appendixB

School Science Lessons
Appendix B, Biology
2012-02-06
Please send comments to: J.Elfick@uq.edu.au

Table of contents
4.0 Biology fixatives
1.0 Biology media and solutions
6.0 Culture media for routine cultivation and identification of fungi
9.0 Glues and pastes, adhesives
4.7 Insect fixing fluids
2.0 Microscopy adhesives
3.0 Microscopy stains
Plants, common names first
Plants, Dicotyledons
Plants, Family names, dicotyledons
Plants, Monocotyledons
7.0 Prepare acids and bases
8.0 Prepare salt solutions
5.0 Standard buffer solutions

1.0 Biology media and solutions
1.1 Acid alcohol, biology solution
1.2 Alcohol (ethanol) absolute alcohol, biology solution
7.1 Andrade's acid fuchsin indicator
9.1.2.24 BAP medium
9.1.2.14 Basal agar medium
9.1.2.15 Basal broth medium
1.3 Benedict's reagent, Benedict's solution, biology solution
9.1.2.25 Buffer reagent, phosphate buffer reagent
1.4 Carbol xylol, biology solution
9.1.2.26 Domestos solution, 20%
1.5 Fluorescein solution, biology solution
9.1.2.16 Glucose nutrient agar
9.1.2.19e Glucose nutrient agar medium
3.12 Gram's iodine solution, microscopy stain
1.6 Iodine solution, biology solution, tests for starch, tests for lignin
9.1.2.12 Liquid broth media
3.15 Lugol's iodine solution, microscopy stain
9.1.2.17 Malt extract agar medium
9.1.2.17a Malt extract broth medium
9.1.2.19d Mannitol yeast extract agar (MYEA)
9.1.2.11 Media or solutions, Prepare sterile media or solutions
3.26 Weigert's haematoxylin, Weigert's iron haematoxylin, microscopy stain
3.27 Methyl cellulose, methocel (low substitution)
9.1.2.29 Microbiology chemicals
9.1.2.19b Milk agar medium
9.1.2.18 Minimal agar medium
9.1.2.23 MS agar medium
9.1.2.19c Nitrogen-free mineral salts agar medium
9.1.2.19 Nutrient agar medium
9.1.2.20 Nutrient broth medium
1.13 Phenylthiocarbamide, PTC, phenylthiourea, PTU, ₇H₈N₂S
9.1.2.27 Ringer solution
1.7 Ringer solution, biology solution
1.8 Saline solution, biology solution
1.9 Scott's blueing solution, biology solution
1.10 Sea water substitute, biology solution
1.11 Sodium thiosulfate solution, biology solution
9.1.2.28 Salt solution
9.1.2.19a Starch nutrient agar medium
9.1.2.13 Sterile solutions
9.1.2.30 Tensides
9.1.2.21 Urea agar medium
9.1.2.22 Vinegar bacteria medium
1.12 Xylene and methylbenzoate, biology solution

9.0.0 Glues and pastes, adhesives
3.100 Casein adhesive, Prepare plastic from milk casein, (See 1.)
1.3 Adhesives, List of adhesives
9.1.0 Flour and milk glues
9.1.1 Flour glue
9.1.2 Milk glue
9.1.3 Wallpaper paste
9.2.0 Commercial glues

2.0 Microscopy adhesives
2.1 Glycerine jelly, adhesive to stick sections to microscope slides
2.2 Haupt's adhesive, adhesive to stick sections to microscope slides
2.3 Meyer's albumen, adhesive to stick sections to microscope slides
2.4 Canada balsam

3.0 Microscopy stains
3.1 Acetic alcohol, microscopy stain
3.2 Aceto-orcein stain, microscopy stain
3.3 Acetocarmine, microscopy stain
3.4 Aniline hydrochloride, microscopy stain
3.5 Aniline sulfate, microscopy stain
3.5.1 Carbol fuchsin, microscopy stain
3.6 Carmine stain, microscopy stain
3.7 Congo red, microscopy stain
3.8 DCIP (2,6-dichlorophenol-indophenol), microscopy stain
3.9 Delafield's haematoxylin, microscopy stain
3.10 Eosin, tetrabrornofluorescein, microscopy stain
3.11 Gram stain, microscopy stain
3.11.1 Crystal violet microscopy stain
3.11.2 Gram stain decolorizing solution
3.12 Gram's iodine solution, microscopy stain
3.13 Haematoxylin, microscopy stain
3.14 Heidenhain iron haematoxylin, microscopy stain
3.15 Lugol's iodine solution, microscopy stain
3.16 Karo syrup mountant, microscopy stain
3.17 Lactophenol, microscopy stain
3.18 Lactophenol cotton blue, microscopy stain
3.19 Leishmann's stain, microscopy stain for white blood cells, Wright's stain
3.19a Methylene blue, microscopy stain
5.6.5.1 Methyl violet, microscopy stain
3.20 Neutral red (0.1%), microscopy stain
3.21 Orange G, microscopy stain
3.22 Phloroglucinol (10% solution), microscopy stain
3.23 Safranin, microscopy stain
3.24 Schulze's solution (chlor-zinc-iodine), microscopy stain
3.25 Toluidine blue, microscopy stain
3.26 Weigert's haematoxylin, microscopy stain
9.1.2.1.6 Resazurin stain, microscopy stain

4.0 Biology fixatives
4.1 Aceto-alcohol, biology fixative, for plant material
4.2 Ethanol solution
4.3 FAA (formalin, acetic acid, alcohol), biology fixative, for plant material
4.4 Formaldehyde, formalin, biology fixative, for animal material
4.5 Formol-saline, biology fixative, for marine animals
4.6 Zenker's fluid, biology fixative, for animal material
4.7 CRAF (chromic acid, acetic acid, formalin), biology fixative, for plant material

4.7.0 Insect fixing fluids
4.15 Barber's relaxing fluid, insect fixing fluid
4.8 Carnoy's fluid, insect fixing fluid
4.9 KAA, insect fixing fluid
4.10 Kahle's fluid, insect fixing fluid
4.12 Lacto-alcohol, insect fixing fluid
4.11 Oudeman's fluid, insect fixing fluid
4.13 Pampl's fluid, insect fixing fluid
4.14 Sugaring mixture, insect fixing fluid

6.0 Culture media for routine cultivation and identification of fungi
6.1 Calcofluor white with 10% KOH, to identify fungi
6.2 Cellotape flag preparations, to identify fungi
6.3 Cornmeal agar, to identify fungi
6.4 Cornmeal glucose sucrose yeast extract agar, to identify fungi
6.5 Czapek Dox Agar, to identify fungi
6.0 Direct microscopic mounts or squash preparations
6.6 Indian ink mounts
6.7 Lactophenol cotton blue (LPCB), to identify fungi
6.8 Malt extract agar, to identify fungi
6.12 Orcinol-Bial's Reagent
6.9 Potassium hydroxide with chlorazol black, to identify fungi
6.10 Potato dextrose agar, to identify fungi
6.11 Rice grain slopes, to identify fungi

1.1 Acid alcohol, biology solution
100 mL 70% alcohol and 1 mL hydrochloric acid, used for cleaning slides and coverslips and decolorizing some stains, e.g. haematoxylin.

1.2 Alcohol (ethanol), biology solution
It is used in various concentrations for preserving and dehydrating. Absolute alcohol bottles must be stoppered at all times, since alcohol readily absorbs water from the atmosphere. Remove water from alcohol by using drying agents, e.g. anhydrous copper (II) sulfate.

1.3 Benedict's reagent, Benedict's solution
1. Dissolve 173 g of sodium citrate and 100 g of anhydrous sodium carbonate in 800 mL of water then filter the solution. Dissolve 17.3 g of copper sulfate in 100 mL of water and add this to the filtered solution.. Make up to 1 L with water.
2. Solution A: Dissolve with heat 173 g sodium citrate, 100 g anhydrous sodium carbonate, Na₂CO₃, in 800 mL water. Filter and dilute to 850 mL.
Solution B Dissolve 17.3 g copper sulfate crystals, CuSO₄.5H₂O in 100 mL water.
Pour Solution B with stirring, into Solution A, and make up to 1 litre.
3. Solution A: Dissolve 17.3 g of sodium citrate and 10 g of anhydrous sodium carbonate in 60 mL of water.
Solution B: Dissolve 1.73 g of copper sulfate crystals in 20 mL of water.
Pour solution B into solution A while stirring, and make up to 1 litre.

1.4 Carbol xylol, biology solution
BE CAREFUL! Use gloves!
Mix 25 g phenol with 100 mL xylene. The phenol dissolves slowly, and cools when dissolving. It is used for dehydrating in staining techniques.

1.5 Fluorescein solution
Dissolve one gram of fluorescein in 100 mL methylated spirit.

1.6 Iodine solution, biology solution, tests for starch, tests for lignin
See 9.132: Tests for starch, iodine tests for starch
Store iodine container inside a second container because the lid of iodine container may deteriorate. Iodine is scarcely soluble in water, so iodine solution is iodine dissolved in potassium iodide solution. Use safety glasses and nitrile chemical-resistant gloves when weighing solid iodine because it is harmful and corrosive. Once in solution the amounts of iodine used are minute. Specific tests for starch giving blue-black colour and general stain. Kills and fixes living material and makes cytoplasm and nucleus more visible. Lignin walls in xylem stained brown, leaving cellulose walls of parenchyma relatively unstained. Stains proteins brown, nuclei dark brown, chloroplasts brown but black if starch present, cellulose cell walls yellow, lignified cell walls deep yellow to brown.
1. Dissolve 2 g potassium iodide in water. Add 1 g iodine crystals.
2. Dissolve 1 g iodine and 4 g potassium iodide in 300 mL water.
3. Add l g iodine crystals and 5 g potassium iodide to 50 mL water. Dilute to 100 mL
4. Dissolve 10 g of potassium iodide in 100 mL of deionized water and add 5 g of iodine crystals.
5. Dissolve 15 g of potassium iodide in 20 mL of deionized and add 3 g of iodine crystals. Dilute to 1 litre.
5. Add iodine flakes to methylated spirit to form alcoholic iodine solution.

1.7 Ringer solution, biology solution
A mixture of salts that dissolve in water to form a physiological saline solution.
Prepare fresh before use. 0.9 g sodium chloride, 0.042 g potassium chloride, 0.025 g calcium chloride, 100 mL deionized water.

1.8 Saline solution, biology solution
Dissolve 9 g sodium chloride in 1 litre of deionized water.

1.9 Scott's blueing solution, biology solution
It is used with haematoxylin to develop blue colour.
2 g sodium bicarbonate, 20 g magnesium sulfate, 1 litre deionized water.

1.10 Sea water substitute, biology solution
Dissolve in 2 litres of water, 45.0 g sodium chloride, 3.5 g magnesium sulfate, 5.0 g magnesium chloride, 2.0 g potassium sulfate.

1.11 Sodium thiosulfate solution, biology solution
It is used to decolorize iodine and wash iodine from tissue.
Dissolve 5 g sodium thiosulfate (hypo) in 100 mL deionized water. Add drops of iodine solution. The colour of iodine vanishes. Add drops of weak solutions of acidified bleaching fluid (calcium hypochlorite) or citric acid or sulfuric acid. The colour of iodine solution returns.

1.12 Xylene and methyl benzoate, biology solution
They are used during staining procedures. These chemicals are highly flammable, toxic and readily absorbed through the skin. Care must be exercised when handling. These are not miscible with water.

1.13 Phenylthiocarbamide, PTC, phenylthiourea, PTU, C₇H₈N₂S
See 9.24.2: PTC tasters and non-tasters
Phenyl thiocarbamide is toxic but is safe at a concentration < 0.13%. Weigh 0.13g of solid phenylthiocarbamide on a watch glass. Dissolve the solid in 100 mL deionized water. Cut strips of absorbent paper, 5 cm x 1 cm. Soak the strips in the solution, remove with forceps, drain and dry the papers on a drying
tray in an incubator < 50^oC. Store the indicator papers in a sealed bottle. Prepare strips of paper soaked in solutions X2 or X10 as dilute after diluting the above solution 50 mL to 100 mL or 10 mL to 100 mL. Rinse the mouth after each tasting trial. The chemical has a very persistent bitter taste, so handle the strips of paper with forceps.

2.1 Glycerine jelly, adhesive to stick sections to microscope slides
Soak 10 g gelatine in 60 mL water for 2 hours. Add 70 mL glycerine and 1 g phenol crystals. Heat the solution gently in a water bath and then cool. To soften the jelly before embedding heat in water bath.

2.2 Haupt's adhesive, adhesive to stick sections to microscope slides
Dissolve 1 g gelatine in 100 mL water at 30^oC. Add 2 g phenol crystals and 15 mL glycerine. Stir, cool and filter.

2.3 Meyer's albumen, adhesive to stick sections to microscope slides
Beat an egg white until well broken up, but not stiff. Pour into a tall cylinder and leave to stand overnight. Add equal volume of glycerine to liquid collected from bottom. Add a thymol crystal to prevent growth of fungus.

2.4 Canada balsam
Refractive index similar to crown glass (n = 1.55), used to conserve microscope samples but nowadays synthetic resins used. From Abies balsamea (Pinus balsamea) balsam fir, natural turpentine, balsam tree, American silver fir, balm of gilead fir, Canada turpentine oil, thin resin in bark used as "Canada balsam" in microscopy, Pinaceae

3.1 Acetic alcohol, biology fixative
Mix 99 mL 70% ethanol with 99 mL concentrated ethanoic acid. Prepare this solution prepared immediately before use.

3.2 Aceto-orcein stain, microscopy stain
Stains chromosomes and nuclei crimson, stains cytoplasm pink.
1. Add l g synthetic orcein to 25 mL concentrated ethanoic acid (glacial acetic acid) and 20 mL deionized water. Boil for 4 to 5 minutes in a narrow neck flask, fitted with a glass filter funnel to act as a condenser. Filter the solution while still hot. Add 5 mL concentrated ethanoic acid and stir to dissolve any orcein appearing on the surface of the mixture after filtration. Add 4 to drops of glycerol to retard evaporation.
2. Heat 50 cc 60% acetic acid until almost boils. Add 0.5 g orcein stain. Stir, cool and filter. Use freshly prepared solution.

3.3 Acetocarmine, microscopy stain
Concentrated ethanoic acid (glacial acetic acid) 45 mL, 0.5 g carmine, 55 mL deionized water.

3.4 Aniline hydrochloride, microscopy stain
BE CAREFUL! Use gloves!
Make a saturated solution in aniline hydrochloride in deionized water. Filter then add a few drops of hydrochloric acid until solution is acid.

3.5 Aniline sulfate, microscopy stain
BE CAREFUL! Use gloves!
Prepare as for the aniline hydrochloride above but use sulfuric acid.

3.5.1 Carbol fuchsin, microscopy stain
Dissolve 1 g of fuchsin in 10 mL of ethanol. Add this solution to 90 mL of 5% aqueous phenol then filter the solution.

3.6 Carmine stain, microscopy stain
4 g carmine, 1 mL concentrated hydrochloric acid, 15 mL deionized water. Boil gently in a fume cupboard for 10 minutes with continuous stirring. Cool then add 95 mL 85% ethanol (alcohol). Filter before using the stain.

3.7 Congo red, microscopy stain
Congo red is blue in acid and red in alkali and is made from coal tar. Used in dialysis diffusion experiments and as a yeast stain.

3.8 DCIP (2,6-Dichlorophenol-indophenol), microscopy stain
This stain is used to show action of enzymes e.g. succinic dehydrogenase, chloroplasts when exposed to light and to test for vitamin C. Add 4 g carmine and 1 mL concentrated hydrochloric acid 15 mL deionized water. Boil gently in a fume cupboard for 10 minutes with continuous stirring. Cool, and add 95 mL 85% ethanol (alcohol). Filter the solution. Add 1 g 2,6-Dichlorophenol-indophenol and 1 litre water.

3.9 Delafield's haematoxylin, microscopy stain
1. Dissolve 4 g powder in 25 mL absolute ethanol. Mix gradually into 400 mL saturated aqueous alum, NH₄Al(SO₄)₂.12H₂O. Leave to stand for 3-5 days with a cotton plug in flask, exposed to direct light. Filter, then add 100 mL glycerine and 100 mL methanol. Leave to stand for at least 6 weeks.
2. Dissolve 1 g haematoxylin in 6 cc absolute alcohol. Add this drop by drop to 100 mL saturated ammonium alum. Leave in light and air for one week. Filter than add 25 mL glycerine and 25 mL methyl alcohol. Leave to stand until dark colour. Filter again.

3.10 Eosin, tetrabrornofluorescein, microscopy stain
Counter stains are used to stain cell walls and cell contents. This is an animal tissue counter stain. Eosin stains cytoplasm
1. Use 1 g Eosin Y powder, 1000 mL 70% ethyl alcohol (ethanol) 5 mL glacial acetic (ethanoic) acid. Dilute 100 mL with 100 mL 70% alcohol, Add 2-3 drops of glacial acetic (ethanoic) acid.
2. Dissolve 1 g eosin in 100 mL 70% ethanol.
3. Demonstrate fluorescence with 1:500 ethanol solution.

3.11 Gram stain, microscopy stain
Solutions required for the Gram stain procedure: 3.12 Gram's iodine solution | 3.11.1 Crystal violet solution | 3.11.2 Gram stain decolorizing solution | 3.23 Safranin, microscopy stain.
Mix 2 g potassium iodide crystals and 1 g iodine crystals in 200 mL of deionized water (Gram's iodine solution). Put the heat-fixed smear on the staining rack, cover with Crystal violet solution and leave for 1 minute. Tilt the staining rack and gently wash with water for 3 seconds. Flood the slide with Gram's iodine and leave for 1 minute. A stain iodine complex forms. Tilt and wash with water for a few seconds. Hold the slide at a 45 angle where the smear is clearly visible and run 95% alcohol, as a decolorizing agent (Gram stain decolorizing solution), down the smear until no more colour runs out after 2-10 seconds. Wash with water. Counterstain by flooding the smear with safranin for 1 minute. Wash, blot dry and examine under an oil immersion lens. Gram stain is used routinely as a differential stain so that you can classify bacteria as "gram positive" or "gram negative". Gram positive bacteria stain a blue purple colour because they retain the stain-iodine complex inside their cells. Gram negative bacteria stain a red colour because they have cell walls which allow the stain-iodine complex to be washed out of the cell by alcohol.

3.11.1 Crystal violet microscopy stain
1. Dissolve 0.4 g crystal violet in 20 mL 95% ethanol. Mix with 80 mL 1% aqueous ammonium oxalate. Leave to stand for 48 hours before use. The solution is stable and can be stored for months.
2. For Solution A, dissolve 2 g crystal violet in 100 mL absolute alcohol. For solution B, dissolve 1 g ammonium oxalate in 100 mL deionized water. Add 25 mL Solution A to 100 mL Solution B.
3. For solution A, dissolve 1 g of crystal violet in 20 mL 95% ethanol. For solution B, add 0.8 g of ammonium oxalate to 80 mL deionized water. Add solution A to solution B and filter.

3.11.2 Gram stain decolorizing solution
1. Acetone/ethanol (50:50 v/v) 0.1% basic fuchsin solution.
2. 50% ethanol, 50% acetone, basic fuchsin solution.
3. Dissolve 0.2 g of safranin or basic fuchsin in 10 mL 95% ethanol. Mix with 90 mL, of deionized water. The solution is stable and can be stored for months.

3.12 Gram's iodine solution, microscopy stain
Dissolve 2 g of potassium iodide crystals and 1 g of iodine crystals in 200 mL of deionized water.

3.13 Haematoxylin, microscopy stain
Stains nuclei in plant and animal cells purple, blue or black. This stain was formerly "logwood" because it was made from the heartwood of the tree Haematoxylon campechianum that has a sweet taste and smells of violets. It is similar to natural dyes natural red 24 from brazilin, C₁₆H₁₄O₅ and brazilein C₁₆H₁₂O₅, from Caesalpinia echinata. Use a stock solution of 10% haematoxylin in 95% alcohol.

3.14 Heidenhain iron haematoxylin, microscopy stain
Part 1. 4 g FeNH₄(SO₄)₂.12H₂O (ferric alum, i.e. iron (III) ammonium sulfate) 100 mL deionized water,
Part 2. 10 g haematoxylin, 100 mL 95% ethanol. Mix equal quantities of Part 1. and Part 2. The mixture is useful for a few hours only. Solution 1. is used as a mordant and 2. is for staining.

3.15 Lugol's iodine solution, aqueous iodine solution, microscopy stain
1. Dissolve 6 g potassium iodide in 1000 mL water. Add 4 g iodine crystals.
2. Dissolve 5 g of iodine crystals and 10 g of potassium iodide in deionized water and make up to 100 mL. For bacterial staining, dilute to 1 / 5 with water.
3. Dissolve 1 g iodine crystals and 2 g potassium iodide in 300 mL deionized water.

3.16 Karo syrup mountant, microscopy stain
40 mL clear Karo syrup, 40 mL deionized water. Add 2 small crystals thymol (or phenol) (about 0.2 g) as a preservative.

3.17 Lactophenol, microscopy stain
BE CAREFUL! Use gloves!
Dissolve 25 g phenol in 50 mL water. Add 25 mL lactic acid. Add 50 mL glycerine. Store away from light.

3.18 Lactophenol cotton blue, microscopy stain
100 mL lactophenol, l g cotton blue. Dilute 5 mL to 100 mL with lactophenol before use.

3.19 Leishmann's stain, microscopy stain for white blood cells, Wright's stain
(Or use the ready made dark green crystalline powder Wright's stain that contains the red acid dye eosin and the blue basic dye methylene blue. Variants of Wright's stain include buffered Wright's stain, Gram's stain, Wright-Giemsa stain, Jenner's stain.)
1. Dissolve 0.15 g of Leishmann's stain in 10 mL absolute alcohol in a flask. Plug the flask with cotton wool and heat in a water bath over an electric heater for 15 minutes.
2. Add 0.115 g Leishmann's stain to 100 mL of pure methanol in a flask. Plug the neck with cotton wool and warm in a water bath for 15 minutes with occasional shaking.

3.19a Methylene blue, microscopy stain
Methylene blue (0.1%) Dilute 0.1 g methylene blue in 100 mL deionized water, dilute 1 in 10.

3.20 Neutral red (0.1%), microscopy stain
red pH < 6.8 to yellow pH > 8.0
1. 0.1 g neutral red, 100 mL deionized water, Dilute 1 in 10.
2. Mix 0.1 g stain + 0.2 mL 1% acetic acid + 100 mL water. Use as 1% solution in 59% ethanol.

3.21 Orange IV, orange G, tropeolin 0, microscopy stain
Botanical tissues counter stain. Dissolve 0.5 g powder in 100 mL 95% alcohol.
0.1% solution as acid-base indicator, pH < 1.4 red to pH > 2.6 orange.

3.22 Phloroglucinol (10% solution), microscopy stain
Acid phloroglucin stains lignin in plant cells bright red.
Dissolve 1 g phloroglucin in 100 mL 50% alcohol.

3.23 Safranin, microscopy stain
C.I. 50240, C.I. Basic Red 2, Toxic if ingested, stain for lignin and cell walls
1. Dissolve 0.2 g safranin in 10 mL 95% ethanol. Mix with 90 mL, of deionized water. The solution is stable and can be stored for months
2. Dissolve 1 g safranin in 100 mL 50% by volume alcohol / water mixture.
3. Dissolve 0.5 g safranin in 100 mL deionized water.
4. For a general contrast stain for lignin and cell walls, dissolve 0.02 g safranin in 10 mL 5% ethanol. Add 100 mL water to form 1% solution.

3.24 Schulze's solution (chlor-zinc-iodine), microscopy stain
Dissolve 20 g zinc chloride in 9.5 mL warm water. Cool, add drop by drop 1.5 mL of the following solution until a persistent precipitate of iodine forms: 0.5 g iodine, 1 g potassium iodide, 20 mL deionized water.

3.25 Toluidine blue
Dissolve o.05 g toluidine blue stain in 100 mL of water. he stain works best in fresh plant material. Lignified plant cell walls stain blue-green. Unlignified plant cell walls stain pink-purple.

3.26 Weigert's haematoxylin, Weigert's iron haematoxylin, microscopy stain
This stain is used for animal tissue
Part A: 2.5 g iron (III) chloride FeCl₃.6H₂O, 4.5 g iron (II) sulfate FeSO₄.7H₂O, 2 mL hydrochloric acid, 298 mL deionized water
Part B: 1 g haematoxylin, 100 mL 95% ethanol. Mix 1 part of B to 3 parts of A just before use. This mixture can be used for up to 3 weeks.

3.27 Methyl cellulose, methocel (low substitution)
Dissolve 10 g in 0 mL water to slow Protozoa for study under the microscope.

9.1.2.1.6 Resazurin stain
Prepare a 0.005% solution of the blue redox indicator dye resazurin by dissolving 1 tablet in 50 mL of deionized water. Resazurin is pink when oxidized and colourless when reduced.

4.1 Aceto-alcohol, biology fixative
Absolute ethanol (alcohol) 30 mL, ethanoic (acetic) acid, glacial 10 mL, mix immediately before use and discard after 1 hour, for animal material.

4.2 Ethanol solution
Use a 50% solution of ethanol / water to preserve biological specimens.

4.3 FAA, biology fixative
Formalin (40% methanal) 5 mL, ethanol (alcohol) 70% 90 mL, ethanoic (acetic) acid, glacial 5 mL.

4.4 Formaldehyde, formalin, biology fixative
Formalin is the commercial name for 40% formaldehyde solution. 1% formalin = 1 mL formalin then dilute to 100 mL with water. Formaldehyde vapour irritates eyes and delicate body tissues. Biological specimens preserved in formaldehyde should be kept in sealed containers.

4.5 Formol-saline, biology fixative for marine animals
Formalin (40% methanal) 100 mL, sodium chloride, 10% solution 7 mL, deionized water 83 mL.

4.6 Zenker's fluid, biology fixative
Potassium dichromate 2.5 g, mercury II chloride 5.8 g, deionized water 95 mL.

4.7 4.15 CRAF, biology fixative
Chromic acid, 1% 40 mL, formalin (40% methanal) 10 mL, ethanoic (acetic) acid, glacial 5 mL, deionized water 5 mL.

4.8 Carnoy's fluid, insect fixing fluid
95% alcohol 75 mL, chloroform 30 mL. glacial acetic acid 10 mL.

4.9 KAA, insect fixing fluid
Kerosene 10 mL (soft body insects 5 mL) 95% alcohol 100 mL, glacial acetic acid 20 mL.

4.10 Kahle's fluid, insect fixing fluid
95% alcohol 100 mL, glacial acetic acid 7 mL, formalin 40 mL.

4.11 Oudeman's fluid, insect fixing fluid
70% alcohol 88 mL, glycerine 4 mL, glacial acetic acid 8 mL.

4.12 Lacto-alcohol, insect fixing fluid
Lactic acid 40 mL, 98% alcohol 37 mL, water 23 mL.

4.13 Pampl's fluid, insect fixing fluid
Glacial acetic acid 4 mL, water 30 mL, 40% formaldehyde solution 6 mL, 95% alcohol 15 mL.

4.14 Sugaring mixture, insect fixing fluid
500 g treacle, 1 kg brown sugar, 300 mL beer, 5 mL rum.
Boil until uniform thickness occurs.

4.15 Barber's relaxing fluid, insect fixing fluid
95% alcohol 50 mL, water 50 mL, ethyl acetate 20 mL, benzol 10 mL.

5.0 Standard buffer solutions
The table below shows how to prepare buffer solutions for a particular pH by mixing pairs of the following four solutions.
Solution A = 0.l M boric acid, prepared by dissolving 3.09 g AR boric acid and 3.73 g potassium chloride in water and making up the solution to 500 mL in a measuring flask.
Solution B = 0.l M sodium hydroxide, prepared by diluting standard sodium hydroxide.
Solution C = 0.l M citric acid, prepared by dissolving 9.60 g AR citric acid in water and making up the solution to 500 mL in a measuring cylinder.
Solution D = 0.2 M disodium hydrogen phosphate, prepared by dissolving 17.82 g pure Na₂HPO₄.2H₂O in water, and making up the solution to 500 mL in a measuring cylinder.

pH	Solution A	Solution B	pH	Solution C	Solution D
.	mL	mL		mL	mL
10.0	25	21.8	6.0	14.6	25
9.6	25	18.0	5.6	18.1	25
9.2	25	13.0	5.2	21.6	25
8.8	25	8.0	4.8	25	24.3
8.4	25	4.0	4.4	25	19.8
8.0	1.4	50	4.0	25	15.7
7.6	3.4	50	3.6	25	11.9
7.2	3.8	25	3.2	25	8.2
6.8	7.4	25	2.8	25	8.2
6.4	11.1	25	2.4	50	3.4

6.0 Direct microscopic mounts or squash preparations
Using sterile technique, remove a small portion of the colony with an inoculation needle and mount in a drop of Lactophenol Cotton Blue on a clean microscope slide. Cover with a coverslip, squash the preparation with the butt of the inoculation needle and then blot off the excess fluid.

6.1 Calcofluor White with 10% KOH, to identify fungi
Use for the direct microscopic examination of skin scrapings, hairs, nails and other clinic specimens for fungal elements. This as a very sensitive method. A fluorescence microscope with the correct ultraviolet filters is required.
Solution A: Potassium hydroxide reagent.
Potassium hydroxide 10 g.
Glycerine 10 mL.
deionized water 80 mL.
Solution B: Calcofluor white reagent
Calcofluor white 0.5 g.
Evans blue 0.02 g.
deionized water 50 mL.
Mix one drop of each solution on the centre of a clean microscope slide. Place the specimen in the solution and cover with a coverslip.

6.2 Cellotape flag preparations, to identify fungi
An excellent technique for the rapid mounting of sporulating fungi because more of the reproductive structures intact.
1. Using clear 2 cm wide cellotape and a wooden applicator stick (orange stick) make a small cellotape flag (2 x 2 cm).
2. Using sterile technique, gently press the sticky side of the flag onto the surface of the culture.
3. Remove and apply a drop of 95% alcohol to the flag, to act as a wetting agent and also dissolve the adhesive glue holding the flag to the applicator stick.
4. Place the flag onto a small drop of Lactophenol cotton blue on a clean glass slide, remove the applicator stick and discard. Add another drop of stain, cover with a coverslip, gently press and mop up any excess stain.

6.3 Cornmeal agar, to identify fungi
Use for routine cultivation and identification of fungi.
Cornmeal agar (Oxoid CM 0103) 8.5 g.
deionized water 500 mL.
1. Mix dry ingredients into 100 mL water, boil remaining water.
2. Add boiling water to mixture and bring to boil.
3. Dispense for slopes.
4. Autoclave for 10 minutes at 120^oC, remove and slope.

6.4 Cornmeal glucose sucrose yeast extract agar, to identify fungi
Use for zygomycete sporulation
Cornmeal agar (Oxoid CM 0103) 17 g.
Dextrose (Glucose) 2g.
Sucrose 3 g.
Yeast extract 1 g.
deionized water 1000 mL.
1. Mix dry ingredients into 100 mL water, boil remaining water.
2. Add boiling water to mixture and bring to boil.
3. Dispense for slopes.
4. Autoclave for 10 minutes at 120^oC, remove and slope.

6.5 Czapek Dox Agar, to identify fungi
Use for routine cultivation of fungi, especially Aspergillus, Penicillium, and non-sporulating moulds.
Czapek Dox Agar (Oxoid CM97) 45.4 g.
deionized water 1000 mL.
1. Soak the ingredients in small amount of water.
2. Bring remaining water to boil, add to soaking ingredients and bring to the boil again, stirring continuously.
3. Dispense for slopes as required.
4. Autoclave at 121^oC for 10 minutes, remove and slope or pour for plates as required.

6.6 Indian ink mounts
For the direct microscopic examination of cerebrospinal fluid, CSF, for Cryptococcus species, place a drop of Indian ink on the specimen, mix well with a sterilized loop, and cover with a coverslip. The best brands to use are "Pelikan" or "Talons" Indian ink.

6.7 Lactophenol Cotton Blue (LPCB), to identify fungi
Use for the staining and microscopic identification of fungi
Cotton Blue (Aniline Blue) 0.05 g.
Phenol Crystals (C₆H₅O₄) 20 g.
Glycerol 40 mL.
Lactic acid (CH₃CHOHCOOH) 20 mL.
deionized water 20 mL.
This stain is prepared over two days.
1. On the first day, dissolve the Cotton Blue in the deionized water. Leave overnight to eliminate insoluble dye.
2. On the second day, wearing gloves add the phenol crystals to the lactic acid in a glass beaker. Place on magnetic stirrer until the phenol is dissolved.
3. Add the glycerol.
4. Filter the Cotton Blue and deionized water solution into the phenol / glycerol / lactic acid solution. Mix and store at room temperature.

6.8 Malt extract agar, to identify fungi
Use for routine cultivation and identification of fungi.
Oxoid Malt Extract 20 g
Bacto Agar 20 g
deionized water 1000 mL
1. Dissolve malt extract in a plastic beaker and pH the solution to pH 6.5 with NaOH.
2. Soak agar in small quantity of solution. Bring remaining solution to the boil, stirring constantly.
3. Add to soaking agar. Bring to boil, stirring constantly.
4. Dispense for slopes as required.
5. Autoclave at 121^oC for 10 minutes, remove and slope or pour for plates as required.

6.9 Potassium hydroxide (KOH) with Chlorazol Black, to identify fungi
Use for the direct microscopic examination of skin scrapings, hairs, nails and other clinic specimens for fungal elements.
Potassium hydroxide 10 g
Coral Azole E Black (0.1%) 10 mL.
Glycerol 10 mL.
deionized water 80 mL.
Using sterile technique, remove a small portion of the specimen with an inoculation needle and mount in a drop of KOH on a clean microscope slide. Cover with a coverslip, squash the preparation with the butt of the inoculation needle and then blot off the excess fluid.

6.10 Potato dextrose agar, to identify fungi
Use for routine cultivation and identification of fungi.
Potato dextrose agar 39 g
deionized water 1000 mL
1. Soak potato dextrose agar in small amount of the water in a stainless steel jug.
2. Boil remaining water, add to soaking ingredients, bring to the boil, stirring constantly.
3. Dispense for slopes as required.
4. Autoclave at 121^oC for 15 minutes. Remove and slope or pour for plates as required.

6.11 Rice grain slopes, to identify fungi
Use to induce sporulation and differentiation
Polished rice grains
deionized water
1. Place 1/2 teaspoon rice grains into wide neck 20 mL lass vials.
2. Add 8 mL deionized water to each vial.
3. Lid, then slope on racks ensuring rice grains are evenly distributed.
4. Autoclave racks at 121^oC for 15 minutes.

6.12 Orcinol-Bial's Reagent
Dissolve 0.2g orcinol in 100 mL concentrated hydrochloric acid (caution, corrosive).

7.0 Prepare acids and bases
molarity 1 X volume 1 = molarity 2 X volume 2, M1V1 = M2V2
Dilute acids
Acetic acid 3 M: Dilute 172 mL of 17.4 M acid to 1 litre of water. (99 -100% acetic acid, ethanoic acid)
Hydrochloric acid 3 M: Dilute 258 mL of 11.6 M acid to 1 litre with water. (35% hydrochloric acid)
Hydrochloric acid 4 M: Dilute 400 mL of 10 M acid to 1 litre of water - for normal class use.
Nitric acid 4 M: Dilute 240 mL of 15 M acid. to 1 litre water - for normal class use.
Nitric acid 3 M: Dilute 195 mL of 15.4 M acid to 1 litre of water. (69% nitric acid)
Sulfuric acid 6 M: Dilute 168 mL of 17.8 M acid to 1 litre of water. (95% sulfuric acid)
Sulfuric acid 2 M: Dilute 112 mL of 35 M in 800 mL water, then add water to 1 litre - for normal class use.
Dilute bases
Ammonia solution 4 M: Dilute 220 mL (28% ammonia) 18 M concentrated solution to 1 litre of water ("ammonium hydroxide")
Ammonia solution 3 M: Dilute 200 mL (28% ammonia) 14.8 M concentrated solution to 1 litre of water.
Ammonia solution 2 M: Dilute 330 mL (10% ammonia) 6 M concentrated solution to 1 litre of water - for normal class use.
Potassium hydroxide 4 M: Dissolve 220 g KOH sticks in water, dilute to 1 litre of water - for normal class use
Sodium hydroxide 3 M. Dissolve 126 g the sticks, 95%, in water and dilute to 1 litre of water.
Sodium hydroxide 4 M Dissolve 160 g NaOH in 500 mL water, then dilute to 1 litre of water - for normal class use
Sodium hydroxide 8.5 M Dissolve 330 g NaOH in water, dilute to 1 litre of water. (For CO₂ absorption)
Calcium hydroxide (limewater)
1. 0.02 M. Saturated solution, 1.5 g Ca(OH)₂per litre, use some excess, filter off CaCO₃, and protect from CO₂ of the air.
2. Add 125 g of slaked lime, Ca(OH)₂, to 3 litres of water, shake, allow precipitate to settle, siphon off clear liquid, and protect from CO₂of the air.

7.1 Andrade's acid fuchsin indicator
Add 5 g of acid fuchsin to deionized water. Add 150 mL of M sodium hydroxide soln.. Mix the soln. and leave to stand for one day. The colour should change from from red to brown. If the soln. is not decolorized enough, add 10 mL of M sodium hydroxide soln.. Mix the soln. and leave to stand for one day. Repeat the process until the soln. has a straw yellow colour.

8.0 Prepare salt solutions
Dissolve amount below then dilute to 1 L with water.
Aluminium chloride, AlCl₃.6H₂O, For 0.1 M soln., 24 g of hydrated salt in 1 L water
Aluminium sulfate, Al₂(SO₄)₃.18H₂O, For 0.l M soln., 66 g of hydrated salt in 1 L water
Ammonia, NH₃(aq) or NH₄OH, For 2 M soln., dilute 330 mL of 10% soln. in 1 L water
Ammonium chloride, NH₄Cl, For 5 M soln., 270 g in water
Ammonium carbonate (NH₄)₂CO₃.3H₂O, For 2 M soln., 300 g in 450 mL 10% NH₃, then dilute in 1 L water
Ammonium iron (II) sulfate, For 0.1 M soln., 39.2 g in water, add 5 mL conc. H₂SO₄ in 1 L water
Ammonium oxalate, C₂O₄(NH₄)₂.2H₂O, For 0.1 M soln., 16 g in 1 L water
Ammonium sulfate (NH₄)₂SO₄, For 0.1 M soln., 13.2 g in 1 L water
Barium chloride, BaCl₂.2H₂O, For 0.1 M soln., 24.4 g in 1 L water
Bismuth chloride, BICl₃, For 0.17 M soln., 53 g in 1 litre of dilute HCl, 1 part conc. HCl to 5 parts water
Bismuth nitrate, Bi(NO₃)₃.5H₂O, For 0.083 M soln., 40 g in 1 litre of dilute HNO₃, 1 part conc. HNO₃ to 5 parts water
Calcium chloride, CaCl_2,anhydrous 0.l M soln., 11 g in 1 L water
Calcium chloride, CaCl₂.2H₂O, For 0.1 M soln., 14.7 g in 1 L water
Calcium hydroxide, Ca(OH)₂Limewater, 10 g in 1 L water, shake, allow it to settle, decant clear liquid
Calcium nitrate, Ca(NO₃)₂, For 0.1 M soln., 16.4 g in 1 L water
Calcium sulfate, CaSO₄.2H₂O, For 0.1 M soln., Shake 10 g in 1 L water, leave to stand, decant the clear liquid
Cobalt (II) chloride-6-water, CoCl₂.6H₂O, For 0.1 M soln., 23.8 g in 1 L water
Cobalt nitrate, Co(NO₃)₂.6H₂O, For 0.1 M soln., 29 g in 1 L water
Copper (II) nitrate, Cu(NO₃)₂.6H₂O, For 0.1 M soln., 29.6 g in 1 L water
Copper (II) sulfate, CuSO₄.5H₂O, For 0.1 M soln., 25 g in 1 L water + 5 mL conc. H₂SO₄
Iron (II) ammonium sulfate, Fe(NH₄SO₄)₂.6H₂O, For 0.5 M soln., 196 g in 1 L water + 10 mL conc. H₂SO₄, dilute to 1 litre
Iron (III) chloride, FeCl₃.6H₂O, For 0.1 M soln., 27 g in 1 L water + 20 mL HCl
Iron (III) nitrate, Fe(NO₃)₃.9H₂O, For 0.1 M soln., 40.4 g in 1 L water
Iron (II) sulfate, FeSO₄.7H₂O, For 0.1 M soln., 27.8 g in 1 L water + 1 mL conc. H₂SO₄ to clear
Iron (III) sulfate, Fe₂(SO₄)₃.9H₂O, For 0.1 M soln., 56 g in 1 L water
Lead ethanoate (CH₃COO)₂Pb.3H₂O, For 0.1 M soln., 38 g in 1 L water + dilute ethanoic acid to clear
Lead nitrate, Pb(N0₃)₂, For 0.1 M soln., 33 g in 1 L water
Magnesium chloride, MgCl₂.6H₂O, For 0.1 M soln., 20.3 g in 1 L water
Magnesium nitrate, Mg(N0₃)₂.6H₂O, For 0.1 M soln., 25.6 g in 1 L water
Magnesium sulfate, MgSO₄.7H₂O, For 0.1 M soln., 24.7 g in 1 L water
Manganese sulfate, MnSO₄.H₂O, For 0.1 M soln., 16.9 g in water
Nickel chloride, NiCl₂.6H₂O, For 0.1 M soln., 24 g in 1 L water
Potassium bromide, KBr, For 0.1 M soln., 12 g in 1 L water
Potassium carbonate, K₂CO₃, For 0.1 M soln., 13.8 g in water
Potassium chloride, KCl, For 0.1 M soln., 7.5 g in 1 L water
Potassium dichromate, For 0.1 M soln., 29.4 g in 1 L water (K₂Cr₂O₇)
Potassium dihydrogen orthophosphate, For 0.1 M soln., 13.6 g in 1 L water (KH₂PO₄)
Potassium hydroxide, KOH, For 2 M soln., 110 g of KOH sticks in 1 L water
Potassium iodide, KI, For 0.1 M soln., 16.6 g in 1 L water
Potassium nitrate, KNO₃, For 0.1 M soln., 10.l g in 1 L water
Potassium permanganate, KMnO₄, For 0.1 M soln., 15.8 g in 1 L water
Potassium sulfate, K₂SO₄, For 0.1 M soln., 17.4 g in 1 L water
Silver nitrate, AgNO₃, For 0.1 M soln., 17 g in 1 L water
Sodium borate, Na₂B₄O₇.l0H₂O, For 0.1 M soln., 38 g in 1 L water
Sodium carbonate, Na₂CO₃.10H₂O, For 0.1 M soln., 28.6 g in 1 L water
Sodium carbonate, Na₂CO₃ (anhydrous), For 0.1 M soln., 10.6 g in 1 L water
Sodium chloride, NaCl, For 0.1 M soln., 5.8 g in 1 L water
Sodium chromate, Na₂CrO₄.4H₂O, For 0.1 M soln., 23.4g in 1 L water
Sodium dichromate, Na₂Cr₂O₇.2H₂O, For 0.1 M soln., 29.8 g in 1 L water
Sodium ethanoate, CH₃COONa.3H₂O, For 00.1 M soln., 13.6 g in 1 L water (sodium acetate)
Sodium hydrogen carbonate, NaHCO₃, For 0.1 M soln., 8.4 g in 1 L water
Sodium iodide, NaI, For 0.1 M soln., 15 g in 1 L water
Sodium molybdate, Na₂MoO₄.2H₂O, For 0.1 M soln., 24.2 g in 1 L water
Sodium nitrate, NaNO₃, For 0.1 M soln., 8.5 g in 1 L water
Sodium nitrite, NaNO₂, For 0.1 M soln., 7 g in 1 L water
Sodium oxalate, Na₂C₂O₄, For 0.1 M soln., 13.4 g in 1 L water
Sodium sulfate, Na₂SO₄.10H₂O, For 0.1 M soln., 32.2 g in 1 L water
Sodium sulfide, Na₂S.9H₂O, For 0.5 M 120 g in 1 L water
Sodium sulfite, Na₂SO₃.6H₂O, For 0.1 M soln., 23.4 g in 1 L water
Sodium sulfite, Na₂SO₃ (anhydrous), For 0.1 M soln., 12.6 g in 1 L water
Sodium thiosulfate, Na₂S₂O₃.5H₂O, For 0.1 M soln., 24.8 g in 1 L water
Strontium (II) chloride, SrCl₂.6H₂O, For 0.1 M soln., 26.7 g in 1 L water
tri-Sodium phosphate, Na₃PO₄.12H₂O, For 0.1 M soln., 38 g in 1 L water
Tin (II) chloride, SnCl₂.2H₂O, For 0.5 M soln., 113 g in 170 mL conc. HCl, dilute to 1 L + add tin foil
Tin (IV) chloride, SnCl₂.5H₂O, For 0.1 M soln., 35 g in 1 L water
Zinc sulfate, ZnSO₄.7H₂O, For 0.1 M soln., 28.8 g in 1 L water

9.0 Glues and pastes, adhesives
9.1.0 Flour and milk glues
9.1.1 Flour glue
Flour glues may be edible but they are not approved foods. They are safe for use by children but they should not be encouraged to consume them.
Combine 1 cup flour, 1.5 cups water, 1/3 cup sugar, 1 teaspoon vinegar. Remove any lumps and keep in a closed container.

9.1.2 Milk glue
Combine 1 cup water, 8 teaspoons powdered milk, 4 teaspoons vinegar, removes curds with a sieve, remove limps combine with 1 teaspoon baking soda solution.

9.1.3 Wallpaper paste
1 cup flour, 3 teaspoons alum, water to form appropriate consistence, 10 drops oil of cloves as preservative. Remove any lumps and keep in a closed container.

9.2.0 Commercial glues and pastes (in plastic containers)
Aerosol spray adhesive, sensitive spray glue, for binding textile, fabrics, paper, cardboard, flat PVC, foils, urethane/rubber foams, cork, pictures, photograph mounting, 330 g can
"Bostik" kids paste, non-toxic, with brush, 250 mL bottle
"Bostik" clear gum, non-toxic, bonds paper and cardboard, 5 litre
"Bostik" kids PVA, non-toxic, non-staining dries clear easily wash off clothes, 1 kg, 5 litre bottle
"Clag" paste, non-toxic, easy to apply with brush applicator, for cut and paste and papier mache, dries after 10-20 minutes, 150 g, 300 g, 5 kg bottle
"Clag", celmix, non-toxic adhesive powder, for finger painting, thickening PVA, hardboard sealer, 500 g container
"EC Mix-a-paste", non-toxic, multipurpose adhesive powder, 500 g
"EC Mix-it", instant papier mache, just add water, 2 kg pack
"EC" Tintex craft paste, non-toxic, non-staining, cellulose powder, adhesive for papier mache or gelling base for finger paint, 500 g tin
PVA glue, witches hat lid, kids school glue, non-staining, non-toxic, general classroom use, 250 mL bottle
PVA glue, Helmar, professional woodworking glue, 5 litre bottle
"UHU" craft glue, PVA, non -toxic. quick setting, washable adhesive, for hobby, arts and craft, general all purpose water resistant, 33 mL
"UHU", WOW glue stick, Goes on Purple, coloured overlay, dries clear, safe non-toxic, acid free, washable PVP glue stick, suit young children, 36 g
Accessories
Double -sided tape, high tack, acid and solvent free, 6 mm wide X 50 m roll
Glue brushes, flat, 15 mm wide, pack / 24
Paste spreaders, plastic paddles, 130 mm, pack / 24
Sellotape sticky dots, removable, clear double sided pack / 64, 1600

9.1.2.11 Prepare sterile media or solutions
Equipment: 1 autoclave or pressure cooker, 1 conical flask, 300 mL to 2.0 L according to quantity, culture tubes or test-tubes, sealed with cellulose bungs, Petri dishes, pipette and filler, 5 mL, sterile, pipette aid, 1 pH meter, 1 spatula, 1 set of scales, 1 piece of weighing paper, 1 pair of insulated gloves
Procedures:
Agar media:
1. Weigh out the individual components used for the manufacture of the culture medium onto a piece of weighing paper and place them into a conical flask. The quantities referred to below are for a 1 litre solution. If less culture medium is required, reduce the proportions of the individual components accordingly. The size of the conical flask depends on the amount to be prepared, 300 mL conical flasks are, for example, required for 200 mL culture medium.
2. Swirl the preparation around. until the soluble components have dissolved. Determine the pH and adjust it if necessary with 1 M NaOH or 1 M HCl, according to the values referred to below. 3. Seal the conical flask with a cellulose bung and autoclave in a pressure cooker at 121^oC and 1 bar excess pressure for 20 minutes. Remove the conical flask from the pressure cooker using protective gloves and cool the flask to 50^o.
3. This process can be accelerated by placing the flask under running water. The temperature has been reached when the bare back of the hand can touch the outside of the vessel without an unpleasant sensation.
4. Fill sterile plastic Petri dishes with the still liquid medium so that the base of the Petri dish is covered. Lift the lid of each Petri dish only for a short period of time so that no germs from the air get into the culture medium.
5. Place the filled Petri dishes in a safe place until the agar has set completely.
6. Inoculate the culture medium plates with material that contains bacteria, according to experimental needs.

9.1.2.12 Liquid broth media
1. Prepare the components required for the production of culture media according to the description for agar media, steps 1 through 3, autoclave, and cool down.
2. Fill culture tubes that have been sterilized in a drying cabinet, 180^oC, 3 hours, with 5 mL of the culture medium, using sterile glass pipette and fillers. 3. Inoculate the culture tubes with material containing bacteria, as described in the experiments.

9.1.2.13 Sterile solutions
Prepare the components required for the production of the solution according to the description for agar media, steps 1 through 4, autoclave, and cool down.

Media and solutions
9.1.2.14 Basal agar medium
Components: glucose 10.0 g / litre, casein peptone 4. 0 g / litre, meat extract 4.0 g / litre, yeast extract 0.5 g / litre, liver extract 0.5 g / litre, NaCl 2.5 g / litre, agar 11.0 g / litre (pH 7.2).

9.1.2.15 Basal broth medium
The composition as for basal agar medium above, without agar. This is a liquid medium for overnight cultures.

9.1.2.16 Glucose nutrient agar
With ready made medium: glucose 1. 0 g / litre, nutrient agar 20.0 g / litre, or from individual component: glucose 1.0 g / litre, peptone from meat 1. 0 g / litre, meat extract 3.0 g / litre, agar 12.0 g / litre (pH 7.0).

9.1.2.17 Malt extract agar medium
Ready made: malt extract agar 48.0 g / litre, or from individual components: malt extract 30.0 g / litre, peptone from meat 3.0 g / litre, agar 11.0 g / litre (pH 1.6), autoclave carefully (10 min.)
Dissolve 15 g malt extract and 18g bacteriological agar in 1 litre of deionized water. Dispense into bottles and sterilize with an autoclave.

9.1.2.17a Malt extract broth medium
The composition is the same as malt extract agar medium without the agar. This is a liquid medium for overnight cultures.

9.1.2.18 Minimal agar medium
Components: K₂HPO₄ 3.5 g / litre, sodium citrate x 2 H₂O 0.5 g / litre, MgSO₄.7H₂O 0.1 g / litre, (NH₄)₂SO₄ 1.0 g / litre, glucose 2.0 g / litre, D, L-histidine 0. 2 g / litre, D, L-arginine 0.2 g / litre, thiamine-HCl 0.05 g / litre, agar 11.0 g / litre (pH 7.2).

9.1.2.19 Nutrient agar medium
Ready made: nutrient agar 20.0 g / litre,
or from individual components: peptone from meat 1. 0 g / litre, meat extract 3. 0 g / litre, agar 12.0 g / litre (pH 7.0).

9.1.2.19a Starch nutrient agar medium
Heat 4 g of soluble starch in 100 mL deionized water. Leave to cool then mix the suspension with 100 mL of hot liquid nutrient agar. Sterilize the mixture at 120^oC for 15 minutes.

9.1.2.19b Milk agar medium
Prepare sterilized nutrient agar, leave to cool to 45-50^oC, add 10% pasteurized milk, skimmed, semi-skimmed or full cream milk. When a milk agar plate is made, it is assumed that the microbial population of the milk will not affect the experiment. However, the uninoculated area of the plate acts as a control.

9.1.2.19c Nitrogen free mineral salts agar
Dissolve 0.50 g of FeCl₃.6H₂O in 500 mL of deionized water. Add 2 g K₂HPO₄ + 0.25 g of MgSO₄.7H₂O + 10 g glucose. Add 0.1 M NaOH until pH = 8.3. Add this solution to a mixture of 7.5 g agar and 1 g CaCO₃. Autoclave the mixture at 121^oC for 20 minutes then pour into Petri dishes to make a nitrogen free mineral salts agar plate.

9.1.2.19d Mannitol yeast extract agar (MYEA)
Heat a mixture of 10 g agar in 1 litre of water until the agar is dissolved. Add 0.5 g K₂HPO₄, 0.2 g MgSO₄.7H₂O, 0.2 g NaCl, 0.2 g CaCl₂.6H₂O, 10 g mannitol and 0.4 g yeast extract. Autoclave the mixture at 121^oC for 20 minutes then pour into Petri dishes to make MYEA plates.

9.1.2.19e Glucose nutrient agar medium
Add 0.5% (w/v) glucose to nutrient agar, dispense in bottles and sterilize

9.1.2.20 Nutrient broth medium
Ready made: nutrient broth 8.0 g / litre, or from individual components: The composition is the same as nutrient agar medium without the agar. This is a liquid medium for overnight cultures.
Nutrient broth medium may also be purchased as a powder.

9.1.2.21 Urea agar medium
Ready made medium: 1. urea agar (Christensen) 21.0 g / litre (pH 6.8)
2. urea 20.0 g / litre, deionized water 50 mL. Sterilize 2. with sterile filter, autoclave 1. and cool to 50^oC, add 1.2 to 1.1) Components of urea agar: peptone from meat 1. 0 g / litre, glucose 1. 0 g / litre, NaCl 1. o g / litre, K₂HPO₄, 1.0 g / l, phenol red 12 mg / litre, agar 12.0 g / litre (pH 6.8).

9.1.2.22 Vinegar bacteria medium
Components of vinegar medium: peptone from meat 3. 0 g / l, yeast extract 1. 0 g / litre, mannitol 21.0 g / litre. This is a liquid medium for overnight cultures.

9.1.2.23 MS agar medium
With ready made medium: MS powder (basal salts with minimal organics) 4.7 g / l, granulated sugar 30.0 g / litre, agar 8. 0 g / litre, phytohormone solution (e.g. BAP, see below) 1 M1 (pH 1.8). Regulate with 1 m KOH.

9.1.2.24 BAP Medium
Individual components: BAP (6-benzylaminopurine) 100 mg, 1 M HCl 20 mg. Fill up to 100 mL with boiling deionized water (stock solution). Add 0.5 to 1.0 mL of this stock solution to culture medium, e.g. MS agar medium.

9.1.2.25 Buffer reagent, phosphate buffer reagent
Individual components for 1 M solution:
1. K₂HPO₄, 17.4 g / 100 mL,
2. KH₂PO_4,12.9 g / 100 mL
Composition of 1 M buffer reagent: Make a solution of bipotassium hydrogen phosphate (solution 1.) and potassium bihydrogen phosphate (solution 2.) separately, each time using 100 mL deionized water. Place 60 mL of solution 1 into a 100 mL conical flask and, using a pipette and filler, adjust the pH to 7.5 with solution 2.
Composition of 0.1 M buffer reagent: Place 5 mL of solution 1. into a 100 mL conical flask with 45 mL deionized water. Place 5 mL of solution 2. into another 100 mL conical flask with 45 mL deionized water. Adjust the pH of dilute solution 1. to 7.5 by adding dilute solution 2 with a pipette and filler.

9.1.2.26 20% Domestos Solution
Composition: "Domestos" to household cleaning fluid 20 mL. Fill up with deionized water to 100 mL. "Domestos" is composed of saturated sodium hypochlorite solution.

9.1.2.27 Ringer Solution
Composition: 1 Ringer tablet. Fill with 500 mL deionized water and autoclave.

9.1.2.28 Salt Solution
Components: K₂HPO₄, 3.8 g / litre, KH₂PO₄ 1.2 g / litre, MgSO₄.7H₂O 1.1 g / litre, NaCl 2.5 g / litre, Fe₂(SO₄)₃.4H₂O 0.05 g / litre, Mn₂(SO₄)₃ 4H₂O 0.05 g / litre, "Tween 80" 1 mL/ litre (pH 7.0).