School Science Lessons
Please send comments to: J.Elfick@uq.edu.au
Table of contents
Warning and Comment
4.1.0 Microbiology cultures
2.1.0 Microbiology safety
188.8.131.52 Microbiology techniques
4.3.0 Pectin, pectinase
See also: 1.0.0 Appendix B. Biology
184.108.40.206 Microbiology techniques
220.127.116.11 Aseptic transfer of bacterial
cultures from a bottle or tube
18.104.22.168 Aseptic transfer of bacterial
cultures from a culture plate
3.1.10 Colony counts using the calibrated drop method
3.1.3 Flaming and cotton wool plugs
3.1.6 Inoculate with a Pasteur pipette
22.214.171.124 Lawn plate technique
126.96.36.199.1 Estimation of aerobic
mesophilic bacteria by the plate count method
3.1.7 Pour a plate
188.8.131.52 Make heat-fixed stained
3.1.9 Prepare spread plates, lawn plates
3.1.4 Prepare streak plates
3.1.5 Prepare pour plates
184.108.40.206 Serial decimal dilution
of a bacterial suspension
220.127.116.11 Spread plate technique
18.104.22.168 Staining rack
22.214.171.124 Streak plate dilution method
for pure cultures from a mixed suspension
3.1.1 Wire loops
4.1.0 Microbiology cultures
4.1.1 Colonies of different micro-organisms
4.1.2 Enrichment of wild yeast strains, Aspergillus
4.3.1 Grow African violet, with in vitro culture
4.3.2 Grow African violet, from pieces of leaf
4.3.3 Grow Gerbera using in vitro culture
4.1.3 Prepare fixed slide preparations
4.1.4 Prepare India ink background stain
3.1.9 Prepare spread plates, lawn plates
4.1.7 Prepare streptomycin using Streptomyces
4.1.9 Presence of bactericidal substances using a
coin and Bacillus mycoides
4.1.5 Safe microscopy of Penicillium camemberti
4.1.8 Streptomycin on Bacillus subtilis, small
4.1.6 Soil bacteria that decompose urea
4.2.0 Fermentation, Food preparation
126.96.36.199 Enzyme technology, pectinase, amylase,
protease, lipase, lactase
188.8.131.52 Enzyme technology, pectinase in the industrial
production of juice
4.2.4 Make wine from grape juice and vinegar from
7.1.1 Pectinase, (pectin), Aspergillus niger
4.2.10 Enzyme technology, industrial uses of pectinase
4.2.7 Microbial decomposition of thin paper, cigarette
4.2.8 Prepare apple juice gel
4.2.5 Prepare cider from apple juice
4.2.3 Prepare lactic acid with sourdough
4.2.9 Prepare pectinase, an enzyme that decomposes
4.2.6 Prepare vinegar with Acetobacter aceti
4.2.1 Prepare yoghurt
4.2.1a Prepare yoghurt, a report from Turkey
4.3.17 Prepare yoghurt, test milk
4.2.2 Prepare sauerkraut
4.2.11 Split lactose from milk or whey using immobilized
4.3.0 Pectin, pectinase
4.3.14 Breakdown of pectin by micro-organisms
4.2.9 Prepare pectinase, an enzyme that decomposes
184.108.40.206 Enzyme technology, pectinase, amylase,lactase
220.127.116.11 Enzyme technology, pectinase in the
industrial production of juice
3.1.1 Wire loops
Sterilize wire loops by heating in a Bunsen burner flame until red.
Hold the handle of the wire loop close to the top, like holding a pen,
at an almost vertical angle, leaving the little finger free to take
hold of the cotton wool plug or screw cap of a test-tube or bottle.
Heat the end of the loop slowly because after use it may hold culture that
may splutter on rapid heating.
Hold the handle end of the wire in the light blue cone of the flame, the
cool area of the flame.
Move the rest of the wire slowly upwards into the hottest region of the
flame above the light blue cone and hold it there until it is red hot.
Heat the full length of the wire. Use the wire loop as soon as it is cool.
Do not put the wire loop down on the desk and do not wave it around in
Sterilize the wire loop again immediately after use.
Use a sterile graduated pipette and filler or dropping (Pasteur) pipette
to transfer cultures, sterile media and sterile solutions.
Remove the pipette from its container or wrapper by the end that contains
a cotton wool plug.
Fit the teat.
Hold the pipette barrel as you would a pen but do not grasp the teat.
Leave free the finger to take hold of the cotton wool plug or cap of a
test-tube or bottle.
Leave the thumb to control the teat.
Depress the teat carefully to take up enough fluid but not enough to wet
the cotton wool plug.
Return any excess fluid if a measured volume is required.
Keep the pipette tip beneath the liquid surface while taking up liquid
to avoid taking up air bubbles.
Immediately after use, put the "contaminated pipette" into a discard pot
of 0.25% v/v sodium chlorate I (sodium
hypochlorite) then remove the teat.
Never use the mouth to "suck up" fluid into a pipette!
3.1.3 Flaming and cotton wool
Flame the neck of bottles and test-tubes.
Loosen the cap of the bottle.
Lift the bottle or test-tube with the left hand.
Remove the cap of the bottle or cotton wool plug with the little finger
of the right hand.
Turn the bottle, not the cap.
Do not put the cap or cotton wool plug down on the desk.
Flame the neck of the bottle or test-tube by passing the neck forwards
and back through a hot Bunsen burner flame.
After the procedure, replace the cap on the bottle or cotton wool plug
using the little finger.
Label test-tubes and bottles with a marker pen where it will not rub off.
Cotton wool plugs are used to plug test-tubes and pipettes to allow the
passage of air but prevent the passage of micro-organisms.
They must be made of non-absorbent cotton wool, be kept dry, and must keep
its shape after being removed and returned to the
3.1.4 Prepare streak plates
See diagram 9.4.13: Streak plate
Streaking causes a progressive dilution of an inoculum over the surface
of solidified agar medium in a Petri dish so that the
colonies of bacteria or yeast grow separated from each other as single
isolated pure colonies.
1. Partially lift the lid of the Petri dish containing the solid medium.
Hold the charged wire loop parallel with the surface of the agar.
Smear the inoculum backwards and forwards across a small area of the agar
medium on the left hand side of the plate.
Remove the wire loop and close the Petri dish.
Flame the wire loop and allow it to cool.
2. Turn the Petri dish through 90o anticlockwise.
Use the cooled wire loop to streak the agar plate across the surface in
three parallel lines.
A small amount of culture must be carried over.
Remove the wire loop and close the Petri dish.
Flame the wire loop and allow it to cool.
3. Turn the Petri dish through 90o anticlockwise again and
streak across the surface of the agar in three parallel lines.
Remove the wire loop and close the Petri dish.
Flame the wire loop and allow it to cool.
4. Turn the Petri dish through 90o anticlockwise then streak
the wire loop across the surface of the agar into the centre of the plate.
Remove the wire loop and close the Petri dish.
Use a marker pen to label the Petri dish at the edge off the plate.
Flame the wire loop.
Seal and incubate the plate in an inverted position so that condensation
cannot occur on the lid and drip onto the culture, causing
colonies to spread into each other.
Professional microbiologists start with the Petri dish inverted on the
Then they lift out the base, invert it, then inoculate the agar facing
3.1.5 Prepare pour plates
Use a pipette to add inoculum from a broth culture to the centre of a
Petri dish, then add previously molten, cooled agar medium.
Rotate the Petri dish to mix the culture and medium thoroughly and ensure
that the medium covers the plate evenly.
Pour plates allow micro-organisms to grow both on the surface and within
Most of the colonies grow within the medium and are small and may be confluent.
The few colonies that grow on the surface of the medium are generally of
the same size and appearance as colonies on a streak
If the dilution and volume of the inoculum, usually 1 mL, are known, the
viable count of the sample can be calculated, i.e. the
number of bacteria or clumps of bacteria per mL.
The dilution chosen should produce 30 to 300 separate countable colonies.
3.1.6 Inoculate with Pasteur
Loosen the cap or cotton wool plug of the bottle containing the inoculum.
Remove the sterile Pasteur pipette from its container, attach the bulb
and hold it in the right hand.
Lift the bottle or test-tube containing the inoculum with the left hand.
Remove the cap or cotton wool plug with the little finger of the right
Flame the bottle or test-tube neck. Squeeze the teat bulb of the pipette
slightly, put the pipette into the bottle or test-tube and
draw up some of the culture.
Always hold the pipette as still as possible.
Do not squeeze the teat bulb of the pipette after it is in the broth because
this could cause bubbles.
Remove the pipette and flame the neck of the bottle or test-tube again,
before replacing the cap or cotton wool plug.
Place a bottle or test-tube on the bench.
When inoculating e a Petri dish, lift the lid with the right hand just
enough to insert the pipette and release the required volume of
inoculum onto the centre.
Replace the lid.
Put the pipette into a discard pot of disinfectant.
Remove the teat while the pipette is pointing into the disinfectant.
3.1.7 Pour plates
Collect one bottle of sterile molten agar from the water bath.
Hold the bottle in the right hand then remove the cap with the little
finger of the left hand.
Flame the neck of the bottle.
Lift the lid of the Petri dish slightly with the left hand and pour the
sterile molten agar into the Petri dish and replace the lid.
Flame the neck of the bottle and replace the cap.
Rotate the Petri dish to mix the culture and the medium thoroughly and
to ensure that the medium covers the plate evenly.
Leave the plate to solidify.
Seal and incubate the plate in an inverted position.
The whole base of the plate must be covered.
Do not let agar touch the lid of the plate.
The surface must of the inoculated medium must be smooth with no bubbles.
Use a sterile spreader to distribute inoculum over the surface of agar
plates with a dry surface.
First, dry the surface of agar plates by either incubating the plates
for several hours, e.g. overnight, or put them in a hot air oven
at 60oC for 60 minutes with the two halves of the Petri dish
separated and the inner surfaces directed downwards.
Sterilize glass spreaders in a hot air oven.
Do not put the spreader down on the bench.
3.1.9 Prepare a spread plate,
The plate should have a growth of culture spread evenly over the surface
of the growth medium.
Use it to test the sensitivity of bacteria to antimicrobial substances,
e.g. disinfectants and antibiotics, and to determine the number
of bacteria or clumps of bacteria per mL, colony count.
For an accurate count, the dilution and volume of the inoculum, usually
0.1 mL, must be known and the dilution chosen must
produce 30 to 300 separate countable colonies.
Loosen the cap of the bottle or test-tube containing the broth culture.
Remove a sterile Pasteur pipette from its container and attach the bulb
held in the right hand.
Hold a sterile pipette in the right hand and the bottle or test-tube containing
the broth culture in the left.
Remove the cap or cotton wool plug of the bottle or test-tube with the
little finger of the right hand and flame the neck.
With the pipette, remove a small amount of broth.
Flame the neck of the bottle or test-tube and replace the cap or plug.
With the left hand, partially lift the lid of the Petri dish containing
the solid nutrient medium.
Place a 5 drops of culture on the surface, an area of 0.1 cm3,
or enough to cover a UK 5 pence piece.
Replace the lid of the Petri dish.
Place the pipette in a discard jar of disinfectant.
Lift the lid of the Petri dish to allow entry of a sterile spreader.
Place the spreader on the surface of the inoculated agar and move the
spreader in a top-to-bottom or a side-to-side motion to
spread the inoculum over the entire surface of the agar.
Do this as fast as possible to reduce contamination.
Replace the lid of the Petri dish.
Put the spreader in a discard jar of disinfectant.
Leave the inoculum to dry.
Seal and incubate the plate in the inverted position.
To produce an agar plate inoculated with mould mycelium inoculated at
the centre, invert the plate, lift the base of the Petri dish
that contains the medium and inoculate onto the centre of the downwards
facing agar surface with a bent wire.
This method avoids the problem of spores falling off the piece of mycelium
and producing unwanted inoculation sites.
3.1.10 Colony counts using
the calibrated drop method
Use this method only for pure cultures of bacteria and yeast.
The procedure is similar to the spread plate procedure but the inoculum
is added as drops from a dropping pipette calibrated to
deliver drops of known volume, e.g. 0.02 mL.
About six drops from different cultures can be put on the same plate, thus
saving the number of plates needed.
The method is not usually suitable for mixed cultures, e.g. soil samples
Incubators are not really necessary for microbiology in schools because
most of the cultures suitable for use in schools grow at
room temperature so can be incubated in a cupboard.
Incubators can be set at a range of temperatures but overlong incubation
of a forgotten mould cultures may result in a massive
formation of spores, which may cause contamination problems and be a health
The internal temperature of incubators may vary to it is best to use a
water baths for accurately controlled temperatures needed
for studying enzyme reactions and growth-temperature relationships.
4.1.1 Colonies of different
See diagram 20.114: Required equipment
Moulds usually form a soft, stringy colony.
Colourless mycelia may also grow below the surface of the agar medium.
The aerial mycelium of Penicillium roqueforti usually has blue-green
Spores of other fungi are also coloured brown or yellow or black.
The diameter of a single colony is usually more than 10 mm.
Yeasts or bacteria, other than Streptomyces, have shiny matt or
slimy colonies often above the surface of the agar.
The colonies are often grey or yellow.
Bacillus subtilis bacteria colonies are usually grey.
Yeasts may be grey, red, orange, yellow, brown.
The diameter of a single colony is usually less than 10 mm.
Streptomyces bacteria have an earthy smell.
In the Petri dish, a truncated, round, single colony less than 3 mm in
diameter forms that grows in the shape of a lens.
The aerial mycelium is often coloured after 6 days incubation.
Streptomyces griseus produces a grey aerial mycelium.
Other Streptomyces stain the surrounding agar brown.
Equipment: 1 Bunsen burner, 1 inoculation loop, 1 conical flask 250 mL
9.2.14 Basal agar medium, 100 mL
9.2.15 Basal broth medium, 50 mL
Petri dishes X 3
Felt tip pen X 1
Cultures of the following micro-organisms, incubated overnight:
Penicillium roqueforti (DSM-No. 1079)
Rhodotorula rubra (DSM-No. 70403)
Streptomyces griseus (DSM-No. 40236, ATCC 23345)
Bacillus subtilis (DSM-No. 1079, ATCC 6051)
1. Prepare overnight cultures in basal broth medium, 45 minutes.
2. Incubation of overnight cultures, 24 hours.
3. Inoculation, 15 minutes.
4. Incubation, 6 days
1. On the day before the investigation, prepare
cultures of the four species of micro-organisms and incubate overnight.
2. Prepare 100 mL of basal agar medium in 3 agar plates.
3. Divide the Petri dishes into 4 sectors on the underside, using a felt
4. Inoculate each of the 4 sectors at one point with one of the 4 types
5. Incubate the Petri dishes at 30oC for 6 days.
6. Identify the colonies.
4.1.2 Enrichment of wild yeast
strains, Aspergillus niger
Produce yeast cultures derived from natural isolates within closed Petri
dishes for demonstration purposes.
However, for safety reasons they should not be used for further experiments.
Some moulds spoil food but others can be used in the production of food,
e.g. Camembert cheese, Indonesian bean cake
tempeh, soya bean cheese.
However, there are considerable safety risks in the open microscopy of
For example if a student who accidentally coughs into a spore culture can
propel large numbers of spores into the air such as
Aspergillus niger that can infect the respiratory tract and may
be fatal for those whose immune system has been weakened.
The Petri slide procedure for safe microscopy of mould avoids these risks.
18.104.22.168 Enrichment of wild
Equipment: Petri dishes
22.214.171.124 Malt extract agar medium
Unwashed apple, or another fruit with sugar content
Time needs: 45 minutes
1. Prepare malt agar plates according to the instructions.
2. Roll a piece of unwashed fruit across the surface of the chilled
agar, close the dishes immediately.
Incubate the Petri dishes for a few days at 30oC.
Observe different yeast types.
Some yeasts are brilliantly coloured.
Some mould cultures can be recognized by their cotton-like texture.
4.1.3 Prepare fixed slide preparations
See Diagram 20.120: 1. Put a cover slide on
a microscope slide, 2. Spreading a drop of liquid
Focussing sharply on living bacteria is difficult, so they are almost
always observed in fixed preparations and stained.
Equipment: 1 microscope slide, 1 coverslip, 1 eye dropper, 1 Bunsen burner
Time needs: 30 minutes
1. Remove grease carefully from a microscope slide with a lint free towel
or a piece of tissue soaked in ethanol.
2. Place a drop of bacteria or yeast suspension in the middle of the microscope
The drop should flow out evenly and must not remain in globular form.
The suspension eventually will flow back together, even when only traces
of grease are present on the slide.
This not only lengthens the drying time, but allows thick layers of bacteria
to develop, as well.
So it may no longer be possible to observe individual micro-organisms.
If attempts to remove grease from the slide are unsuccessful, use a drop
of extremely diluted bacterial suspension and leave to
air dry in place of the smear technique.
This method is easier than executing a smear with the delicate coverslip.
3. Place a coverslip on the microscope slide at an angle of 45o
so that the solution is collected in the space between the slide and
slip and held by the properties of adhesion and cohesion.
It is important to pull and not push the suspension across the slide with
the coverslip to ensure that the thickness of the coating
4. Push the coverslip evenly across the entire surface of the microscope
This spreads the suspension across the slide, and the film of liquid becomes
This drying step must not be accelerated by heating.
Bacterial structure changes when bacteria are heated in water.
5. Allow the smear to dry.
The first method is recommended.
Because of the poor conducting qualities of glass, it is difficult to estimate
the effect of heating on the bacteria with the second
Organisms and the protein coagulated in the cells by heating adhere to
the slide surface.
6. Fix the bacteria to the slide by briefly heating the slide in a flame.
This can be done in one of two ways: with a low flame such as the pilot
flame of a Bunsen burner and with the coated side of the
slide oriented downwards, or with the high flame of a Bunsen burner and
the coated side of the slide up.
Pass the slide through the flame three times at a speed of roughly 30 cm
4.1.4 Prepare India ink background
If you stain the background uniformly black by the use of India ink, only
the organisms are illuminated in the microscope, and
they appear in bright contrast to the background.
Equipment: 2 microscope slides, 1 coverslip, 1 eye dropper
Materials: India ink, ethanol
Time needs: 30 minutes
1. Thoroughly clean two microscope slides by wiping them with a lint free
towel or a tissue soaked in ethanol.
2. Place a small drop of water on the microscope slide.
The drop must spread out, otherwise further measures are necessary to remove
grease from the slide.
3. Use a glass rod to mix a drop of India ink with the evenly spread drop
4. Place a coverslip at a 45o angle on the microscope slide
in such a manner that the solution is collected in the space between
the slide and slip and held by the properties of adhesion and cohesion.
5. Push the coverslip evenly across the entire surface of the microscope
The suspension is thus spread across the slide.
The thickness of the film of liquid decreases.
6. Allow the smear to air dry.
7. Prepare a second smear, using a drop of bacterial or yeast suspension
instead of a drop of water.
8. Compare both smears are then compared under a microscope set at 400
Open the condenser completely and use the brightest possible light source.
9. If observing bacteria with a microscope for the first time, prepare
a control slide for comparison.
4.1.5 Safe microscopy of Penicillium
camemberti and Mucor mucedo using the Petri slide technique
| See diagram 20.120 (3): Inoculate a culture
medium with fungal spores
| See diagram 20.120 (4): Remove fungal spores
from a pure culture
| See diagram 9.202: Penicillium, Mucor
Also, Penicillium chrysogenum in a malt extract agar is safe for
use in schools.
Petri slides are extremely flat, disposable Petri dishes that can be
sealed tightly, height = 6 mm, inner diameter = 47 mm,
area of the base plate = 52 X 75 mm.
They are designed for counting micro-organisms.
Unknown micro-organisms are placed on nutrient cardboard discs or membrane
filters, and colonies are cultivated inside
The construction of the chambers also enables both the safe microscopy
of sealed fungal cultures and the microscopy of
sporangia from the side.
A packet of Petri slides containing 100 slides can be ordered from laboratory
Petri slide cultures can be kept for several weeks.
Basal agar medium is suitable for the cultivation of various moulds.
Fungi do grow on other nutrients such as glucose nutrient agar, but in
such cases they do grow more slowly.
Equipment: 1 autoclave or pressure cooker, 2 Petri slides, 1 Bunsen burner,
1 sterile Pasteur pipette, 1 wide-necked 300 mL
conical flask, 1 inoculation needle, household aluminium foil
126.96.36.199 Basal agar medium 200 mL
188.8.131.52 Glucose nutrient agar 200
mL 1 M HCl,
Pure culture of Penicillium camemberti (DSM-No. 1995)
Pure culture of Mucor mucedo (DSM-No. 809, ATCC 38693)
Time needs: 45 minutes, inoculation of the Petri slides: 15 minutes
1. Place the culture medium in a conical flask, seal with aluminium foil,
and autoclave in a pressure cooker.
The time required for sterilization is 20 minutes after the sealing of
the pressure valve.
The Petri slides do not have to be sterilized, as they remain germ free
inside during the production process.
Sterilize Pasteur pipettes by wrapping them in aluminium foil and heating
them to 180oC for 30 minutes in a drying cabinet.
2. Remove the aluminium foil from the conical flask.
Fill a sterile Pasteur pipette with 5 mL of sterilized culture medium.
Do not touch the tip of the pipette.
Hold the Petri slide chamber upright between the thumb and index finger
of the left hand.
Lift the lid up far enough to reveal the side of the base of the filling
Introduce the tip of the pipette through the hole in the side into the
middle of the chamber.
The tip must not touch the outer parts of the chamber.
Carefully pipette 3 mL of culture medium into the vertically held chamber
without smearing the upper part of the chamber with
3. Remove the Pasteur pipette from the chamber and replace the lid of
Ensure that the chamber remains vertical as the agar sets.
Refill the pipette and prepare additional chambers in a similar manner.
Pass the tip of the pipette through a Bunsen buner flame periodically.
4. After the agar has set inside the chambers for about 30 minutes, inoculate
the chambers with different fungi.
Use an inoculation needle that is sterilized by heating in the Bunsen
burner flame and held to the sporangia of a pure culture of mould.
5. Open a Petri slide in the usual way.
Insert the needle through the hole until the pointed end transfers the
spores that adhere to it by contact with the surface of the agar.
6. Close the chamber again and incubate it for about a week in a vertical
position at room temperature.
Individual sporangia can be seen clearly even with microscopy using transmitted
One can "take an optical walk" through the about 5 mm wide sporangia "wood"
by using the fine focussing apparatus.
Differences in the sporangia at the edge of the colony and in the middle
can be seen clearly, and the mass of hypha in the nutrient
agar can be examined up to its finest traces.
The Petri slide cultures can be kept for several weeks in a dark cabinet
at room temperature.
It is inadvisable to keep them in a refrigerator, as the cold chambers
become slightly steamed up if they are used again.
The cultures hardly dry out at room temperature, and the fungi stops growing
one or two weeks after they are inoculated because
they lack oxygen and nutrients.
7. Place the Petri slides flat onto the stage of the microscope once the
cultures have grown, examine with one of the two low
power objectives (10 X, or 40 X).
1. Mouldy fruit or bread should not be examined using open microscopy
because the types of mould that grow on them are
often of the genera Penicillium and Aspergillus.
The spores of Aspergillus may be harmful if students inhale them.
Also, Aspergillus flavus (and Aspergillus parasiticus) produce
aflatoxins, toxic and carcinogenic mycotoxins.
2. It is impossible to find the genus of a fungal colony and compare various
species of mould by microscopic examination from
above because the sporangia, which may be used to distinguish one genus
from another, can only be seen from the side.
3. The tip of the pipette must not touch the outside of the Petri slide
when the slide is being filled with agar because the pipette is
contaminated with bacteria or fungal spores from the environment so the
spores soon begin to sprout and grow in the sterilized agar.
Any additional experiment with a nutrient base that has been contaminated
in this way is invalid.
4.1.6 Soil bacteria that decompose
See diagram 20.120 (5): Proper use of Drigalski
This experiment shows the importance of soil bacteria as decomposers of
184.108.40.206 Basal agar medium, 10
220.127.116.11 Urea agar medium, 5 plates,
18.104.22.168 Ringer solution, sterile,
22.214.171.124 Salt solution, to dilute
the micro-organism suspension
Beaker, 400 mL, Bunsen burners, X 2, Drigalski spatula, Eye dropper, sterile,
Glass plate, Graduated cylinder, 10 mL,
Incubator, Test-tubes, sterile, X 6, Waterproof felt tip marking pen
Materials: Soil, 1g
Time needs: steps 1 to 8: 45 minutes
1. Label six sterile test-tubes in series as follows: 10-1,
10-2, 10-3, 10-4, 10-5, 10-6.
2. Label the undersides of the agar dishes, not the lids, in series as
follows: Nutrient plates: K, 10-1, 10-2, 10-3,
10-4, 10-5, 10-6,
Urea agar plates: K, 10-1, 10-2, 10-3,
3. Pipette 10 mL of the Ringer solution into the test-tube labelled 10-1.
It is important to begin with the control sample because the sterile water
is later needed for rinsing and so becomes contaminated.
4. Sterilize the Drigalski spatula with ethanol.
Apply a drop of sterile Ringer solution to the Petri dish labelled "K".
Spread the solution evenly over the surface of the dish with the sterilized
5. Add one gram of soil to the test-tube labelled 10-1.
Mix thoroughly by shaking and by rolling the test-tube back and forth
between the palm of the hands.
Apply one drop of the 10-1 suspension to each of the two plates
labelled 10-1, spread the suspension evenly with the
resterilized spatula (according to step 5).
Finally, rinse the pipette with sterile water.
6. Transfer one drop from the test-tube labelled 10-1 to the
test-tube labelled 10-2, mix carefully again.
Distribute one drop of the 10-1 suspension evenly on each of
the two Petri dishes labelled 10-2.
7. Proceed in the same manner until all other test-tubes are filled, each
with one drop of suspension from the previous tube and
nine drops of water, and until all Petri dishes are inoculated according
to their respective labels.
8. Incubate the dishes for seven days at 30oC.
9. Count the colonies as soon as they can be easily recognized.
Prepare complete counts of plates with 50 to 200 colonies.
10. To find the titre of the suspension, one must find the volume of
For this purpose, fill a small graduated cylinder with drops (n) to a volume
of 2 mL.
The volume (V) of a drop is derived according to the following formula:
V = 2 or m mL.
For example if 32 drops are necessary to attain a volume of 2 mL, the volume
of a drop, V = 2 or 32 mL = 0.06 mL.
11. Divide the number of counted colonies by the dilution factor.
The results indicate the number of viable cells in one drop of the suspension.
When this number is later divided by the drop volume (V) the number of
viable cells in 1 mL of the suspension is obtained.
For example: 186 colonies were counted on the Petri dish labelled 10-1.
The volume of a drop is 0.06 mL.
The number of colonies is divided by the dilution factor: N = 186 cells
Divide by the volume (V) of drops: N = 1.86 X 10-6 cells or
0.06 mL = 3.1 X 10-7 cells per mL.
As with yeasts, bacteria are transferred in conglomerates.
They must be separated from each other because they will otherwise not
disperse evenly in the suspension.
The bacterial colonies on plates that have been diluted very little (10-3,
10-4) are smaller than colonies on plates that have been
diluted a great deal (10-6, 10-7) because the bacterial
colonies compete for the nutrients in the agar.
On plates where the concentration is high, more colonies develop but they
are more prone to "starve" than few colonies on
plates that are very dilute.
Fewer colonies grow on urea agar plates than on basal medium plates at
a comparable concentration because only very few
micro-organisms possess urease and can use urea as a source of carbohydrates
Most can, however, use glucose, which is present in large quantities in
the basal medium.
The decomposition of urea is visible by means of a red indicator zone.
During the decomposition of urea, a basic ammonia is excreted from the
bacterial cells and released to the environment.
4.1.7 Prepare streptomycin
using Streptomyces griseus
The following investigation employs Streptomyces griseus that produces
the antibiotic streptomycin.
Only those test organisms that are not sensitive to streptomycin can grow
on the same culture medium plate as Streptomyces griseus.
The use of Streptomyces or streptomycin is possible in school experiments
because this antibiotic is no longer used in medicine,
and the possible spread of resistant strains is no longer problematic from
a medical point of view.
Equipment: 1 autoclave, 1 incubator, 1 Bunsen burner, 8 disposable Petri
dishes, 2 sterile 300 mL conical flasks with sterile
stoppers, 1 inoculating loop, 6 culture tubes, 5 X 5 mL sterile pipettes,
Materials: Pure culture of Streptomyces griseus (DSM-NO. 40236,
ATCC 23345) a selection of the following pure culture
strains as test organisms: Bacillus mycoides (DSM-No. 10, ATCC 6051)
Candida utilis (DSM-No. 2361, ATCC 9950)
Escherichia coli K-12 (DSM-No. 498, ATCC 23716) Micrococcus luteus
(DSM-No. 20030, ATCC 4698)
Pseudomonas fluorescens (DSM-No. 50090, ATCC 13525) distilled water,
basal broth medium for overnight cultures
100 mL basal agar medium for Petri dishes 200 mL
Time needs: preparation and autoclaving of the solutions: 45 minutes,
preparing the precultures: 15 minutes, waiting time: 24 hours,
distribution of the test organism cultures: 16 minutes
Preparation: Prepare and autoclave the culture media.
Place the basal agar medium into eight Petri dishes, 20 to 25 mL per plate.
Suspend again the culture of Streptomyces griseus in 1 mL of sterile
basal broth medium according to the manufacturer's
instructions, pipette into a test-tube into which 5 mL sterile culture
medium has been placed. Incubate for 24 hours in an incubator at
30oC, overnight culture.
1. Inoculate the basal agar media in the Petri dishes with the overnight
culture of Streptomyces griseus.
Sterilize an inoculation loop by passing it through a Bunsen burner flame,
allow the loop to cool, then dip into the culture.
Streak the germs that adhere to the loop onto the culture medium as a
vertical line as far over to the right as possible so that the
left part of the surface of the culture medium remains sterile.
Sterilize the inoculation loop by passing it through a flame once more.
2. Incubate the Petri dishes for two days at 30oC.
3. In the interim, after incubating for 24 hours, prepare the overnight
cultures of the selected organism, Bacillus mycoides,
Candida utilis, Escherichia coli K-12, Micrococcus luteus,
Pseudomonas fluorescens, using the same procedure as for the
overnight culture of Streptomyces griseus.
4. Once the incubation time has elapsed, inoculate the culture media from
four Petri dishes on which the Streptomyces cultures
are now visibly growing with the test organism, use an inoculation loop,
as in step 1, to make horizontal streaks on the empty
part of the medium, the left side of the dish.
The streaks must always be vertical to the Streptomyces culture
and be drawn close up to the edge of it, but it must not be
touched or contamination will occur.
Keep cultures overnight.
5. Incubate the Petri dishes for two days at 30oC and keep
them in a cool place for a day longer.
6. In the interim, after incubating for 24 hours, inoculate the culture
media of the other four Petri dishes, on which the
Streptomyces cultures are now even more vigorous, with the test
organisms using an inoculation loop as in step 4.
7. Incubate the Petri dishes for two days at 30oC.
8. Compare and assess the first and second set of four plates.
Some test organisms grow in the vicinity of the Streptomyces sp.
culture and some do not.
Streptomyces griseus produces an antibiotic, streptomycin, that
diffuses into the culture medium.
Some organisms, e.g. Bacillus mycoides, Escherichia coli,
Micrococcus luteus, are sensitive, i.e. they are killed off by the
antibiotic at a certain concentration.
Other organisms are resistant, e.g. the yeast Candida utilis, are
not susceptible to streptomycin because they are eukaryotic.
The distance between sensitive test organisms and the Streptomyces
culture is larger in older streptomyces cultures because
production of antibiotic increases in the older cultures.
4.1.8 Streptomycin on Bacillus
subtilis, small disc test
The effectiveness of species of fungus to release antibiotics into the
environment can be tested by using the small disc test.
Equipment: Petri dishes: 1 paper punch, or pair of scissors, 1 pair of
tweezers, 1 Bunsen burner, 1 small glass beaker,
1 large glass beaker as a water bath, needles, corks and glass beaker or
test-tubes with rubber stoppers, filter paper, aluminium foil
126.96.36.199 Nutrient agar medium, 200
188.8.131.52 Malt extract agar medium,
Pure culture of Bacillus subtilis (DSM-No. 10, ATCC 6051) a yeast
suspension that was incubated overnight,
Streptomycin or another antibiotic
Yeast, 1 g to 100 mL water
1. Prepare nutrient agar as a bacterial culture medium and malt agar as
a yeast culture medium
2. Sterilize the culture media and cool them in a water bath to 40oC,
keep them at this temperature.
These resistant micro-organisms cannot damage your health because Streptomycin
is no longer used in medicine.
3. Add 1 mL of a culture of Bacillus subtilis or yeast suspension
(which was incubated overnight) per 200 mL to the still liquid
culture medium, close the conical flask and mix the contents vigorously.
Cooling is necessary because the hot agar would damage the organisms.
If the agar is left to cool without putting it into a water bath, it sets
4. Pour the culture medium that is inoculated in this way into Petri dishes
and allow it to set.
5. Use a hole punch or a pair of scissors to produce small discs (0 =
5 mm) from a sheet of filter paper.
6. Sterilize two small discs by placing each one on a needle and then
into 96% ethanol overnight, or by sticking them into a cork
and placing them in a glass beaker sealed with aluminium foil and then
into a drying cabinet at 135oC for three hours.
Exposure to ultraviolet light for ten minutes will also work.
7. Produce a solution of streptomycin in sterile water.
The concentration of the pure agent should be about 50 mg or mL.
8. Dip the sterile small discs into the antibiotic solution and dry them
in a drying cabinet at 10o3.
Dip at least one small disc into sterile water free of antibiotic, as a
If the damp paper discs were placed onto the agar, the agents would begin
to diffuse uncontrollably.
9. Use a sterilized pair of tweezers to place each of the dried antibiotic
plates onto an inoculated labelled agar plate.
As a control, use at least one petri dish without a small disc.
The addition of small paper control discs shows that filter paper itself
does not contain any substances that inhibit the growth
The control plate without the small paper discs shows that a completely
uniform bacterial lawn develops on an untreated plate.
10. Incubate the closed Petri dishes at 30oC for two days.
A uniform bacterial or yeast lawn should be present on all of the control
plates, and a circular zone free of bacteria should be
visible around the small discs that contained antibiotics.
The yeast is not affected by the antibiotics.
Compare the streptomycin sensitivity of microbial strains:
10.1 Bacillus subtilis
10.2 Saccharomyces cerevisiae.
The antibiotic gradually diffuses into the agar.
The diameter of the zone in which no bacteria grow is a measure of the
concentration of the agent.
If two different biocatalyst solutions of a similar concentration were
used, the diameter of the bacteria free zone.
The significance of colonies that grow within an antibiotic bacteria free
zone is that each of them is formed from one mutant of the
bacterium that is resistant to the antibiotic.
4.1.9 Presence of bactericidal
substances using a coin and Bacillus mycoides
If a coin is placed onto a culture medium that is uniformly inoculated
with Bacillus mycoides, a
bacterial lawn grows with a bacterial free zone around the coin.
The coin may consist of German silver, an alloy of copper, nickel, and
Their metal anions kill cells of Bacillus mycoides by inhibiting
growth and division.
From the side, it is obvious that the colony is more dense at the edge
of the zone than in the rest of the bacterial lawn.
The metal anions encourage growth in small quantities.
An area of resistant micro-organisms is often formed in the immediate
vicinity of the coin.
These micro-organisms can be traced to the coin and have become enriched
in the course of time.
They are resistant to ions of heavy metals.
The demonstration is therefore also indicative of the pressure of selection
that bactericides exert on a population of
This problem occurs quite frequently in hospitals, where certain micro-organisms
suddenly occur in large numbers,
e.g. the bacterium Serratia marcescens.
Inhibiting and encouraging growth of micro-organisms by the use of bactericidal
the formation of resistance mutation selection
Equipment: 1 autoclave, 1 incubator, 1 Bunsen burner, 4 disposable Petri
dishes, 1300 mL sterile conical flask, stopper,
6 culture tubes, adhesive tape for sealing the Petri dishes, 5 X 5 mL
184.108.40.206 Basal broth medium, for
cultures incubated overnight, 100 mL
220.127.116.11 Basal agar medium, 100
Distilled water 100 mL,
Pure culture of Bacillus mycoides (DSM-No. 2048, ATCC 6462)
Time needs: preparation and autoclaving of the nutrient solution: 45
minutes, preparing the
overnight culture: 15 minutes, waiting time: 48 hours
Preparation: Prepare and autoclave the basal broth medium.
Suspend again the culture of Bacillus mycoides in 1 mL of sterile
liquid basal medium according to the manufacturer's instruction,
pipette this into a test-tube previously filled with 5 nil of the sterile
liquid basal medium.
Incubate for 24 hours at 30oC, culture incubated overnight.
1. Place the basal agar medium into conical flasks, autoclave and cool
to 45oC under running water.
The approximate temperature has been reached if you can hold the warm
conical flasks to the back of your hand with no
unpleasant sensation, back-of-hand test.
2. Add the culture that was incubated overnight and mix well with the
culture medium by swirling the contents of the flask.
3. Pour the inoculated culture medium into four Petri dishes.
4. Once the agar is set, place a coin onto the surface of the culture
medium in the middle of the Petri dish.
5. Close the Petri dishes and seal them with adhesive tape.
The Petri dishes must be protected against accidental opening, and must
be sealed because micro-organisms that may grow on
the coin, and possibly on the culture medium, are unknown, any risk that
wild strains may pose are avoided in this way.
4.2.1 Prepare yoghurt, (activity
for primary grade 4 students, about 9 years old)
Yoghurt is made from milk inoculated with a mixed culture of Lactobacillus
bulgaricus and Streptococcus thermophilus, then
inoculated for hours then left to cool so that the milk proteins coagulate
at about pH 4.3.
Equipment: 1 one whisk or one wooden spoon, 20 cups or glasses, 1 oven
ring, 1 saucepan, 20 teaspoons, 1 thermometer (100oC),
incubator or insulated box made out of polystyrene foam (dimensions:
20 cm high X 35 cm long X 30 cm wide, thickness of the
polystyrene: 6 cm) or "yoghurt machine".
Materials: 3 litres of milk, 3 containers of yoghurt made from whole milk,
1. Heat the milk to 72oC to kill any harmful bacteria in the
2. Allow the milk to cool to 45oC or body temperature, 37oC.
3. Place a teaspoon, (5 mL), of unpasteurized yoghurt and lactic acid
bacteria starter into a plastic container.
4. Add the cooled milk to the container.
5. Mix all of the ingredients.
6. Cover the container with cling film.
7. Place the yoghurt mixture into an insulated box to let the contents
incubate at 45oC for 3-4 hours or overnight at room
8. After the milk has thickened, the yoghurt is ready.
It tastes acidic.
9. If yoghurt is made from whole milk, the product is smoother but, unless
homogenized, a fat layer may separate.
To avoid this separation, add 3% of skim milk powder to the fresh milk
before adding the yoghurt starter.
4.2.1a Prepare yoghurt, a
report from Turkey
Yoghurt and kefir are your national foods and the genus Pediococcus
(Lactobacillus), [family Lactobacillaceae], and yeasts
presently pose no risk, (no GMO, i.e. Genetically Modifed Organism, no
extreme food additives) since they are consumed in
high amounts in Turkey.
Therefore one of the safest ways is to suggest using these microorganisms
in the classroom.
Most secondary high schools (grades 8-12) in Turkey have laboratories
for performing experiments and when we go to
pre-service teacher candidate supervision in high school biology courses
we do the Gram staining at your research laboratory at
the university then take the prepared slides to the school.
Another word of caution could be looking for mould growth, petri dish covers
and possible contamination from oral, nasal flora.
These can be pre-checked by the biology science teacher and contaminated
Schools may afford to get small size (desktop) autoclaves and do their
own after the experiment sterilization.
From: Figen Erko, Gazi University, Department of Biology Education,
4.2.2 Prepare sauerkraut
Activity for primary grade 4 students, about 9 years old
Equipment: 1 bowl, diameter 30 cm, chopping board, Kilner jar, 2 mL, with
rubber ring, lid, and clasp, 4 kitchen knives,
1 wooden cylinder, 43 cm, or 1 egg cup
Materials: 1 large cabbage, 20 to 40 g salt
1. Cut a white cabbage into strips on a chopping board.
2. Put the chopped cabbage into a bowl, together with the salt.
3. Mix together well the cabbage and the salt.
4. Put the salted cabbage into a Kilner jar.
5. Press the cabbage together well with your fist.
6. Press a wooden cylinder or an egg cup onto the cabbage, attach the
lid, and close the Kilner jar with a clasp.
Tell the students that the cabbage must remain like this for about two
weeks, until sauerkraut has been formed.
They can taste the sauerkraut at that time.
4.2.3 Prepare lactic acid with
See diagram 20.162: Making sourdough in glass
(a) Egyptians invented sour dough bread 3,500 years ago.
They observed that dough made from rye flour can ferment and be used to
bake light piquant bread.
They could produce large quantities of sourdough from a small amount so
they always saved a small amount of dough for next time.
The souring of the rye flour is caused by consecutive fermenting of the
dough by two groups of micro-organisms: yeasts and
lactic acid bacteria.
(b) Yeasts of the genera Saccharomyces and Kluyveromyces,
together with lactic acid bacteria of the genera Lactobacillus
Lactococcus, stick to the grain and get into the flour in this
Sourdough is made by mixing rye flour with water.
The organisms take up their activity and enrich the "dough" in their substrate.
Repeat inoculation of fresh dough with this culture encourages the yeasts
to grow first and the lactic acid bacteria to grow later.
During the growth of the yeast, the volume of the dough greatly increases
and the dough smells of alcohol.
After the third inoculation, you can measure the souring of the dough,
pH 4.5. The sour dough contains mainly lactic acid bacteria.
(c) People with problems digesting gluten may benefit from eating sourdough
bread because the gluten becomes partially broken
down to make wheat and rye more digestible and more easily assimilated.
Equipment: aluminium foil, 1 measuring cylinder, 100 mL, 1 felt tip pen,
waterproof, 6 glass beakers, 400 mL, 1 shallow plastic
bowl, 15 X 30 cm, 1 set of scales, 1 spatula, 1 thermometer, 50oC,
1 wooden spoon
Materials: rye flour (type 1250), warm tap water 40oC, pH
paper (3.5 pH to 5.5 pH)
Time needs: mixing of the dough: 15 minutes, inoculating: 2 X 5 minutes
waiting time: 3 X 24 hours
1. On the first day, mix dough made from 100 g of rye flour and 100 mL
water, 40oC, with the spoon.
Put the dough into the first glass beaker (dough 1) seal the beaker with
aluminium foil and place it in a safe place at room temperature.
Step 1. Spontaneous growth of the bacteria contained in the flour requires
the addition of sufficient water of the right temperature,
40oC, and standing time.
Constant humidity and temperature also are necessary.
The lactic acid bacteria can develop their activity in the rye flour particularly
well because rye contains very little gluten protein,
in contrast to wheat.
Dough made from wheat flour only "ferments" if bakers' yeast is added to
2. On the next day, prepare another dough as described in step 1, put
it into a glass beaker (dough 2).
Mix a quarter of dough 1 (from the day before) with 75 g of rye flour and
75 mL warm tap water, 40o
3. Place this into a glass beaker (dough 3).
Seal the glass beakers (dough 2 and dough 3) with aluminium foil and place
them in a safe place.
The rest of dough 1 is no longer required and can be put on the compost
3. Three glass beakers are required on the third day.
Fill the first with fresh dough prepared as described in step 1 (dough
Place a mixture of 75 g rye flour and 75 mL water, 40oC, into
the washed plastic bowl, add 50 g of dough 2.
Place this mixture into the second glass beaker (dough 5).
Finally, place a mixture of 75 g rye flour and 75 mL water, 40oC,
into the washed plastic bowl, add 50 g of dough 3.
Place this mixture into the third glass beaker (dough 6).
Seal the three (dough 4, dough 5, dough 6) glass beakers with aluminium
foil and put them in a safe place.
So dough 2 and dough 3 are no longer required and can be discarded.
Steps 2 and 3. Fresh dough is prepared repeatedly because sourdough of
various ages should be available for comparison on
the third day.
Water must be at the right temperature because the dough being prepared
requires a specific temperature to promote the growth
of yeasts and lactic acid bacteria.
Dough 5 take up more space than dough 4 and dough 6 when the pH continuously
increases from dough 4 to dough 6 because
of the activity of the yeast cells that form carbon dioxide gas.
The growth of the yeast is reduced as the dough becomes increasingly acidic.
The lactic acid bacteria become increasingly more enriched after the dough
has been inoculated several times.
4. After three hours, measure the volume of dough 4, 5, and 6, appraise
the smell, and measure the pH with pH paper.
4.2.4 Make wine from grape
juice and vinegar from wine
See diagram 4.2.4: Alcohol production test
The types of yeast that cause alcoholic fermentation belong to the genus
Saccharomyces and can always be isolated from ripe fruit.
Nowadays, the production of wine employs strains of Saccharomyces ellipsoideus,
which is closely related to the brewers' yeast or
bakers' yeast Saccharomyces cerevisiae.
During this process, the fruit sugar is converted to ethanol and carbon
Certain bacteria, e.g. the genera Acetobacter and Gluconobacter,
can oxidize ethanol to acetic acid, via intermediate stages.
In the past, vinegar was produced at home.
An industrial procedure for the production of vinegar was developed in
the fourteenth century in the area of Orleans, France.
One part of mash and one part of fresh wine vinegar were put into wooden
casks, lying on their sides, as a "starter".
In later techniques, the vinegar bacteria were placed onto wooden lattices
or beech shavings to encourage them to expand.
These techniques, Fessel procedure, were eventually developed to such a
degree that a solution containing alcohol was dripped
onto the container that was filled with beech shavings from above, while
a counter current of air was guided over the shavings
A common method used to produce vinegar is in dilute alcohol solutions
from fermenting wine or malt infusion wash with Mycoderma
4.2.5 Prepare cider from apple
Pure culture yeasts must be used for wine making because the fermentation
of wild wine yeasts is unpredictable.
The demijohn must not be filled to the top because the carbon dioxide
produced by the fermentation of alcohol can form several
litres of foam together with the yeast cells.
These can be pushed through the air lock and out of the demijohn.
As a safety precaution, the demijohn should never be kept on a surface
that must remain clean.
During fermentation, the formation of carbon dioxide creates excess pressure
in the demijohn.
The water contained in the air lock prevents large amounts of oxygen from
entering the demijohn and encouraging the growth of
Equipment: 1 rubber tube, internal diameter 5 mm, 1 rubber stopper with
air lock, 1 household funnel, 1 demijohn, 2 litres
Materials: 250 g granulated sugar, 2 X 0.7 litre bottles of unclarified
apple juice, 1 package of wine yeast
Time needs: starting and inoculating the wine: 5 minutes, fermentation
time: 6 months
1. Place 240 g granulated sugar into the demijohn, dry funnel, add a
bottle of apple juice.
Dissolve the sugar by carefully swirling the demijohn from side to side.
2. Add the wine yeast with the second bottle and swirl it round.
3. Seal the demijohn with a rubber stopper and an air lock that is filled
4. A cloudy development in the fermentation gases is visible after three
Vigorous fermentation recedes after ten days.
5. After about six months the yeast has sunk to the base and the fresh
wine appears clear.
Use a piece of rubber tubing to siphon the wine off from the yeast.
Step 1. The alcohol content determines the life of wine to a large degree.
In Germany, table wines with an alcohol content of 8% by volume generally
have to be preserved by the addition of sulfurous
acid or potassium pyrosulfate but the wine called "port" with 15% alcohol
by volume has a disinfecting effect so it preserves itself.
Sugar must be added to achieve a high concentration of alcohol.
Cider can be made from industrially produced apple juice in an extremely
simple way, as it contains almost no pectin, but
Pectin might cause the fermenting wine to set or might prevent the deposition
of particulate matter.
Wines that contain very little acid, e.g. pear wine, often taste insipid
and do not produce the esters necessary for good bouquet.
So you do not need to clarify the wine to remove particulate matter that
is linked to the pectin or to acidify the wine artificially.
Also, you do not need to add yeast nutrient salt that contains nitrogen
because apple "must" contains sufficient nitrogen compounds.
Step 2. At the beginning of the fermentation process, the respiration
processes of micro-organisms creates negative pressure in
This must not be allowed to last for longer than three days.
If the yeast culture does not grow, the juice must be inoculated again.
4.2.6 Prepare vinegar with Acetobacter aceti
19.1.4: Prepare vinegar from wine
9.2.22 Vinegar bacteria medium
Equipment: aluminium foil, 1 glass tube, 1 aquarium pump, 1 glass tube,
right angled, 1 one way tap, right angled,
2 X 5 mL sterile pipettes, 1 conical flask, 500 mL pipette aids, 1 culture
tube, 1 rubber stopper, single bored, 2 glass bottles,
1.5 litres with stopper attachments at their bases, 1 rubber stopper, double
bored, rubber tubing, stand material
Materials: pure culture of Acetobacter aceti (DSM-No. 3508) beech
tree shavings, cotton wool, distilled water, sterilized
distilled water, 750 mL, wine (cider or unsulfured port) 250 mL, 1 M NaOH
Time needs: preparation and autoclaving of the solutions: 45 minutes,
preparing the culture: 15 minutes, waiting time: 48 hours,
constructing of fermenter and preparing main cultures: 45 minutes
Preparation: Suspend again the culture of Acetobacter aceti according
to the manufacturer's instructions, inoculate the culture
with 100 mL vinegar bacteria medium in a 300 mL conical flask.
To ensure sufficient addition of oxygen, place the flask on to a magnetic
stirrer for 48 hours.
The stirring rods should be autoclaved with the culture medium before
1. Attach a 1.5 litre bottle that has a fixture for a stopper at its
base about 40 cm above the table, using the stand.
A single bore rubber stopper with an angled, one way tap seals the lower
outlet of the bottle where the stopper is attached.
2. Attach a second bottle of this kind directly beneath the outlet of
the one way tap, or "fermenter".
Seal its lower outlet with cotton wool inside, fill its interior with beech
The beech shavings immobilize the vinegar bacteria to the fermenter.
The cotton wool should retain coarser particles that can be separated from
the wood shavings.
3. Close the lower outlet of the second bottle with a stopper that has
been bored through twice.
In one of those openings, attach a glass tube as an attachment for the
In the other, insert a right-angle glass tube as a product outlet.
4. Use an aquarium pump to blow air constantly into the inside of the
fermenter through the stopper, the cotton wool filter keeps
the system sterile.
5. As medium, use a mixture of unsulfured port and sterile distilled water
in the ratio of 1 to 1.
The pH value must be adjusted to 7.0 using 1 M NaOH.
Pour 200 mL of the medium over the beech shavings in the fermenter.
Allow the contents to stand for 48 hours.
The wine must be unsulfured so that the vinegar bacteria do not die off.
This is the case in home-made wines and is usually true of ports, as well.
The pH value of the medium must be adjusted to 7.0 so that the reduction
of the pH value because of the formation of vinegar
can be monitored.
6. Place the other 800 mL of the medium in the upper container.
Adjust the tap so that it releases one drop per 5 minutes.
7. The product is continuously caught in a 500 mL conical flask at a rate
of 1 drop per five minutes.
Test the product once a day with indicator paper to monitor the development
Air is blown into the bioreactor because without oxygen, the vinegar bacteria
The air must be filtered so that it is sterile because the air in the room
contains fungal spores that develop in the fermenter and
may cause the formation of mould on the beech shavings.
4.2.7 Microbial decomposition
of thin paper, cigarette paper
Equipment: 1 autoclave or pressure cooker, 2 glass Petri dishes, 1 large
dish that can be covered as a damp chamber, 1300 mL
conical flask with cellulose stopper, 1 drying cupboard or a Bunsen burner,
tripod, pipe clay triangle, and crucible
Materials: 80 g soil, absorbent paper (approx.
90 cm2) sterile tap water, 6 strips of cigarette paper
Time needs: sterilization of the water: 30 minutes, sterilization of
a part of the soil sample: 180 minutes, preparing the experiment
in the damp chamber: 15 minutes, waiting time: about three to four weeks
Preparation: Sterilize 100 mL tap water in a closed conical flask for
Sterilize half of the soil sample in a glass Petri dish in a drying cupboard
at 180oC for three hours, or use a Bunsen burner, tripod,
pipe clay triangle, and crucible for 30 minutes.
1. Place the unsterilized part of the soil sample into a glass Petri dish.
Dampen the sterilized soil sample with sterile tap water in the other sterile
Ensure that both of the experiments in preparation are equally damp.
The sterile soil serves as a control, no micro-organisms should grow on
the cigarette paper during the four weeks.
2. Place three strips of cigarette paper, 1 cm wide, on the dampened soil
sample in each of the Petri dishes.
For health reasons, cigarette paper does not contain lignin and is therefore
more suitable for this investigation than is filter paper,
which does contain lignin.
Lignin prevents enrichment of organisms that decompose cellulose.
3. Cover both Petri dishes, seal the edges with adhesive tape, and place
them into the larger dish, which has been covered with
dampened absorbent paper.
Cover the large dish.
The Petri dishes must be sealed so that micro-organisms do not accidentally
escape and dangerous micro-organisms do not develop.
The damp chamber prevents the soil from drying out.
4. Allow the experiment to stand in a safe place for about four weeks.
Only micro-organisms that decompose cellulose be enriched on cigarette
paper because cellulose is the only source of
carbohydrate in the paper.
micro-organism that grow on the paper also live off cellulose.
18.104.22.168 Enzyme technology,
pectinase, amylase, lactase
See diagram 9.56.1: Cell walls and membranes
Cellulases, amylases, proteases, and lipases are enzymes that are released
by cells into the environment to help breakdown the
large polymer food molecules that cannot be taken up into the cells whole.
7.1.1 Pectinase, (pectin),
Juice from oranges and lemons and from tropical fruits such as mango,
papaya, and passion fruit is concentrated in the land of
origin where the water that has been removed.
The water is replaced in the country where it is to be consumed.
However, fruit juice contains pectin so jelly usually forms when fruit
juice is concentrated.
In the juice of fleshy fruit such as papaya when water is removed, pectin
polymerizes and causes setting.
To prevent this setting the fruit juice industry adds pectinase, an enzyme
that splits pectin.
A form of pectinase can be extracted from the mould Aspergillus niger.
The enzyme amylase, which is also extracted from mould, is used in the
textile industry to remove starch from cotton.
Starch naturally adheres to cotton and inhibits the uptake of dye when
textiles are being dyed.
The baking industry mixes amylase with flour and supplements the naturally
occurring amylase in flour.
This enzyme is necessary to prepare the dough because it breaks down a
small proportion of the starch in the flour to glucose,
which serves the yeast as food.
Manufacturers of liquid and powder detergents use amylase to breakdown
the starch that forms as dirt on cutlery or in clothes.
Protease is used in washing powder to decompose protein stains and lipases
are used to break up fat stains.
Cellulases are used in the processing of fruit and vegetables to destroy
the cell walls.
Lactase works inside the organism where it decomposes lactose molecules
to α-glucose and β-galactose.
Whey contains relatively large quantities of lactose.
Many adult humans cannot breakdown lactose in the digestive tract because
they no longer produce the "infant's enzyme" lactase.
Undigested lactose removes water from the intestinal wall, which results
Bacteria in the intestinal flora that can split lactose decompose the
products of splitting, developing gas in the process, the gas
Lactase is used to decompose lactic acid and to produce glucose.
In the following investigations, the students describe the effect of pectinase,
explore the splitting of lactose, and decompose
starch with Bacillus subtilis.
A protease, (peptidase, proteinase), is an enzyme that hydrolyzing peptide
bonds in proteins and peptides.
Proteases ares used to degrade proteins, to study protease inhibitors and
to study thermal inactivation kinetics.
Proteases may be isolated from bacteria.
Alcalase is a protease from Bacillus licheniformis, Savinase is a protease
from Bacillus species.
Subtilisin is a protease from Bacillus subtilis.
Proteinase K is a protease from Tritirachium album.
Protease from Aspergillus oryzae contains both endoprotease and
A protease usually begins the hydrolytic breakdown of proteins by splitting
them into polypeptide chains.
An endopeptdase catalyzes the hydrolysis of a peptide chain within the
chain, not near either terminus, e.g. pepsin, trypsin.
An exopeptidase catalyzes the hydrolysis of the terminal amino acid of
a peptide chain, e.g. carboxypeptidase.
22.214.171.124 Enzyme Technology,
pectinase in the industrial production of juice
1. Pectins are vegetable polysaccharides, their main components are galacturonic
acid and its methyl ester.
The multiplicity of pectin is determined by the various degrees of polymerization
Together with cellulose, they are reticulum substances of vegetable cell
walls, especially as a sort of "putty"
in the middle lamella between the cells.
They occur in solution in the cell sap.
2. Pectins have a great ability to combine with water, which accounts
for the high gelling capacity of jams and jellies.
For this reason, pectin are extracted from slices of sugar beet and from
the remains of apples and lemons
that have been used for making juice.
They are then used as gelling agents in the food, cosmetic, and pharmaceutical
industries, and in medicine.
3. Pectinases destroy the pectin in the cell wall and in the plasma so
that it no longer retains juice in the chopped fruit.
Fruit can therefore be pressed more effectively, resulting a high yield
Pectinases also are used to clarify fruit juice.
Pectin retains substances that make the juice cloudy.
Once pectin has been destroyed, those substance can easily be precipitated
4.2.8 Prepare apple juice gel
Equipment: 1 glass beaker, 800 mL, 1 chopping board, 2 glass beakers,
400 mL, 1 watch glass, 1 tripod, 1 plastic bowl,
1 ceramic net, 1 piece of muslin, 1 Bunsen burner, 1 gas lighter, 1 wooden
spoon, 1 kitchen knife
Materials: 1 apple, sugar, tap water
Time needs: production of the juice: 20 minutes, production of the jelly:
1. Cut an unpeeled apple into eight equal pieces, leaving the core intact.
Place the pieces into the larger glass beaker, and just cover them with
2. Boil the mixture for ten minutes, stirring all the time.
The pieces of apple must become mushy.
Cool the coarse puree and press it through a muslin cloth into the plastic
3. Weigh the empty glass beaker in advance.
Place the juice into the small glass beaker and weigh it.
4. Add an equal amount of sugar and heat the juice again, stirring all
5. After the juice has simmered for about five minutes, do a gelling test
by observing the drops that fall from the wooden spoon.
If the drops are thick and remain on the wooden spoon, you can allow the
jelly to cool down.
Divide the jelly and pour it into two glass beakers for this purpose.
Do the gelling test by placing a little of the boiling juice onto a cold
watch glass with a wooden spoon.
If the juice gels on cooling, the boiling can be stopped.
Do not boil the juice for longer than ten minutes, otherwise it will no
4.2.9 Prepare pectinase, an
enzyme that decomposes pectin
Freshly pressed apple juice is replaced with equal parts of pure alcohol.
The pectin in the juice forms an insoluble gel with the alcohol.
Juice to which pectinase has been added does not produce gel, however,
and also deposits substances that make the juice cloudy
Equipment: 6 test-tubes with stoppers, 3 pipettes, 5 mL, 1 test-tube
stand, 2 pipette aids, 1 measuring cylinder, 1 kitchen grater,
1 glass beaker, 50 mL, 2 bowls, plastic, 1 glass beaker, 100 mL, 1 cotton
napkin, 1 glass rod
Materials: 1 apple, 5 mL 5% pectinase solution, 30 mL 96% alcohol or
Time needs: 45 minutes
1. Grate the unpeeled apple into a plastic bowl.
Squeeze the juice vigorously out of the puree over the second plastic
bowl through a napkin folded double.
Transfer the juice to a glass beaker.
A medium sized apple such as a Granny Smith produces about 50 mL of juice.
2. Pour 10 mL of the juice and 2 mL of the pectinase solution into a glass
beaker, shake the mixture.
Position the glass beaker so that it will stand completely still so that
substances that make the juice cloudy are deposited.
3. Put 5 mL of the juice and 5 mL of alcohol into a test-tube.
Close the tube with a stopper, shake it carefully twice, and let it stand.
4. Add 3 mL of pectinase solution to the remaining juice, 35 mL, stirring
Start the stop watch.
At three, six, nine and twelve minutes, pipette 5 mL of the juice out of
the glass beaker into a test-tube, mix with 5 mL alcohol,
seal, and rotate carefully twice.
Place each of the test-tubes as still as possible in the test-tube racks.
5. After each test-tube has stood for at least five minutes, swirl it
carefully to see whether the flocculation remains as a clump of
gel on the surface or whether they collect as loose components at the bottom
of the test-tube.
4.2.10 Enzyme technology,
industrial uses of pectinase
In this investigation, the students find the amount of juice produced
from apple mash with and without the addition of pectinase.
The fact that pectinase increases the juice yield indicates the significance
of the use of pectinase for the fruit juice.
Pectinase is used to increase yield, clarify juice and reduce transport
Trade terms include "naturally unclarified," "clear," concentrated," and
However, some juice is still produced by the normal pressing technique.
Reasons for the use of pectinase include increased yield, energy savings
because the juice is easier to press, more economic
methods of transport, and the ability to transport juice over longer distances.
Reasons against the use of pectinase include interference with the natural
flavour and consistency of the juice, and less wild fruit
is processed, inability of small cider companies to compete with producers
of cheap juices.
Equipment: 2 tea strainers, 2 funnels, 1 kitchen grater, 1 plastic bowl,
2 glass beakers, 100 mL, 2 glass rods, 2 spoons, 2 stands,
sleeves, and stand clamps, 2 measuring cylinders, 50 mL, 1 set of scales
Materials: 2 apples, 10 mL pectinase solution, freshly made, 5%, tap water
Time needs 25 minutes
1. Grate both unpeeled apples over a plastic bowl, using a household grater.
2. Divide the apple mash equally between the two glass beakers, A and
B, with the help of the scales.
3. Pipette 10 mL pectinase solution into glass beaker A, and 10 mL water
into glass beaker 2.
Allow the glass beakers to stand as they are for ten minutes.
Stir the mash at one minute intervals, using glass rods.
4. In the meantime, attach the funnels to the stands, using clamps.
Place a tea strainer into each funnel and place the measuring cylinders
under the funnels.
Do not forget to stir the mash!
5. After the time has elapsed, tip out of the glass beakers the apple
mash A and B into tea strainers.
You may need spoons to do this.
The mash may not be pressed into the tea strainer.
6. After five minutes, measure the quantity of juice in the cylinders.
4.2.11 Split lactose from
milk or whey using immobilized lactase
See diagram 20.190: Split lactose
See 126.96.36.199: Buffer reagent, phosphate
Enzymes that are not released to the environment but that are active in
the inside of cells are formed by micro-organisms in
relatively small quantities.
The industrial production of such enzymes, of which lactase is one, is
also quite tedious.
The cells must first be broken open before enzymes of this kind can get
into the culture medium from which they are produced.
Dairies that use lactase for the treatment of whey therefore treat the
expensive lactase with due care.
It is immobilized before use, that is, it is bound to a vehicle.
This allows several consecutive uses of the enzyme because it does not
have to be thrown away with the waste products after it
has been used the first time.
The opposite is true of amylase, which is used in washing powder, this
enzyme is naturally active outside the cell.
Immobilized lactase can be used as often as desired for school experiments.
It can be preserved with isopropanol and kept for six months in the refrigerator.
Equipment: 1 filter tube (Duran, pore size 40 to 100 mu, 20 mm) 1 suction
flask with rubber stopper attachment, 1 Woulfe bottle,
3 conical flasks, 500 mL, 1 water jet vacuum pump, 1 measuring cylinder,
100 mL, 1 glass beaker, 100 mL, 2 Pasteur pipettes,
2 glass beakers, 50 mL, 3 rubber caps for Pasteur pipettes, 1 conical flask,
100 mL, 1 pipette, 10 mL, with pipette aids,
3 conical flasks, 300 mL, 1 stand with 2 clamps and 2 nuts
Materials: 1 piece of tubing, aluminium foil, 5 mL isopropanol lactase,
sugar test strips, e.g. Diabur to Test 500, BOEHRINGER,
Eupergit C, e.g. ROHM PHARMA Ltd whey from health food shop or skimmed
UHT milk, 20 mL 1 molar phosphate buffer
reagent, 150 mL 0.1 molar phosphate buffer reagent Time needs: Immobilization
of the lactase: 10 minutes, waiting time: 2 days,
splitting of lactose: 25 minutes.
1. Several days before conducting the investigation, immobilize the enzyme
lactase as follows: dissolve 0.1 g lactase in 20 mL 1
molar phosphate buffer reagent in a 100 mL glass beaker.
Add 1 g eupergit C to this solution. Shake the suspension for a short while.
Finally, seal the glass beaker with aluminium foil and allow it to stand
for at least 2 days at 20oC, room temperature.
Shake the beaker now and again about 2 to 3 times daily to facilitate the
immobilization of lactase in eupergit.
2. After two days, place the suspension in the filter tube and place the
tube onto the suction flask.
Attach both to a stand and connect them to the Woulfe bottle with a piece
of rubber tubing attached to a water jet vacuum pump.
3. Rinse the suspension in the filter tube with 40 mL 0.1 molar phosphate
buffer reagent by rinsing it several times and removing
the liquid by suction.
4. Finally, remove the suction flask.
Place a 50 mL beaker under the glass beaker.
5. Add skimmed milk or whey drop by drop, using a Pasteur pipette. There
should be a surplus 1 to 3 cm high above the eupergit.
The milk products that drip out of the filter tube are caught in a glass
6. Test the milk products that have dripped through for glucose. The presence
of glucose can be ascertained using glucose test
strips that can be purchased from a supplier.
The "untreated milk" or whey can be used as a control.
7. After the experiment has been completed, purify the immobilized enzyme
with about 100 mL 0.1 molar phosphate buffer
reagent until the filtrate is clear.
In the last rinse, add 2% isopropanol by volume (preservation buffer reagent)
to the phosphate buffer reagent to preserve the enzyme.
8. Seal the lower end of the filter tube with a rubber cap that is pushed
over the end.
9. Add a preservation buffer reagent that contains isopropanol to the
eupergit lactase compound until there is a surplus of 1 to 2 cm 10.
Close the filter tube with aluminium foil at the upper end and keep the
tube in the refrigerator.
4.3.1 Grow African violet with
in vitro culture
See diagram 20.194 Construct a sterile tunnel
Saintpaulia ionantha, African violet, Usambara violet, Gesneriaceae
Numerous shoots develop from pieces of shoot or leaf of the Usambara violet
after 2 to 4 weeks if the pieces are placed onto a
medium containing cytokinin.
Prepare about 50 pieces from a piece of leaf 0.5 cm2.
If the shoots are then transferred to a medium that does not contain hormones,
it produces roots after about one week.
The small plants can be cultivated further in plant pots.
All of them produce flowers of the same colour and otherwise possess similar
Here the students experience the conspicuous production of clones.
The work must be done in sterile conditions or other micro-organisms might
be produced that would overrun the pieces of plant
tissue in a very short time.
Equipment: blowtorch, up to 600oC, with jet, paint stripper
blow torch, 2 screw clamps, 1 wooden lath 1 cm X 1 cm X 50 cm
Materials: plexiglass 30 X 45 cm, 3 mm thick
1. File down the sharp edges of the plexiglass plate.
Mark points A, B and C, D.
Put the piece of plexiglass on a wooden table so that line A-B is exactly
on the edge of the table.
Place the wooden lath onto the plexiglass exactly on the edge of the table,
secure the lath on both sides with screw clamps so
that the plexiglass is between them.
Heat the A-B line with the blowtorch until the plexiglass softens.
After 1 minute at about 600oC, bend the plexiglass that juts
out beyond the A-B line upwards at the desired angle.
Hold the plexiglass until it cools.
2. Bend the plexiglass along the C-D line.
To achieve the desk form of the tunnel, the and angles should be 90o
and 110o, respectively.
Several sterile tunnels can be piled on top of one another.
4.3.2 Grow African violet,
(Usambara violet), (Saintpaulia ionantha) from pieces of leaf
See diagram 20.197: Cultivate a tissue culture
in a sterile tunnel
See 9.2.23: MS agar medium | See 9.2.24: BAP medium
| See 9.2.25: Buffer reagent, phosphate
buffer reagent | See 9.2.26: 20% Domestos
solution 10 mL
70% alcohol, 100 mL, (or use denatured alcohol); 96 96% alcohol, 100 mL,
(or use denatured alcohol); sterile tap water 100 mL
Before tissue cultures are prepared, prepare Petri dishes, using MS agar
medium as a culture medium.
Before this is done, add BAP medium in the ratio of 0.5 mL per litre of
(BAP: 6-benzylaminopurine, a cytokinin).
Pour the medium into sterile, disposable Petri dishes (diameter 9 cm)
while it is still hot (> 50oC).
One litre is sufficient for 35 dishes.
Stack the dishes to avoid the formation of condensation in the lids of
the Petri dishes and to protect the surface of the table.
Label five empty Petri dishes with the date and type of medium and pile
the dishes on top of one another.
Lift up the whole pile with the lid of the lowest dish, pour the medium
into the lowest dish, cover it with the lid and the rest of the pile.
Lift the lid of the next dish, together with the rest of the pile, place
the medium into the next dish, and so on.
After one week, place pieces of the leaf onto the sterile culture media.
Plates that should be kept for longer periods of time are packed in cling
film or plastic bags to prevent their drying out and to protect them from
Equipment: 1 bent pair of tweezers (sterile) 1 scalpel (sterile), 1 kitchen
timer, 1 container for decanting liquids, 4 glass beakers,
200 mL, sealed by a glass Petri dish (sterile) 1 sterile tunnel, 1 Bunsen
burner, adhesive tape, transparent freezer bags, wooden sticks ca.
1. Rinse a leaf of an Usambara violet in 70% alcohol in a sterile glass
beaker for about 1 minute.
The glass lid must only be opened for as short a time as possible and
must be replaced immediately!
Carefully decant the alcohol without removing the lid so that the objects
do not slip out.
The rinsing of the leaf increases the wetness of the surface.
2. Add dilute "Domestos" solution and shake the glass beaker.
Sterilization time: 1 to 2 minutes.
Decant the solution as in 1.
3. Rinse the leaf in sterile tap water three times per 5 to 10 minutes,
shake slightly with the lid closed.
Carefully decant the last water used for rinsing.
4. Sterilize tweezers in 96% alcohol then use them to take the leaf out
and place it on an empty sterile Petri dish.
5. Cut away the tissue at the edge of the leaf that was damaged during
the process of sterilization.
Also, use the scalpel, sterilized in 96% alcohol to cut away the larger
micro-organisms that were not killed during the sterilization of the surface
may be present in the vascular tissue.
6. The freezer bag is waterproof, but porous to air.
The wooden stick prevents the plastic of the bag from pressing on the
7. Cut the leaf into strips about 5 mm wide and 1 cm long.
8. Gently press the strips of leaf onto the culture medium that contains
Close the Petri dish and seal it with adhesive tape.
Place the Petri dish into a well lighted place for about 2 to 4 weeks at
9. When shoots form, transfer them to culture medium without cytokinin
so that they form roots.
10. After two weeks, as soon as small roots have been formed, transfer
the plants to plant pots.
These in turn are placed into freezer bags that are tied at the top.
Place a small wooden stick into the soil.
4.3.3 Grow Gerbera using
in vitro culture
Gerbera jamesonii, gerbera, Barberton daisy, Transvaal daisy, Asteraceae
How much profit does a gardener make if she plants 1 000 plants that have
been produced in vitro?
How much profit does she make if she sows seedlings?
The gardener must buy both the young plants that have been produced in
vitro and the seed.
The Gerbera seeds do not germinate very well so she must buy an
average of 1 430 seeds if she wishes to grow 1 000 young plants.
While they are being grown, there are losses, and a number of young plants
do not blossom, so the gardener is only able to sell 700
of the 1 000 young plants grown from seed.
The losses made from the young plants grown in vitro were less, 950 of
1 000 young plants could finally be sold as pot plants.
"Overheads" refers to the financial cost of the required area in the greenhouse
multiplied by the number of days during which this area is occupied by plants.
The plants that have been produced in vitro can be compared to seedlings
that are seven weeks old.
The former come into flower within one to three weeks, while eight weeks
elapse between the flowering of the first and last plants produced from
So the overhead for the plants produced in vitro is considerably less.
Gerbera plants produced in vitro result in higher profit is higher
than if the plants were sown from seed.
The experiments in this document were devised by
an international group of science educators and tested with students from
schools in Germany.
However, some or all of the experiments may be illegal in your country.
Before planning to teach any of the experiments below, you must get permission
from the head of your school science department,
the principal or head teacher of your school, and the Ministry of Education
in your country.
Also, check that you can follow the safety precautions below.
Do not attempt any of the experiments below, apart from J1 or J2, if you
have no experience of teaching biotechnology.
Comment: The biosafety advice given to schools
in Germany, USA and the UK is significantly different in some aspects to
guidelines and legislation that apply in Australia for working with microbiological
(including bacteria, protozoa, fungi or yeast and mould) and genetically
In Australia, see the following:
1. Australian or New Zealand Standard, Safety in laboratories, Part 3:
Microbiological aspects and containment facilities (AS or NZS 2243.3:2002).
2. Gene Technology Act 2000, passed by the Federal Government In December
The legislation came into force on 21 June 2001.
The legislation is the Commonwealth's component of a new national scheme
for the regulation of genetically modified organisms,
(GMOs), which will include legislation in every Australian jurisdiction.