Working with microbes: Safety, aseptic technique and use of equipment

The microbial experiments in this lab will involve Escherichia coli K12. E. coli K12 is a Risk Group 1 organism that can be worked with in an open lab, as long as proper aseptic technique is followed.

Disposal of waste

Disposable materials (e.g. micropipette tips, toothpicks, disposable plastic inoculation loops, gloves, plastic cuvettes and plastic test tubes) contaminated with Risk Group 1 organisms can be disposed of in the buckets containing clear plastic bags on your bench. Do not put any glass in the buckets on the bench. Contaminated broken glassware and syringes should be disposed of in the “sharps” containers at the far end of the bench. Gloves do not have to be disposed of in biohazard waste containers unless they have become contaminated with microbial cultures. Gloves that have become contaminated with hazardous chemicals should be discarded in the designated containers. Contaminated pipettes should be disposed of in the rectangular white boxes on your bench. Culture supernatants contaminated with microbes should be poured or pipetted into an Erlenmeyer flask. Contaminated flasks, tubes, and plates that have had microbes in them should be discarded in the appropriate containers located near the door of the lab. Tape labels should be removed from glassware before disposal. Do not pour contaminated media, buffers, etc. out of flasks or tubes unless it is necessary for the experiment.

Aseptic Technique

Aseptic technique is a set of procedures that must be followed when working with microbes to prevent the contamination of the experimenter, the environment, or the experimental cultures. Bacteria worked with in research labs may be potential pathogens, so it is important not to expose yourself accidentally to these bacteria. It is also important to avoid the release of experimental cultures into the lab environment. These strains may contain genetic markers such as antibiotic resistance, which should not be released into the environment. Finally, it is important for the experimental cultures themselves not to become contaminated with microbes from the environment, which will result in confusing and invalid experimental results. Microbes are widely distributed in dust, skin, dandruff, and nasal secretions, so it is very easy for cultures to become contaminated unless steps are taken to minimise the risks of these agents coming in contact with experimental materials.

Procedures

1. Lab coats must be worn.

2. Wash hands at beginning and end of lab.

3. Avoid touching your face or mouth while in the lab. This will reduce the risk of infecting yourself, and also the risk of contaminating your cultures with dandruff.

4. Do not touch any contaminated materials or the contaminated parts of equipment. Also avoid touching any materials or equipment parts that are supposed to be sterile or that will come into contact with your experimental cultures.

5. If you have any cuts, scrapes, burns etc. let the TAs know about it before or during the lab.

6. Wipe down bench with disinfectant (ethanol) before and after lab work.

7. Wipe up any spills or drips (including drips on the lips or sides of a flask or tube resulting from pouring) using a paper towel freshly soaked in ethanol. Soak any contaminated broken glassware with ethanol, and notify one of the TAs about it.

8. Avoid creating aerosols by spraying liquid out of a pipette. The pipette tip should be placed against the side of the flask or tube while liquid is being ejected.

9. When inoculating, pipetting, or pouring material from a glass tube or flask, reduce the risk of contamination by passing the rim of the flask or tube briefly through a Bunsen burner flame before and after the transfer. If you are pouring from a tube or flask, allow the rim to cool for a second before letting liquid come into contact with it. If a drop remains on the rim of the tube or flask after pouring, wipe it off using a paper towel freshly soaked in ethanol before flaming the rim. This avoids the creation of aerosols by the splattering of boiling liquid. Do not flame pipettes, plastic tubes, or other plastic equipment.

10. Glass or metal spreading rods are used to spread liquid cultures evenly over the surface of agar plates. These rods are sterilized by dipping in ethanol, tapping off the excess, and then igniting the ethanol in a Bunsen burner flame. Let the rod cool for a couple of seconds before touching the agar surface. This will avoid creating aerosols from the liquid cultures that the plate has been inoculated with. Do not hold the burning rod over the ethanol stock, and avoid lifting the burning end above the end you are holding. (This may allow burning ethanol to run down onto your fingers.) If you do ignite the ethanol stock, quickly cap it to smother the flames.

11. The longer that experimental cultures or equipment are in contact with the open air, the greater the chance that they will become contaminated by dust or aerosols. Because of this, transfers between tubes and/or plates should be carried out as quickly as safety and accuracy allow. This can be accomplished by understanding what you have to do before opening the tube or plate, and by not leaving the tube or plate open between transfers. In addition, sterile equipment should be kept sterile until right before use. The best way to do this is to leave the equipment in its package (e.g. wrapper or closed micropipette tip box) until you are ready to carry out the transfer. Do not set sterile or contaminated equipment down on your lab bench or other surfaces. This includes caps or plugs for culture tubes or flasks, which should be held with the little finger of your transfer hand during transfers. (The transfer hand is the one holding the pipettor or inoculating loop.)

Equipment (per group)

There are 14 workstations set up in the lab. Each group of 2-3 students will use the same workstation in each lab period. Under the counter at each station, there will be a cabinet containing the following supplies, which will be shared by 3 groups (working on different days):

1 p1000, 1p100, 1p20, 1p10

1 bottle ethanol and “hockey stick” spreader

labelling tape and pen

1 jar of sterile 0.5 ml Eppendorf tubes

(These are for the PCR lab)

Each individual group will also have one of the drawers next to the cabinet. The drawer should contain the following supplies:

boxes of sterile p1000, p100/p20, and p10 tips

1 jar of sterile 1.5 ml Eppendorf tubes

Each group’s equipment will be reused in subsequent labs, so handle it properly and report any problems with the equipment. You will be expected to return empty tip boxes to the designated tray when you obtain fresh sterile tips. Empty Eppendorf tube jars should be refilled (by you), labelled (group and day) and placed on the designated “To Be Autoclaved” tray.

Micropipettor use

Each type of micropipettor has a range in which it can be used to measure volumes accurately. Pipettors should not be used to measure volumes outside of this range. Volumes larger than 1 ml should be measured using disposable 5 ml pipettes.

p1000 = 200 microlitres to 1.000 ml

p200 = 20.0 microlitres to 200.0 microlitres

p20 = 2.00 microlitres to 20.00 microlitres

p10 = 0.50 microlitres to 10.00 microlitres

When using micropipettors, depress the plunger to the first stop, and insert it in the solution being transferred. Smoothly and gradually draw back the plunger to draw solution up into the tube. (If the plunger is released rapidly, aerosols will be created and an inaccurate volume will be transferred.) Transfer the solution to the new tube, and eject the solution by gradually depressing the plunger. The remaining drop can be ejected by depressing the plunger to the second stop. Aerosols can be minimized by touching the tip to the side of the tube before blowing out the final drop. (This procedure will be demonstrated in the lab.)

Observe the volumes in the tip during transfer – with practise, you will be able to recognise incorrect volumes and repeat the transfer properly. This is especially important when pipetting viscous solutions like genomic DNA, which will be drawn up into the tip at a slower rate. Liquids with low surface tension (such as organic solvents) may drip from the tip during transfer. This can be minimized by holding the pipettor at a 45° angle while transferring the solution.