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Basic microbiology practical manual

Microbiology basic practical

Course

Pharmaceutical Microbiology - Practical (BP307P)

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Annamalai University

Academic year: 2022/2023

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Microbiology Society

Basic

Practical

Microbiology

A MANUAL

About this resource

Copyright

Microbiology is a popular option for practical work in schools. This manual, which explains the basic techniques necessary to carry out microbiology experiments safely and effectively, is intended as a guide for teachers and technicians. The Microbiology Society, in association with the Microbiology in Schools Advisory Committee (MiSAC) has also produced a book of 21 practical investigations which complements the manual – Practical Microbiology for Secondary Schools is suitable for use with Key Stages 3, 4 and post-16 and the equivalent Scottish qualifications. This resource can be downloaded from the Microbiology Society’s dedicated education website: microbiologyonline.org The Microbiology Society offers schools membership and runs an advice service on microbiology teaching. For further details, email education@microbiologysociety, telephone +44 (0)20 7685 2682 or write to the address at the bottom of the page. Other useful sources of advice: x CLEAPSS – cleapss.org The Gardiner Building, Brunel Science Park, Uxbridge UB8 3PQ x Microbiology in Schools Advisory Committee (MiSAC) – misac.org c/o NCBE (see address below) x National Centre for Biotechnology Education (NCBE) – ncbe.reading.ac Science and Technology Centre, The University of Reading, 2 Earley Gate, Whiteknights, Reading RG6 6AU x Scottish Schools Equipment Research Centre (SSERC) – sserc.org 2 Pitreavie Court, South Pitreavie Business Park, Dunfermline KY11 8UB

Published by the Microbiology Society, Charles Darwin House, 12 Roger Street, London WC1N 2JU, UK

x Kath Crawford (SAPS Scotland) x John Richardson (SSERC) x Members of MISAC x John Schollar (NCBE) x John Tranter (CLEAPSS) Editors: Dariel Burdass, John Grainger & Janet Hurst Design and Production Editor: Ian Atherton Photographs: Dariel Burdass & Faye Jones Front cover illustration: TEK Image / Science Photo Library ISBN 0 95368 383 4

Basic Pract Cover 2009 16/1/09 12:01 Page 4

Credits & Acknowledgements

x x x x x

Wladimir Bulgar/Science Photo Library

Educational use: Electronic or paper copies of the resource or individual pages from it may be made for classroom and bona fide educational uses, provided that the copies are distributed free of charge or at the cost of reproduction and that the Microbiology Society is credited and identified as copyright holder.

Basic Practical Microbiology – A Manual is copyright. The Microbiology Society asserts its moral right to be identified as copyright holder under Section 77 of the Designs, Patents and Copyright Act (UK) 1988.

The Microbiology Society gratefully acknowledges the support from the following sources:

MiSAC

Essential methods for maintaining, preparing and using cultures

Part 2: Microbiology in Action

Obtaining suitable cultures
Pure cultures
Maintaining stock cultures
Checking cultures for contamination
Preventing contamination of cultures and the environment
Aseptic transfer of cultures and sterile solutions
Preparing cultures for class use
1 sensitivity to antimicrobial substances Practical activities
- Using the microscope 2
- Stained preparations
- Making a smear
- A simple stain
- A differential stain: Gram’s staining method
1 guidelines Appendices
2 micro-organisms
3 resources
4 of cultures and equipment
5 of the autoclave/pressure cooker
6 serial dilutions

Part 1: The Basics

An introduction to microbiology, aseptic technique and safety

As well as causing a familiar range of diseases in animals and plants and problems in food spoilage and deterioration of other materials, microbes are also our ‘invisible allies’. Indeed, life on Earth would not be sustainable without the benefits that many of them provide. The teaching of such an important subject as microbiology cannot be achieved effectively without enhancing the theory with ‘hands on’ experience in the laboratory. The purpose of this manual is to provide teachers and technicians with good techniques in practical microbiology to ensure that investigations proceed safely and achieve the required educational aims successfully. This manual has been written for a right-handed person.

Preparation

Safety guidelines The small size of microbes and the consequent need to deal with cultures that contain many millions of microbial cells require special procedures for their safe use. Activities involving micro-organisms are controlled by the Control of Substances Hazardous to Health (COSHH) Regulations and teachers and technicians have a duty under the Health and Safety at Work Act to comply with any safety instructions given by their employers. These include using model risk assessments. The publications normally used in secondary schools are: x Topics in Safety , 3rd edition (ASE, 2001) x Safeguards in the School Laboratory , 11th edition (ASE, 2006) x CLEAPSS Science Publications, CD-ROM (latest edition) x Safetynet (SSERC) x Safety in Science Education (DfEE, available from the ASE website) The guidelines are straightforward and largely common sense and, as such, are not an obstacle to conducting interesting microbiological investigations in a school laboratory. Planning ahead is essential when embarking on practical microbiology investigations. There are five areas for consideration. x Preparation and sterilisation of equipment and culture media. x Preparation of microbial cultures as stock culture for future investigations and inoculum for the current investigation. x Inoculation of the media with the prepared culture. x Incubation of cultures and sampling during growth. x Sterilisation and safe disposal of all cultures and decontamination of all contaminated equipment. [Appendix 1: Safety guidelines] [Appendix 3: Safety resources]

Basic Pract Book 2006 2/11/06 11:17 am Page 1

Training in GMLP is aimed at developing proficiency in containing any uncontrolled spread of microbes in order to protect:

x practical investigations from becoming contaminated with microbes from external sources

x the operators (students, teachers and tech- nicians) from the very small possibility of infection. ( The teacher supervising the practical session must make themselves aware of any medical condition that could cause the student to be at greater risk than average in the laboratory, e. treatment with immunosuppressive drugs etc. )

It is important to arrange the workplace care- fully to ensure safe and effective operations.

[Appendix 1: Safety guidelines]

Spillage management

Spills

Spillages of cultures must be reported immediately to the teacher or technician to be dealt with quickly. The keeping of a record of all such incidents is recommended. Spilled cultures and surrounding debris (e. glass, cotton wool plugs), if any, must not be touched with unprotected hands. Wearing disposable gloves, disinfect the area by covering the spill with several layers of paper towel/cloth soaked in a suitable disinfectant (see Commonly available disinfectants and their uses , page 7) and leave for 15–30 minutes. Spill debris should then be swept into a dustpan using paper towels. All disposable material should then be transferred to a suitable container, e. an autoclave/ roasting bag, for autoclaving and disposal. The dustpan must be decontaminated either by autoclaving or by soaking (at least 24 hours) in hypochlorite (sodium chlorate I).

Broken glass

Observe an appropriate disposal procedure for broken glass if present. It should be swept carefully into a suitable container, autoclaved and disposed of in a puncture proof container.

Splashes on clothing and the skin

Contaminated clothing should be soaked in disinfectant. Splashes on the skin should be treated as soon as possible; washing thoroughly with soap and hot water should be sufficient, but if necessary the skin can be disinfected.

Hint It is useful to have a spillage kit always at hand ready for use. Suggested components: x beaker for making fresh disinfectant x disposable gloves x dustpan x paper towel/cloth x autoclave/roasting bag

Aerosols Spillages also carry a risk of generating aerosols (an invisible ‘mist’ of small droplets of moisture) which may contain microbes and might be inhaled. The risk of spillages occurring is lessened by using cultures grown on agar instead of in liquid media whenever possible. Care should also be taken to avoid generating aerosols during practical work. The risk is minimised by adhering to GMLP with special attention to the correct use of pipettes (see Inoculation and other aseptic procedures page 8).

Good microbiological laboratory practice (GMLP)

A carefully arranged laboratory bench

Resources

Equipment

Equipment Loop (wire/plastic)

Straight wire

Spreader (glass/plastic) Forceps (metal/plastic)

Pipette (calibrated/dropping; glass/plastic)

Teat Test tube

Universal bottle (wide neck); McCartney bottle (narrow neck)

Bijou bottle

Medical flat

Conical flask

Petri dish (plastic/glass)

Marker pen Personal protective equipment [Level 2, Level 3, Topics in Safety , 3rd edition (ASE, 2001), Topic 15; or Appendix 1: Safety guidelines ]

Use Routine inoculation of agar slopes/deeps and small volumes of liquid media (up to ca 10 cm 3 ); making streak plates Inoculation from very small colonies; transfer of small inocula from liquid media for nutritional work Making spread/lawn plates Transfer of sterile paper/antibiotic discs; also plant material, e. short lengths of root with nodules Transfer of measured volumes/drops of culture/sterile solutions (dry, non-absorbent cotton wool plug in neck prevents contamination) Filling and emptying pipettes safely ( never pipette by mouth) Small volumes (ca 5–10 cm 3 ) of liquid media/agar slopes/sterile solutions for inoculation (held in test tube rack; dry non-absorbent cotton wool plug or plastic cap prevents contamination) Volumes of liquid and agar media/sterile solutions up to ca 20 cm 3 for inoculation or for storing sterile media or stock cultures on agar slopes (stay upright on bench; plastic screw cap prevents contamination and reduces evaporation during long storage) Very small volumes (up to ca 3 cm 3 ) of sterile solutions (stay upright on bench; plastic screw cap prevents contamination) Large volumes of sterile media/solutions for storage; available in range of capacities, 50–500 cm 3 (plastic screw cap prevents contamination and reduces evaporation during long storage) Large volumes of liquid media for inoculation and liquid/media for short-term storage (non-absorbent cotton wool plug prevents contamination but does not reduce evaporation during long storage) Plastic : pre-sterilised for streak/spread/lawn/pour plates; Glass : only for materials for sterilisation by hot air oven, e. paper discs Labelling Petri dishes, test tubes, flasks, bottles and microscope slides Clean laboratory coat/apron : protection of clothing, containment of dust on clothing; Safety spectacles : not considered essential when dealing with suitable cultures and observing GMLP, but may be required by local regulations and for dealing with chemicals

Media, sterilisation and

disinfection

Preparation of culture media Rehydrate tablets or powder according to manufacturer’s instructions. Before sterilisation, ensure ingredients are completely dissolved, using heat if necessary. Avoid wastage by preparing only sufficient for either immediate use (allowing extra for mistakes) or use in the near future. Normally allow 15–20 cm 3 medium per Petri dish. Dispense in volumes appropriate for sterilisation in the autoclave/pressure cooker. Agar slopes are prepared in test tubes or Universal/McCartney bottles by allowing sterile molten cooled medium to solidify in a sloped position. Bottles of complete, sterile media are available from suppliers but are expensive. [Appendix 4: Suppliers of cultures and equipment]

Pouring a plate 1. Collect one bottle of sterile molten agar from the water bath. 2. Hold the bottle in the right hand; remove the cap with the little finger of the left hand. 3. Flame the neck of the bottle. 4. Lift the lid of the Petri dish slightly with the left hand and pour the sterile molten agar into the Petri dish and replace the lid. 5. Flame the neck of the bottle and replace the cap. 6. Gently rotate the dish to ensure that the medium covers the plate evenly. 7. Allow the plate to solidify. The base of the plate must be covered, agar must not touch the lid of the plate and the surface must be smooth with no bubbles. The plates should be used as soon as possible after pouring. If they are not going to be used straight away they need to be stored inside sealed plastic bags to prevent the agar from drying out.

Storage of media Store stocks of prepared media at room temperature away from direct sunlight; a cupboard is ideal but an open shelf is satisfactory. Media in vessels closed by cotton wool plugs/plastic caps that are stored for future use will be subject to evaporation at room temperature; avoid wastage by using screw cap bottles. Re-melt stored agar media in a boiling water bath, pressure cooker or microwave oven. Once melted, agar can be kept molten in a water bath at ca 50 °C until it is ready to be used. Sterile agar plates can be pre-poured and stored in well-sealed plastic bags (media-containing base uppermost to avoid heavy condensation on lid).

Sterilisation vs disinfection Sterilisation means the complete destruction of all the micro-organisms including spores, from an object or environment. It is usually achieved by heat or filtration but chemicals or radiation can be used. Disinfection is the destruction, inhibition or removal of microbes that may cause disease or other problems, e. spoilage. It is usually achieved by the use of chemicals.

Step 4

Sterilisation using the autoclave/pressure cooker

The principle of sterilisation in an autoclave or pressure cooker is that steam under pressure is used to produce a temperature of 121 °C which if held for 15 minutes will kill all micro-organisms, including bacterial endospores.

[Appendix 5: Use of the autoclave/pressure cooker]

Sterilisation of equipment and materials

xWire loop

Heat to redness in Bunsen burner flame.

xEmpty glassware and glass (not plastic!) pipettes and Petri dishes

Either : hot air oven, wrapped in either greaseproof paper or aluminium and held at 160 °C for 2 hours, allowing additional time for items to come to temperature (and cool down!). Or : autoclave/pressure cooker. Note: plastic Petri dishes are supplied in already sterilised packs; packs of sterile plastic pipettes are also available but cost may be a consideration.

xCulture media and solutions

Autoclave/pressure cooker.

xGlass spreaders and metal forceps

Flaming in alcohol (70 % IDA).

Choice, preparation and use of disinfectants

Specific disinfectants at specified working strengths are used for specific purposes. The choice is now much more straightforward as the range available from suppliers has decreased.

Commonly available disinfectants and their uses

Disinfectant Use Working strength VirKon Work surfaces, discard pots for 1 % (w/v) pipettes and slides, skin disinfection Spillages Powder Hypochlorite Discard pots for pipettes and 2,500 p.p. (0 %, v/v) (sodium chlorate I) slides available chlorine Alcohol Skin disinfection 70 % (v/v) industrial denatured alcohol (IDA)

When preparing working strength solutions from stock for class use and dealing with powder form, wear eye protection and gloves to avoid irritant or harmful effects.

Disinfectants for use at working strength should be freshly prepared from full strength stock or powder form. Activity of VirKon solution may remain for up to a week (as indicated by retention of pink colour) but less, e. 1 day, after use. Use working strength hypochlorite on day of preparation.

Using a pipette

Sterile graduated or dropping (Pasteur) pipettes are used to transfer cultures, sterile media and sterile solutions.

1. Remove the pipette from its container/ wrapper by the end that contains a cotton wool plug, taking care to touch no more than the amount necessary to take a firm hold.

2. Fit the teat.

3. Hold the pipette barrel as you would a pen but do not grasp the teat. The little finger is left free to take hold of the cotton wool plug/cap of a test tube/ bottle and the thumb to control the teat.

4. Depress the teat cautiously and take up an amount of fluid that is adequate for the amount required but does not reach and wet the cotton wool plug.

5. Return any excess gently if a measured volume is required. The pipette tip must remain beneath the liquid surface while taking up liquid to avoid the introduction of air bubbles which may cause ‘spitting’ and, conse- quently, aerosol formation when liquid is expelled.

6. Immediately after use put the now contaminated pipette into a nearby discard pot of disinfectant. The teat must not be removed until the pipette is within the discard pot otherwise drops of culture will contaminate the working surface.

Hints x A leaking pipette is caused by either a faulty or ill-fitting teat or fibres from the cotton wool plug between the teat and pipette. x A dropping (Pasteur) pipette can be converted to delivering measured volumes by attaching it to a non-sterile syringe barrel by rubber tubing. x Commercial dispensing systems are available such as measuring Pasteur pipettes. [Appendix 4: Suppliers of cultures and equipment]

Step 1a Step 1b Step 2

Step 3 Step 4 Step 6

Converting a Pasteur pipette by attaching a syringe barrel

Flaming the neck of bottles and test tubes

1. Loosen the cap of the bottle so that it can be removed easily.

2. Lift the bottle/test tube with the left hand.

3. Remove the cap of the bottle/cotton wool plug with the little finger of the right hand. (Turn the bottle, not the cap.)

4. Do not put down the cap/cotton wool plug.

5. Flame the neck of the bottle/test tube by passing the neck forwards and back through a hot Bunsen flame.

6. After carrying out the procedure required, e. withdrawing culture, replace the cap on the bottle/cotton wool plug using the little finger. (Turn the bottle, not the cap.)

Hints x Label tubes and bottles in a position that will not rub off during handling. Either marker pens or self-adhesive labels are suitable. x Occasionally cotton wool plugs accidentally catch fire. Douse the flames by immediately covering with a dry cloth, not by blowing or soaking in water.

Step 5

Step 6

Pour plate

A pour plate is one in which a small amount of inoculum from broth culture is added by pipette to the centre of a Petri dish. Molten, cooled agar medium in a test tube or bottle, is then poured into the Petri dish containing the inoculum. The dish is gently rotated to ensure that the culture and medium are thoroughly mixed and the medium covers the plate evenly. Pour plates allow micro-organisms to grow both on the surface and within the medium. Most of the colonies grow within the medium and are small in size and may be confluent; the few that grow on the surface are of the same size and appearance as those on a streak plate.

If the dilution and volume of the inoculum, usually 1 cm 3 , are known, the viable count of the sample, i. the number of bacteria or clumps of bacteria, per cm 3 can be determined. The dilutions chosen must be appropriate to produce between 30 and 100 separate countable colonies. [Appendix 6: Preparing serial dilutions]

Inoculation using a Pasteur pipette

1. Loosen the cap/cotton wool plug of the bottle containing the inoculum.

2. Remove the sterile Pasteur pipette from its container, attach the bulb and hold in the right hand.

3. Lift the bottle/test tube containing the inoculum with the left hand.

4. Remove the cap/cotton wool plug with the little finger of the right hand.

5. Flame the bottle/test tube neck.

6. Squeeze the teat bulb of the pipette very slightly, put the pipette into the bottle/test tube and draw up a little of the culture. Do not squeeze the teat bulb of the pipette after it is in the broth as this could cause bubbles and possibly aerosols.

7. Remove the pipette and flame the neck of the bottle/test tube again, before replacing the cap/cotton wool plug.

8. Place bottle/test tube on bench.

At all times hold the pipette as still as possible.

Inoculating the Petri dish

1. Lift the lid of the Petri dish slightly with the right hand and insert the pipette into the Petri dish and gently release the required volume of inoculum onto the centre of the dish. Replace the lid.

2. Put the pipette into a discard pot. Remove the teat while the pipette is pointing into the disinfectant.

Step 1

Pouring the plate

1. Collect one bottle of sterile molten agar from the water bath.

2. Hold the bottle in the right hand; remove the cap with the little finger of the left hand.

3. Flame the neck of the bottle.

4. Lift the lid of the Petri dish slightly with the left hand and pour the sterile molten agar into the Petri dish and replace the lid.

5. Flame the neck of the bottle and replace the cap.

6. Gently rotate the dish to mix the culture and the medium thoroughly and to ensure that the medium covers the plate evenly.

7. Allow the plate to solidify.

8. Seal and incubate the plate in an inverted position.

The base of the plate must be covered, agar must not touch the lid of the plate and the surface must be smooth with no bubbles.

Hints x Use a water bath at 50 °C to store bottles of molten agar. x Ensure that the temperature of the molten agar is cool enough for mixing with the culture. x Take care not to contaminate the molten agar in the bottles with water from the water bath. x To avoid contamination ensure: that the water in the water bath is at the right depth the bottles are kept an upright position that the outsides of the bottles are wiped before they are used

Step 2 Step 3 Step 4

Using a spreader

Sterile spreaders are used to distribute inoculum over the surface of already prepared agar plates.

Wrapped glass spreaders may be sterilised in a hot air oven (see Media, sterilisation and disinfection page 6). They can also be sterilised by flaming with alcohol.

Hint It is advisable to use agar plates that have a well-dried surface so that the inoculum dries quickly. Dry the surface of agar plates by either incubating the plates for several hours, e. overnight, beforehand or put them in a hot air oven (ca 55–60 °C) for 30–60 minutes with the two halves separated and the inner surfaces directed downwards.

Hints Ensure that the spreader is pointing downwards when and after igniting the alcohol to avoid burning yourself. Keep the alcohol beaker covered and away from the Bunsen flame.

Sterilisation using alcohol

1. Dip the lower end of the spreader into a small volume of alcohol (70 % IDA) contained in a vessel with a lid (either a screw cap or aluminium foil).

2. Pass quickly through a Bunsen burner flame to ignite the alcohol; the alcohol will burn and sterilise the glass.

3. Remove the spreader from the flame and allow the alcohol to burn off.

4. Do not put the spreader down on the bench.

Hint The calibrated drop (Miles & Misra) method for colony counts of pure cultures of bacteria and yeast is a more economical method than pour and spread plates. The procedure is as for the spread plate but fewer plates are needed because: (1) the inoculum is delivered as drops from a dropping pipette that is calibrated (by external diameter of the tip) to deliver drops of measured volume e. 0 cm 3 ; (2) many drops (six or more) can be put on one plate. The method is not usually suitable for mixed cultures obtained from natural samples, e. soil.

9. Replace the lid of the Petri dish.

10. Place the pipette in a discard jar.

11. Dip a glass spreader into alcohol (70 % IDA), flame and allow the alcohol to burn off.

12. Lift the lid of the Petri dish to allow entry of spreader.

13. Place the spreader on the surface of the inoculated agar and move the spreader in a top-to-bottom or a side-to-side motion to spread the inoculum over the surface of the agar. Make sure the entire agar surface is covered.

This operation must be carried out quickly to minimise the risk of contamination.

14. Replace the lid of the Petri dish.

15. Flame spreader using alcohol.

16. Let the inoculum dry.

17. Seal and incubate the plate in the inverted position.

Steps 7–8 Step 10

Step 11 Steps 12–

Working with moulds

It is sometimes appropriate to prepare a mould inoculum as a spore suspension (particular care is necessary to prevent them from escaping into the air), but often the inoculum is a portion of the mycelium taken with a loop or straight wire with the end few millimetres bent at a right angle. When an agar plate with a mould inoculated at the centre is required, it is easy to inoculate accidentally other parts of the plate with tiny pieces of mould, usually spores, that fall off the loop or wire. This can be avoided by placing the Petri dish on the working surface lid down, lifting the base (containing medium) vertically above the lid and introducing the inoculum upwards onto the centre of the downwards- facing agar surface with a bent wire.

Incubation

Note the previous comments on labelling (see Inoculation and other aseptic procedures page 11). For guidance on incubation temperatures see Appendix 1: Safety guidelines. The lid and base of an agar plate should be taped together with 2– short strips of adhesive tape as a protection from accidental (or unauthorised!) opening during incubation. Agar plates must be incubated with the medium-containing half (base) of the Petri dish uppermost otherwise condensation will occur on the lid and drip onto the culture. This might cause colonies to spread into each other and risk the spillage of the contaminated liquid. The advantages of incubators are that they may be set at a range of temperatures and reduce the possibility of cultures being interfered with or accidentally discarded. However, many cultures suitable for use in schools will grow at room temperature in the interval between lessons and can be incubated satisfactorily in a cupboard. The temperature of an incubator varies from the set temperature, oscillating by several degrees in the course of use. Water baths are used when accurately controlled temperatures are required, e. for enzyme reactions and growth-temperature relationships, when temperature control of incubators is not sufficiently precise. They should be used with distilled or deionised water to prevent corrosion and emptied and dried for storage.

Hint Overlong incubation of mould cultures will result in massive formation of spores which readily escape, particularly from Petri dishes, and may cause contamination problems in the laboratory and be a health hazard. This can occur in an incubator, at room temperature and even in a refrigerator.

Labelling a plate

Taping a plate

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