General Protocols

Agarose Gel Electrophoresis

1. Add TAE Buffer (100 mL) and Agarose (1 g) to a flask [for a 1% gel]
2. Cover with plastic wrap, poke hole in top. Then microwave flask until agarose is fully dissolved; avoid boiling
3. Take flask to fume hood, and allow to cool to touch. Add REDsafe (4 μL) to agarose, gently swirl to mix
4. Gently pour agarose into assembled gel casting tray, removing any bubbles with a pipette tip
5. Allow gel to solidify until translucent. Then transfer to running apparatus filled with TAE buffer
6. Load samples containing 3 μL loading dye and ~10-15 μL of DNA
7. Run gel at 110 V until the dye is ~2/3 of the way down the gel (approx. 40 mins)

Preparing Chemically Competent E. coli Cells

1. Inoculate 5-10 mL LB with Top10 E. coli culture at 37 °C shaking over night
2. Subculture 1 mL of bacteria in 50mL LB at 37°C shaking until OD₆₀₀ is 0.4-0.6 (~2.5 hr)
3. Centrifuge the subculture at 13000xg for 10 min at 4 °C. Discard supernatant.
4. Resuspend pellet in 12.5 mL of cold 50 mM CaCl₂. Incubate on ice for 10 min.
5. Repeat centrifugation and discard supernatant.
6. Resuspend pellet in 2 mL of cold 50 mM CaCl₂, 15% glycerol.
7. Aliquot competent cells and store at -80°C

Preparation of Antibiotic Agar

• LB agar
• Plates

1. Weigh 35 g of LB-Agar powder mix per litre of media desired. One litre makes 40-50 plates. Ensure that that the mixture volume does not exceed half of the volume of the flask/contained used, as it will boil over in the autoclave.
2. Dissolve LB-Agar, using water from one of the wall mounted Nanopure filters. Add a stir bar and use a magnetic stirrer to facilitate mixing.
3. Cover the flask with aluminum foil, and secure the foil with autoclave tape. The foil should be somewhat loose (to avoid building pressure in the flask while sterilizing and blowing the foil off), but not so loose that lots of liquid can escape.
4. Put the flask in a plastic autoclave tray, load into the autoclave, and sterilize using the 20 minute liquid program.
5. Once the autoclave finishes venting (which can take twice as long as the sterilization proper), check that the foil covering is still in place. If it is not, the media is contaminated! Unload using the insulated oven gloves.
6. Allow the media to cool until it can be handled without the oven mits. The cold room can be used to speed this up. Alternatively, if a large batch of media is prepared flasks may be kept hot in the prep lab water bath, to avoid all of them cooling at once. Agar polymerization cannot be reversed once it starts, but media can be kept from solidifying by keeping it hot.
7. Once media is cool, add other desired ingredients. Use the magnetic stirrer to mix, but do NOT add a stir bar now, or the media will be contaminated. (If one wasn't added before, you must do without.)
8. Pour agar into plates

Common additions include:

1. Ampicillin (stock 100 mg/ml, final 100 μg/ml)
2. Kanamycin (stock 50 mg/ml, final 50 μg/ml)
3. Chloramphenicol (stock 50 mg/ml, final 10 μg/ml)
4. Spectinomycin (stock 100 mg/ml, final 100 μg/ml)
5. To achieve final concentrations, add 1 mL of stock per 1 L of media, except for chloramphenicol, where 0.6 mL per 1 L of media is added instead.
6. Pour directly from the flask into sterile petri plates. Use a quick pass with a Bunsen burner flame to eliminate any bubbles that form during pouring. Do not subject the plate to continuous heat or the plate will melt, and the heat sensitive ingredients added in the previous step will be destroyed. Bubbles can allow cells to access nutrients without being exposed to the plate's antibiotic, and should be blown out immediately before the gel can set. It's a good idea for one person to pour while another flames bubbles.
7. Allow the plates to stand right side up overnight, or until the gel sets if they are needed sooner. Plates should be stored upside down to prevent condensation from falling on the media. Store petri plates in the plastic bags they ship in, in the 4 degree cold room.

Overnight Cultures (per culture tube)

• 10 mL culture tube (16 mm x 160 mm or 16 mm x 125 mm) or 15 mL Falcon tube
• 5 mL LB
• 5μL 1000X antibiotics
• Single colonies on a plate (best not to start an overnight from a glycerol stock)

1. Add 3 mL sterile/autoclaved LB in a 15 mL Falcon tube
2. Pipet 3μL of 1000X antibiotic into the LB
3. Select a single colony using a sterile toothpick or pipette tip
4. Place toothpick or pipette tip in the culture tube and stir
5. Remove toothpick, or in the case of a pipette tip, leave in the tube
6. Place culture tube in incubator at 37 °C overnight shaking vigorously (250 RPM)

Glycerol Stocks

• Overnight bacterial growth
• Screw cap tubes
• Glycerol

1. Pipet 0.5 mL of glycerol into two 1.5 mL screw cap tubes
2. Add 0.5 mL of overnight culture to each tube
3. Pipet up and down to gently mix
4. Place one tube in -20 °C freezer
5. Place the other tube in -80 °C freezer

Plating a Liquid Culture onto Agar

• Agar plate
• Liquid bacterial culture
• 100% ethanol
• Plating rod

1. Suspend culture in ~100 μL of LB media to produce concentrated liquid culture
2. Pipette liquid culture onto antibiotic agar plate
3. Dip plating rod into ethanol and burn any excess fluid. Cool rod on agar, making sure to avoid bacteria
4. Use plating rod to evenly spread the liquid culture throughout the plate.
5. Sterilize plating rod in between each plate
6. Store plates in 37 °C incubator until adequate bacterial growth is observed.

Streaking Plates

• Agar plate
• Incubation loop

1. Touch desired colony with a sterilized incubation loop until adequate sample is acquired
2. Gently streak onto surface of agar plate in a zig-zag motion until 1/4 of the plate has been streaked
3. Sterilize streaking loop with fire until glowing hot
4. Repeat streaking motion on other 1/4 of plate
5. Sterilize and streak until entire plate is covered in a circular direction. Do not let the last two streaking patterns come into contact.
6. Place in 37°C incubator until adequate bacterial growth is observed.
7. When picking colonies from streaked plate, take advantage of the gradient in colony density and use only isolated colonies

Bacterial Smear

• Glass microscope slides

1. Draw a one inch circle on the back of a glass microscope slide
2. Pipet 50 μL of distilled water in the circle and use a sterile inocculation loop to transfer a colony of bacteria into this water
3. Fix bacteria on to the slide by running it through a flame a few times
4. Allow the smear to dry before performing any stains

Gram Stain

• Crystal violet
• Lugol's iodine
• Safrinin

1. Flood bacterial smear with Crystal Violet for one minute
2. Rinse with distilled water
3. Flood with Lugol's Iodine for one minute
4. Rinse with distilled water
5. Decolourize smear using alternating applications of 95% ethanol for ten second and distilled water for ten seconds until the water running off of the slide runs clear
6. Counter-stain with Safrinin for one minute
7. Rinse one last time with distilled water and pat dry with Kim-wipes

Gram-negative cells will stain pink or red, and Gram-positive cells will stain purple or blue.

Spore Stain

• Malachite green
• Safrinin

1. Collect the spores for the smear from the suspended spores by centrifuging them for two minutes at 14,000 rpm, and use an inocculation loop to collect the spores
2. Place slides onto a hot plate at 150 °C
3. Place a piece of paper towel over the smear to allow for complete flooding and steaming with Malachite Green
4. Continuously flood the paper towl to prevent drying for five minutes
5. Rinse with distilled water
6. Counter-stain with Safrinin for one minute
7. Rinse one last time with distilled water and pat dry with Kim-wipes

Cells will stain pink and spores will stain green.