Aseptic Technique and Culture Handling

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ASEPTIC TECHNIQUE

HANDLING TUBED AND BOTTLED MEDIA

USE OF PIPETTES

• AUTOMATIC PIPETTES

• USE OF AUTOPIPETTES

USE OF THE LOOP

STREAKING A PLATE FOR SINGLE COLONIES

USE OF THE SPREADER

GENERAL TIPS ON HANDLING CULTURES

ASEPTIC TECHNIQUE

Aseptic technique is a method for working with cultures, media, specimens and equipment such that the item does not become contaminated with extraneous organisms and organisms are not transferred from the item to the environment. It is a way of handling materials which is essential in the Microbiology laboratory and is increasingly necessary in other laboratories, particularly those where fresh specimens of human origin are handled (Haematology, Clinical Chemistry, Histopathology) and also to prevent chemical contamination where molecular techniques such as PCR are performed.

There are several basic procedures which must be followed to prevent contamination:

• Keep all media, reagents and cultures closed at all times except when actually being manipulated.
• Leave all sterile instruments (pipettes, tips etc.) in containers until needed.
• Do not allow anything (especially your fingers) to come into contact with the instruments except the culture or reagent.
• Do not put closures (lids) on the bench.
• Items such as pipettes and tips should be used once only and a fresh item used for each subsequent manipulation.
• When mixing sterile reagents or media and liquid cultures, place the sterile reagent, media or diluent in the tubes first and then the liquid culture.
• Instruments such as loops and spreaders should be flame sterilized between each manipulation.
• Closures on containers should be briefly passed through the Bunsen burner flame before opening. The neck of the container should also be briefly flamed once opened. Do not allow the open part of the container to come into contact with anything. When the manipulation is completed, the neck and closure should be flamed again.
• Avoid the production of aerosols e.g. do not pipette up and down repeatedly in the same specimen and do not vigorously shake cultures before opening.
• Dispose of contaminated materials promptly into suitable containers, do not leave them lying around on the bench.
• Keep the work area clean and tidy and spray with 70% ethanol at the end of the working period. Dispose of soiled bench covers promptly (place in biohazard bins).
• In the event of a spill, notify demonstrators immediately.

HANDLING TUBED AND BOTTLED MEDIA

Handling tubed and bottled media - aseptic transfer video

Closures on sterile containers of media or other reagents should never be placed down on the bench. This means that you must be able to open the container and hold the closure in your hand such that it does not become contaminated while carrying out the required manipulation.

• Loosen but do not remove the closure of the container.
• Flame the closure by briefly passing it through the Bunsen burner flame.
• If you are right handed, pick up the loop or other instrument in your right hand and hold the container in your left hand. With the little finger of your right hand, remove the closure and keep it held by this finger. Carry out whatever manipulation is required.
• Flame the top of the container again and replace the closure.
• If you are left handed, reverse these instructions.
  
  

USE OF PIPETTES

Several different types of pipettes are used in the laboratory:

• Graduated pipettes
• Pasteur pipettes
• Transfer pipettes
• Autopipettes (e.g.Gilsons or Finns)

GRADUATED PIPETTES

Plastic disposable graduated pipettes are used primarily for tissue culture.. These pipettes should always be used with an automatic pipette filler such as an autopump (in the biosafety cabinets). Always hold these pipettes vertically when measuring the volume. Dispose of in contaminated waste bins.

PASTEUR PIPETTES

Pasteur pipettes are packed in groups of 5 into paper bags for sterilization. When required, feel the bag to determine which is the pointed end of the pipettes (arrow on the bag also indicates this, but check) and then tear the top of the bag at the opposite end. Place a teat over one pipette and carefully remove it, making sure not to touch the tip or allowing it to come into contact with anything else. To avoid aerosol production, depress the teat before placing the pipette into liquid, then draw up the liquid and transfer it. Do not allow bubbles to break in air, the tip of the pipette should be under the surface of the liquid when the material is released, unless individual drops need to be counted. Any excess should remain in the pipette and should not be put back into the original container. Discard the pipette into a sharps container, removing the teat as you do so. Do not use pipettes which have a wet cotton wool plug.

TRANSFER PIPETTES

These pipettes are similar to Pasteur pipettes, but with a built in bulb or teat and they are made of plastic rather than glass. They can be either sterile or non sterile. Sterile transfer pipettes are packed into single paper containers. Use as for a Pasteur pipette. Discard into the plastic lined beakers on the bench.

AUTOMATIC PIPETTES

A range of autopipettes is available in the laboratory. They can be found on the rack in the centre of the benches.

Laboratory Autopipettes. Click on the image to open a larger view.

USE OF AUTOPIPETTES

These pipettes must always be used in conjunction with a tip. The tip must be the appropriate size to deliver the correct volume. Note that 20 μl is the smallest volume available in these pipettes, so you will need to adjust any volumes in dilutions to account for this. Tips are supplied presterilised either in boxes or in paper bags.

• Set the required volume in the window on the side of the pipette, by turning the knob on the top of the pipette.
• Fit the appropriate tip to the pipette.
• Do not touch the tip or allow it to come into contact with anything else.
• Press the button on the top of the pipette down to the FIRST stop, place the tip into the liquid and holding the pipette vertically, SLOWLY draw up the required volume by releasing pressure on the button.
• Transfer to the new container and SLOWLY expel the liquid, pressing down to the SECOND stop on the top of the pipette. Release pressure on the button.
• Remove the pipette and hold over the discard beaker.
• Eject the tip into the beaker by pressing the bar on the side of the pipette, just below the top.

Do all manipulations slowly and carefully to avoid aerosol production and contamination of the barrel of the pipette. If liquid enters the barrel, inform the demonstrators. For sensitive techniques such as PCR, tips with cotton wool filters can be used.

Use of Autopipettes video

USE OF THE LOOP

Sterilising the loop video

The wire loop is the basic tool of microbiology. It is used for many manipulations of bacteria including subculturing, streaking for single colonies, emulsifying colonies on slides for stains and biochemical tests etc. The loop is simply a loop of wire (usually nichrome) attached to a heat proof handle. Other types of loops include platinum wire (for performing some oxidase tests), standard loops (standard 3mm diameter, used in urinalysis) and presterilised, single use plastic loops.

The wire loop must always be sterilized prior to use. It is sterilized by heating in a Bunsen burner flame until it is red hot.

• Adjust the Bunsen flame until 2 cones are visible.
• Insert the loop into the flame:

• Gradually draw the wire through the flame until the loop is just below the tip of the lower cone.
• SLOWLY raise the loop until almost in the vertical position (without burning your hand) and in the outer cone:

When the loop and the lower 2-3 cm of the wire are red hot, remove the loop and place upright in a rack to cool. Alternatively, you can cool the loop by touching it onto a sterile part of an agar plate or the moisture in the lid of a plate. Use only when cooled as a red hot loop may kill your culture. Inserting a hot loop into a liquid culture may cause it to sputter and produce aerosols. Additionally, placing a wet loop into a flame may also cause sputter, hence the need to slowly place the loop into the flame.

Flaming the loop video

STREAKING A PLATE FOR SINGLE COLONIES:

When working with cultures, it is usually desirable to produce single colonies on a plate as these represent a clone of a single organism and therefore represent a “pure” culture, i.e. one type only. Most cultures contain organisms in very high concentrations and so must be plated in such a way as to gradually reduce the number until individual organisms remain. There are many different ways of streaking a plate for single colonies but the aim is to reduce the numbers streaked out with each subsequent step. The most important thing to consider is that the loop must be flamed between each streak or an unused portion of the loop should be used for each streak.

1. With a sterilized loop, pick up a colony (or part of a colony) from an agar plate or a loopful of liquid culture.
2. Inoculate to a third of a fresh agar plate by rubbing the loop back and forth across the agar. Take care not to scratch up the surface of the agar.

3. Flame the loop. (This step between the initial inoculum and subsequent steps is very important as the highest concentration of organisms is now on the loop.)
4. From one end of the initial inoculum, draw out 4-5 streaks with the loop.

5. Now either flame the loop again or turn it over, rotate the plate and do another 4-5 streaks from the previous ones.

6. Repeat the flaming and streaking once more. Finish with a final “squiggle” on the remaining portion of the plate.

7. A swab can be similarly inoculated to an agar plate by using the swab to produce the initial inoculum (rolling the swab as it is being inoculated to the plate) and then using the loop to streak out for single colonies.
8. A loop can also be used to transfer liquid inocula from one medium to another, to inoculate sloped or tubed media (streak the slope and stab the butt), and to prepare smears for staining (see section on microscopy).

USE OF THE SPREADER

A spreader is a triangular shaped piece of thick wire, larger than a loop, with the base of the triangle on the bottom and the apex forming a handle.

The spreader is considerably thicker than a loop and cannot be sterilized by heating in the Bunsen flame as the handle would become too hot. It is sterilized by dipping in ethanol, passing through the flame to ignite the ethanol, and then holding while the ethanol burns off. Care must be taken not to allow the burning ethanol to drip onto the bench or onto fingers.

The aim of using a spreader is to produce a uniform “lawn” of growth on an agar plate which consists of a confluent growth of the organisms i.e. no spaces between the colonies. Lawn plates are used for sensitivity testing, certain growth factor testing and for some bacteriophage work. It is difficult to produce the uniform confluent growth required using the small surface area of the loop, hence the larger area of the spreader.

1. Place an aliquot of a liquid inoculum ready for spreading on an agar plate.
2. Dip the base of the spreader into a small beaker of 95% ethanol. Drain excess ethanol back into the beaker.
3. Pass the spreader briefly through the flame of the Bunsen and allow the ethanol to burn off. Do not allow the ethanol to drip off and do not touch anything with the spreader once sterilized.
4. Place the base of the spreader into the inoculum on the plate and spread it by rubbing the spreader back and forth across the plate, rotating the plate as you do so. Spread in at least three different directions to ensure good coverage. Finish with a sweep around the rim of the plate.

5. Replace the spreader into the beaker of ethanol ready for the next use. Do not pass through the flame again until required. Do not heat the spreader in the flame and then place it in ethanol as a hot spreader can ignite the beaker of ethanol. If this occurs, place a heat proof mat over the beaker until the flame goes out. DO NOT TOUCH THE BEAKER – IT WILL BE HOT!

 Making a lawn plate with the spreader video

GENERAL TIPS ON HANDLING CULTURES

1. ALWAYS label the container of media and not the lids. Lids can be lost or mixed up. Labels should include:

• Your bench number or name
• Date
• Organism
• Any special conditions e.g. anaerobic/25^oC
• Any special procedures e.g. a dilution factor
• If a clinical specimen, the number of the specimen

2. Keep media closed at all times unless performing some manipulation.

3. Incubate all agar plates upside down except if they contain liquids. Organisms, antibiotic discs etc. will not fall off. Incubating this way prevents condensation falling onto the culture.