carried
out using digital images on learnJCU prior to entry into the lab. The same
demonstration slides will be available in the lab for assistance with any
queries only.
Complete all mycology manipulations in a certified Biological Safety Cabinet
(BSC) II.
Terminology: Division(phylum) – order – genus – species (if known)
A1. Zygomycota – Entomophthorales – Entomophthora
Entomophthora muscae is a member of the Zygomycota that rarely produces
zygospores. It more commonly produces a series of sporangia with a single
spores at the tip, pushing out from the cuticle of the infected host. These are
often described as conidiophores bearing a single conidium at the top. The
demonstration slide of Entomophthora shows these fungal fruiting bodies
bursting through the integument of the fly. No sexual stage is present. The
fungal infection coming through the exoskeleton of the insect instigates the
production of packed masses of sporangiospores.
A1.1 Label from the demonstration slide, the conidiophores bearing a single
apical conidium, hyphae inside the host and the host exoskeleton (use figure
6.3 for support).
39
A3 Zygomycota – Mucorales – Rhizopus stolonifer
A3.1 Draw and label from the demonstration slide of Rhizopus stolonifera
(NB 2 images to use here), the sporangium, columnella, colarette and
sporangiospores (asexual stage), suspensors and zygospores (sexual stage)
(use figure 6.4 for support)
A3.2 Within a BSC II, mount some of the mycelium from the prepared
culture of Rhizopus in trypan blue, using a sterile toothpick to
manipulate the specimen.
Tease apart the mycelium and add a coverslip.
Examine your prepared slide, draw relevant images and label any
sexual and asexual structures you observe. Note any any differences
between the slide that you have prepared and the demonstration slide
provided.
Note: Sexual structures may not be present in prepared cultures as sexual
stages often require very particular conditions to develop
A4 Ascomycota – Schizosaccharomycetales – Schizosaccharomyces
octosporus
A4.1 Draw and label from the demonstration slide of
Schizosaccharomyces octosporus. sexual reproductive features: ascus
containing 8 ascospores. This needs to be done at maximum magnification to
see spores. Select an ascus where you can see most or all spores to draw.
(compare with figure 6.5).
40
A5 Ascomycota – Eurotiales – Aspergillus nidulans/flavis
A5.1 There are two images to view (sexual and asexual) Draw and label from
the demonstration slides of Aspergillus nidulans, the cleistothecium
containing multiple asci, each with 8 ascospores (sexual), NB. Hulle cells are
not obvious. Also draw and label the asexual conidiophore, phialides (these
look like fingers that the conidia come off the top of) and conidia. (compare
with figure 6.6a and b).
A5.2 Within a BSCII, Use a sterile toothpick to mount some of the mycelium
from the prepared culture of Aspergillus nidulans/flavis (or A. flavus) in trypan
blue.
Tease apart the mycelium and add a coverslip.
Gently put pressure on the coverslip to release the asci.
Examine your prepared slide, and observe the sexual and asexual structures
and record all of your observations, including noting any differences between
the slide that you have prepared and the demonstration slide provided.
Note: Sexual structures may not be present in prepared cultures as sexual
stages often require very particular conditions to develop.
A6 Ascomycota – Pezizales – Tuber aestivum (black summer truffle)
A6.1 Draw and label from the demonstration slide of Tuber aestivum, the
thin walled asci containing between 1 and 3 spiky ascospores. Also note the
surrounding thick hyphae. Compare with the laminate provided on LearnJCU
and draw your observations.
A7 Basidiomycota – Ustilaginales – Ustilago cynodontis
A7.1 Draw and label from the demonstration slide of Ustilago cynodontis
infecting plant tissue ustilospores alongside plant tissues. (compare with
figure 6.8).
41
A8 Basidiomycota – Agaricales – Chlorophyllum molybdites & Agaricus
bisporus
A8.1 Draw and label from the demonstration slide of Chlorophyllum
molybdites the hyphae and basidiospores
A8.2 Draw and label from the specimens of Agaricus bisporus provided in the
lab the stipe, pileus, gills, and annulus (compare with figure 6.9).
A9 Basidiomycota – Agaricales – Lycoperdon
A9.1 Draw and label from the demonstration slide of Lycoperdon, hyphae
with cavities (lacunae) lined with basidia (compare with figure 6.10).
42
Part B
Environment and Microbes
Biological factors and physico-chemical conditions influence the activity of
microbes in the environment and their general survival. The interaction among
the environment, host and microbe is significant in disease development.
The temperature tolerance range of mesophiles and thermophiles
dramatically illustrates the significance of temperature in microbial activity.
1. Within a BSCII, use a sterile toothpick to inoculate Fusarium onto 4
potato dextrose agar (PDA) plates labelled 5°C, 25°C, 37°C and 45°C
on both base and lid. Note: Use non-sporing fungi if possible.
2. Within a BSCII, use a sterile toothpick to inoculate Humicola insolens
onto 4 PDA plates labelled 5°C, 25°C, 37°C and 45°C. (Figure 6.11).
Figure 6.11 Sub-culturing from a plate cultured with fungi: cut out a piece of
agar with fungi(less than 1cm square). Move the lid off the sub-culturing
plate just enough to place the agar on to edge of the plate, tap down
with the toothpick and replace the lid.
3. Seal the agar plates with parafilm. On the lid, draw around the edges
of the subculture square/s with a fine marker, to assist with the growth
measurements in the following days. Place a line across the lid through the
plug of agar to aid in measuring and place a dot to measure your zero point at
the edge of the agar (see figure 6.12) and incubate the plates at the relevant
temperatures right side up.
4. Measure colony radius on day 2 (Wednesday), day 3 morning and day
3 afternoon(Thursday). Please see Figure 6.12. Make observations regarding
contamination and other issues that might affect reliability of your results.
43
Figure 6.12: Measuring radius of fungal growth
5. Graph these results as per the instructions below.
Process your measurements
1. Plot radius (y axis) vs time (x axis) for the different temperature plates
for Fusarium and Humicola (you should have up to 8 separate lines
and time should be reported in hours to take into account different
return times)
2. Calculate the linear growth rate (change in radius over time in a linear
area of the curve – the gradient of the curve) for each temperature and
species
3. Plot growth rate (y axis) vs temperature (x axis) for each species
4. Answer the following as part of the discussion
a. What temperature provides the best conditions for growth rate
for each species?
b. What relevance does this have to people?
44
Total Assessment (10%) – see LearnJCU list for due dates
Full report of PART B will be graded as per rubric (5%)
Drawings (5%) will be marked based on;
Correct Name (Phyla, genus), total magnification eg 1x, 400x,
All required labels correctly identify structures (any structure that was
requested but not found should be noted below the image as not being
found),
Drawing of correct structures (this is not an art competition, draw what
you see), magnification too low to see appropriate detail of structure
can affect your mark.
BEFORE LEAVING THE LABORATORY!
PLACE ALL CULTURES TO BE INCUBATED IN THE CONTAINERS
PROVIDED.
ENSURE ALL BOTTLES AND PLATES ARE LABELLED
CORRECTLY AND SEALED APPROPRIATELY.
REMOVE LABELS FROM GLASSWARE AND PLASTICWARE
ALL USED MATERIALS SHOULD BE PLACED IN THE CORRECT
RECEPTACLES, AS DETAILED IN, BIOSAFETY PRECAUTIONS FOR
PRACTICAL CLASSES IN MICROBIOLOGY AND IMMUNOLOGY;
“DISPOSAL OF WASTE MATERIAL”.
RETURN YOUR MICROSCOPE TO ITS PLACE OF ORIGIN.
WIPE DOWN THE BENCH WITH 70% ETHANOL AND PAPER
TOWEL. DO NOT USE KIMWIPES.
REMOVE YOUR LAB COAT AND RETURN IT TO THE HOOKS.
LASTLY WASH YOUR HANDS, PRIOR TO LEAVING THE
LABORATORY.
45
Part A: identification information
Division / Phyla: Zygomycota
The organisms present in this division can reproduce sexually by the fusion or
conjugation of 2 compatible hyphae to form gametangia which mature into
zygosporangium (zygospore). This is known as the teleomorphic phase and is
illustrated in Figure 6.1.
Figure 6.1
Zygomycota can also reproduce asexually. This is known as the anamorphic
phase and is illustrated in Figure 6.2. This diagram demonstrates the
anamorph and teleomorph phases of Rhizopus stolonifer. The smaller circles
illustrate the anamorphic phase of compatible strains. The phase moving
through the centre of the diagram is demonstrating the teleomorphic phase
which occurs when the two compatible strains approach and achieve contact
with each other.
Figure 6.2
46
Entomophthorales
Entomophthorales (Figure 6.3) can be parasitic on insects or merely
saprophytic. Several organisms are pathogens of humans and animals such
as Basidiobolus which are typically isolated from toad or frog faeces.
Entomophthora is primarily recognized as a fungal disease of adult Diptera
insects, but may also be saprobic in soil. The fungus develops inside the body
of the insect and proliferates to the point where it pushes the abdominal
segments apart and bursts through causing the fly to appear banded in
appearance.
Figure 6.3: Entomophthorales Figure 6.4: Murorales
Mucorales
Mucorales (Figure 6.4) are typically saprophytic. The asexual spores are
produced in sporangia.
Rhizopus stolonifer is characterised by the presence of stolons and
pigmented rhizoids, and the formation of sporangiophores. Rhizopus
stolonifer colonies are fast growing and form a dense cottony growth that is at
first white becoming grey or yellowish brown with sporulation.
47
Division / Phyla: Ascomycota
Ascomycota form sexual non-motile spores, ascospores, in structures
resembling sacs identified as asci (or singularly as an ascus). Ascospores are
formed within the ascus by an enveloping membrane system packages each
nucleus with its adjacent cytoplasm and provides the site for ascospore wall
formation.
1. Endomycetales
Endomycetales is a faction of fungi which possess ascocarps; numerous
yeasts belong in this group. Endomycetales fungi possess a zygote or a single
cell, developing directly into an ascus. The asci occur unprotected in the
suspending medium.
Schizosaccharomyces octosporus (Figure 6.5) is a saprophytic yeast. Cell
division in the somatic cells is by a form of budding in which the fertile apex of
the mother cell grows marginally before the next daughter cell is blown out.
The fertile tip develops annellations. Schizosaccharomyces octosporus can
be differentiated from other Schizosaccharomyces species by its eight-spored
asci.
Figure 6.5: Schizosaccharomyces octosporus
48
2. Eurotiales
Eurotiales fungi contain the sexual spores in completely closed ascocarps,
identified as cleistothecia. The asci are globose or broadly oval and are
scattered at various levels within the ascocarp (Cleistothecia), rather than
gathered in a hymenium. Eurotiales members possess distinctive asexual
structures.
Aspergillus nidulans/flavis (Figure 6.6) colonies are typically plain green in
colour with dark red-brown cleisothecia developing within and upon the
conidial layer. The conidial heads are short and columnar. Aspergillus
nidulans/flavis possess thick-walled hülle cells; which often surround the
cleisothecia.
Figure 6.6a: Aspergillus nidulans sexual structures
49
Figure 6.6b: Aspergillus nidulans asexual structures
Pezizales
The order Pezizales is characterized by asci that generally open by rupturing
to form a terminal or eccentric lid or operculum. The ascomata are apothecia
or are closed structures of various forms that are derived from apothecia.
Note the transition from open to closed in the various genera (Figure 6.7).
Apothecia range in size from less than a millimeter to 15cm and may be
sessile or stalked. This order includes both the truffles and morels. The
truffles (family Tuberaceae, genus Tuber) produce closed, solid ascomata
which do not liberate spores.
http://www.mycolog.com/CHAP4b.htm
Figure 6.7: Tuberaceae sp.
50
Division/Phyla: Basidiomycota
The most conspicuous and familiar Basidiomycota are those that produce
mushrooms, which are sexual reproductive structures. The Basidiomycota
also includes yeasts (single-celled forms) and asexual species.
Spores of the Basidiomycota fungi are basidiospores and are located upon
club-shaped end cells identified as basidia that typically bear external
meiospores (usually 4). The basidium can develop free and unconnected or
attached to hymenium. Sexual spores may or may not be held in a complex
fruiting body termed a basidiocarp or basidiomata.
Ustomycetes
The Ustomycetes species are composed entirely of fungi that are parasitic on
flowering plants producing sori containing dark spores identified as
ustilospore. The Ustomycetes are commonly identified as the smuts due to
the characteristic black, powdery teliospores that give the infected plants a
dirty, soot-like appearance. Two spore stages, the teliospore and
basidiospores are produced. Basidiocarps are not produced. Additional spore
stages are absent.
Ustilago cynodontis (Figure 6.8) is a couch grass smut, a loose smut, with no
protective covering present over the sorus (fruiting structure). The ovary
tissue of the plant is invaded and displaced by fungal spores. The spores
germinate to provide a germ tube, termed a metabasidium and meiosis
occurs. The metabasidium becomes septated and often branches irregularly.
Sporidia bud from the cells.
Figure 6.8: Ustilago cynodontis
51
Agaricales
The members of this order have basidia arranged in a relatively well-defined
fertile layer or hymenium.
Agaricus bisporus (Figure 6.9) has different appearances at different stages of
maturity and is a common edible mushroom with different common names
depending on colour (white/brown) and maturity
Figure 6.9 Agaricus bisporus (a) mature mushroom; (b) immature mushroom
with a closed cap; (c) longitudinal slice of immature mushroom. (Zhang et. al.
2018)
Lycoperdon (Figure 6.10) is a typical puff ball organism. The gleba (sporing
tissue in an angiocarpous fruiting body) at maturity is powdery. In the
immature structure, cavities (lacunae) develop in the gleba and basidia line
these cavities. Lycoperdon produce within the ball vast numbers of dust-like
spores mingled with elastic threads. When the ball is compressed, the rind or
peridium bursts at the summit to form a single mouth, and the elastic threads
cause the spores to fly out in puffs like smoke.
52
Figure 6.10: Lycoperdon
53
Appendix 1: Macroscopic Examination of Bacteria
Colonial Morphologies
Colonial morphology is occasionally sufficiently characteristic to provide diagnostic
value. The growth medium, growth conditions, visible pigmentation, age and
diameter of colonies should be recorded in conjunction with the colony description.
Consistency
1. Butyrous (the consistency of butter)
2. Viscous (can be noticed by the growth trailing from the loop when withdrawn
from the plate)
3. Membranous (markedly thin)
4. Brittle (dry/friable)
Optical Characteristics
1. Opaque (not permitting light to pass through the growth)
2. Translucent (some light passes through the growth)
3. Opalescent (opal/milky like in appearance)
4. Iridescent (exhibiting changing colours in reflected light, similar to a rainbow)
5. Dull (not glossy)
6. Glistening (glossy)
7. Rough
8. Smooth
9. Wrinkled
10. Dry
Whole Colony Characteristics.
1. Punctiform
2. Circular
3. Rhizoid
4. Irregular
5. Filamentous
Colony Edge Characteristics
1. Entire
2. Undulate
3. Lobate
4. Filamentous
5. Curled
Elevation Characteristics
1. Flat
2. Raised
3. Convex
4. Pulvinate
5. Umbonate
54
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