An introduction to. Practical Microbiology

Transcription

1 An introduction to Practical Microbiology

2 A DVD video recording has been produced to accompany this booklet, illustrating the procedures described. It is available from: Professor Joanna Verran Manchester Metropolitan University Faculty of Science and Engineering Faculty Development Unit John Dalton Building Chester Street Manchester Ml 5GD Telephone: Web: Copyright notice This booklet is copyright. Extracts may be photocopied, provided they are used exclusively within the institution for which this work has been purchased. For reproduction for any other purpose, written permission must first be obtained from Manchester Metropolitan University. Copyright Manchester Metropolitan University, 2003

3 Contents Microbiology in schools... 4 The variety of microorganisms... 5 Risk assessment... 6 General safety precautions... 7 Sources of and preparation and maintenance of cultures.. 8 Principles of aseptic techniques... 9 Sterilisation Dealing with spills Growth media Equipment Preparing plates Streak plates Pour plates Confluent lawns Slopes, stabs and liquid cultures Techniques for cultivating fungi Incubation Disposal Serial dilution The antibacterial action of toothpastes Appendices 1 Media recipes Using freeze-dried cultures Suppliers of microbial cultures Useful addresses... 35

4 Microbiology in schools With their small size, rapid growth rate and relatively low cost, microbes are ideal for practical work in schools. Apart from their intrinsic importance, microorganisms can also be used to demonstrate many of the properties of larger plants and animals. About this booklet This booklet explains the basic techniques needed to handle fungi and bacteria safely, including methods of sterilization and disposal. The cultivation of algae, protozoa and other microbes is not covered readers should consult a more specialist reference for appropriate details e.g., CCAP publications see Appendix 3. Additional support Two UK organisations, the SGM (Society for General Microbiology) and MISAC (Microbiology in Schools Advisory Committee) work together to support microbiology education both pre- and post-16. The SGM has a teachers' membership scheme and runs a microbiology Web site for schools. The National Centre for Biotechnology Education (The University of Reading) also provides advice about school microbiology. Appendix 4 at the back of this booklet provides useful addresses, including those of organisations that specialise in safety advice for schools and colleges. Safety in school microbiology More information and charts suggesting microorganisms suitable for school use are provided in Topics in safety (Third edition, 2001) Association for Science Education. ISBN: This book, which deals with numerous aspects of safety in school science, is available from the Bookselling Department, ASE, College Lane, Hatfield, Herts. AL10 9AA. 4

5 The variety of microorganisms Individual microorganisms are too small to see without the aid of a microscope, but many of their effects are familiar from our everyday experience. Although disease-causing microbes such as Salmonella and the human immunodeficiency virus (HIV) capture the headlines, most microorganisms are relatively benign or even beneficial. Bacteria and fungi were used in the production of bread, cheese and alcoholic drinks for thousands of years before their role was properly understood. The invention of the microscope was a crucial development in revealing the great diversity of the microbial world. Scientists gradually developed methods of obtaining pure cultures of microbes so that they could be studied and classified. At least five different groups of microorganisms are recognised today. Whilst some bacteria may be responsible for life-threatening diseases others are essential for our well-being. For example, Clostridium tetani, introduced into a cut or scratch can rapidly lead to death through tetanus. However, another common soil bacterium, called Rhizobium, plays a vital role in the recycling of nutrients by fixing nitrogen in the root nodules of leguminous plants. The cyanobacteria (or blue-green bacteria) share with other bacteria a relatively simple internal structure. They are often the dominant organisms in green blooms on lakes and some, such as Spirulina, are collected, dried, then used as single cell protein. The fungi are familiar both as foodstuffs and food spoilage organisms. This group includes mushrooms, yeasts and moulds. Several species cause diseases such as athlete s foot and potato blight but others are used to manufacture valuable medicines and enzymes. Perhaps the best known protozoan is Amoeba which feeds on other microbes in fresh water. Many other protozoans, like Plasmodium the cause of malaria, have adapted to a parasitic way of life. Algae are photosynthetic. Algal blooms in rivers, ponds and on shorelines may cause environmental damage. Some algae also produce toxins that can accumulate in the bodies of fish and shellfish and cause poisoning when eaten. Unlike other microbes, viruses are unable to reproduce by themselves. To do this, they must infect other organisms. In so doing they are responsible for many human diseases such as influenza, measles and AIDS, as well as diseases of plants and other microorganisms. 5

6 Risk assessment Like all work in the school science laboratory, educational activities using microbes are governed by the Control of Substances Hazardous to Health (COSHH) regulations. The purpose of such regulations is not to restrict the practical work that is done in schools but simply to make it safer to carry out. Teachers and technicians in the United Kingdom have a duty under the Health and Safety at Work Act (1988) to comply with safe working practices and to carry out risk assessments for any practical work undertaken. General (model) risk assessments should be made following the guidance in publications such as Topics in safety (2001) Association for Science Education (Third edition). Following their professional judgement, teachers and technicians may modify such assessments based on the abilities of their students, their own abilities and experience and the precise circumstances in which the work is to be undertaken. Such modifications should be recorded appropriately. In addition to the guidance suggested in other publications, here and on the accompanying DVD video recording, teachers and technicians will need to comply with any additional regulations imposed by their local educational authority and/or school or college governing body. Factors to be considered in risk assessments Adapted from basic practical microbiology: A manual (2001) Society for General Microbiology (this publication is available from Factor Level of practical work as defined by Topics in Safety Choice of microorganisms (ACDP Hazard group 1) Sources of cultures Type of investigation/activity Composition of culture media Volume of culture Laboratory facilities Equipment Incubation conditions Disposal procedures Expertise of teachers and technicians Student age, ability and discipline 6

7 General safety precautions The microbes recommended for practical work in schools* pose a minimal health hazard given good practice. This means treating any microorganisms as though they were potentially harmful and using aseptic techniques when handling them. Hazards The major potential sources of infection in laboratories are aerosols: tiny droplets of liquid laden with microbes. They are released into the air by accidental spills or careless aseptic technique. Aerosols remain in the air for long periods of time and may be inhaled. Breakages and spills may lead to skin and eye infections. Ingestion of microorganisms is likely if cultures are pipetted by mouth. The safety rules listed below are designed to minimise such risks. Good microbiological laboratory practice Any exposed cuts or abrasions should be protected with waterproof dressings before the practical work starts. Everyone involved teachers, technicians and students should always wash their hands before and after practical work. The laboratory door and windows should be closed when work is in progress. This will reduce air movements and consequently the risk of accidental contamination of plates, etc. It is strongly recommended that laboratory coats are worn, and where necessary (e.g., when heating liquids), eye protection. High standards of cleanliness must be maintained. Non-porous work surfaces should be used and they should be swabbed with an appropriate laboratory disinfectant before and after each practical session. To reduce the risk of ingesting microbes, no hand-to-mouth operations should occur (e.g., chewing pencils, licking labels, mouth pipetting). For the same reason, eating, drinking and smoking must not be allowed in the laboratory. Cultures should not be removed from the laboratory. *a list of microorganisms thatare recommended for school use is maintained by the ASE, SGM, CLEAPSS, NCBE and SSERC. It can be downloaded from their Web sites, e.g., 7

8 Sources of and preparation and maintenance of cultures Cultures of microorganisms should only be obtained from reputable school suppliers or 'approved' samples from the environment. Several organisations (see Appendix 4) maintain lists of microbes that are suitable for school use. These organisms present minimum risk given good microbiology laboratory practice. It is important to refer to an up-to-date list, as new information and regulations can lead to alterations in the safety status of different species and strains. It is essential to maintain pure cultures of microorganisms, not only for safety reasons, but also to ensure the integrity and success of a scientific investigation. Maintenance and storage of cultures In schools, microbial cultures are often maintained on agar slopes. With these, it is easier to detect contamination than if liquid cultures are used. Slope cultures should be transferred onto fresh medium every 8 12 weeks or so, and incubated until the organisms have grown. They should then be stored in a cool (10 15 C), dark place, not in a refrigerator. from for class use; the other is a reserve or longer-term stock from which the next pair of stock cultures are taken. New cultures should be obtained regularly, as repeated subculturing can lead to contamination or genetic changes in the cells or colony morphology. Contamination of cultures may be difficult to remedy it is better to obtain a new culture than to try to isolate a pure one from a stock that seems to be contaminated. If in doubt, obtain a fresh supply of the microorganism or return to the long-term stock culture. Preparation of cultures for class use Microbes should be subcultured once or twice a few days before they are required to ensure that they are growing actively and are suitable for use in a practical investigation. Two cultures are usually prepared: one is a 'working' stock for taking sub-cultures 8

9 Principles of aseptic techniques The aims of aseptic techniques are: To obtain and maintain pure cultures of microorganisms; To make working with microorganisms safer. put back on the bottle (see drawing). This prevents contamination of the bench and the culture. A pure culture contains only one species of microorganism, whereas a mixed culture contains two or more species. Contamination of cultures is always a threat because microbes are found everywhere: on the skin, in the air and on work surfaces and equipment. To obtain a pure culture, sterile growth media and equipment must be used and contaminants must be excluded. These are the main principles of aseptic techniques. Growth media must be sterilised before use by autoclaving them. Sterile containers (flasks, Petri dishes, etc.) should be used. Lids must be kept on these containers to prevent contamination. It is essential to prepare the work area carefully before you start. All necessary equipment and materials should be arranged so that they are readily- athand. Work should be done near a lighted Bunsen burner. Rising air currents from the flame will help to carry away any microbes that could contaminate growth media and cultures. When cultures are transferred between containers, tops and lids should not be removed for longer than necessary. After a lid has been taken from a bottle, it should be kept in your hand until it is A blue flame about 5 cm high should be used for sterilising loops or wires and flaming the necks of bottles. After removal of the top, the neck of the culture bottle should be flamed for a few seconds. This will kill any microbes present there and cause convection currents which will help to prevent accidental contamination of the culture from the atmosphere. Bottles should not be heated until they become hot and dangerous to handle. Wire loops, however, must be heated until they glow red hot along the entire length of the wire part. This should be done both before and after cultures are transferred. Heat the stem of the loop first as it is brought into the Bunsen burner flame, to reduce sputtering and aerosol formation. Allow the loop to cool before you use it to transfer a culture (some people like to cool the loop after flaming it, by touching 9

10 it briefly onto the agar at the edge of a culture plate). Before and after use, glass spreaders should be dipped in alcohol, then passed quickly through the flame and the alcohol allowed to burn off. Great care must be taken to keep the alcohol away from naked flames! When the burner is not in use, it should be kept on a visible yellow flame. 10

11 Sterilisation Sterilisation means the complete destruction of all microorganisms, including their spores. Disinfection is the destruction, inhibition or removal of microbes that may cause disease or other undesirable effects. All equipment should be sterilised before starting practical work so that there are no contaminants. Cultures and contaminated material must also be sterilised after use for safe disposal. The four main methods of sterilisation are: Red heat (flame): Instruments like inoculating loops and wires are sterilised by holding them in a Bunsen flame until they glow red hot. By introducing the loop or wire slowly, spluttering and aerosol formation are avoided. Dry heat (hot air oven): This is not often used in schools due to the very high temperatures and time it requires to sterilise equipment. Moist heat (autoclaving): This is the preferred method of sterilisation for culture media, aqueous solutions and discarded cultures. Autoclaving uses high pressure steam, usually at 121 C. Microbes are more readily killed by moist heat than dry heat as the steam denatures their protein. Normally autoclaving is done using a domestic pressure cooker or purpose-built autoclave or steam steriliser. Chemicals: Glassware can be sterilised and contaminated materials rendered safe by soaking them in a suitable disinfectant. Autoclaving Two factors are critical to the effectiveness of this process. Firstly, all air must be removed from the autoclave. This ensures that high temperature steam comes into contact with the surfaces to be sterilised: if air is present the temperature at the same steam pressure will be lower. The materials to be sterilised should be packed loosely so that the air can be driven off. Screwcapped bottles and jars should have the lids loosened slightly to allow air to escape and to prevent the dangerous build-up of pressure inside them. Secondly, sufficient time must be given for heat to penetrate (by conduction) to the centre of the media in Petri dishes or other containers. The times for which medium or apparatus must be held at various temperatures for sterilisation are shown below: Sterilisation time (minutes) Temperature 100 ϒC 110 ϒC 115 ϒC 121 ϒC 125 ϒC 130 ϒC Holding time 20 hours hours 50 minutes 15 minutes minutes minutes Domestic pressure cookers can be used in school laboratories but their small capacity can be a disadvantage Temperature (ϒC)

12 Notice that just a small difference in temperature can result in a great difference in the time required for sterilisation. It is also important that these temperatures are reached by all materials to be sterilised for the specified time e.g., even the broth in the very centre of a flask or fermenter vessel. Three factors therefore contribute towards the duration of the autoclaving process: penetration time the time taken for the least accessible part of the autoclave's contents to reach the required temperature; holding time the minimum time in which, at a given temperature, all living organisms will be killed; safety time a safety margin; usually half the holding time. Domestic pressure cookers operate at 121 C. Thus the total sterilization time might typically be: penetration time, say 5 minutes; plus 15 minutes holding time; plus a safety margin of 5 or so minutes, giving a total of 25 minutes. Purpose-built autoclaves often operate at other temperatures, and whilst the savings in time they offer can seem attractive, it should be remembered that higher temperatures are detrimental to certain media. Glucose solution, for example, is caramelized at high temperatures, forming compounds which may prove toxic to microorganisms. In the case of glucose this reaction can be avoided by adjusting the ph of the medium to 4.0. After sterilisation the ph can be readjusted as necessary. A browning reaction (the Maillard reaction) can also be caused by the interaction of nitrogenous compounds and carbohydrates in the medium at high temperatures here too the compounds formed are toxic to some microbes, so it may in some circumstances be necessary to autoclave the carbohydrate and the remainder of the medium separately. The manufacturer s instructions should always be followed when using either a pressure cooker or an autoclave. Particular care should be taken to ensure that there is enough water in the autoclave so that it does not boil dry during operation. Most domestic pressure cookers require at least 250 cm 3 of water larger autoclaves may need considerably greater volumes. The use of distilled or deionised water in the autoclave will prevent the build-up of limescale. Before the exit valve is tightened, steam should be allowed to flow freely from the autoclave for about one minute to drive off all the air inside. After the autoclave cycle is complete, sufficient time must be allowed for the contents to cool and return to normal atmospheric pressure. The vessel or valves should not be opened whilst under pressure as this will cause scalding. Premature release of the lid and the subsequent reduction in pressure will cause any liquid inside the autoclave to boil. The agar or broth will froth up and it may boil over the outside of the containers. Autoclaves should be dried carefully before storage to prevent corrosion ('pitting') of the pressure vessel. 12

13 The autoclave/pressure cooker should always be checked before use to ensure that it is in proper working order. CLEAPSS produces (for its members) a series of invaluable guides on the routine maintenance of pressure cookers and autoclaves. These guides are essential reading for teachers and technicians in the UK. The effectiveness of an autoclave can be checked by placing a Steriliser Control Strip in the middle of the pressure vessel. Typically, the strips contain a yellow pigment which upon autoclaving changes to brown. Such test strips are available from school science suppliers. Chemical sterilisation Today, relatively few disinfectants are used for sterilisation in school microbiology. Each type of disinfectant has a specific use. Some examples are listed below: Microsol 3 (also known as VirKon) has a wide range of uses and is relatively safe for school use. It is suitable for sterilising work surfaces, placing in discard pots for pipettes and slides and for skin disinfection. It should be used according to the manufacturer's instructions (usually diluted about 1% v/v). In powder form Microsol 3 may also be used for dealing with spillages. Hypochlorite (sodium chlorate I, or bleach) is not ideal for sterilisation of used Petri dishes etc. as it can be inactivated by protein and plastic materials. It can however, be used in discard pots at 2,500 ppm available chlorine (0.25% v/v). Ethanol is suitable for skin disinfection (70% v/v industrial methylated spirit). Most disinfectants must be freshly prepared to ensure their effectiveness. Eye protection and gloves must be worn when preparing disinfectants from full strength concentrates for class use. 13

14 Dealing with spills A basic safety recommendation is that in schools microbes should be cultured on solid media rather than in liquid, wherever possible. There are several reasons for this suggestion, a principal one being the reduction in the likelihood of accidental spillage from flasks or test tubes. A microbial spills kit A microbial spills kit is an essential requirement for any science department wishing to conduct practical work using microorganisms. It is relatively cheap and simple to stock such an emergency kit. The contents should include: a plastic screw- capped bottle containing a measured quantity of a suitable undiluted disinfectant e.g., Microsol 3 ; a small measuring jug to which the bottle of disinfectant can be added, with the correct volume of water needed to dilute it marked on the side; disposable cloths which can be soaked in disinfectant and applied to spilt cultures; a small plastic dustpan and some paper towels; a pair of disposable plastic gloves; an autoclavable waste disposal bag. Even though they may not be required to deal with spills themselves, pupils as well as teachers and technicians should know what to do in the event of a microbial spill. Aerosols The major potential source of infection in a microbiology laboratory is aerosols, small microbe-laden droplets which may be released into the air, persist for half an hour or more, and be inhaled. In the event of a spill This should be dealt with by a teacher or technician, who should wear disposable gloves whilst clearing up the mess. The broken container and / or spilled culture should be covered with a cloth soaked in disinfectant. After not less than 10 minutes (to give the disinfectant time to work), it must be cleared away using paper towels and a dustpan. The contaminated material must be placed in an infected waste container or disposal bag. This, and the gloves, must be autoclaved before disposal. The dustpan should also be autoclaved or placed in freshly-diluted disinfectant solution for 24 hours. All accidents of this type should be recorded. Accidental contamination of skin or clothing As soon as possible, anyone who has been splashed should wash. Severely contaminated clothing should be placed in disinfectant before it is laundered (a hot wash in a domestic washing machine is sufficient for any of the organisms recommended for use in schools). 14

15 Growth media The culture medium must supply all the essential nutrients needed by the microorganism. Moisture must be available and the ph of the medium should be adjusted to that preferred by the organism under study. If these requirements are met, and the incubation conditions such as temperature and the degree of aeration are favourable, good growth should result. There are both liquid and solid growth media. Liquid media A liquid medium is used for boosting growth. The convection currents in a liquid prevent overcrowding and the build up of waste products by dispersing the organisms. If a mixed culture is grown in liquid media, it is difficult to isolate an individual species, so it is impossible to obtain a pure culture from such a sample. It is necessary to subculture onto a solid medium in order to isolate the pure culture. Solid media Solid media contain a gelling base (gelatine, agar or silica gel) plus the nutrients needed for growth. Agar is the most commonly-used gelling agent. It dissolves in water at 100 C and sets at around 48 C. Once set however, it must be reheated to 100 C before it will melt again, so it should to be kept in a water bath at 50 C until ready for use, and then used quickly once removed. For this reason, molten growth media should not be pipetted. Microorganisms deposited on solid media will multiply under appropriate incubation conditions. One organism will eventually produce a colony. Preparation of media Tablets or powders should be rehydrated according to the manufacturer's instructions. The solid should be allowed to dissolve fully before dispensing in appropriate volumes and autoclaving. Stocks of prepared, sterilised media in screw-topped bottles may be stored at room temperature, out of direct sunlight. Solid media may be re-melted in a water bath or microwave oven (tops should be loosened before melting the media). IMPORTANT Culture media designed to select for pathogens (such as those which might be used in hospital laboratories e.g., blood agar, McConkey s agar) should not be used in schools. Recipes for various growth media are given in Appendix 1. 15

16 Equipment Containers These are needed to store sterile media and to grow cultures in. The most frequently used are: a) Petri dishes are used to cultivate microbes on solid nutrient media. Sterile plastic dishes are used once, then destroyed along with their contents after use. Glass Petri dishes can be re-used after autoclaving, but any cost savings are likely to be offset by the time needed to clean and resterilise them. b) McCartney bottles/universal bottles are plastic or glass screw capped jars of cm 3 capacity. They are used for holding stock cultures, deeps, slopes and molten agar for pouring plates. (A Petri dish holds cm 3 of medium.) c) Bijoux bottles are the same as Universal bottles but only have about 5 cm 3 capacity. They are used for holding cultures. d) Bacteriological tubes are test tubes with a metal cap or cotton wool bung. They are used for the same purposes as bottles, but care must be taken because spills occur more easily. Unless the tubes have screw caps, they are unsuitable for longer-term storage of cultures, as the agar medium tends to dry out. e) Medical flats are flat glass medicine bottles. The most useful sizes are 100 cm 3 and 250 cm 3 capacity. They are mainly used for storing large quantities of prepared sterile media. f) Conical flasks are used for growing microbes in liquid culture media and also for holding large volumes of prepared sterile media. Inoculating Instruments Because (non-disposable) wires and loops have to be sterilised to red heat in a Bunsen flame it is best to have instruments with metal handles. Glass handles crack very easily when inoculating loops and wires are heated. a) Loops are used for most inoculations. A platinum or nickel wire (0.5 mm in diameter) is twisted round to make the loop on a metal handle. Single-use disposable plastic loops are also used. 16

17 b) Wires are used for preparing stab cultures and for picking off individual colonies when they are too close together to use a loop. The same type of wire is used as for the loop. Note: disposable plastic loops designed specially for picking individual colonies are also available. c) Thick wires are useful for cutting and transfer of fungal mycelium. Nichrome wire of 1 mm diameter is used with a flattened tip. A thick wire must be flamed in ethanol, rather than heating to red heat. d) Spreaders are used to prepare confluent lawns or plates. They are bent glass or plastic rods. The glass type are flamed with alcohol to sterilise them; the plastic, disposable versions are supplied ready-sterilised. e) Sterile pipettes. Plastic disposable sterile pipettes can be purchased. Glass pipettes must be sterilised by autoclaving before use. All pipettes should have a small piece of cotton wool pushed into the mouth (i.e., the end away from the tip) to act as a filter. Small pipettes with a fine drawn out tip are generally known as Pasteur pipettes. Incubators Incubators are used to maintain cultures at a known temperature. In school laboratories incubation should generally be below 30 C. Incubation can also take place in a suitable cupboard or on a window sill or bench at room temperature. Transfer cabinets These protect the operator during transfer and minimise the risk of contamination of cultures from airborne particles. Despite the fact that many suppliers list them in their catalogues, such chambers are not necessary for work in schools. 17

18 Preparing plates Pouring a plate Assuming right-handedness 1. Wipe the bench with disinfectant, and have a Bunsen burner on with a roaring flame. A sterile Petri dish and a bottle of molten nutrient agar (15 20 cm 3 ), perhaps in a 50 C water bath, should be ready for use. 2. Put the sterile Petri dish near your right hand with the lid uppermost. Do not open the Petri dish until ready to pour the agar. Loosen, but do not remove, the top of the agar bottle. 3. Hold the bottle containing the nutrient agar in the right hand and, using the little finger of your left hand, remove the top and keep hold of it against the palm of your left hand. 4. Flame the neck of the bottle by passing it briefly through the flame. Using the left hand, lift the lid of the Petri dish slightly and pour the molten agar onto the base. 5. Replace the lid of the Petri dish and place the empty bottle and top into a disinfectant container ('discard pot'). 6. Gently rotate the base of the Petri dish so that the agar forms a continuous layer over the base. Leave the plate on the bench to set. (Should air bubbles form on the agar surface, quickly and carefully wave a yellow Bunsen flame over the surface while it is still liquid.) Drying a plate Sometimes condensation may collect on the lids of prepared Petri dishes, particularly if the agar medium is poured whilst it is still too hot. The plate can be dried, upside-down, on the bench or in a suitable cupboard if an incubator is not available. This is how to do it: 1. Once the agar has set, lift the Petri dish, holding the base in the right hand and the lid in the left. 2. Quickly, to avoid contamination, turn the dish over and open it, putting the lid down first and the base on top at an angle. Make sure the surface of the medium does not touch the lid. 3. Leave the plate to dry where it won t be disturbed until no droplets of moisture are visible on the lid or the agar. 18

19 Streak plates The purpose of a Streak Plate is to dilute the small amount of bacterial culture put on the plate down to individual cells; each of which will form an isolated pure colony during incubation. A streak plate is used to separate mixed cultures into pure ones. The individual colonies that grow can be used to inoculate fresh plates or tubes. Each colony grows from a single cell. Streak plates can also be used to check whether a culture is pure or not because any contaminating microorganisms will form different colonies on the medium. 1. Label the base of the Petri dish with your name, the date and the microorganism being used. 2. It may initially be helpful to draw four sectors on the base of the plate and number them. 3. Hold the loop in the right hand and sterilise to red heat in the Bunsen flame. Let it cool for a few seconds. 4. With the loop in the right hand hold the bottle containing the inoculum in your left. Hold the top with the little finger of your right hand, and unscrew the bottle using your left hand. Keep the top held against your palm with your little finger. 5. Flame the neck of the bottle. 6. Put the loop in the bottle and pick up some inoculum. Flame the neck of the bottle again and replace the top. Put the bottle down. 7. Lift the Petri dish lid with your left hand. 8. Streak the agar surface with the loop as shown in the diagram. (Do not dig into the agar, as this may lead to microbes growing in anaerobic conditions beneath the agar surface.) 9. Remove the loop and close the Petri dish. Flame the loop and allow it to cool. 10. Turn the plate anti-clockwise. 11. With the cooled loop, streak the plate as shown in the diagram, making sure that a small amount of material from the first set of streaks is carried over. 12. Repeat steps 9, 10 and 11 for the second, third and fourth segments. Remove and flame the loop, replace the lid and rotate the plate again. 13. Replace the lid and flame the loop. Seal and incubate the plate in an inverted position single colonies here 19

20 A pour plate is prepared by mixing a small amount of bacterial culture with molten nutrient medium. Pour plates allow microorganisms to grow both on the surface and within the medium. With the molten medium at 50 C, the majority of bacteria are not killed by heat and can grow once the medium has set. Pour plates If the dilution is known, the viable count of the bacterial sample i.e., the number of live bacteria per cm 3, can be determined. 1. Hold the bottle containing the bacterial culture in your left hand, and a sterile Pasteur pipette in your right. Do not touch the trunk of the pipette if possible, and especially not the tip. 2. Hold the top of the culture bottle with the little finger of your right hand. Remove the bottle by unscrewing it with your left hand. Flame the bottle neck. 3. Squeeze the teat bulb of the pipette very slightly, put the pipette into the bottle and draw up a little of the culture about 1 cm 3. Remove the pipette and flame the neck of the bottle again before replacing the top. 4. Put the bottle down and pick up the molten nutrient agar bottle in your left hand. 5. Remove the bottle top as before and flame the bottle neck. 6. Insert the pipette vertically into the bottle and release the inoculum onto the agar. Do not touch the sides of the bottle or the agar with the pipette. 7. Put the pipette in a discard pot. Remove the teat while the tip is pointing into the disinfectant. Flame the neck of the bottle and replace the top. 8. Gently roll the bottle between your hands to mix the culture and medium. Avoid making air bubbles. 9. Hold the bottle in your left hand; remove the top with the little finger of your right. Flame the neck of the bottle and lift the lid of the Petri dish slightly with your right hand. 10. Pour the agar mixture into the Petri dish and replace the lid. 11. Flame the neck of the bottle and replace the top. 12. Gently rotate the dish to ensure that the medium covers the plate evenly. 13. Allow the agar to solidify; invert the plate and incubate it. 20

21 Confluent lawns Confluent lawns are also known as bacterial lawns. There are two ways of preparing a lawn: spread plates and flood plates. Ideally, these should result in a culture spread evenly over the surface of the growth medium. out quickly to minimise the risk of contamination. 8. Replace the lid. Put the spreader in alcohol. 9. Let the lawn dry, seal the Petri dish, then incubate it in an inverted position. Preparing a spread plate 1. Hold a sterile pipette in your right hand and a bottle containing broth culture in your left. Remove the top with the little finger of the right hand and flame the bottle neck. 2. With the pipette, remove a small amount of broth. Flame the neck again and replace the top. 3. Using the left hand, slightly lift the lid of a Petri dish containing a nutrient medium and place a few drops of broth onto the surface. 4. Close the lid and dispose of the pipette in a discard jar. 5. Dip a glass spreader in alcohol and pass it through a Bunsen flame. The alcohol will burn and sterilise the glass. Keep the alcohol beaker away from the Bunsen flame. 6. Take the lid off the Petri dish and place it on the bench facing downwards. 7. Place the spreader onto the surface of the inoculated agar and, rotating the dish with the left hand move the spreader in a top-to-bottom or a side-to-side motion to spread the inoculum over the surface of the agar. This operation must be carried Preparing a flood plate 1. Hold a bottle containing broth culture in your left hand; remove the top using the little finger of your right and flame the bottle s neck. 2. Lift the lid of a set agar plate and pour a little broth onto the medium to cover it. Replace the lid. 3. Flame the neck of the bottle and replace its top. 4. Gently rotate the plate; keeping your fingertips on the lid and the base of the dish against the bench, until the agar surface is covered with broth. 21

22 5. Carefully tip any excess broth into a disinfectant pot and replace the lid of the dish. 6. After allowing the plate to dry, seal it and incubate it in an inverted position. This method is wasteful of culture! Uses of pour and spread plates Because they produce an even lawn of bacterial growth, pour and spread plates can be used to test the sensitivity of bacteria to many antimicrobial substances, e.g., mouthwashes, tomato puree, soaps, garlic, disinfectants and antibiotics. The effect of these substances on the bacteria can be measured from the diameter of the zone of inhibition that forms around the tested substance. The larger the zone, the more bacterial growth is inhibited. The substances tested can be diluted to find the minimum concentration needed to inhibit bacterial growth (see page 28). 22

23 Slopes, stabs and liquid cultures Inoculation of slopes Slopes are used to maintain stock cultures. Once the culture has grown it will keep in the dark at C for 3 4 months. Some bacteria such as E.coli will keep for much longer. The low temperature slows the metabolism of the microorganism down, with growth and reproductive cycles being extended over long periods. 1. The top of the slope bottle is removed with the little finger of the right hand, and the bottle neck flamed. 2. A loop charged with inoculum is placed at the bottom of the slope and a zig-zag line made up to the top. 3. The loop is taken away, the neck of the bottle flamed and the top replaced. 4. The loop is sterilised in the Bunsen flame. 5. The inoculated slope is incubated at an appropriate temperature. Inoculation of stab cultures (Deeps) Stab cultures are used to cultivate anaerobes or to separate a mixed culture into aerobes and anaerobes. Anaerobic culture methods such as these are not recommended for use in schools due to the risk of cultivating unwanted anaerobic pathogens. 1. Holding a wire with bacteria on the tip in the right hand, remove the top of the bottle held in the left hand with the little finger of the right. 2. Flame the neck of the bottle and stab the wire gently into the medium in a straight vertical line. 3. Take the wire out, flame the neck of the bottle and replace the top. Incubate. 4. Re-sterilise the wire in the Bunsen flame. Inoculation of liquid media To inoculate using a wire or loop: 1. Hold the loop carrying inoculum in the right hand and with the little finger remove the top of a bottle containing sterile liquid medium. 2. Flame the neck of the bottle and hold it at an angle. 3. Rub the loop against the side of the tube. 23

24 4. Remove the loop; flame the neck of the bottle and replace the top. Incubate. 5. Sterilise the loop in the Bunsen flame once more. To inoculate using a pipette: 1. With the right hand hold a pipette containing a few drops of the inoculum. The little finger of the same hand should be used to remove the top of a bottle containing sterile growth medium. 2. Flame the neck of the bottle and drop the inoculum from the pipette into the medium. Do not let the pipette touch the sides of the bottle as this might contaminate the medium with unwanted microbes. 3. Remove the pipette; flame the neck of the bottle and replace the top. 4. Place the used pipette in a discard jar. 5. Incubate the culture medium. 24

25 Techniques for cultivating fungi Fungi include the yeasts (unicellular) and moulds (mycelial). They have different growth requirements from bacteria and need to be grown on media that have a high carbohydrate to nitrogen ratio, and a low ph. Malt extract agar is normally used. Growth of fungi takes around 7 days and is at a lower incubation temperature than that used for bacteria. Fungal plates are incubated the right way up i.e., with the lid uppermost. Inoculating fungi onto agar plates 1. Dip the end of a 5 mm cork borer in alcohol and then burn the alcohol off to sterilise it. Hold the borer horizontally: be careful that the flames do not pass up the centre of it! 2. Allow the borer to cool, then push the tip into the edge of the fungal colony to be subcultured. 3. Dip a thick wire in alcohol and ignite the alcohol to sterilize it. Allow the wire to cool. Use it to pick up the disc formed by the borer. 4. Put the disc in the centre of a fresh Petri dish containing a suitable sterile growth medium. 5. Return the borer and wire, once they have cooled, to the ethanol beaker. 6. Seal the plate and incubate it. N.B. The techniques used for cultivating yeasts are similar to those employed with bacteria (as yeasts too, are unicellular), but they are still grown on fun gal medium. 25

26 Incubation For safety reasons, plates used in schools are generally sealed after inoculating them. After inoculation, use self-adhesive tape to seal the Petri dishes as shown below. Do not seal the plate entirely, as this will create anaerobic conditions within the dish. Fungi Petri dishes containing fungi do not need to be inverted. Fungal cultures should be incubated for 7 days or so. Room temperature is sufficient to allow their growth. Cultures can be incubated in any suitable cupboard, although a proper laboratory incubator allows greater control of temperature. Incubation at 37 C Label the plate, on the base, before inoculation. Write your name, the date and the organism used, so that it can be identified. Bacteria Bacterial cultures in Petri dishes should be incubated with the base uppermost, so that any condensation that forms falls into the lid and not on the colony. (If there is heavy condensation in the sterile Petri dish before inoculation, it should be dried as described in the Pouring a Plate section.) Generally, cultures in schools should not be incubated above 30 C, and temperatures approaching 37 C should be avoided to minimise the risk of growing human pathogens, which thrive at this temperature. However, the delicate strains of E. coli used in some investigations (e.g., bacterial transformations) should be incubated at 37 C for speedy growth. Good microbiological practice will ensure that human pathogens are not inadvertently cultivated at this temperature. Some organisms, such as the bacteria that are used to make yoghurt, require much higher incubation temperatures of between C. After 2 3 days of incubation at C, bacterial colonies will be seen. 26

27 Disposal It is very important to dispose of all the materials used in the practical class properly to avoid contamination of the laboratory and people. Any containers used for storing and growing cultures must be autoclaved, then washed in disinfectant and rinsed, before being reused. Two autoclave bags should be available in the laboratory; one for reusable glassware and another for disposable materials. You should have a large and a small discard jar near your work area for pipettes and microscope slides, and there should be a metal bucket for disposal of any broken glassware. Glass Petri dishes should be placed in the reusable glassware autoclave bag. Plastic Petri dishes are put in the autoclave bag for disposable items. in the disinfectant, otherwise aerosols can be created. Pipettes are then autoclaved, washed and rinsed before being used again. Contaminated paper towels/cloths are put in the autoclave bag for disposable items. Any contaminated glassware should be put in the autoclave bag for glassware. Glassware that is not contaminated can be washed normally. Broken glassware should be put in a bucket reserved exclusively for that purpose. If the glassware is contaminated it must be autoclaved before disposal. Uncontaminated glassware can be disposed of straight away. Spreaders, loops and wires do not need to be autoclaved; they should simply be sterilised using a Bunsen flame. The same practices are applied to glass and plastic culture bottles. Disposable plastic pipettes, loops, spreaders, microscope slides and any liquid from cultures are put in the small disinfectant pot. Plastic pipettes etc. are then autoclaved and disposed of. Microscope slides are soaked in disinfectant for 24 hours then washed and rinsed before being reused. Glass pipettes should be put carefully in the larger pot. Do not take the pipette filler off the end of the pipette until the tip is 27

28 Serial dilution 1. Place 9 cm 3 of sterile water or saline solution into each of a series of sterile containers and label them for the dilution factor (e.g., 10-1, 10-2, 10-3, 10-4 and so on). 2. To the first container add 1 cm 3 of the substance to be diluted e.g., broth culture, milk, water etc. and mix the contents well by rotating the container gently. This provides an initial 10-1 dilution. 3. Using a fresh pipette, remove 1 cm³ of the 10-1 dilution and place it in the second container. Mix well. This gives a 10-2 dilution. 4. Again using a fresh pipette, transfer 1 cm 3 of the 10-2 dilution into the third container. Mix. This yields a 10-3 dilution. 5. Continue this process of removing 1 cm 3 from the previous dilution and adding it to the next 9 cm 3 of diluent until the end of the dilution series is reached (10-8 is usually sufficient for an overnight broth). The antibacterial action of toothpastes This method uses two species of microorganism and two types of toothpaste. A pair of additional plates without any toothpaste are used as controls. 1. Label the Petri dishes so that you know which treatment will be applied to each. 2. Put a small amount of toothpaste in the centre of each of two sterile Petri dishes. 3. Repeat for the other tube of toothpaste. 4. Inoculate three of the medium bottles with a few drops of the Escherichia coli culture. Roll the bottles gently to mix their contents. 5. Carefully pour two of them over different types of toothpaste, and the third one into an empty Petri dish as a control. 6. Repeat Steps 4 and 5 using Micrococcus luteus. 7. Allow the medium in the six plates to solidify. 8. Incubate the plates in the inverted position. Observe the effects of the two types of toothpaste on the bacteria by comparing their growth in the control plate to that with the toothpastes. This method can be used to test a range of different antibacterial substances. 28

29 Appendix 1 Media recipes All these recipes are made up to 1 litre with distilled water unless otherwise stated. Most of these media can be made up as broths by omitting the agar. Beef extract Peptone NaCl Agar MEDIA FOR BACTERIA Nutrient agar Adjust ph to 7.4 before autoclaving. Milk agar (for milk bacteria) l0 g l0 g 5 g 20 g Make up nutrient agar as above, but using only 900 cm 3 of water. Dissolve 20g of dried skimmed milk in 100 cm 3 of water. Autoclave separately. Mix together aseptically once the liquids have cooled to C. Yeast extract Malt extract Acetobacter broth (for Acetobacter aceti) 5 g 10 g Dissolve in 1 litre of water, autoclave, then add 30 cm 3 of ethanol once cool. Mannitol yeast extract agar (for Rhizobium) K 2 HPO g MgSO 4.7H 2 O 0.2 g NaCl 0.2 g CaCl 2.6H 2 O 0.2 g FeC1 3.6H 2 O 0.01 g Mannitol 10 g Yeast extract 0.4 g Agar 10 g Photobacterium broth (for Photobacterium phosphoreum) Yeast extract Peptone 3 g 5 g Dissolve the above in 250 cm 3 distilled water, then add 750 cm 3 of sea water (use artificial sea water from school science suppliers or an aquarist s shop). Autoclave and dispense as usual. 15 g of agar can be added to make a solid medium if required. Vibrio natriegens broth (Suitable for growth experiments) Nutrient broth NaCl 13 g 20 g After autoclaving, adjust ph to 7.5. For a solid medium, add 15 g of agar. Note: this recipe is a suitable alternative to Heart Brain infusion broth. 29

30 Methylophilus broth (for Methylophilus methylotrophus) (NH 4 ) 2 SO g MgSO 4.7H 2 O 0.2 g NaH 2 PO 4. 2H 2 O 1.4 g K 2 HPO g FeC g Trace metal solution 1 cm³ Water 960 cm³ Trace metal solution (per litre) CuSO 4.5H 2 O 0.02 g MnSO 4.4H 2 O 0.01 g ZnSO 4.7H 2 O 0.01 g CaCO g 1M Hydrochloric acid 36.6 g Sterilize by autoclaving before adding 0.5% by volume of methanol. 15 g of agar can be added if a solid medium is required. Malt extract Peptone Agar Malt Extract Agar (for fungi) 30 g 5 g 15 g Yeast complete medium (suitable for Saccharomyces spp.) Peptone Yeast extract Glucose Agar 10 g 10 g 20 g 15 g Tributyrin agar (for detection of lipase activity) Nutrient agar recipe Tributyrin as above 10 g Potato Glucose Agar MEDIA FOR FUNGI Potato dextrose agar (for mould fungi) 250 g 20 g 20 g This can be made using reconstituted dried potato or boiled and peeled fresh potatoes. Starch agar (for detection of amylase activity) NaNO 3 KCl MgSO 4. 7H 2 O Na 2 HPO 4.12H 2 O FeSO 4. 7H 2 O Agar 3 g 0.5 g 0.5 g 1 g 0.01 g 15 g Dissolve the above in 900 cm 3 water. Dissolve 2 g starch in 100 cm 3 hot water. Add to the mineral solution. Autoclave and dispense as usual. 30

31 Appendix 2 Using freeze-dried cultures Microorganisms obtained from culture collections are often supplied in glass ampoules. The sealed ampoules contain freeze-dried (lyophilized) cultures in a high vacuum. It is advisable to use the dried culture to inoculate a Petri dish, agar slope and McCartney bottle of nutrient broth at the same time. In this way, the Petri dish can be checked for visible signs of contamination, and if it is found to be pure, the broth can be used as an inoculum, whilst the slope can be incubated then stored for future use. The following procedures describe how to safely open such ampoules and resuscitate the microbes they contain with minimum risk of accidental contamination. It is recommended that all instructions should be read and understood first. If possible, the various manipulations should initially be practised using non-sterile equipment. If you are uncertain of your ability to juggle the equipment smoothly, it may be helpful to enlist the assistance of a partner. Opening an ampoule 1. Score the ampoule over the cotton wool plug using a diamond glass cutter. 2. Heat the end of a glass rod until it glows red hot, then hold it on the score mark on the ampoule. This should cause the glass to crack along the line of the score. 3. Wait for air to slowly seep into the ampoule (this takes seconds). IMPORTANT If this procedure is not followed, then when the glass is broken the plug may be forced to the bottom of the ampoule, spraying freeze-dried culture into the laboratory. 4. Carefully snap off the tapered end of the ampoule, and discard it into disinfectant. 5. Since the cotton wool plug may be contaminated with dried microbes, it should be removed using fine forceps and placed in disinfectant. 6. Flame the neck of the ampoule and use fine forceps to place a new, sterile cotton wool plug in the mouth of the ampoule. 7. The ampoule can now be placed on the bench or upright in a small beaker whilst the materials necessary for resuscitating the culture are assembled. 31