By Joni Frederick
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1.1. General Lab Safety and Rules
- Fire & emergency procedures & exits
- Eyewash stations
- First aid kits
- Labcoats and protective gear
- Laminar flow/ biological safety and fume-hoods
A number of chemicals used in any molecular biology laboratory are hazardous. All manufacturers of hazardous materials are required by law to supply the user with pertinent information on any hazards associated with their chemicals. This information is supplied in the form of Material Safety Data Sheets or MSDS. This information contains the chemical name, CAS#, health hazard data, including first aid treatment, physical data, fire and explosion hazard data, reactivity data, spill or leak procedures, and any special precautions needed when handling this chemical. A file containing MSDS information on the hazardous substances should be kept in the lab. In addition, MSDS information can be accessed on World Wide Web. You are strongly urged to make use of this information prior to using a new chemical and certainly in the case of any accidental exposure or spill.
The instructor/lab manager must be notified immediately in the case of an accident involving any potentially hazardous reagents.
The following chemicals are particularly noteworthy:
- Phenol – can cause severe burns
- Acrylamide – potential neurotoxin
- Ethidium bromide – carcinogen
These chemicals are not harmful if used properly: always wear gloves when using potentially hazardous chemicals and never mouth-pipet them. If you accidentally splash any of these chemicals on your skin, immediately rinse the area thoroughly with water and inform the instructor. Discard the waste in appropriate containers.
1.3. Ultraviolet Light
Exposure to ultraviolet light can cause acute eye irritation, and “sunburn” on exposed skin. Since the retina cannot detect UV light, you can have serious eye damage and not realize it until 30 min to 24 hours after exposure. Therefore, always wear appropriate eye protection when using UV lamps.
Exposure to ultraviolet light can cause acute eye irritation, and “sunburn” on exposed skin. Since the retina cannot detect UV light, you can have serious eye damage and not realize it until 30 min to 24 hours after exposureThe voltages used for electrophoresis are sufficient to cause electrocution. Cover the buffer reservoirs during electrophoresis. Always turn off the power supply and unplug the leads before removing a gel.
1.5. General Housekeeping
All common areas should be kept free of clutter and all dirty dishes, electrophoresis equipment, etc should be dealt with appropriately. Since you have only a limited amount of space to call your own, it is to your advantage to keep your own area clean. Since you will use common facilities, all solutions and everything stored in an incubator, refrigerator, etc. must be labeled. In order to limit confusion, each person should use his initials or other unique designation for labeling plates, etc. Unlabeled material found in the refrigerators, incubators, or freezers may be destroyed. Always mark the backs of the plates with your initials, the date, and relevant experimental data, e.g. strain numbers.
1.6. Disposal of Buffers and Chemicals
- Any uncontaminated, solidified agar or agarose should be discarded in the trash, not in the sink, and the bottles rinsed well.
- Any media that becomes contaminated should be promptly autoclaved before discarding it. Petri dishes and other biological waste should be discarded in Biohazard containers which will be autoclaved prior to disposal.
- Organic reagents, e.g. phenol, should be used in a fume hood and all organic waste should be disposed of in a labeled container, not in the trash or the sink.
- Ethidium bromide is a mutagenic substance that should be treated before disposal and should be handled only with gloves. Ethidium bromide should be disposed of in a labeled container.
- Dirty glassware should be rinsed, all traces of agar or other substance that will not come clean in a dishwasher should be removed, all labels should be removed (if possible), and the glassware should be placed on the dirty dish counter. Bottle caps, stir bars and spatulas should not be placed in the bins but should be washed with hot soapy water, rinsed well with hot water, and rinsed three times with distilled water.
2. Buffers and Solutions
2.1. Calculation of Molar, % and “X” Solutions
- A molar solution is one in which 1 liter of solution contains the number of grams equal to its molecular weight. Eg. To make up 100 ml of a 5M NaCl solution = 58.456 (mw of NaCl) g/mol x 5 moles/liter x 0.1 liter = 29.29 g in 100 ml of solution
- Percent solutions. Percentage (w/v) = weight (g) in 100 ml of solution; Percentage (v/v) = volume (ml) in 100 ml of solution. Eg. To make a 0.7% solution of agarose in TBE buffer, weight 0.7 of agarose and bring up volume to 100 ml with TBE buffer.
- “X” Solutions. Many enzyme buffers are prepared as concentrated solutions, e.g. 5X or 10X (five or ten times the concentration of the working solution) and are then diluted such that the final concentration of the buffer in the reaction is 1X. Eg. To set up a restriction digestion in 25 μl, one would add 2.5 μl of a 10X buffer, the other reaction components, and water to a final volume of 25 μl.
2.2. Preparation of Working Solutions from Concentrated Stock Solutions
Many buffers in molecular biology require the same components but often in varying concentrations. To avoid having to make every buffer from scratch, it is useful to prepare several concentrated stock solutions and dilute as needed. Eg. To make 100 ml of TE buffer (10 mM Tris, 1 mM EDTA), combine 1 ml of a 1 M Tris solution and 0.2 ml of 0.5 M EDTA and 98.8 ml sterile water. The following is useful for calculating amounts of stock solution needed: Ci x Vi = Cf x Vf , where Ci = initial concentration, or conc of stock solution; Vi = initial vol, or amount of stock solution needed Cf = final concentration, or conc of desired solution; Vf = final vol, or volume of desired solution.
Concentrated solutions, e.g. 1M Tris-HCl pH=8.0, 5M NaCl, can be used to make working stocks by adding autoclaved double-distilled water in a sterile vessel to the appropriate amount of the concentrated solution.
2.3. Glassware and Plastic Ware
Glass and plastic ware used for molecular biology must be scrupulously clean. Dirty test tubes, bacterial contamination and traces of detergent can inhibit reactions or degrade nucleic acid. Glassware should be rinsed with distilled water and autoclaved or baked at 150°C for 1 hour. For experiments with RNA, glassware and solutions are treated with diethyl-pyrocarbonate to inhibit RNases which can be resistant to autoclaving. Plastic ware such as pipets and culture tubes are often supplied sterile. Tubes made of polypropylene are turbid and are resistant to many chemicals, like phenol and chloroform; polycarbonate or polystyrene tubes are clear and not resistant to many chemicals. Make sure that the tubes you are using are resistant to the chemicals used in your experiment. Micro pipette tips and microfuge tubes should be autoclaved before use.
Biological functions are very sensitive to changes in pH and hence, buffers are used to stabilize the pH. An accurate pH reading depends on standardization, the degree of static charge, and the temperature of the solution. pH is the quantitative measure of the acidity or basicity of aqueous or other liquid solutions. The pH is determined by measuring the concentration of the hydrogen ion, denoted [H+]. [H+] ranges between approximately 1 and 10-14 gram-equivalents per litre. In pure water, which is neutral, the [H+] is 10-7 gram-equivalents per litre, which corresponds to a pH of 7. A solution with a pH less than 7 is considered acidic; a solution with a pH greater than 7 is considered basic, or alkaline. pH is calculated as the negative log of the H+ concentration ( pH = -log[H+] ).
Calibrating a pH Meter and Measuring the pH of a Solution:
Calibration of the pH meter – Remove the pH meter probe from the pH 7 Standard Solution, rinse the probe with a gentle stream of ddH2O, remove any excess liquid from the probe tip, place the probe in the pH 7 Standard Solution, adjust the meter to read a pH of 7.0, remove the probe from the solution, rinse the probe with a gentle stream of ddH2O and remove any excess liquid from the probe tip. Place the probe in the pH 4 Standard Solution, adjust the meter to read a pH of 4.0 (If the solution is basic, read the second at pH 9).
Measuring the pH of a Solution – Rinse the probe with a gentle stream of ddH2O and remove any excess liquid from the probe tip, place the probe into the solution, use the meter to measure the pH of the solution and record the measured pH.
- An undiluted solution is termed the 100 dilution. This may be a chemical solution, or a suspension of bacteria or bacteriophage
- Typical diluions are 10-fold dilutions; each dilution is 1/10th the concentration of the previous dilution. The first 10-fold dilution is termed the 10-1 dilution
- Each subsequent dilution is made from a previous dilution
Example: Preparation of Phosphate Buffered Saline (PBS)
4.3 mM Sodium Phosphate, Dibasic (Na2HPO4)
137 mM Sodium Chloride (NaCl)
2.7 mM Potassium Chloride (KCl)
1.4 mM Potassium Phosphate, Monobasic (KH2PO4)
- Measure a volume of 800 ml of ddH2O with a graduated cylinder and transfer to an Erhlenmyer flask.
- Add a magnetic stir bar to the Erhlenmyer flask and place the flask on a magnetic stir plate. Adjust the speed of the magnetic stir bar so that oxygen is not introduced into the solution while it is rapidly mixed.
- Transfer to the flask:
8 g of NaCl, 0.2 g KCl, 1.44 g of Na2HPO4 and 0.25 g of KH2PO4
- Allow the solutes to dissolve for 3 to 5 min.
- Ensure that there are no remaining particles of undissolved salts in the solution before adjusting the pH. If particles are present, continue stirring.
- Reduce the speed of the magnetic stir bar so that the solution is gently mixing.
- Ensure that the pH meter has been properly calibrated and rinse the pH probe with ddH2O. Remove the excess water from the probe tip (without touching the probe tip) with clean paper towel. Place the pH probe into the solution.
- Slowly add 1 M HCl dropwise with a transfer pipette and allow the HCl to fully dissolve into the solution.
- Measure the pH with the pH meter.
- Repeat Steps #8 through #10 until the pH of the solution is 7.4.
- Pour the solution into a fresh graduated cylinder and adjust the final volume to 1 liter with ddH2O.
- Transfer the solution to a container that is suitable for autoclaving.
- Autoclave the solution.
- Allow the solution to cool to room temperature.
- Store the PBS solution at room temperature. The PBS solution is sterile; when using the PBS Solution, ensure that sterile techniques are employed.
3. Aseptic/Sterile Technique
Aseptic procedures include the use of surgical gloves, masks, sterile instruments and aseptic technique.
- An agent or action that kills or inhibits the growth of micro-organisms.
- A chemical agent that is applied topically to inhibit the growth of micro-organisms.
- Prevention of microbial contamination of living tissues or sterile materials by excluding, removing or killing micro-organisms.
- A steam sterilizer consisting of a metal chamber constructed to withstand the pressure that is required to raise the temperature of steam to the level required for sterilization. Early models were termed “autoclaves” because they were fitted with a self-closing door.
- A chemical or physical agent that kills vegetative (non-spore forming) bacteria.
- An agent that prevents multiplication of bacteria.
- Non-pathogenic micro-organisms that are living and reproducing as human or animal parasites.
- Introduction of micro-organisms to sterile articles, materials or tissues.
- An agent that is intended to kill or remove pathogenic micro -organisms, with the exception of bacterial spores.
- A process that kills nonspore-forming micro-organisms by hot water or steam at 65-100oC.
- A species that is capable of causing disease micro-or ganism in a susceptible host.
- A process that reduces microbial contamination to a low level by the use of cleaning solutions, hot water or chemical disinfectants.
- An agent that kills all types of micro-organisms.
- Free from micro-organisms.
- The complete destruction of micro-organisms.
3.2. Autoclaving/steam sterilization
Steam sterilization/autoclaving depends on the use of steam above 100°C. Temperatures ranging from 121-134°C at pressures of 15-30 psi are generally recommended. The biocidal action of moist heat is a denaturation of major cell constituents. In the first stage of sterilization, air is evacuated and the chamber brought to the pre-set sterilizing temperature, which is maintained for a holding period sufficient to kill all microbial contaminants. Minimum holding times for the sterilization of medical equipment for example are 15 minutes at 121°C, 10 minutes at 126°C, and 3 minutes at 134°C. Most broths and microbial cultures are held at 120°C for 20 minutes. The steam is then removed and the contents allowed to dry or cool. The chamber is then pressured by the introduction of filtered air.
The colour change on a chemical dye impregnated indicator strip placed within the pack can provide a convenient and rapid visual check that the appropriate sterilization conditions were reached.
4. Media Preparation
Bacteria and fungi are grown on or in microbiological media of various types. The medium that is used to culture the microorganism depends on the microorganism that one is trying to isolate or identify. Other added ingredients may be growth factors, NaCl, and pH buffers which keep the medium from straying too far from neutral as the microbes metabolize.
4.1. General Procedure
- Carry out all culturing operation in a laminar flow hood.
- Disinfect all surfaces prior to use with a disinfectant solution.
- Swab down the working surface liberally with 70% ethanol.
- Periodically spread a solution of 70% ethanol over the exterior of gloves to minimize contamination. Replace them if torn.
- In case of any spill, spread a solution of 70% alcohol and swab immediately with non-linting wipes.
- Discard gloves after use and do not wear them when entering any other lab area.
- Bring into the work area only those items needed for a particular procedure.
- Leave a wide clear space in the center of the hood (not just the front edge) to work on. Do not clutter the area to prevent blockage of proper air flow and to minimize turbulence.
- Swab with 70% alcohol all glassware (medium bottles, beakers, etc.) before placing them inside the hood.
- Arrange the work area to have easy access to all of it without having to reach over one item to get at another (especially over an open bottle or flask).
- Use sterile wrapped pipets and discard them after use into a biohazard waste container.
- Check that the wrapping of the sterile pipet is not broken or damaged.
- Inspect the vessels to be used:
- T-flask – Must be free from visible contamination or breakage, or lack container identification. The plastic covering the flask must be intact.
- Bottles – Check for cracks, expiration dates.
- Spinner flasks – Check for cracks, expiration dates, and proper assembly.
- Discard any biohazard or contaminated material immediately.
- Never perform mouth pipetting. Pipettor must be used.
- When handling sterile containers with caps or lids, place the cap on its side if it must be laid on the work surface.
- Make sure not to touch the tip of the pipette to the rim of any flask or sterile bottle.
- Clean the work area when finished by wiping with 70% alcohol.
4.2. Luria Broth
Minimal media has no added sugars whilst defined media have particular growth factors, nutrients and vitamins added in certain amounts.
- rehydrate the powder form of the medium
- stir and possibly boil the medium to get the agar dissolved well
- distribute the media into tubes
- autoclave to sterilize the tube media
- autoclave the agar medium for plate production and then pour into sterile petri dishes
4.2.1. Inoculation of Media
Using aseptic technique, media can be inoculated using a sterile loop after flaming. A single colony may be picked from a streak or spread plate and the loop immersed in the media. A measured volume of another cultured bacterial sample may be added to the media. Media is generally incubated at 37°C for 12-16 hours, depending on the type of culture.
Set the petri dishes out, RIGHT SIDE UP (small side down), tops covering the dishes. Cover the bottom of each plate approximately 2/3 of the way across, put on the lid and gently rotate the plate to distribute the agar across the entire dish bottom. Allow all plates to stand until they are completely solidified. Once they have solidified, place them upside down for storage. Agar plates are always placed UPSIDE DOWN (bottom dish with agar on top) for two reasons. Although there may be air contaminants in the incubator, it will be more difficult for them to get into the plates, and then move up to the agar to settle since gravity is working against this movement. Often you will see a bit of water condensation on the top of the agar plate, particularly when they have been at 37°C. The water molecules are cohesive and tend to run together. If the plate was sitting right side up, the water droplets could then fall onto the agar, creating a kind of little lake on the agar plate and messing up your plates. Upside down plates prevents the condensation from dropping on the agar surface.
4.3.1. Spread Plates:
- Add a measured aliquot of inoculum to a sterile medium plate
- dip the glass spreader in ethanol and flame. Make sure all the ethanol has burnt off. Allow to cool for a few seconds
- while lifting the plate lid and rotating with the same hand, use the glass spreader to evenly distribute the inoculum over the media
- close the plate and dip the spreader in ethanol
4.3.2. Streak Plates:
- Pick up a loopful of your inoculum from either a broth or an agar culture. Using a sterile agar medium plate (lift the lid just enough to insert the loop), streak a vertical line straight down
- When streaking the agar, keep the loop horizontal and only streak the surface of the agar: DO NOT DIG INTO THE AGAR
- Move the loop in a zig-zag pattern across the agar until 1/3 of the plate is covered, finishing the first section
- Sterilize the loop in the flame and let it cool before continuing to spread the bacteria. You can do this by 1) sticking the hot loop in the agar at the edge of the agar away from the bacteria, or 2) just holding the loop for a few seconds while it cools
- Rotate the plate about 90 degrees and spread the bacteria from the first streak into a second area using the same zig-zag spread technique
- Sterilize the loop again. Rotate the plate about 90 degrees and spread the bacteria from the second streak into the 3rd area in the same pattern
- Sterilize the loop again. Replace the lid and invert the plate. Incubate the plate
Ampicillin: Generally a 50 mg/mL working stock solution in water. Used at a final concentration of 20 μg/ml. Filter sterilize; or prepare from the sterile water and the sterile powder, store at -20°C in small aliquots (avoid more than 10 freezing-thawing cycles), it is possible to use 50% glycerol instead of H2O. The advantage – solution is liquid at -20°C, some laboratories use 96% (or 50%) EtOH for ampicillin solubilization. There are two advantages in this method: (i) solution is liquid at -20°C, (ii) the solution is self sterilizing. But the disadvantage is very serious: ethanol is a weak catalyst of β-lactam ring disintegration. So, solution is unstable.
Chloramphenicol: Generally a 34 mg/mL working stock solution in ethanol or methanol. Used at a final concentration of 25 μg/ml. Store at -20°C in small aliquots.
- Biohazard bins
- Bleach or other bacteriocide
4.6. General safety rules when working with microbes
- NO EATING OR DRINKING IN THE LAB!
- ALWAYS disinfect the tables BEFORE and AFTER
- Wash your hands with soap both BEFORE and AFTER, and, in addition, when you have a spill
- Be sure to have nonessential materials (the ONLY essential thing is the lab handouts and a notebook to write in) off of the table
- ALWAYS use the proper aseptic technique when transferring cultures from one medium to another
- Heat the entire piece of metal of the inoculation instrument in the flame: it should be RED HOT. Be sure to COOL your inoculation instrument before picking the inoculum
- Use an inoculating needle for agar deeps and an inoculating loop for the agar plate and the broths
- Label all test tubes and petri plates with your name (initials), date, and name of organism
- Do not dump ANY microbial suspension down the drain
- Keep test tube caps and petri dish covers on media to reduce contamination (whether it is sterile media or already cultured)
Most of the experiments you will conduct in the laboratory will depend on your ability to accurately measure volumes of solutions using micropippettors.
5.1. General Tips
RULE OF THUMB: Always select the SMALLEST size pipette that will handle the volume you wish to move to achieve the greatest accuracy. Accuracy decreases as you use unnecessarily large pippets for small volumes.
LOADING: Load a sterile tip (blue for P1000; yellow for P200) and then reclose the tip box to maintain sterilitiy.
- Push the plunger down slowly to the point of first resistance: this sets the load volume.
- While holding the plunger at the load volume set point, put the tip into the solution so that it is immersed just enough to cover the end (3-4 mm).
- Slowly release the plunger to draw up the liquid making sure to keep the tip immersed. Visually inspect the load to make sure it is correct.
DELIVERY: To deliver the volume, place the tip into the receiving vessel and press the plunger all the way to the bottom – this expells all the liquid and gives a little extra volume to get the last droplet out – AND THEN, WITHOUT RELEASING THE PLUNGER, withdraw the tip.
CHANGE TIP? Repeat as necessary using the same tip if NOT changing solutions. Use a fresh tip for every change of solution, or whenever it is prudent to maintain sterile conditions and avoid cross contamination of solutions or cultures.
TIP DISCHARGE: While holding the tip over an appropriate waste receptacle, press the white tip discharge slider on the back of the grip.
SMALL VOLUMES TECHNIQUE: With small volumes, especially the 1-10 ul range used in molecular biology protocols, you must keep track of the droplets you pipet. Carefully expell the liquid droplet on the side wall of the tube so that you can see it. If adding to a larger volume, flush the tip with the solvent liquid after expelling the droplet to make sure you get all the delivery liquid. With small volumes you’ll usually need to centrifuge and then vortex the tube to get a good mixing of the reagents.
Check the calibration of your micropipet by using the fact that 1 ml of deionized (or distilled) water has a mass of 1 g. Pipet a range of volumes spanning the micropipet’s usuable range and mass them on a top loading balance having at least 3 decimal place accuracy. Pipets having greater than 5% error should be recalibrated.