Certificate III in Laboratory Skills
MSL30118 BDV6
Certificate IV in Laboratory Techniques
MSL40118 BDV3
Microscopic & aseptic techniques cluster
Training Manual
Laboratory Operations
© North Metropolitan TAFE 2020
This publication is copyright to North Metropolitan TAFE. Apart from fair dealing for the
purpose of private study, research, criticism or review, as permitted under the Copyright
Act 1968 no part may be reproduced without written permission.
Whilst every precaution has been taken to supply complete and accurate information,
North Metropolitan TAFE assumes no responsibility for any liability, loss or damage
caused or alleged to be caused directly or indirectly by the instructions contained in or
accompanying this publication.
Cluster
MSL973019 Perform microscopic examinations
MSL973016 Perform aseptic techniques
Laboratory Operations
Version
02/2020
Author(s)
Elizabeth Fitzgerald, Chris Williams
Microscopic examinations & aseptic techniques
Contents
1. Occupational Safety and Health (OSH) ............................................................................................. 5
2. Hand Washing Instructions .............................................................................................................. 6
3. Procedures to follow in a biological spill: ......................................................................................... 7
4. Auto pipette Operation .................................................................................................................... 8
5. Receiving, handling and storing samples – Microscopic Techniques ............................................. 12
6. The Microscope .............................................................................................................................. 13
7. Use of Oil Immersion ...................................................................................................................... 17
8. Measuring with the microscope: Micrometry................................................................................ 18
9. Temporary Slides, Simple Staining ................................................................................................. 23
10. Structure and function of cells, micrometry .................................................................................. 25
11. Construction materials testing ....................................................................................................... 28
12. Cell function: Water movement in and out of cells ........................................................................ 30
13. Microenumeration the Haemocytometer...................................................................................... 32
14. Temporary Mounts: Darkfield and Phase Contrast Microscopy ..................................................... 36
15. Serial dilution pipetting practice .................................................................................................... 39
16. Percentage viability of yeast cells .................................................................................................. 41
17. Aseptic Techniques......................................................................................................................... 43
18. Transfer materials aseptically ......................................................................................................... 48
19. Sterilization..................................................................................................................................... 50
20. Receiving, handling and storing samples– Aseptic Techniques ...................................................... 53
21. Throat swab.................................................................................................................................... 54
22. Aseptic Pipetting ............................................................................................................................ 56
23. Producing Confluent Plates ............................................................................................................ 57
24. Gram Stain ...................................................................................................................................... 59
25. Preparation of Microbiological Media ............................................................................................ 60
26. Micro-organisms in the Environment ............................................................................................ 62
27. Food processing-Quality Control Test ............................................................................................ 65
28. Bacterial Cultures ........................................................................................................................... 69
29. Producing Purity Plates from Mixed Cultures................................................................................. 71
30. Culturing Fungi ............................................................................................................................... 72
31. Membrane Filtration ...................................................................................................................... 76
32. The effect of UV light on bacterial growth. .................................................................................... 79
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Training Manual Record
Page Yes No
Auto pipette Operation
8
Receiving, handling and storing samples – Microscopic
Techniques
12
The Microscope
13
Use of Oil Immersion
17
Measuring with the microscope: Micrometry
18
Temporary Slides, Simple Staining
23
Structure and function of cells, micrometry
25
Construction materials testing
28
Cell function: Water movement in and out of cells
30
Micro-enumeration the Haemocytometer
32
Temporary Mounts: Dark Field and Phase Contrast Microscopy
36
Serial dilution pipetting practice
39
Percentage viability of yeast cells
41
Transfer materials aseptically
48
Sterilization
50
Receiving, handling and storing samples– Aseptic Techniques
53
Throat swab
54
Aseptic Pipetting
56
Producing Confluent Plates
57
Gram Stain
59
Preparation of Microbiological Media
60
Micro-organisms in the Environment
62
Food processing-Quality Control Test
65
Bacterial Cultures
69
Producing Purity Plates from Mixed Cultures
71
Culturing Fungi
72
Membrane Filtration
76
The effect of UV light on bacterial growth.
79
Appendix A: Central Laboratories Specimen Receipt
81
By observation the student satisfactorily completed the following this manual
Comments
Yes
No
Lecturer Sign & Date
Student Name
ID
© North Metropolitan TAFE 2020
Signature
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Microscopic examinations & aseptic techniques
Occupational Safety and Health (OSH)
It is very important to understand that you are now training for industry where you will be
exposed to a variety of hazards. In order to be trained effectively, you will need to handle
hazardous substances and carry out potentially hazardous procedures while in the TAFE
laboratory. Obviously, the TAFE staff members take care for your safety, but you will be
exposed to higher risks than you perhaps faced at school or in other workplaces.
Consequently, the Science team have put in place a series of protocols which you must
follow. Failure to follow these guidelines may result in termination of your study, just as
breaches of safety rules in industry can result in dismissal. In this biologically-orientated
unit, the major hazards include;

handling of infectious materials

handling chemicals which may be volatile, poisonous or carcinogenic

danger of cutting yourself.
Our basic methods of preventing OSH incidents in this unit involve:
Wearing PPE, basically wearing a Lab coat, long trousers, covered non-slip shoes (made of a
non-permeable substance), eye protection and in some cases protective gloves. It is your
responsibility to provide yourself with everything except the gloves, i.e. make sure you come
to TAFE suitably dressed and with your PPE and long hair tied back. You will not be allowed
to enter a lab unless you are suitably attired and you will not be allowed to carry out a
practical without appropriate PPE. You may be excluded from the TAFE lab if you are not
suitably attired and you will not be allowed to carry out a practical without appropriate PPE.
It is part of the elements of competency of this unit that you cannot gain competency in
this unit unless you are always compliant with these protocols.
Fume cupboards and Biological Safety Cabinets (Safety Level II) are to be used to protect you
from being splashed by or inhaling poisonous substances or infectious material.
The requirement for sensible behaviour in the laboratory, absolutely no eating and drinking
in the laboratory (don't even chew your pens and certainly don't put your fingers in your
mouth or eyes) and always wash your hands before leaving the laboratory. Keep your bag in
a safe place to prevent tripping accidents.
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Hand Washing Instructions
The skin is your hands first defence against infection from pathogenic organisms. Whilst it’s
intact, it’s impermeable to organisms such as HIV and hepatitis so its care and hygiene are
crucial.
As organisms acquired during a session of laboratory work are in the category of surface
contaminants (as distinct from the less accessible resident skin flora), they may be removed
by washing the hands thoroughly whenever leaving the laboratory. Anti-bacterial skin
cleansers are not essential for routine use, but there are several reasons for not using bar
soaps (e.g., can act as reservoirs of contamination).
Hand washing is covered by Australian Standard AS/NZS 4815:2001. The recommended
procedure is as follows:
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Procedures to follow in a biological spill:
1. Pour disinfectant solution: 70% isopropyl alcohol, 10% Lysol, or household bleach on all
broken glass and contaminated surfaces. Extend coverage over 10cm around original
(contaminated) area.
2. Cover the spill area with paper towels. Add additional disinfectant to fully saturate
them. Wait 30 minutes.
3. Wearing rubber gloves and using tongs, pick up all glass or residue along with paper
towels and place in Bio-hazard bag.
4. Disinfect the area again by following steps 1–3.
5. Seal the Bio-hazard bag and autoclave the contents. Dispose.
6. Wash hands thoroughly.
I have read and understand the safety information in this manual
Date
Overview of conducted by:
(Name of Lecturer )
Student Name
Student ID
Student signature
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Microscopic examinations & aseptic techniques
Auto pipette Operation
Automatic pipettes are used in laboratories to dispense volumes from 1L to more than
5000L (5mL). There are different makes and models available. Some are a set volume only
while others are adjustable for a range of volumes. If the auto pipette is calibrated and used
correctly it will dispense accurate volumes of liquid. Two different methods are used in this
exercise
Element of Competency: 1, 2, 6
Pipetting Techniques
Practice dispensing various volumes of water with traditional pipettes, eg plastic disposable,
pasteur, glass bulb type, and glass graduated, as demonstrated.
The first step in auto-pipetting is to choose the pipetting mode best suited to the type of
work. These pipetting modes are forward, reverse, and repetitive. Pipetting whole blood
also requires a special technique.
The pipetting sequence consists of three actions: aspirate, to draw up the sample; dispense,
to deliver the sample; and blow-out, to empty the tip completely.
Figure: Parts of the automatic pipette
Forward pipetting
Forward pipetting is the standard technique for pipetting aqueous liquids.
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Microscopic examinations & aseptic techniques


Press the operating button to the first stop.
Dip the tip into the solution to a depth of 1 cm, and slowly release the button. Withdraw the tip
from the liquid, touching it against the edge of the vessel to remove excess liquid.

Dispense the liquid into the receiving vessel by gently pressing the operating button to the first
stop. After one second, press the button down to the second stop. This action will empty the
tip. Remove the tip from the vessel, sliding it along the wall of the vessel.

Release the operating button to the ready position.
Reverse pipetting
The reverse technique is used for pipetting solutions with a high viscosity or a tendency to
foam. This method is also recommended for dispensing small volumes. It is only possible
with air displacement pipettes.

Press the operating button to the second stop.

Dip the tip into the solution to a depth of 1 cm, and slowly release the button. This action will
fill the tip. Withdraw the tip from the liquid, touching it against the edge of the vessel to
remove excess liquid.

Dispense the liquid into the receiving vessel by pressing the button gently and steadily down to
the first stop. Hold the button in this position. Some liquid will remain in the tip, and this should
not be dispensed.

The liquid remaining in the tip can be pipetted back into the original solution or thrown away
with the tip

Release the operating button to the ready position
Materials
Auto pipettes (a range of volumes) & tips, small beakers, distilled water,
analytical balance
Method
Forward Pipetting: Follow the directions for ‘forward’ pipetting.
1. Set the volume on a pipette to 100L or 200L or 500L.
2.
Place a clean empty beaker on the pan of the analytical balance and tare it (zero).
Remove the beaker from the balance.
3.
Using the auto pipette dispense distilled water of your chosen volume, (e.g. 100L)
into the tared beaker. Replace the beaker on the balance and note the weight of the
water. Record the weight to 3 decimal places.
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4. Tare the beaker again. [Note: the water already in the beaker does not need removing
as the 'tare' function allows for contents.]
5. Repeat this procedure another 7 times, recording the weight each time. Tare the
beaker between each addition.
Reverse Pipetting: Follow the directions for ‘Reverse’ pipetting.
1.
Set the volume on a pipette to 100L or 200L or 500L.
2. Place a clean empty beaker on the pan of the analytical balance and tare it (zero).
Remove the beaker from the balance.
3. using the auto pipette, dispense distilled water of your chosen volume, (e.g. 100L) into
the tared beaker. Replace the beaker on the balance and note the weight of the water.
4. Tare the beaker again. [Note: the water already in the beaker does not need removing
as the 'tare' function allows for contents.]
5. Repeat this procedure another 7 times, recording the weight each time. Tare the
beaker between each addition.
Results:
1.
Tabulate the readings recorded.
Pipette serial number
Forward pipetting
Auto pipette set at
0
Temperature of water
C
Reverse Pipetting
L
Auto pipette set at
L
Weight of distilled water (g)
1
2
3
4
5
6
7
8
TOTAL
Average
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2. Calculate the % error (inaccuracy) for both methods.
% error =
Calculations
3. Comment on the accuracy and reproducibility of your technique with the auto pipette.
4. Why might one method (forward or reverse) be used in preference to another?
5. List some of the ways that incorrect volumes may be dispensed.
6. What safety feature(s) does the auto pipette have to prevent the handling of
contaminated tips?
__________________________________________________________________________________________
__________________________________________________________________________________________
__________________________________________________________________________________________
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Receiving, handling and storing samples – Microscopic Techniques
Introduction
The quality of the specimen has an effect on the tests that are performed and their results.
In all routine laboratories, the way in which specimens are collected and transported to the
laboratory will vary. On receipt of the specimen, many laboratories have a central area
where the specimens are receipted, checked and distributed to the appropriate
department.
Many different types of specimens are received in a routine laboratory, and it is necessary
to observe certain details to ensure an accurate report with the minimum of delay. Most
laboratories have written procedures for staff to follow.
Materials
Examples of laboratory request forms, Examples of specimens /samples
Method
Study as many of the completed examples of request forms & specimens /samples as you
can. Record details in table below. Correctly fill out the example of a lab form and specimen
label.
Results
1. Correctly complete the specimen label and request form. Your lecturer will give these
to you.
2. Study the forms and samples provided. List any errors on the forms and tubes and state
what action should be taken.
Request Form ID
Error
Action
Competency Record: By observation the student satisfactorily completed the following Section/s
Yes No
Comments
Auto pipette Operation
Receiving, handling and storing samples – Microscopic
Techniques
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Lecturers signature:
Date:
The Microscope
Element of Competency: 2, 4, 6
Introduction
The compound light microscope is used widely in biological laboratories and it is essential to
know how to set up and use a microscope correctly.
In this exercise, you will become familiar with microscope components, their functions, and
learn how to set up a microscope for optimal viewing of specimens.
Do not rush this practical. You will make extensive use of the microscope in this course and
you must take the time to learn to use it properly. Practice setting up your microscope
properly, using both eyes to look down both oculars, finding the right plane of field and
focusing on a slide. If you do not get these basics right now, you will struggle through the
rest of the course. These skills are essential in a laboratory and you will use them over again
in several other units in your studies,
Materials
Nikon microscope, stage micrometres, slides & cover slips, small transparent plastic ruler,
prepared stained slide of plant tissue, Lens cleaner, Lens tissue, Immersion oil, prepared
slides for observation
Examine the objective lenses. You will find that they are colour-coded with a ring of colour.
List the colour against the magnification of the lenses in the table above.
Observe that the ocular lenses have x10 magnification.
You can calculate the total magnification of the image you will see down the microscope
when using each of the objective lenses by multiplying the ocular magnification by the
objective magnification. Do this now and enter your results in your table.
Ocular
Magnification
Objective
Magnification /colour code
Total Magnification
10x
10x
10x
10x
Focusing an Image & Moving the Slide and Image
Before switching on your microscope, ensure the light control is turned down. The light
globe is more likely to blow if you light it up on high immediately. This also prevents eye
damage from looking at too strong a light source.
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Put the slide you have on the stage, and swing the x4 objective into position.
Use the coarse focus knob to bring the letter 'e' into focus. Remember to use both eyes! You can
adjust the oculars to suit your own eyes, by moving them apart or together to suit the width of
your face.
Look at the 'e' on your stage, and then look at the 'e' down your microscope.
Draw the 'e' image you see. How is the image different from a regular 'e'?
Stage
Microscope
You should see that the image is both laterally and vertically inverted
Move your slide from left to right and up and down. How does the image move in respect to
how the slide is moving on the stage? In which direction does the image move?
See that the image moves in the opposite direction to what you are doing to the stage.
Using the microscope stage is a bit like reversing a car. You move the slide in the opposite
direction to how you want the image to move. This might take a bit of practice to get used
to!
For example, if I wanted to move the black dot into the middle of the field of view, I would
need to move the image up. To do this I would move the slide on the stage down!
Look at the distance between the x4 objective lens and the slide while the image is in focus. Estimate
the distance in centimetres. Record your observation in the table below.
Objective
Working Distance (mm)
4x
10x
40x
100x
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Swing the x10 objective lens into position using the nose piece,
Your microscope is fairly sophisticated, as it is both par-focal and par-central. This means
that when you change from a low to a higher magnification (e.g. from x10 to x40
objective), you won't have to make much adjustment to bring the slide into focus.
Look at your 'e' image, and you should find that it is nearly in focus. Refocus your 'e'
image using the fine focus only.
Estimate the distance between the x10 objective lens and the slide. Note that it is
much closer than when you were using the x4 objective.
Swing your microscope now from x10 to x 40 objective lens.
Notice that this lens is almost sitting on the slide - estimate this distance in millimetres.
If you look at the slide it should not be far from being in focus, although it will be difficult
to focus the newspaper image now on such a high magnification.
Do not use the coarse focus on this lens as you may ram the objective through the slide,
this can damage the rather expensive objective lens and obviously the slide. To focus you
need only use the fine adjustment knob.
Always focus a specimen on low power (x4 or x10 objective) before using the higher
power objectives. It will stop you from damaging the high-power objectives, and it is also
much easier to find the plane of field on a lower magnification.
Swing the x4 or x10 objective into place before removing your slide, so that you
don't damage an objective lens that is too close.
You can now return your slide to the tray provided.
Adjusting the brightness and contrast
Your lecturer will provide you with a stained slide for you to look at under the microscope.
Set up your microscope as before to focus on the prepared stained slide using the x10
objective.
Describe what happens as you slide the light control from low to high intensity.
_________________________________________________________________________________________
_________________________________________________________________________________________
Describe what happens as you move the condenser lens up and down.
If you now change to x40 objective, is the microscope still set up optimally or do you
need to adjust it?
It is best to have the light only as bright as you need it. If it is too bright not only will it
'wash the image out' (you won't see the stain colours properly), but it may also damage
your eyes.
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After you have adjusted the light intensity, then you should adjust the condenser until
the image is as sharp as you can get it. In general, stained slides are best observed with
the condenser up high (close to the stage), and wet mounts (unstained) are seen in
better contrast when the condenser is dropped down.
Try to remember to check these settings (light intensity and condenser) whenever you
use the microscope. Failure to do these checks is the commonest cause of students
complaining about image quality!
Replace slides in correct position in boxes as directed by your lecturer. If you have time,
you might like to look at more stained slides, to practice focusing some more (and to see
some other interesting things!)
Use the Nikon Microscope YS100 instruction manual for swift setup.
Things to Remember:
Before turning the microscope on, check that the light intensity is on low. Also turn the
light down before switching the microscope is turned off and replacing the cover. Always
focus on a low power objective first. If you have trouble focusing on low power, it will be
even harder on high power, and you risk damaging your slide or objective lens.
If you lose focus when changing from a lower power magnification to a higher power, just
drop back to the low power objective and re-focus. This will be quicker in the long run
than persisting with focusing on a higher magnification. Swing the objective to a lower
magnification before removing slides. The microscope is to be kept clean, so that images
can be best focused. Use a blower brush, lens tissue and lens cleaning fluid if necessary.
Make sure there is no immersion oil residue left anywhere.
IMPORTANT IF IN DOUBT - ASK FOR HELP.
Before placing the microscope in the cupboard, check the following:  the light is switched off on the microscope
 the mains cord is wrapped carefully around the microscope neck
 the slide is removed from the stage
 the low power objective or the blank is in the light path
 lenses are wiped clean with lens tissue (especially the oil immersion)
 the stage is in its lowest position, and dry
 the mechanical stage is centred
 the dust cover is in place
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Use of Oil Immersion
Element of Competency: 1, 2, 4, 5
Introduction
Most microscopes have an objective lens that is used with oil or another type of liquid
between the lens and the slide. In this exercise, you will learn how to use the oil immersion
lens.
Materials
Nikon microscope, Prepared Slides (ants, fungi, and plant cells), Lens cleaner Kim wipes
Immersion oil
Method
1. Select a slide and set up your microscope following the instructions on page 12 in
the Nikon microscope manual.
2. Select an area on the slide that is relatively thin and looks interesting place this area in
the centre of the field of view. Observe your area under both low power and high
power dry objectives to make sure it will focus well.
3. Set up for oil immersion by following the instructions from your lecturer also you can
refer to pages 22-23 of the Nikon instruction manual
Results
4. Draw labelled diagrams of your specimen under x 40 objective high power (using the dry
objective). Make sure you record the details of your sample e.g. sample name etc.
5. Draw labelled diagrams and write notes about the same specimen observed under oil
immersion, and compare the images. Use the areas below to record your drawings
Figure 1: Image at 400X magnification no oil
Sample details:
© North Metropolitan TAFE 2020
Figure 2: Image at 1000x magnification oil
Sample details:
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Microscopic examinations & aseptic techniques
Measuring with the microscope: Micrometry
Element of Competency 1, 2, 3, 4, 5, 6
Introduction
The millimetre is not a very suitable unit for microscopic measurement, most cells are only a
very small fraction of a millimetre in size, and for instance a red blood cell is approximately
0.007mm in size. This makes for unwieldy expression of size, so the preferred international
unit for microscopic measurement is the micrometre, written ‘µm’.
Note: 1000m = 1mm.
The size of a cell can be estimated if the diameter of the field of view is known. Using the
stage micrometre measure the diameter of field of view (FOV) for each objective. Record
your results in table below.
Objective
Diameter of FOV (mm)
Diameter of FOV (m)
4x
10x
40x
100x
This estimation isn't very accurate so an ocular micrometre is used.
Measurements of cells are made using a scale called an ocular micrometre. This is inserted
into the ocular lens of the microscope.
The spacing or distance between the lines on the ocular micrometre is arbitrary, i.e. it has
no units of measurement, so the exact distance between lines of graduations on the ocular
micrometre must be determined by calibration against a stage micrometre which has units
of measurement, for each objective lens.
Micrometre slides may have etched lines exactly 0.1mm (100m) apart or 0.01mm (10m)
apart. This information will be marked on the slide.
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Ocular micrometer
Ocular micrometer superimposed on
slide micrometer
Materials
Microscope, Ocular micrometer, Stage micrometer, Slides with cells suitable for measuring
Method
1. Place the stage micrometer on the stage beneath the low power objective. Use the
stage micrometer to measure the diameter of the field of view for the three dry
objective lenses (i.e. 4x, 10x and 40x). Fill in the results table
2. If not already fitted, place the ocular micrometer in the eyepiece. This may be done by
unscrewing the lens and inserting the ocular micrometer. The exact location of the
ocular insertion depends on the microscope model. As demonstrated
3. Superimpose the ocular micrometer lines over those of the stage micrometer by
adjusting the focus, and mechanical stage. Using the 4x objective, get the scales into
clear focus and determine how many ocular graduations are equivalent to one stage
micrometer division/unit. Note that this will be either 10µm or 100µm depending on
the stage micrometer used.
4. Repeat this for each of the objective lenses used and fill in the table provided. Keep a
record of these measurements for use in later exercises.
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5. Now measure selected cells in the slides provided to determine the average diameter
of 10 or more cells.

s=
number of divisions on stage micrometer

e=
number of ocular divisions (eye piece unit, epu)

c=
length of each stage micrometer division
6. e.g. 100 ocular divisions or eye piece division/unit (epu)

each ocular division =
=
? Stage divisions
s/e x c mm
Example:
Objective
lens
Number of Length of stage Number of eye
stage
micrometer
piece
micrometer division (mm) division/unit
divisions (s)
(c)
(epu) (e)
4x
1
Or
Objective Number of
lens
stage
micrometer
divisions (s)
4x
25
0.1mm
Calculation
4
One individual
ocular division
(1 epu x1000)
= µm
0.025 x 1000
=25µm
Length of
stage
micrometer
division (mm)
(c)
Number of eye Calculation
piece
division/unit
(epu) (e)
One individual
ocular division (1
epu x1000) =
µm
0.1mm
100
0.025 x 1000 =
25µm
Notes
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Results
1.
Calibrate the ocular micrometer for your microscope using the 0.lmm
stage micrometer. Record your results in table below
Make
Objective
lens
Model
Number of
stage
micrometer
divisions (s)
Length of stage
micrometer
division (mm)
(c)
Number of eye
piece
division/unit
(epu) (e)
Calculation
One individual
ocular division
(1 epu x1000)
=µm
2. Measure ten cells of the same type with the 40x objective. In the table below
record the diameter of the cells and determine the average size. Show all
calculations.
Cells
1
Example Calculation:
Using 40X objective: epu value x number of epu across the cell
= size of cell
25 µm x 10 =250 µm
2
3
4
5
6
7
8
9
10
Average
3. Explain why the ocular micrometer must be calibrated.
____________________________________________________________________________________
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Competency Record: By observation the student satisfactorily completed the following Section/s
Yes
No
Comments
The Microscope
Oil Immersion
Measuring with the microscope:
Micrometry
Lecturers signature:
Date:
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Temporary Slides, Simple Staining
Elements of Competency: 1, 2, 3, 4, 5, 6, UPK
Introduction
The functions and activities of cells (both plant and animal) are the distinguishing features
between living and non-living things. Cells can be seen under the light microscope but
detailed structure can only be observed using the electron microscope.
Temporary slides are prepared, viewed and then discarded afterwards. They are usually
rapidly prepared for a specific purpose and they are not intended to last for more than a
short time. They may be used for counts or to check if the material is suitable prior to
lengthy processing and the preparation of permanent slides.
Prepare the following slides as given in the method below. For each slide prepared, focus on
the specimen with LP then with HP. Draw and label a simple diagram of the cells or
structures and add a measurement to the diagram (use µm). If you have not already
calibrated an eyepiece graticule it will be necessary to do this first.
Example:
Materials
1% methylene blue stain, 1% neutral red stain, Saline solution, Microscopes Slides & Cover
slips, Tooth picks, labels,
Method
Cheek smear
1. Make two slides of cheek cells (as demonstrated). Label the slides; add stain and then the
cells.
2. Stain one slide with methylene blue and the other with neutral red.
3. Draw a fully labelled diagram of cells from each of the slides. Use table below.
4. Comment on the two staining techniques. Which stain gives a clearer image of the cells?
Notes
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Epithelial cells
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Structure and function of cells, micrometry
Materials
Saline solution, Microscopes Slides & Cover slips, Tooth picks, labels, Tomatoes, Pears, Iodine
solution, Aniline sulphate, Slides & cover slips, Gem blades, needles, labels, 1% Toluidine blue
Epidermal cells of onion
1. Place a drop of water on a clean slide.
2. Peel one layer of ‘skin’ from the onion. Carefully place the layer in the drop of water,
keeping the tissue flat. Add a cover slip.
3. Add iodine by the irrigation method, as explained.
4. Draw and measure the cells using 100x or 400x magnification
Parenchyma cells
1. Remove a small amount of the tomato pulp (tissue immediately below the skin) and place
in a drop of water on a slide. Tease out the tissue to separate the cells. Place a cover slip
on top.
2. Irrigate with iodine solution. Take care not to get this on top of the cover slip.
3. Observe under both low and high power on a microscope and make a fully labelled
diagram of three parenchyma cells.
4. Draw and measure the cells using 100x or 400x total magnification.
Sclerenchyma cells
1. Remove a small amount of the white tissue of a pear and place in a drop of water on a
slide. Squash the tissue thoroughly to separate the cells.
2. Remove excess water and add one or two drops of aniline sulphate. TAKE CARE as this
solution is corrosive.
3. Observe under the microscope and make a fully labelled diagram of sclerenchyma
sclereids (stone cells) and surrounding parenchyma cells.
4. Draw and measure the cells using 400 x magnifications
Results
1. Well-labelled diagrams with a scale should be recorded for each of the preparations
examined and compare the staining techniques Scales should be rounded up values e.g.
100m not odd numbers or fractions. The size of the diagram, the scale, and the actual
measurement should all match. Also state where they are found and their function.
Epidermal cells of onion
Where is it found, and what is its function?
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Parenchyma cells
Where is it found, and what is its function?
Sclerenchyma cells
Where is it found, and what is its function?
1.
Staining technique comparison
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2. Record the correct method for labelling slides
3. How difficult was it to see the unstained onion and tomato cells on the slide? What
effect is achieved by adding stains?
4. For how long would you expect the preparation mounted in water to last? Explain.
____________________________________________________________________________
____________________________________________________________________________
____________________________________________________________________________
____________________________________________________________________________
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Construction materials testing
Elements of Competency 1, 2, 3, 4, 5, 6, UPK
The supply of river sand is running out and quarries are accessing alternative sources of sand for use
in concrete mixes in construction. The sand should not be an aggregate that is likely to break down
into smaller particles. A technician in a quarry company is required to analyse samples of crushed
rock using a light microscope. The technician looks for characteristics of the sample, such as grain
size, angularities, roundness, sharpness, cracks, presence of organic matter, mineral structure and
whether the particles are a conglomerate. If the sample does not meet the characteristics, the
company will need to treat it to make it suitable for use in concrete mixes (for example by washing,
crushing and sieving).
Conglomerate-rock comprising pieces of other rock, geology coarse-grained sedimentary rock
containing fragments of other rock larger than 2 mm/0.08 in. in diameter, held together with
another material such as clay
In this exercise you will use methods to analyse the components of a sand sample and record
measurements.
Materials
Assorted sand samples, cello tape, ocular micrometres, balance, labels, weigh boats, spatulas, slides &
cover slips
Method
1.
Using one of the purified sand samples, accurately weigh 0.5 g into a white plastic
weigh boat.
2. Sprinkle some sand on a piece of clear cellotape; stick the tape to a microscope slide,
check the shape and size of the sand grains.
3. Compare the particles in your sand samples with the Sand Chart (provided). Record
your observations in the table provided
4. From the same slide select and measure the different sizes of sand grains - use your
calibrated ocular micrometre. Measure at least 5 of the smaller size and 5 of the
larger size sand grains to get a range of sizes. Fill in the table in the results section.
5. Repeat the above for a different sand sample
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Results: Sample 1
Sample ID
1.
Shape (roundness, angular)
Sizes of sand grains.
Colour
Sorting
Magnification_______________________
Sample ID
Size range (µm)
1
2
3
Average
4
5
Small
Large
Results: Sample 2
Sample ID
Shape (roundness, angular)
2. Sizes of sand grains.
Sample ID
Colour
Sorting
Magnification ________________________
Size range (µm)
1
2
Average
3
4
5
Small
Large
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Cell function: Water movement in and out of cells
Elements of Competency: 1, 2, 3, 4, 5, 6, UPK
Introduction
The functions and activities of cells (both plant and animal) are the distinguishing features
between living and non-living things. The effects of placing plant cells in different
concentrations of saline solution can be observed under the light microscope.
The results can be explained in terms of water movement in and out of cells
Materials
Spirogyra or onion, distilled water, 0.8% saline solution, 5.0% or 8% saline solution,
microscopes, slides & cover slips, watch glasses, forceps, labels
Method
1.
Place 2 small strands of Spirogyra or onion epidermis peel into 0.8% saline solution
for 2 minutes
2.
Remove the strand/onion peel and mount in 0.8% saline solution on a slide. Ensure
that it is flat and not folded.
3.
Cover with a cover slip
4.
Remove excess saline solution with a paper towel, ensuring that the saline solution
does not come into contact with the microscope components.
5.
Observe under the microscope using x40 and x100
6.
Draw a cell, labelling the cell wall, cell membrane and cell contents
7.
Repeat the procedure for 5% or 8% saline solution and distilled water.
Notes:
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Results
1.
Use the following table to record the effect on the plant cell when placed in 0.8%
saline, 5% or 8% saline and distilled water. Include the size of cell & nucleus
Solution used
Diagram of observation of plant cell
Explanation of your observations
Competency Record: By observation the student satisfactorily completed the following Section/s
Yes
No
Comments
Temporary Slides, Simple Staining
Construction materials testing
Cell function: Water movement
in and out of cells
Lecturers signature:
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Microenumeration : the Haemocytometer
Elements of Competency 1, 2, 3,4,5,6
Introduction
For counting smaller microscopic particles a specially engraved microscope slide is used. It
was originally designed to count numbers of red and white blood cells, hence the name
haemocytometer. Most blood counts are carried out with automatic counters but the
haemocytometer is still used for a variety of manual counts. The types of things which may
be counted include yeast cells, algal cells and monoclonal antibodies.
Materials
Microscope, Haemocytometer, algae culture, yeast culture, capillary tubes, automatic
pipette & tips, formalin 10%, test tubes &racks
Method
1.
Set up your microscope for optimal illumination. Examine the haemocytometer
under the microscope with the 4x objective and locate the grid on the central
platform. Use the 40x objective for counting the sample. Practice moving
around the grid systematically.
2.
Using an ordinary slide, make a wet mount preparation of the algal culture provided
to check the shape and size of the cells and a suitable concentration for counting if
there are too many cells your count is subject to error.
3.
If necessary, dilute the algal sample carefully, using carefully measured volumes. Mix
the cells well prior to removing a sample. Immobilise motile cells if necessary by
adding a drop of 10% formalin to the diluted sample. This must be done in the fume
hood. Record the dilution factor (if used) either 1 in 2 or 1 in 10.
4.
Load a sample into the haemocytometer as demonstrated. Make sure there are no
air bubbles and that the space under the cover slip is evenly filled without overflow.
Handle the slide with care, scratches make counting difficult.
5.
Examine the slide under low power and check that the cells are distributed evenly
over the counting area. If the cells are clumped or poorly distributed, clean the slide
(use only lens tissue and alcohol) and load it again.
6.
Count all the cells in the 25 small squares of the central large square. Use the correct
counting convention to avoid ‘double’ counting.
7.
Calculate the number of cells per mm3 and convert to cells per ml (cm3).
8.
After use, clean the slide with water and rinse it with alcohol. Wipe dry with lens
tissue and store in a box to prevent scratching the grid.
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Calculation
Let N equal the number of cells counted in the centre mm 2.
The depth of the chamber is 0.1mm.
The volume of fluid over the centre mm 2 is therefore
1mm x 1mm x 0.1mm = 0.1 mm3 (0.1 l).
If there are N cells present in 0.1mm3 of solution then 1mm 3 contains N x 10 cells.
The dilution factor must also be accounted for. If 1ml of algal culture had 1 ml of diluent
added then multiply the final number of cells by 2.
To convert cells per l to cells per ml, multiply by 1000.
1.
Results for algae sample
Enter your own results. This diagram shows the 25 squares in the central mm 2
Calculations:
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2. Results for yeast sample
Enter your own results. This diagram shows the 25 squares in the central mm 2
Calculations:
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Discussion
1.
List the major sources of error in:

sample preparation

loading the chamber

counting the cells under the microscope

Estimating the final number of cells.
2. Which of the errors are inherent?
3. Which of the errors are technical and may be overcome with practice and care?
If a student diluted a sample 1 in 3 (1ml of sample and 2 ml of diluent) and counted an
average of 232 cells in the centre mm2, what is the cell count per

ml of culture?

100 ml of culture?

litre of culture?
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Temporary Mounts: Dark field and Phase Contrast Microscopy
Sometimes biologists want to examine live specimens that are difficult to see without
staining the specimens. Although there are a few vital stains available they are not always
suitable for the purpose and with the methods described here it is possible to see colourless
live specimens without staining. However your microscope does need a couple of
attachments. Darkfield is relatively cheap and simple, requiring only a 'stop' to be inserted
in the light path. Phase contrast requires a special lens and a special matching attachment
on the condenser.
The Nikon site, www.microscopvu.com/ has a lot of information and tutorials on this topic.
Go to the site and then navigate to the relevant parts. Much of the information is quite
technical, but there is a fair amount of introductory information and lots of illustrations.
Darkfield
As a result of placing a 'stop' in the light path, all the direct rays are cut out and only light
reflected or scattered from the surface of the specimen passes through the microscope, the
background remains dark, and the specimen is sharply outlined.
Phase contrast
This technique is a little more complex and shows much more detail of the internal structure
of cells than darkfield can. It exploits the fact that cells, organelles and the media they are
mounted in have different refractive indices (refraction causes light to bend at different
angles to the normal as light passes between media). You need to understand that light is
said to have a waveform and that the light passing through a specimen consists of many
waves and that the organelles in a cell retard any waves passing through them by
approximately 1/4 of a wavelength, so now you have 'out of phase' waves passing through
the microscope. If you can further retard the waves which have passed through the
organelles by another 1/4 of a wavelength (by using a phase plate) they will be completely
out of phase with the waves which don't pass through the specimen and in fact these waves
will cancel each other out resulting in full interference and making some parts of the cell
much darker than others. Nikon site www.microscopvu.com
Temporary Mounts (simple and unstained), and Dark Field and Phase Contrast Microscopy
We are going to make some basic unstained slides of cells that we can view under the
microscope. We can look at them using light microscopy, dark field and phase contrast
microscopy to see which is the best method for observing these cells. Because these slides
are unstained, the cells are still 'alive' (most staining kills the cells, although it makes them
easier to see). This means the motile cells will be moving around for us to observe.
Materials
Glass slides and cover slips, sterile physiological saline (0.85% NaCI), plastic pipettes,
toothpicks, onion, knife and chopping board, gem blade, forceps, dissecting needle,
Protozoa cultures - Paramecium and Amoebae
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Wet Prep Slide Procedure
Make your slide, and then make microscopic observations on that slide before making your
next slide. Prepared slides dry out quickly, so there is no point making more than one at a
time.
Cheek Cell Prep:
Place a few drops of saline on a clean slide. Remember to always check your slides are clean
before use, even if they are brand new. Take a toothpick and gently scrape some cheek cells
off. Immerse the cells in the saline on the slide. Dispose of the toothpick immediately in the
discard jar.
Gently lower a cover slip over the preparation excluding air bubbles.
Onion Cell Prep:
Cut up the onion to expose the layers. Find the membranous layer between the thicker,
fleshy layers - this is the best one to make your slide from. Slice a small portion of the thin
layer off and place on your slide, being careful not to fold it over. Place a drop or two of
saline onto your onion slide, and lower a cover slip over it.
Protozoa Cell Prep:
Get a clean slide ready, and label it with the name of the protozoa you are going to put on it,
Take a pipette and suck up a small amount of the protozoa culture. Put one or two drops of
the culture onto your slide. Cover with a cover slip.
Bright Field Microscopy
Observe your first slide using light microscopy as you have been doing so far; starting on a
low power objective and going up to x40 objective. Draw a diagram of the cells you see, and
label it with any cellular components that are visible (e.g. cell membrane, nucleus, and
cytoplasm),
Now we will look at the slide using darkfield and phase contrast microscopy, and see if it is
any easier to see the cells and their cellular components.
Dark Field Microscopy
Check that your microscope has the correct condenser assembly attached, it should look like a
largish wheel in the place of the normal condenser, but different models of microscopes may
have different appearances. Ask your supervisor if you are not sure. Focus your slide as
normal on low power {x4 or x10 objective).Now rotate the wheel on the condenser until a
letter 'D' appears in the window (as opposed to the '0' which should have been there when
you started). Some microscope models also require a green filter to be placed over the light
source. Examine the image and refocus the condenser until you get the best image. Try to see
if you can get the darkfield effect to work on high power (x400).Diagrammatically represent
your observations in your Prac Book.
Phase Contrast
Set up your slide and focus it as best you can on bright field illumination on low power (x100),
Rotate the condenser wheel until '10' appears (or 'Ph1', depending on your microscope
model). When you move to x400, change the wheel to '40' (see picture above) or 'Ph2'.
•
Make your observations as before.
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•
Clean up and dispose of your slide safely in the sharps container.
•
Repeat the process with the other slides to prepare.
Observations & Discussion
Which method did you find was the best to view these unstained cells?
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Serial dilution pipetting practice
Element of Competency: 1, 2, 6
Serial dilutions are regularly used in microbiology when, for instance, initial concentrations
of bacteria are orders of magnitude too high to perform a plate count, or for producing a
series of regular dilutions as in titrating serum. It has two advantages:

It allows for rapid achievement of a very high dilution factor.

It requires a relatively small volume of diluent.
Material
Test tubes &racks, 1ml autopipette & tips, vortex mixer, cuvettes, marking pen, 20ml
distilled water per student, 0.5 % methylene blue, and spectrophotometer
Method
1. Turn on spectrophotometer to warm up. Set up your work bench with required
equipment.
2. Label the test tube with the original solution of methylene blue "1x." pipette 5ml into this
tube. Make sure you also put your initials on the tubes.
3. Set up four more empty dilution tubes in a test tube rack. Label the tubes 10-1, 10-2, 10-3,
10-4 to indicate the relative concentration of dye which they will contain.
4. Aliquot 9 ml of dH2O into each of these four labelled dilution tubes, using an
autopipettte.
5. Using a clean tip, transfer 1ml of the original ‘1x methylene blue’ solution from tube
#1 into tube #10-1, and mix well. (Vortex or use pipette to mix as demonstrated)
6. After mixing, use a clean tip to withdraw 1ml from tube #10-1, add it to #10-2. Mix as
before.
7. Using a clean tip, withdraw 1ml from tube #10-2; add it to #10-3. Mix as before.
8. Using a clean tip, withdraw 1ml from tube #10-3; add it to #10-4. Mix as before.
9 When dilutions are done, 9mL should remain in tubes #10-1, #10-2 and #10-3. Tube
#10-4 should contain 10mL.
10. Also prepare a blank tube containing only distilled water
READ AND PLOT THE % ABSORBANCE OF THE DILUTION SERIES
11. Read the A609 of each dilution against a blank of distilled water.
12. Begin with tube #10-4, and work your way up. In this way, you need not wash the
cuvette each time, or change the tip, but touch off the last drop before adding the
next dilution.
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Record your results below
Dilution
Absorbance
13. Plot a graph with the relative concentration (dilution) of methylene blue (indicated by
the tube label) as the X axis and % absorbance at 609 nm as the Y axis. Use the blank
tube (zero methylene blue with an A609 = 0.000) as your first (zero) point. Staple your
graph onto this page
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Percentage viability of yeast cells
Introduction
Yeasts are used in industries such as bread making (to promote 'rising' of the dough) and in
wine production (to ensure the right type of fermentation). It is sometimes necessary to
establish the percentage of yeast cells which are viable - that is they are able to grow and
reproduce.
Using a stain which is taken up by either the living cells (vital stain) or the dead ones makes it
easy to count the cells and establish viability.
Materials
Yeast suspension, 0.1% Rhodamine B stain, Microscopes, Auto pipettes and tips (200 -1000µL),
Small vials, Haemocytometer
Method
In this exercise, Rhodamine B, a red stain which is taken up by the dead cells ('red means
dead") is used with a yeast sample and the number of stained and non-stained cells are
counted.
1. Gently mix your original yeast suspension and take a small subsample (2ml) into a vial,
add 1 one drop of stain.
2. Stand for one minute. Set up your microscope.
3. Load the haemocytometer
4. Focus on the cells using the 4 x objective, then move to the 10x objective and change to
40x count the number of living cells and the number of dead cells in a total of greater
than 100 cells (i.e. make sure you count about 120 cells in total).
5. Repeat your count in the other chamber. Calculate the % of viable cells in the sample.
Results
Slide number
Number of Live Cells Number of Dead Cells Total
1
120 cells
2
120 cells
Average Number of Live Cells
Average Number of Dead Cells
Average Total Number of Cells
120
Percentage of Average Live Cells (%
viability)
Average no. live cells x 100
= Average total no. of cells
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Discussion
1. What is your % viability for the sample? Alternatively you can count the number of
dead cells, and live cells per ml.
2. What is the purpose of counting more than 1 slide preparation for the same sample?
3. List two causes of error in this practical. Comment on whether they are random or
systematic errors.
4. List two (2) quality control procedures used in this practical that aim at reducing the
above errors
Competency Record: By observation the student satisfactorily completed the following Section/s
Yes
No
Comments
Temporary Mounts: Dark field and
Phase Contrast Microscopy
Serial dilution pipetting practice
Percentage viability of yeast
cells
Lecturers signature:
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Date:
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Aseptic Techniques
Transfer materials aseptically
Aseptic transfer of bacteria involves a number of steps that must be carried out without
contaminating the specimen.
General Method for Aseptic transfer of bacteria from a broth to an agar plate:
1.
Assemble all equipment. Work near your Bunsen burner.
2.
Flame the loop including the shaft, as shown below in A. Using your left hand (or if
you are left-handed, use your right hand), hold the tube at an angle so
contamination is minimised.
3.
Remove the top with the little finger of your right hand while holding the loop with
the curved palm of the same hand (if you are left-handed reverse the process).
4.
Flame the neck of the tube, as shown below in B
5.
Carefully remove a sample of culture with your cooled loop (that is, a loopful), as
shown below in C.
6.
Flame the neck of the tube again.
7.
Bring the tube towards the plug. This keeps your loopful steady. Replace the cap (see
Figure C) and put the tube back in the rack.
8.
Place a loopful of culture onto an area of the plate (see Figure D) as the initial
inoculum.
9.
Flame your loop to sterilise it.
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Streak plate method
Streak plate: the specimen is streaked onto the plate with a sterile loop.
The purpose of streaking plates is to distribute the inoculum over the surface of the medium
in such a manner as to separate the bacterial cells from each other. Done properly, this will
result in well-isolated colonies on at least a portion of the plate.
1.
Using the plate provided, label the base with appropriate details.
2.
Hold the loop with a pencil grip and sterilize in the Bunsen flame. Allow to cool.
3.
Pick up a loopful of your inoculum from either a broth or an agar culture, (remove a
small amount of culture).
4.
Pick up the base of the Petri plate to be inoculated with the left hand.
5.
Hold the plate at about 45° to the horizontal (as in Figure 1) and spread the
specimen (see Figure 2) over the primary inoculation site (1) and streak as in (2).
Return the base of the plate to the lid.
6.
Flame the loop. Allow to cool.
7.
Again, pick up the base of the plate and streak as in (3) and (4) of the diagram,
flaming the loop between each change in direction.
8.
Return the plate to the lid, flame the loop and return it to the block.
9.
Place the plate in the correct incubator box in the inverted position.
Figure 1: Holding the plate and the
loop for streaking
Figure 2: Streak pattern
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Tube to tube transfer (slope to slope)
Slope; the sample is streaked onto a slope or slant agar medium in a tube set at an angle.
This increases the surface area for the culture.
1.
Label the slope to be inoculated with appropriate details.
2.
Place the slope to be inoculated, the slope of culture and the block with loop in front
of the Bunsen and convenient to the right hand (if you are right-handed, or to the left
hand if you are left-handed).
3.
Loosen the caps of both slope bottles.
4.
Sterilize the loop by heating it in the Bunsen flame to red heat, starting from the
hand end and drawing the wire through the flame towards the top of the loop.
5.
Allow to cool.
6.
Still holding the loop in your right hand, remove the cap from the slope of culture
with the little finger of this hand.
7.
Keep the bottle as nearly horizontal as possible and flame the neck of the bottle.
8.
With the cooled loop, remove a small amount of the culture.
9.
Flame the neck of the bottle.
10. Replace cap and put the slope of culture back into test tube rack.
11. Now remove the cap of the NA slope, flaming the neck.
12. Inoculate the slope by drawing the loop over the surface of the agar in a snake like
pattern starting at the bottom of the slope.
13. Re-flame the neck of the slope bottle before replacing the cap.
14. Re-flame the loop as in before returning it to the block.
15. Place your inoculated slope in the appropriate incubation box.
Pour plate method
1.
Using the plate provided, label the base with appropriate details
2.
Loosen the cap/ cotton wool plug of the bottle containing the inoculum.
3.
Aseptically attach the sterile tip to your pipette and hold in your right hand.
4.
Lift the bottle/ test tube containing the inoculum with your left hand.
5.
Remove the cap/ cotton wool plug with the little finger of your right hand.
6.
Flame the bottle/ test tube neck.
7.
Put the pipette into the bottle/ test tube and draw up the required volume (1ml) of
the culture.
8.
Remove the pipette and flame the neck of the bottle/ test tube again. Replace the
cap/ cotton wool plug.
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9.
Place the bottle/ test tube on the bench or in its rack.
10. Inoculating the Petri dish
11. Lift the lid of the Petri dish slightly with your right hand and insert the pipette into
the Petri dish. Gently release the required volume of inoculum onto the centre of
the dish. Replace the lid.
12. Put the pipette tip into a discard bin.
13. Collect a bottle of sterile molten agar from the water bath
14. Hold the bottle in your right hand. Remove the cap with the little finger of your left
hand.
15. Flame the neck of the bottle.
16. Lift the lid of the Petri dish slightly with the left hand and pour the sterile molten
agar into the Petri dish. Replace the lid.
17. Flame the neck of the bottle and replace the cap.
18. Move the dish gently to mix the culture and the medium thoroughly and to ensure
that the medium covers the plate evenly move the dish in a figure of eight, or rotate
it until the medium and inoculum are well-mixed and cover the base of the dish.
19. Allow the plate to solidify.
20. Tape the plate closed and incubate in an inverted position.
Spread plate method
1.
Using the plate provided, label the base with appropriate details
2.
Obtain appropriate broth culture.
3.
Mix the broth culture.
4.
Aseptically put the pipette tip into the bottle/ test tube and draw up the required
volume (100 µl) of the culture.
5.
Open the lid of the sterile nutrient agar plate at an angle, like a clam shell.
6.
Carefully deliver the inoculum in the centre of the plate.
7.
Close the lid of the plate.
8.
Dispose of the pipette tip in a biohazard bag.
9.
Obtain a glass spreader and immerse the short end in the alcohol.
10. Flame the glass rod in the Bunsen burner to evaporate the alcohol and sterilize the
spreader.
11. Let the spreader cool 20 seconds.
12. Open the lid of the inoculated dish at an angle and evenly spread the inoculum over
the surface of the media.
13. Rotate the plate and spread again.
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14. Close the lid and let the inoculum soak into the agar for 5 minutes.
15. Immerse the glass spreader in the alcohol and flame to sterilize.
16. Once the inoculum has soaked into the agar, tape the lid shut and turn the plate
upside down.
17. Tape all plates together and label with the initials of someone in the group.
Clean-up Procedure
1.
Place the plates in the tray for incubation upside down to prevent condensation that
may form on the lid from dropping down onto the surface of the media.
2.
Make sure all used pipette tips are placed in a biohazard bag
3.
Return the Bunsen burner, marking pen, alcohol dish, glass spreader, inoculating
loop, test tube rack, and original cultures to the appropriate place.
4.
Spray and wipe the table top with disinfectant.
5.
Remove laboratory coat & gloves
6.
Wash your hands
Source: http://classes.midlandstech.edu/carterp/courses/bio225/chap06/lecture5.htm
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Labelling of plates





The base should be labelled, not the lid
Use a fine-tipped waterproof marker
Written label around the perimeter of the plate
Include any barcode or sticky labels of Lab ID number
Labels to include : The agar medium, Date, Incubation Temperature, Name of specimen /
inoculum, Your initials, any special conditions eg anaerobic, dilution
Transfer materials aseptically
Elements of competency1, 2, 3, 4,
In this experiment, you will review the techniques required for successful aseptic transfer of
microorganisms
Materials
Bunsen burner, inoculating loops, marker pens, Hockey sticks, Flaming alcohol, Nutrient agar
plates, Nutrient broth, Auto pipette and tips (sterile) assorted sizes, Sabouraud's agar, Streak
plate culture of Saccharomyces sp., Broth culture of S.epidermidis, & Bacillus subtilis, Streak
plate culture of S.epidermidis, & Bacillus subtilis
Method
Subcultures: streak, spread plates, tube to tube transfer inoculation
NOTE: use careful aseptic technique for all methods; always label containers PRIOR to
performing the subculture. Keep work area free of clutter and have all equipment on hand
before commencing work.
1.
Label the base of the SAB agar plate. As demonstrated, prepare a streak plate using
Saccharomyces sp. Incubate culture at 25°C for 24 to 48 hours.
2. Repeat the streak in step 1 but use S.epidermidis culture and nutrient agar
3. Use 100 l of the same broth culture to complete a spread plate on nutrient agar
4. Transfer 100 l Bacillus subtilis broth into sterile nutrient broth
5. Incubate S.epidermidis, Bacillus subtilis cultures at 37°C for 24 to 48 hours.
Results
1. Examine the streak plates are the colonies well separated in the last three sections of
the plate? If not, what can you do to improve your technique next time?
2. Examine all the broth cultures for the presence of growth, which is indicated by
turbidity in the broth. Record your observations in the table below.
3. Examine the cultures for the possible presence of a contaminant that may be indicated
by the presence of growth that exhibits a different pattern. Record your observations in
table.
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4.
Record your observations in the table
Culture
Growth (+)or(-)
Contamination (+) or (-) How
do you know?
Culture
Growth (+)or(-)
Contamination (+) or (-) How
do you know?
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Sterilization
Elements of competency1, 2, 3, UPK
To illustrate the effectiveness of an autoclave
Introduction
One method of sterilization is the steam autoclave, which accomplishes the sterilization
process by a combination of high temperature, pressure and time.
Materials (work in pairs)
Working autoclaves, labels, Unsterilized Nutrient broth, 4 x McCartney bottles per group,
Bunsen burner, heat sensitive tape, spore strips/chemical strips, racks, Auto pipette and tips
(sterile) assorted sizes measuring cylinder. 2x Nutrient agar plates per group (part 2)
Procedure
Part 1
1.
2.
3.
4.
5.
6.
7.
Transfer 10ml of nutrient broth into each McCartney bottle
Screw the lid tightly on one McCartney bottle, label autoclaved closed (AC).
Leave the lid loose on the remainder, label autoclaved open (AO).
Screw the lid tightly on one McCartney bottle, label not autoclaved closed (NAC).
Leave the lid loose on the remainder, label not autoclaved open (NAO).
Make sure you also include your group ID on the labels
Autoclave both the open and closed bottles at 121oC for 15 minutes, your lecturer will
put some autoclave tape onto the racks and place a chemical indicator strip in the
autoclave
8. Place the ‘not autoclaved’ bottles into the class tray as directed.
9. This practical will be completed next class
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Part 2
1. Check the steam strips and record result. Strip result
2. Examine all your tubes for signs of turbidity. Record your observation in the table
below. Accurately record your observed changes in tubes.
Tubes
Observations
McCartney bottle,
autoclaved closed (AC)
McCartney bottle,
autoclaved open (AO).
McCartney bottle, not
autoclaved closed (NAC)
McCartney bottle, not
autoclaved open (NAO).
3. Divide the NA plates in two. Label nutrient agar plates as directed and streak a
sample from the previously prepared tubes onto the plates. Incubate 37oC, 24hrs.
These will be kept for next class. Record results in the table below
Observations
Treatment
Observations
Nutrient agar streaked with
Autoclaved closed (AC)
Nutrient agar streaked with
Autoclaved open (AO).
Nutrient agar streaked with not
autoclaved closed (NAC)
Nutrient agar streaked with, not
autoclaved open (NAO).
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Discussion
1. Is steam penetration important? Explain
_____________________________________________________________________________
_____________________________________________________________________________
_____________________________________________________________________________
2. How can the temperature in the autoclave go above boiling temperature of 100 0C?
3. List the biohazard waste containers available in the lab. Why are they different?
4. Could you use the contaminated nutrient broth if you resterilised it? Explain your
answer.
5. Describe how the contaminated reusable samples are disposed of.
Notes
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Receiving, handling and storing samples– Aseptic Techniques
Introduction
The quality of the specimen has an effect on the tests that are performed and their
results. In all routine laboratories, the way in which specimens are collected and
transported to the laboratory will vary. On receipt of the specimen, many laboratories
have a central area where the specimens are receipted, checked and distributed to the
appropriate department.
Many different types of specimens are received in a routine laboratory, and it is
necessary to observe certain details to ensure an accurate report with the minimum of
delay. Most laboratories have written procedures for staff to follow.
Materials
Examples of laboratory request forms, Examples of specimens /samples
Method
Study as many of the completed examples of request forms & specimens /samples as you
can. Record details in table below. Correctly fill out the example of a lab form and
specimen label.
Results
1. Correctly complete the specimen label and request form. Your lecturer will give
these to you.
2. Study the forms and samples provided. List any errors on the forms and tubes and
state what action should be taken.
Request Form ID
Error
Action
Competency Record: By observation the student satisfactorily completed the following Section/s
Yes No Comments
Transfer materials aseptically
Sterilisation
Receiving, handling and storing samples – Microscopic
Techniques
Lecturers signature:
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Throat swab
We are going to now process a sample and perform microscopy and culture on it. This is a
simulation of how a microbiology laboratory works. We will be reporting on a swab, which is
to detect if a patient's has an infection. If you go on to study Microbiology further (e.g. in the
Diploma course), you will learn about how the pathogen is Identified and how to treat the
infection.
Materials
Throat swabs with matching request forms, as provided, Sample log sheets, Lab accession
number stickers, SAB agar plates, Inoculating loops, slides& coverslips crystal violet or
methylene blue, grams iodine. Staining racks, slide dryer.
Procedure
Method for performing a streak plate & slide using a clinical swab
1. Collect specimen, check details and record on specimen record sheet.
2. If correct, attach a laboratory label to the form and to the lid of the swab. (Check with
your lecturer).
3. Correctly label the agar plate provided (you can use one of the patient labels for some of
this). Also label a clean glass slide.
4. Carefully remove the swab from the holding tube. Take care; don’t touch anything with
the end of the swab.
5. Inoculate the agar plate by gently rolling the swab over the agar (see diagram below).
Then roll the swab onto the center of the glass slide. Carefully return the swab to the tube
and put aside (do not discard the swab; it is to be returned at the end of the class).
6. Sterilise by incineration, an inoculating loop. Cool and then streak from the initial
inoculation area across the plate (see below).
 After this streak, the inoculating loop is sterilized again, cooled again and the
 Inoculum for the second sector is obtained from the first sector.
 The same steps are followed for the subsequent streaks.
 This is followed by incubation of the plates under favourable conditions.
 After incubation, i.e. generally 24h, isolated colonies appear.
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Microscopy
1.
Heat-fix the smear by placing it in the slide dryer for a minute @ 50oC to fix the
cells onto the slide. Do not overheat the slide as it will distort the cells.
2.
Cover the smear with methylene blue or crystal violet and allow the dye to remain
in the smear for approximately one minute (Staining time is not critical here;
somewhere between 30 seconds to 2 minutes should give you an acceptable stain,
the longer you leave the dye in it, the darker will be the stain).
3.
Using distilled water wash bottle, gently wash off the excess stain from the slide by
directing a gentle stream of water over the surface of the slide.
4.
Wash off any stain that got on the bottom of the slide as well.
5.
Saturate the smear again but this time with Iodine. Iodine will set the stain
6.
Wash of any excess iodine with gently running tap water. Rinse thoroughly.
7.
Wipe the back of the slide and blot the stained surface with bibulous paper or with
a paper towel.
8.
Place the stained smear on the microscope stage smear side up and focus the
smear using the 10X objective.
9.
Choose an area of the smear in which the cells are well spread in a monolayer.
Centre the area to be studied, apply immersion oil directly to the smear, and focus
the smear under oil with the 100X objective.
Record your results on the specimen form
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Aseptic Pipetting
Elements of competency1, 2, 3, 4, UPK
Materials: Auto pipettes a range of volumes & sterile tips, sterile broth, Bunsen burner, sterile
container
Procedure
1. Choose the appropriate pipette and sterile tips for the volume required.
2. Do not open the pipette box until you are ready to use the tip
3. Ensure that the auto pipette is clean before use. Some auto pipettes are able to be
sterilised
4. It should be obvious that you NEVER touch the tips because they go into samples, cultures
or sterile solutions
5. Carefully mix the sample as demonstrated
6. Take of the top of tube or bottle or flask with your little finger of the pipetting hand.
 This is a little tricky at first, but practice, practice, practice. If the cap must be screwed
off, then catch the cap in the little finger and twist the container.
 NEVER set the top of the container down. However if this is difficult for you place the
cap upwards near the Bunsen burner this will minimize unwanted material falling into
the cap.
 Also keep the container held at an angle to minimize unwanted material fall into tube.
7. F lame the lip of the container.
 Most of the time airborne contaminants will have landed near the mouth of the
container. The flame will incinerate them. The flaming will also heat the walls of the
container so that contaminants that have settled on the inside will also likely be killed.
However, be careful! Too intense a heating might crack a glass container or melt a
plastic one.
8. Open the lid of the tip box and attach the tip.
 Close the tip box once you have attached the tip. Take care not to touch anything with
the tip. Draw up the required volume from the container, PLEASE practice pipetting
every chance you can.
9. Flame the lip of the container before closing.
 Replace the lid by pushing the container into the cap that is still being held by the little
finger, or by twisting the container into a screw-on cap. But be careful! The lip of the
container is hot, and if your aim is faulty you might burn yourself
10. Deliver the sample to the appropriate container & appropriately discard the tip
11. Incubate the transferred broth 37oC, 24-48 hrs
Record your observations next class
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Producing Confluent Plates
Elements of competency1, 2, 3, 4, UPK
Introduction
In this practical, you are going to learn how to produce a basic type of plate, the 'confluent' or
spread plate. The idea is to produce a continuous 'lawn' of bacteria {of one species) on the
surface of the plate. Such a lawn can then be used in antibiotic and disinfectant testing or
simply to quickly grow large quantities of bacteria.
A lawn inoculum is achieved by transferring a small amount of liquid culture onto on to the
surface of an agar plate and spreading it our using a glass 'hockey stick'. The culture is
transferred using a pipette with a sterile tip.
Materials
Auto pipettes a range of volumes & sterile tips, Bunsen burner, tips, nutrient agar plates,
broth culture of suitable bacterium, e.g. E. coli, glass 'hockey' sticks, flaming alcohol
Procedure
1. Label the agar plates, your name, date and culture used.
2. Sterilise the hockey stick by dipping it into the alcohol and then flaming. Keep them well
away from the alcohol while flaming.
3. Set the transfer-pipettes to dispense 500 µl (This is a reasonable amount for producing a
lawn).
4. Aseptically attach a tip to the pipette.
5. Carefully remove 500 µl of culture and dispense onto the surface of your plate.
6. Discard tip into biohazard container
7. Flame the hockey stick, allow to cool, then use it to spread the culture across the
surface of the plate.
8. Re-flame the hockey stick to sterilise it.
9. Incubate your plate in the normal way (agar on top) for 24-48 hours. Record your
observations.
Observations & discussion
Record your nutrient broth and plate results. Is there any contamination several days after
transfer? Record the reasons for contamination. Did you achieve confluent growth?
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Urine Sample
Materials
Urine samples with matching request forms, as provided, Sample log sheets, Lab accession
number stickers, NA plates, Inoculating loops, hockey stick, slides& coverslips, Gram staining
sequence staining racks, slide dryer. Centrifuge, Centrifuge tubes, Slides & cover slips,
Timers, Forceps, pipettes
Method for performing a spread plate & slide using a clinical sample
1. Collect specimen, check details and record on specimen record sheet.
2. If correct, attach a laboratory label to the form and to the urine sample. (Check with your
lecturer).
3. Correctly label the agar plate provided (you can use one of the patient labels for some of
this). Also label a clean glass slide.
4. Follow the procedure for producing confluent growth, however you do not have to flame
the neck of the sample container. Use 1ml of the urine sample
Cell smears from urine samples
Method
1. Collect all urine from a fully voided sample (this is important as most cells are in the
first and last parts of the full volume).
2. Gently mix the sample and divide into a number of centrifuge tubes.
3. Centrifuge at 2500rpm for 5 minutes.
4. Remove the tubes carefully from the centrifuge and check that you can see a
precipitate (the ‘cell button’).
5. Very carefully pour off the supernatant into a jar of hypochlorite and the ‘button’
should remain intact.
6. Re-suspend the ‘button’ into a few drops of supernatant by flicking the side of the
tube with your finger.
7. Sterilise by incineration, an inoculating loop. Cool and obtain a loopful of the
concentrated urine spread this carefully onto a glass slide
8. Allow the slide to air dry
9. Heat-fix the smear by placing it in the slide dryer for a minute @ 50oC to fix the cells
onto the slide. Do not overheat the slide as it will distort the cells.
10. Follow the staining instructions for the gram staining technique below
Record your results on the specimen form
Competency Record: By observation the student satisfactorily completed the following Section/s
Yes
No
Comments
Throat swab
Aseptic Pipetting
Producing Confluent Plates
Urine Sample
Lecturers signature:
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Gram Stain
The Gram stain is the most commonly used stain in the microbiology lab. It is used to
differentiate between two basic types of bacteria, and is dependent on the cell wall
components. In the Gram stain, crystal violet dye is used to stain the bacterial cells purple.
The cells are then treated with iodine to help bonding between the stain and the cell wall.
Acetone is used to wash away crystal violet that has not bonded to a cell wall.
Gram positive cell walls have a thick peptidoglycan layer that is cross-linked, and this
prevents the crystal violet stain from being washed away. The bonding of the stain is aided
by the iodine. Gram negative cell walls have only a thin layer of peptidoglycan layer without
cross-linking, so the crystal violet is washed out as it doesn't bond with the cell wall.
After acetone treatment Gram, positive bacteria will be stained purple, and Gram-negative
bacteria will be unstained. To make it easier to see the Gram-negative cells, the slide is
counter-stained with Safranin, so that the Gram-negative bacteria are stained pink.
Materials
Culture plates of bacteria agar plates you cultured in previous practicals loops Glass slides
sterile, physiological saline (0.85% NaCI) Gram’s staining chemicals : 1 % Crystal violet,
Gram's iodine, Acetone, 1% Safranin
Procedure
Label a slide with an ID for the bacterium, your name, 'Gram stain', and today's date. Place a
loopful of saline onto your slide.
Pick up some colony using a sterilised loop, and emulsify it into the saline. A small number of
bacteria goes a long way! You do not need to use a lot of bacteria (one colony will be more
than enough), or there will be too many cells on top of each other for you to be able see
their shapes easily. Allow the saline to dry out, and heat-fix your slide.
1. flood the slide with crystal violet and leave for 30 seconds. Rinse off this stain with
water
2. then add iodine to the slide. Leave for 30 seconds. Rinse again
3. then use acetone to decolourise the slide. This should occur in less than 10 seconds,
and you will see purple stain wash away.
4. Quickly wash the acetone off so as not to over-decolourise the slide
5. Counterstain the slide with Safranin for 30 seconds. Rinse.
6. Gently blot your slide dry before observing under the microscope.
Observe the cells using oil immersion technique. Go through the normal focusing procedure,
but after you have focused with the x40 objective, swing the objective out of the way and add
a drop of immersion oil between the lens and slide. Swing the x100 objective into position,
and refocus using the fine adjustment knob only. Your cells will be either bacilli (rods) or cocci
(spheres), and be stained Gram positive (dark purple) or Gram negative (pink). Be careful to
not get oil on the other objective lenses.
Clean off the oil from the x100 objective using lens tissue with lens cleaning fluid. Dry with
clean lens tissue.
Dispose of your slides into the sharps container.
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Preparation of Microbiological Media
Elements of competency1, 2, 3, UPK
Materials (per group)
Working autoclaves, labels, distilled water, Bunsen burner, heat sensitive tape, spore
strips/chemical strips, autoclave bottles, pH strips / meter, measuring cylinders,
stirrer/hotplate,1M NaOH, & 1M HCL, balance, nutrient agar powder, Sterile Petri plates,
culture tubes
Introduction
In order to obtain bacteria in pure cultures they must be supplied with artificial media
containing all their growth requirements. They must also be incubated under suitable
conditions of temperature and atmosphere.
Microbiological laboratories may make up their own media or purchase them from media
supply companies. Most media are available in dehydrated form and merely require rehydrating, sterilising and dispensing.
Procedure
1.
Turn on hot plate to get it heated.
2.
Measure distilled water into a beaker and place onto hot plate. Carefully lower a
magnetic stirrer into the beaker and centre it on the hot plate.
3.
Weigh out then dissolve the nutrient broth powder and agar powder into the beaker
of distilled water. Leave the beaker on the hot plate until the powders has fully
dissolved, letting the mixture boil approx. 1min.
4.
Transfer the mixture into an autoclave bottle. Leave the lid loose, and place
autoclave tape on the bottle. Sterilise by autoclaving.
5.
Once sterile and let to slightly cool, pour about 15mL of agar aseptically into sterile
petri dishes in the biosafety cabinet, remember to pour on a flat surface, rotating
the dish to get an even spread of the agar, but not to form bubbles. Make as many
plates as to use the rest of the agar.
6.
Leave the agar to set with the lid slightly covering it to prevent aerial contamination,
but not fully covering it so as not to form condensation.
7.
Once the agar is set, label each plate with "NA" and the date of production. The
plates can be stored in the refrigerator until use.
8.
Label one plate and place it in the incubator overnight as a quality control - if
nothing grows, then the batch is not contaminated. If contamination is shown, the
whole batch of agar is not to be used.
Keep all media for use in future classes.
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Observations
1.
What nutrients are supplied in nutrient agar (NA)?
2. What kinds of organisms can grow on this medium?
3. Were your plates uncontaminated several days after preparation? Does this mean the
plates are safe to use for pure cultures? Explain.
4. Why is it important to ensure the medium does not cool below 50 oC before dispensing
onto plates?
5. Briefly explain how an autoclave works.
6. Describe the procedures and precautions that should be followed when loading and
unloading an autoclave.
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Micro-organisms in the Environment
Elements of competency1, 2, 3, UPK
Introduction
Microorganisms exist in mixed populations in nature and pure cultures grown in the laboratory must
often be isolated from samples containing mixed cultures. In order to grow these microorganisms, they
must be provided with suitable nutrients and environmental conditions.
This exercise will examine some different sources of microorganisms in the environment
and collection of them using the Swabbing technique. Sterile swabs are used to collect
bacterial samples.
The larger the area, the more bacteria you are likely to collect, so it is important to swab the
same area each time. The area should be specified in the procedure manual. A template
such as a 10cm square is often used to collect swabs.
Materials (per person)
Sterile swabs, sterile water, 2 x NA plates, marking pens, 2 x blood agar plates
Procedure
1. Correctly completed the sample request form before commencing the testing
2.
Ensure the plates are labelled correctly. Seal with parafilm.
3.
Incubate all plates at 37oC overnight.
4. Your lecturer will give out a few control plates to students to label if you are given one
please label ‘control’ include your name on this plate as well. Do not open it. Seal
with parafilm and incubate with other plates
Environment Fingers & Hand washing
Body & Mouth
5. Using sterile water and sterile swab aseptically take a sample of the laboratory with a
damp swab. (your lecturer will demonstrate this)
Suggestions include: Floor, bench, windowsill, soap & other areas a discussed with your
lecturer
6. Streak the swab onto a plate of NA that you have labelled prior to taking the swab. Draw
a line across the base of a NA or BA plate to divide into two. Label one side of the plate
‘uwf’ and other ‘wf’. Briefly remove the lid of the plate and press three fingers lightly onto
the ‘uwf’ side.
7. Wash your hands thoroughly and press the same three fingers onto the ‘wf’ side.
Draw a line across the base of a blood agar plate to divide into two (or more).
Label one side of the plate mouth and the other body.
8. Use a sterile swab on the inside of your mouth and inoculate the 'mouth-half' of the
blood agar plate with this swab.
9. Take a swab from one other damp body surface e.g. toes, armpit, ear and streak the
other half of the plate.
10. Ensure that you label your samples correctly and record on the laboratory form
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provided
11. Record area swabs taken. Use a key if required
12. Area swabs taken
Area swabs taken
Area swabs taken
Observations
1. Examine all of the plates at your next practical session and record the results in a
table
DO NOT OPEN THE PLATE
Sample source
Appearance of & Number of
Colonies
Number of different types
2. What type of growth did you expect from the different areas?
3. Compare with your actual results.
4. Make sure that you describe the differences in colony appearance from the different
sources. Why are colonies different in appearance?
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5. Describe how you would obtain a pure culture from your plates – briefly list the steps
required.
Competency Record: By observation the student satisfactorily completed the following Section/s
Yes
No
Comments
Gram Stain
Preparation of Microbiological Media
Micro-organisms in the Environment
Lecturers signature:
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Microscopic examinations & aseptic techniques
Food processing-Quality Control Test
Elements of competency 1, 2, 3, UPK
Microbiological safety and quality of sprouts in Western Australia
Recent culinary influences on the Australian diet have seen sprouted seeds such as bean
sprouts, alfalfa and snow peas etc. become part of our everyday meals. With their stores of
vitamins, minerals, protein, fibre, delicate subtle flavours and crisp textures, sprouted seeds
make a valuable contribution to a healthy, nutritious and varied diet.
There have been increasing numbers of food poisoning outbreaks associated with the
consumption of sprouted seeds like alfalfa, radish, mung bean, and clover sprouts. Sprouts
have been identified as a potential food poisoning problem.
As part of the quality assurance program for Central Sprout Growers Association samples of
sprouts were removed from the production line, placed in sterile bags and then stored in a
refrigerator, in the microbiology laboratory. Later in the morning, the laboratory assistant
removed the samples from the fridge, registered the samples with the date received and
test code, and signed the register book.
Using aseptic techniques she/he carefully prepared a suspension of sprouts in sterile
deionised-water, prepared appropriate dilutions and transferred 1 ml of sprout mix onto the
required number of total plate count agar (or nutrient agar). The plates were then placed in
the incubator. The final results were noted and recorded.
Materials per group
Sterile water diluent - 10ml with glass beads in McCartney bottles, food blender /
homogeniser, sterile spatulas/forceps, sterile petri plates, dilution blanks (9ml) (0.1 %
peptone or Di-water) 6 for each group, tube racks for blanks, water bath at 50C, melted
nutrient agar tubes (about 20 ml in each) in water bath, autopipettes & sterile tips, vortex
mixer
Procedure: The plate count
1. Make sure you have correctly completed the sample request form before
commencing the testing.
2. Using sterile forceps, aseptically weigh 1.0g ± 0.1 g of the sample into a sterile
container
3. Add the food sample to 10 ml of the diluent-containing glass beads and shake until the
food is well mixed and broken up. An alternative to this is a ‘stomacher’ or to use a
homogeniser or hand-held food blender (without the glass beads!) for larger food
samples / volumes. At this point you may need to compress the debris in the tube, or
allow it to settle for 5 minutes. Maintain aseptic technique throughout
4. Set up the 9 ml. sterile blanks in a rack and label them 10-1, 10-2, 10-3, 10-4, 10-5, 10-6
5. Now add 1ml of the blended food sample to the 9 ml 10 -1 tube - use a sterile pipetteand mix well use a vortex mixer for this.
6. Transfer 1 ml of the mixture from the 10-1 tube to the 10-2 tube and mix well (vortex).
7. Repeat this transfer procedure from 10-2 to 10-6.
8. Using a fresh pipette, transfer 1 ml from three of these dilutions to empty petri dishes
labelled to match the dilutions (e.g. sprout 10-2, 10-3, and 10-6.) Your lecturer will tell you
which dilutions you are plating out.
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9. Remove one tube of the melted nutrient agar (make sure it isn’t too hot from the water
bath and pour it carefully into the first plate. Gently mix and allow it to set. Repeat this step
for the other pour plates.
10. Repeat the process using the spread plate method with 100µl of a dilution.
11. Incubate the plates inverted at 30°C ± 1°C for 72 h ± 3 h.
Assessment criteria
For the purpose of this test, sprouts are considered as a ready-to-eat product and the
microbiological results are compared against the WA Guidelines for Ready-To-Eat Foods, for
other organisms. Samples are considered unacceptable if they exceeded the following
criteria:

Total Plate Count- more than 100 colony forming units per gram (cfu/g)
Results
Examine the pour and spread plates. The ‘best’ plates to count are ones with colonies
between 30- 300. Choose the best dilution and count the number of colonies present.
Record colony count:
Assume each colony is derived from a single bacterium, and is therefore a colony – forming
unit (cfu). Calculate the number of bacteria per gram of food sample (1 ml of diluted food
sample = 1g food; 1ml or 0.1ml sample plated out).
Calculation:
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Tabulate your observations of the colonies and cfu / counts on the different media.
Treatment
Observations
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Discussion
1.
What numbers did you get for the plate counts? Does this mean the food should not
be eaten?
2. Why is it important that all the procedures are carried out aseptically?
3. In a microbiological laboratory you may have to take different types of samples for
example: food, water, waste water, surface swabs, air samples, urine, faeces, blood
etc. Make a list of equipment required to take a food sample and what you should
check.
Notes
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Bacterial Cultures
Elements of competency1, 2, 3, UPK
Introduction
To determine the growth requirements of several different types of bacteria
Materials
Various media: Nutrient agar, Blood agar, CLED agar, SAB plates plate cultures of
Streptococcus pyogenes, Staphylococcus epidermidis, Escherichia coli, Saccharomyces,
loops, flaming alcohol, bunsen burner & bacticinerator, auto pipette and 1mL tips (sterile),
Method
1. Culture each of the bacteria using the 'streaking out' method onto agar plates as
directed by your lecturer.
2. Incubate the nutrient and CLED agar plates and one of the blood agar plates for each
organism in the 35oC incubator.
3. Put one blood agar plate of each organism into a CO2 jar. Light a tea light candle, and
place in the jar. Seal the Jar's lid, and watch the flame die out. Place the jar into the
incubator.
4. Seal the remaining blood agar plates with parafilm. Place one for each organism
onto the windowsill of the biology lab. Place the rest of the plates into a dark
cupboard.
5. After 24 hours incubation, remove the plates from the incubator. Seal them with
parafilm and refrigerate until next lesson. Leave the plates on the windowsill and in
the cupboard to incubate until next lesson.
Observations and Discussion
1.
Make observations on each of the agar plates for each organism.
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2. How well did the organisms grow in each of the conditions and on the different plates?
3. Are the colonies well separated in the last three sections of the plate? If not what can
you do to improve your technique next time.
4. Is there a difference in the growth you observed on the two types of media?
5. Record your streak/spread pattern in the circles below.
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Producing Purity Plates from Mixed Cultures
Materials
Agar plate of mixed growth, NA plates, Bacterial loops, Glass slides, Sterile physiological
saline (0.85% NaCI), Gram's staining chemicals: 1% Crystal violet, Gram's iodine, Acetone,
1 % Safranin
Procedure
1.
Make observations on the colony morphology of each organism in the mixed growth
2.
Label a plate for each organism.
3.
Aseptically pick a single colony of one organism type and streak out for isolation.
4.
Repeat for other types of organisms, using a separate plate for each.
5.
Incubate then observe if you were able to produce pure plates of each organism.
The following week:
1.
You can now make a Gram stain of these cultured bacterial colonies. Label a slide
with an ID for the bacterium, your name, 'Gram stain', and today's date.
2.
Place a loopful of saline onto your slide. Emulsify some colony into the saline. You do
not need to use much bacteria (less than one colony will be enough), or there will be
too many cells on top of each other for you to be able see their shapes easily.
3.
Allow the saline to dry out and heat-fix your slide.
4.
Stain the smear using the Gram procedure
5.
Observe the cells using oil immersion technique. Record a diagram of what you see,
and label as Gram positive (purple), Gram negative (pink), and the cell shape – bacilli
(rods) or cocci (spheres).
Competency Record: By observation the student satisfactorily completed the following Section/s
Yes
No
Comments
Food processing-Quality Control Test
Bacterial Cultures
Producing Purity Plates from Mixed Cultures
Lecturers signature:
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Culturing Fungi
Elements of competency1, 2, 3, UPK
Introduction
Fungi have a much higher requirement for carbohydrates than bacteria; therefore agar for growing
fungi needs a different formulation from that suitable for growing bacteria. Just as there are many
different formulations for bacteria, so there are several (but not as many) for fungi. One basic agar is
known as Sabouraud's Dextrose Agar. Dextrose is another term for glucose, and glucose is the
fundamental 6 carbon sugar found in living things. The fungi also require protein compounds; this is
supplied via peptone which is a short chain of amino acids.
Week 1: To 'capture' moulds by exposing bread
Materials: Petri dishes (need not be sterile), water, bread
Procedure
1. Moisten the bread, leave it exposed on the Petri dish until the end of your class (at least
30mins) and then incubate 7 days at room temperature.
Week 2: To transfer fungi from bread to Sabouraud's Dextrose Agar.
Materials
Source of fungi (exposed bread from previous week), 2x Sabouraud's agar plates, Lactophenol cotton
blue stain, slides &coverslips, spirit burner, biosafety cabinets, needles, flaming alcohol, discard jar,
cellotape, microscopes
Procedure to take place in biosafety cabinets.
1. Label two SAB plates.
2. Use flame sterilised forceps or dissecting needle to try to remove a sample of fungus, 2-3mm3 in
size, from your source. Try to remove only one type of fungus. You may be able to set up plates of
different species, but only put one sample on each plate.
3. Gently poke the sample into the surface of the agar. Replace and seal lid.
4. Incubate room temperature.
5. Check growth after 24 and 48 hours.
6. Ensure that you wipe down the inside of the biohazard cabinet, spray with acetic acid (vinegar) for
fungi, ethanol for other organisms.
Week 3:
This procedure is to be carried out in the biohazard cabinet with no fan running. Make sure
the cabinet is cleaned, wipe down the inside of the biohazard cabinet, spray with acetic
acid (vinegar) for fungi & UV light switched on after practical is completed
All slides are to be disposed of into a separate labelled container not into the normal
sharps bin
On a glass slide place a drop of stain.
Take a small piece of sticky tape, and gently place it over the fungal growth. Place the
sticky tape (sticky and fungus-side down) over the stain on the slide.
Observe the stained slide under the microscope with a x40 objective and draw diagrams of
your observations in your logbook. Use a textbook to help you label your diagram of the
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microscopic structures you can see, such as spores, conidiophores, conidia, hyphae, etc.
include measurements. Can you identify the type of fungus present?
Results
Examine your plates for growth.
Do you have a pure culture? _______________________________________
Describe the type of growth observed. Include colour and size of colonies on the plate.
___________________________________________________________________________________
___________________________________________________________________________________
___________________________________________________________________________________
___________________________________________________________________________________
___________________________________________________________________________________
Draw a diagram of the fungal growth.
Diagram from plate
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Diagram from slide
Ensure you label correctly.
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Membrane Filtration
Elements of competency1, 2, 3, UPK
Introduction
Solutions that are thermolabile may be sterilised by passing the fluid through a
microbiological filter. This filter has minute pores (typically of 0.2 – 0.4 m) so bacteria
are retained on the filter and the filtrate is sterile. In this exercise a broth containing
micro- organisms is filtered and the filtrate is plated out to check the efficiency of this
process. A sample of the broth before it has been filtered is also plated out as a control.
Materials (each)
Broth culture of S.epidermidis OR Saccharomyces cerevisiae in a wide mouthed container,
sterile container with lid (for filtrate) test tube rack, marker pen, Inoculating loop or
sterile cotton swab, burner, microbiological filter to fit syringe, sterile 5mL syringe,
sabouraud agar for Saccharomyces cerevisiae, blood agar for S.epidermidis
Procedure
1. Draw a line on the base of the agar plate to divide it into two halves. Label one side ‘F’
for filtered and the other ‘UF’ for unfiltered. Also add your initials and the date and the
culture used. Keep the labels near the edge of the plate so that any growth can be seen
easily.
2. Using aseptic techniques remove the cap from the broth and aspirate about 1-2 mL of
broth into the syringe (before the filter is attached to the syringe). Replace cap on
broth.
3. Again, using aseptic technique, open a filter and attach it to the syringe.
4. Using slight positive pressure, slowly filter the fluid into a sterile container making sure
that contamination is avoided.
5. Use a sterile loop to streak a small amount of the filtered fluid onto the side labelled ‘F’.
6. Repeat step 5 above using the culture that has not been filtered and streak it onto the
unlabelled side of the plate.
7. Seal the plates and incubate at room temperature 25 oC Saccharomyces cerevisiae or at
37oC for S. epidermidis and examine for growth after 24 hours. The plates may be
refrigerated if they cannot be examined the next day.
8. The cultures and the filtrate should also be kept for examination.
Observations
Filtered side
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13. Record the growth of the culture streaked onto the two sides of the plate in the table
above.
14. Examine the broth culture and the filtrate. What differences can you see?
15. How can growth in broth cultures be detected?
16. What effect does microfiltration have on the culture you used?
17. Would this method ensure that viruses were filtered? Explain.
18. List the different methods that were used to sterilise all of the equipment and media
used in this exercise.
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19. Why is the same method of sterilisation not suitable for each item?
20. Are there any colonies on your plates that are visibly different from the
microorganisms that were used in the exercise? Where might they have come from?
21. Why are different temperatures suggested for the incubation of the Saccharomyces
cerevisiae and the Staphylococcus cultures?
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The effect of UV light on bacterial growth.
Elements of competency1, 2, 3, UPK
Materials (per student)
Nutrient agar plates x1, aluminium foil, blood agar plates x1, safety cabinet with UV light, E.
coli or S.epidermidis broth cultures, marking pens, auto pipettes, sterile tips (100µL -1µL),
foil covered platforms
Procedure
1. In this method the liquid sample is added with a sterile pipette and spread as evenly as
possible over the surface of the plate using a sterile spreader. A confluent growth over the
entire plate is produced. No separate colonies should be visible.
2. Label one half of NA plate ”UV 30 minutes” and the other half, “No UV. “
3. Aseptically transfer 100µL (0.1ml=100µL) of culture onto NA plate.
4. Spread the culture with a sterile hockey stick spreader, for confluent growth.
5. With the lids off cover the no UV half of each plate with aluminum foil
6. Place plates in the Biological Safety Cabinets with the UV lights on for the designated time.
Ensure that the plates are placed 22.5cm away from the UV source (use the foil covered boxes
provided). Be sure to put the cover on the cabinet! Observe safety precautions for U.V. light
7. Cover the plates and incubate at 37°C for 24-48 hours. Record results in the table below
Observations: Record patterns of growth on your plates.
Culture
Not UV exposed
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Discussion
8. What effect (if any) did UV exposure have on the samples?
9. What are the limitations of ultraviolet light for sterilization and disinfection?
10. Discuss the results you might expect and compare them with the results you
obtained.
Competency Record: By observation the student satisfactorily completed the following Section/s
Yes
No
Comments
Membrane Filtration
Culturing Fungi
The effect of UV light on bacterial growth
Lecturers signature:
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Appendix A: Central Laboratories Specimen Receipt
Date/Time
Patient Identification
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Sample type
No. received
Test Ordered
Lab Section/s send to
81
Problems with form or sample
Action
taken
Lab ID
No
Tech
Init.
Microscopic & aseptic techniques
Date/Time
Patient Identification
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Sample type
No. received
Test Ordered
Lab Section/s send to
82
Problems with form or sample
Action
taken
Lab ID
No
Tech
Init.