UNBiology4

School Science Lessons
Microbiology
2012-01-31 SP
Please send comments to: J.Elfick@uq.edu.au

Microbiology experiments
Table of contents
Warning and Comment
4.2.0 Fermentation food preparations
4.3.0 In vitro culture techniques
4.3.4.0 Micro-organisms
4.1.0 Microbiology cultures
9.1.2.3 Microbiology media and solutions
2.0 Microbiology safety
9.1.2.1 Microbiology stains
3.0 Microbiology techniques 1
9.1.2.0 Microbiology techniques 2

3.0 Microbiology techniques 1
3.1.10 Colony counts using the calibrated drop method
3.1.3 Flaming and cotton wool plugs
3.1.11 Incubators
3.1.6 Inoculate with a Pasteur pipette
3.1.2 Pipettes
3.1.7 Pour a plate
3.1.5 Prepare a pour plate
3.1.9 Prepare a spread plate, lawn plate
3.1.4 Prepare a streak plate
3.1.8 Spreaders
3.1.1 Wire loops

9.1.2.0 Microbiology techniques 2
9.1.2.3 Aseptic transfer of bacterial cultures from a bottle or tube
9.1.2.4 Aseptic transfer of bacterial cultures from a culture plate
9.1.2.8 Lawn plate technique
9.1.2.5 Prepare heat-fixed stained bacterial smear
9.1.2.1 Prepare stains
9.1.2.10 Serial decimal dilution of a bacterial suspension
9.1.2.9 Spread plate technique
9.1.2.2 Staining rack
9.1.2.7 Streak plate dilution method for pure cultures from a mixed suspension

4.1.0 Microbiology cultures | Acknowledgements
[based on "Teaching Biotechnology in Schools", UNESCO / International Union of Biological Sciences, (IUBS), Germany / USA]
4.1.1 Colonies of different micro-organisms: fungi, yeasts, protists and Streptomyces bacteria
4.2.0 Fermentation processes in food production
4.1.2 Enrichment of wild yeast strains
4.3.0 In vitro culture techniques
4.1.3 Prepare fixed slide preparations
4.1.4 Prepare India ink (China ink) preparations
4.1.7 Prepare streptomycin using Streptomyces griseus
4.1.9 Presence of bactericidal substances using a coin and Bacillus mycoides
4.1.5 Safe microscopy of Penicillium camemberti and Mucor mucedo using the Petri slide technique
4.1.8 Streptomycin on Bacillus subtilis and Saccharomyces cerevisiae using the small disc test
4.1.6 Soil bacteria that decompose urea

4.2.0 Fermentation food preparations
4.2.10 Enzyme technology, industrial uses of pectinase
4.2.7.1 Enzyme technology, pectinase, amylase, protease, lipase, lactase
7.1.1 Pectinase, (pectin)
7.1.2 Amylase
7.1.3 Lactase
4.2.7.2 Enzyme technology, pectinase in the industrial production of juice
4.2.7 Microbial decomposition of thin paper, cigarette paper
4.2.8 Prepare apple juice gel when it is boiled
4.2.5 Prepare cider from apple juice
4.2.3 Prepare lactic acid in sourdough
4.2.9 Prepare pectinase, an enzyme that decomposes pectin
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 quality
4.2.2 Prepare sauerkraut
4.2.4 Prepare wine from grape juice and make vinegar from wine
4.2.11 Split lactose from milk or whey using immobilized lactase

4.3.0 In vitro culture techniques
4.3.1 Grow African violet, (Usambara violet), (Saintpaulia ionantha) with in vitro culture
4.3.2 Grow African violet, (Usambara violet), (Saintpaulia ionantha) from pieces of leaf
4.3.3 Grow Gerbera using in vitro culture

3.1.1 Wire loops
Sterilize wire loops by heating in a Bunsen 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 the air. Sterilize the wire loop again immediately after use.

3.1.2 Pipettes
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 plugs
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 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 test-tube.

3.1.4 Prepare a streak plate
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 yeasts 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 90^o 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 90^o 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 90^o 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, cause colonies to spread into each other.
Professional microbiologists start with the Petri dish inverted on the desk. Then they lift out the base, invert it, then inoculate the agar facing up.

3.1.5 Prepare a pour plate
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 the medium. 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 plate. 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 a Pasteur pipette
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 hand. 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 a plate
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.

3.1.8 Spreaders
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 60^oC 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, lawn 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 oil.

3.1.11 Incubators
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 hazard. 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 micro-organisms: fungi, yeasts, protists and Streptomyces bacteria
See diagram 20.114:
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. 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
Materials:
9.1.2.14 Basal agar medium, 100 mL
9.1.2.15 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)
Time needs:
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 tip pen.
4. Inoculate each of the 4 sectors at one point with one of the 4 types of micro-organism.
5. Incubate the Petri dishes at 30^oC for 6 days.
6. Identify the colonies.

4.1.2 Enrichment of wild yeast strains
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 mould. 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.
Equipment: Petri dishes
Materials:
9.1.2.17 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 30^oC. 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
Materials: Ethanol
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 slide. 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 45^o 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 decreases evenly.
4. Push the coverslip evenly across the entire surface of the microscope slide. This spreads the suspension across the slide, and the film of liquid becomes thinner.
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 method. 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 per second.

4.1.4 Prepare India ink preparation
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 of water.
4. Place a coverslip at a 45^oangle 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 slide. 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 X magnification. 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
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 chamber. 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 suppliers. 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 Pasteur pipette, sterile, 1 conical flask, 300 mL, wide necked, 1 inoculation needle, household aluminium foil
Materials:
9.1.2.14 Basal agar medium 200 mL or
9.1.2.16 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 180^oC 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 hole. 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 culture medium.
3. Remove the Pasteur pipette from the chamber and replace the lid of the chamber. 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 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 light. 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).

Notes:
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 urea
See diagram 20.120 (5): Proper use of Drigalski spatula
This experiment shows the importance of soil bacteria as decomposers of urea.
Equipment:
9.1.2.14 Basal agar medium, 10 plates
9.1.2.21 Urea agar medium, 5 plates,
9.1.2.27 Ringer solution, sterile, 500 mL
9.1.2.28 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, 10^-4.
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 spatula.
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^-1to the test-tube labelled 10^-2, mix carefully again. Distribute one drop of the 10^-1suspension 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 30^oC.
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 one drop. 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 or 10^-4. Divide by the volume (V) of drops: N = 1.86 x 10^-6 cells or 0.06 mL = 3.1 x 10^-7cells per mL. As with yeast, 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 for biosynthesis. 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 bungs, 1 inoculating loop, 6 culture tubes, 5 x 5 mL sterile pipettes, pipette aid
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 30^oC, 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 30^oC.
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 sp. 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 sp. 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 30^oC 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 30^o.
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 using the 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 bungs, filter paper, aluminium foil
Materials:
9.1.2.19 Nutrient agar medium, 200 mL
9.1.2.17 Malt extract agar medium, 200 mL
Ethanol
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 40^oC, 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 too quickly.
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 135^oC 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 10^o3. Dip at least one small disc into sterile water free of antibiotic, as a control. 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 of bacteria. 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 30^oC 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 zinc. 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 micro-organisms. 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 substances. the formation of resistance mutation selection
Equipment: 1 autoclave, 1 incubator, 1 Bunsen burner, 4 disposable Petri dishes, 1300 mL sterile conical flask, bung, 6 culture tubes, adhesive tape for sealing the Petri dishes, 5 x 5 mL sterile pipettes
Material:
9.1.2.15 Basal broth medium, for cultures incubated overnight, 100 mL
9.1.2.14 Basal agar medium, 100 mL
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 30^oC, culture incubated overnight.
1. Place the basal agar medium into conical flasks, autoclave and cool to 45^oC 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 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 balloon whisk or one wooden spoon, 20 cups or glasses, 1 oven ring, 1 saucepan, 20 teaspoons, 1 thermometer (100^oC) 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 beakers of yoghurt made from whole milk, cling film
1. Heat the milk to 72^oC to kill any harmful bacteria in the milk.
2. Allow the milk to cool to 45^oC.
3. Place a teaspoon of yoghurt and lactic acid bacteria into a beaker.
4. Add the cooled milk to the beaker.
5. Mix all of the ingredients.
6. Cover the beaker with cling film.
7. Place the yoghurt mixture into an insulated box.
8. After several hours the milk has thickened. The yoghurt is ready. It tastes acidic.

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 specimens discarded. Schools may afford to get small size (desktop) autoclaves and do their own after the experiment sterilization.
From: Prof. Dr. Figen Erko, Gazi University, Department of Biology Education, Ankara, Turkey

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 in sourdough
See diagram 20.162: Making sourdough in glass beakers
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.
Yeasts of the genera Saccharomyces and Kluyveromyces, together with lactic acid bacteria of the genera Lactobacillus and Lactococcus, stick to the grain and get into the flour in this way. 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. People with problems digesting gluten my benefit from eating dourdough 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, 50^oC, 1 wooden spoon
Materials: rye flour (type 1250), warm tap water 40^oC, 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, 40^oC, 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, 40^oC, 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 it.
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, 40^o3. 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 heap.
3. Three glass beakers are required on the third day. Fill the first with fresh dough prepared as described in step 1 (dough 4). Place a mixture of 75 g rye flour and 75 mL water, 40^oC, into the plastic bowl which has been washed, 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, 40^oC, into the plastic bowl which has been washed, 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 Prepare wine from grape juice and make 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 dioxide. 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, which were 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 from below.
A common method used to produce vinegar is in dilute alcohol solutions from fermenting wine or malt infusion wash with Mycoderma aceti (Ascomycota).

4.2.5 Prepare cider from apple juice
Pure culture yeasts must be used for wine making because the fermentation of wild wine yeasts is unpredictable.
Disinfecting effect
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 vinegar bacteria.
Equipment: 1 rubber tube, internal diameter 5 mm, 1 rubber bung 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 bung and an air lock that is filled with water.
4. A cloudy development in the fermentation gases is visible after three days. 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 sufficient acid. 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 the demijohn. 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
See 9.1.2.22: Vinegar bacteria medium
Equipment: aluminium foil, 1 glass tube, 1 aquarium pump, 1 glass tube, right angled, 1 one way tap, right angled, 2 pipettes, 5 mL, sterile, 1 conical flask, 500 mL pipette aids, 1 culture tube, 1 rubber bung, single bored, 2 glass bottles, 1.5 litres with stopper attachments at their bases, 1 rubber bung, 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 use.
1. Attach a 1.5 litre bottle that has a fixture for a bung at its base about 40 cm above the table, using the stand. A single bore rubber bung with an angled, one way tap seals the lower outlet of the bottle where the bung 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 shavings. 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 bung that has been bored through twice. In one of those openings, attach a glass tube as an attachment for the aquarium pump. 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 bung, 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 11. 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 of acid. Air is blown into the bioreactor because without oxygen, the vinegar bacteria would die. 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 bung, 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 30 minutes. Sterilize half of the soil sample in a glass Petri dish in a drying cupboard at 180^oC 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 Petri dish. 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.

4.2.7.1 Enzyme Technology, pectinase, amylase, protease, lipase, lactase
See diagram 9.56.1: Cell walls and membranes
Cellulases, arnylases, 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.

7.1.2 Amylase
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.

7.1.3 Lactase
Lactase works inside the organism where it decomposes lactose molecules to alpha glucose and beta 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 in diarrhoea. Bacteria in the intestinal flora that can split lactose decompose the products of splitting, developing gas in the process, the gas causes flatulence. 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.

4.2.7.2 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 and esterification. 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 of juice. 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 out.

4.2.8 Prepare apple juice gel when it is boiled
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: 25 minutes
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 tap water.
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 bowl.
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 the time.
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. The gelling test can also be carried out 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 longer gel.

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 bungs, 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 denatured alcohol
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 bung, shake it carefully twice, and let it stand.
4. Add 3 mL of pectinase solution to the remaining juice, 35 mL, stirring constantly. 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 costs. Trade terms include "naturally unclarified," "clear," concentrated," and "concentrate." 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 9.1.2.25: Buffer reagent, phosphate buffer reagent
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 bung 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 20^oC, 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 beaker.
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, (Usambara violet), (Saintpaulia ionantha) with in vitro culture
See diagram 20.194 Construct a sterile tunnel from Plexiglas
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 characteristics. Here the students experience the conspicuous production of clones. The work must be carried out 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 600^oC, 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 600^oC, 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 90^o and 110^o, 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
Materials:
See 9.1.2.23: MS agar medium | See 9.1.2.24: BAP medium | See 9.1.2.25: Buffer reagent, phosphate buffer reagent | See 9.1.2.26: 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 culture medium. (BAP: 6-benzylaminopurine, a cytokinin). Pour the medium into sterile, disposable Petri dishes (diameter 9 cm) while it is still hot (> 50^oC). 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 contamination.
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. 10 cm
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 vascular tissue. 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 plants.
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 cytokinin. 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 room temperature.
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
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 seed. 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.

WARNING!
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, you should 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 the guidelines and legislation that apply in Australia for working with microbiological organisms (including bacteria, protozoa, fungi or yeast and mould) and genetically modified organisms.
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 2000. 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. Copies of the Office of the Gene Technology Regulator's Handbook may be obtained from the OGTR.