Team:British Columbia/Notebook

Welcome to our wiki notebook! We have organized our notebook according to sub-teams. Each page will provide you with a link to our actual notebook on OpenWetWare. To deliver the essentials here on the wiki (so you don't have to read through 6 months of experiments to get our message), we discuss the protocols, experimental outline, troubleshooting and optimization, and potential implications for iGEM.

Standard Operating Protocols (SOPS)

Colony PCR

Supplies Needed:

1. PCR tubes
2. BioBrick PCR primers (G1004, G1005) or (VF2, VR)
3. Taq Polymerase
4. 10x Reaction Buffer
5. 10mM dNTPs
6. sdH2O
7. Colonies to be PCR’ed
8. Agar plate for indexing

Steps:

1. Make master mix of primers and other PCR components EXCEPT Taq polymerase. Keep on ice.

PCR Master mix

Reagent	1x rxn volume (uL)		Master Mix
5x rxn buffer	5	xn
10mM dNTP	0.5	xn
sdH2O	9.15	xn
Phusion polymerase	0.1	xn
MgCl2	2	xn
DMSO - 5%	1.25	xn
gDNA	3	xn
Total	25

N = number of PCR tubes/samples Make sure to add about 2 extra samples to account for pipetting error and 1 extra sample for water control. Example: for 20 colonies, let N be: 20 (colonies) + 2 (extra)+ 1 (water) = 23
2. Add Taq polymerase.
3. Aliquot 25uL per PCR tube. Keep the PCR tubes on ice.
• *Following steps must be done near flame*
4. Touch toothpick/pipet tip/loop to colony, then index plate, then swirl around in PCR tube.
• *Turn flame off*
5. Run PCR:
• Turn machine on
• Load samples on the machine
• Select iGEM
• Select screen
• Make appropriate adjustments to the temperatures and times
1. 95°C for 15 mins
2. 94°C for 30 secs
3. 56°C for 30 secs
4. 68°C for 1 min per kb of expected product

*Round up for this step. i.e. For a 3.6kb construct, use 4 min elongation time.
5. Repeat 2-4 30 times.
6. 68°C for 20 mins
7. 10°C forever
8. Once finished, remove the PCR tubes from the machine.
6. Verify PCR products on agarose gel OR store the PCR tubes in the 4°C walk-in fridge until they can be verified on agarose gel.

Gel Verification (adapted from ‘Molecular Biology Laboratory 2010 iGEM Training Workshop’)

Supplies needed:

1. Agarose (1g/100mL for a 1% gel)
2. 1X TBE Buffer (dilute 10X TBE into diH2O)
• * Can use 0.5X TBE buffer instead to prolong supply of TBE buffer
3. Gel casting trays, combs, and box
4. Loading buffer
5. GelStar Stain
6. Samples to be loaded
7. DNA ladder: 1/20 100bp ladder or 1/20 1000bp ladder (depends on the size of desired product)
• Can add dye directly into the aliquot
8. Graduated cylinder
9. Erlenmeyer flask
10. Parafilm
11. Pipetman of appropriate size

Steps:

1. Pour a 0.8% agarose gel. Weigh out 0.8g of agarose and transfer to a 250mL Erlenmeyer flask.
2. Add 100mLs of 1X or 0.5X TBE buffer and swirl gently to disperse the agarose.
3. Plug flask lightly with a scrunched paper towel, then start by microwaving on high power for 30 seconds. Continue at your own discretion - make sure agarose is completely dissolved.
• *The dissolving step is a fine line between boiling your sample enough to dissolve your material, but not boiling it too much so that liquid starts to evaporate.
4. Allow the solution to cool to 60C by incubating in a 60C waterbath for about 10 minutes.
5. Add GelStar Stain (1/10000, e.g. 5uL/50mL) and swirl to mix.
• *In this case, add 10uL.
6. Remove casting tray and use the thicker tape to tape the sides of the casting tray.
• Make sure it is sealed tightly so no liquid will escape (especially near the base)
7. Pour into gel casting tray and insert comb.
• *Make sure liquid is not too hot when it is being poured - can melt the tape and leak
• *Gel will take approximately 20 minutes to set.
• *Make sure there are no air bubbles.
8. After gel has solidified, remove combs. Fill box with 1X or 0.5X TBE (if only taking a picture of the gel, TBE can be reused 2 or 3 times, but use new TBE if performing gel extraction).
• *Fill box with 1X if you used 1X to dissolve agarose; fill box with 0.5X if you used 0.5X to dissolve agarose
9. Load samples and ladder into wells.
• *Quick and dirty method of loading samples:
• Pipet 2uL of loading dye onto the parafilm N times with enough spacing inbetween, where N is the number of samples
• Pipet 10uL of sample, touch loading dye on parafilm, pipet up and down without pushing to the second stop on the PipetMan to allow dye to mix, load sample into well.
• Repeat until finished loading samples
10. Run gel. The following is based on the machine in the Lagally Lab.
• Machine conditions: 110V, 45min (can be varied depending on number of samples)
• Remember that DNA is negative, so that bottom electrode should be positive, and top should be negative
• Remember to set the middle button to be constant at Volts and not Amps
• If machine indicates error, there may be too much TBE Buffer in the box. Remove some using a 10mL or 25mL pipet.
• Double check that the machine is running by making sure bubbles are rising (at the top of the box)
11. Remove gel and take to transilluminator/imager on the 3rd floor of MSL, right across from the elevator.
• Wipe down transilluminator surface with water
• Place gel on surface and follow instructions on the door of the transilluminator/imager
• Save file under iGEM in the appropriate folder
• Print picture. *Don’t label the bands directly on the picture
12. Cross-reference sample bands with ladder bands to determine band size.
13. Rinse gel box a few times with water, making sure there are no gel bits. Dry with paper towel and return to original place.

BioBrick Restriction Digests

Supplies Needed:

• Restriction digest supermix (5*n μl of Buffer 2, 0.5*n μl of BSA, 37*n μl of ddH2O where n = # of 50uL reactions) stored at -20°C
• Enzymes
• DNA

Steps:

1. Grab an ice bucket and get some ice.
2. Retreive the appropriate digestion enzyme and digest supermix from -20 fridge and place on ice bucket.
3. Thaw restriction digest supermix (42.5μl aliquots).
4. Add 0.3-0.5μg DNA (1-5uL)
• *Can add up to 10uL of DNA if [DNA] is very low or if stuff doesn’t work
5. Add 1uL of each restriction enzyme.
6. (a). Incubate at 37°C for 2 hours. OR
7. (b). Digest overnight by adding 0.5uL of each restriction enzyme.
8. Return the enzyme and and supermix into the -20 fridge.
9. Discard the ice in the sink.

BioBrick Ligations

Supplies Needed:

• T4 DNA Ligase
• Ligase buffer (1uL aliquots)
• diH2O
• Insert and vector DNA (of approximately known concentration)

Note: Label.

Steps:

1. 1. Grab an ice bucket and get some ice.
2. Retrieve ligase and ligation buffer from -20 fridge and place on ice bucket.
3. Calculate insert/vector amounts. Various ratios have been recommended, most commonly 3:1 or 6:1 (molar ratio). Add the calculated insert volume amounts. For simplicity, you can use 1 uL of vector volume as a basis of calculations and then scale up or down. Formula: 1 insert mass (ng) = ratio x (insert length/vector length) x vector mass (ng) Formula 2: volume of insert (uL)= (total plasmid length/insert length)* insert mass /plasmid concentration Example: For a 150bp insert and a 2200bp vector, at a ratio of 6:1: insert mass = 9/22 * vector mass. If you add 1 uL of vector at 30 ng/uL, you would need 9/22*30 of insert, 12.27 ng. If the plasmid DNA concentration were 60 ng/uL and the total plasmid length were 2000, about 3 uL of DNA is needed (2000/150 * 12.27 / 60).
4. Add 1uL ligase buffer (vortex, and make sure it still smells like wet dog). During the process of using the ligase buffer, minimize freeze-thaws.
5. Add sdH2O to 9.5uL and vortex.
6. Add 0.5uL T4 ligase.
7. Incubate at 16°C for at least 1 hour (leave overnight after transforming in case it needs to go for a bit longer).
8. Throw away ice.
Note: Vortexing causes beads of liquid to stick on the walls of the microcentrifuge tube. Spinning in a centrifuge for a few seconds (even less) will cause the liquid to come back to the bottom. A mixed product is the result.

Making Plates

Supplies Needed:

• Petri dishes
• diH2O
• Autoclavable container (500mL glass bottle)
• Antibiotics (Stock solutions: Amp - 100mg/mL in 50% ethanol, Kan - 50mg/mL in H2O, Chl - 35mg/mL in 100%? ethanol, Tet - 15 mg/ml in 50% ethanol - all stored at -20°C)
• Pipet gun + disposable 25mL pipet
• Tinfoil (if using Tet)
• LB Agar Powder
• Colour scheme: Amp - red, Kan - blue, Chl - black, Tet - green

Steps:

1. Calculate amount of media needed (~20mL/plate).
2. Pour 400mL of diH2O into autoclavable container, make sure you remove any autoclave tape on the bottle.
3. Add LB Agar powder (check amount on stock container).
4. Mix as well as you can by swirling and gentle shaking.
5. Autoclave.
• *Mark bottle with autoclave tape, make sure the lid is loose*
• *Follow directions on autoclave*
• *Set to 30 min*
• *Before removing from autoclave make sure the pressure is at 0*
• *Use heat-resistant gloves to remove from Autoclave*
6. Cool until you can hold your wrist against the bottle for 5 seconds without pain.
7. Add antibiotics (final concentrations: A-100-150μg/mL, K-50μg/mL, C-25μg/mL, Tet-10-15μg/mL); if using recommended stock solution concentrations, add 1μL of antibiotic stock per 1mL of media.
8. Swirl gently to mix antibiotics
• *Steps 9-11 are to be done in the Biosafety cabinet in the biohazard room*
• *Ensure the fan and light is on*
• *Ensure no-one else is currently using the Biosafety cabinet*
• *Ethanol everything (unopened plates package, Pipet gun, Pipets, Your arms up to the elbow, bottle of LB Agar) before bringing them inside the Biosafety cabinet*
9. Quickly pipet into plates.
• *Avoid bubbles*
• *Save the plate sleeve, don’t remove from cabinet, must remain sterile*
• Leave to dry 20-30 min before using. Alternatively, leave to solidify 20-30 min before spread plating with appropriate volume of antibiotic.
• For storage, place back in plate sleeve, label according to the colour scheme. If using a light-sensitive antibiotic (Tet), wrap in foil as well.
• *Turn off the light and fan in the biohazard cabinet*
• Keep at 4°C.
• Notes: Ampicillin degrades quickly, so don't make plates more than 2-3 weeks in advance and never leave them at 37°C for more than 16 hours. Kanamycin can last 2-3 months. I don't know about chloramphenicol and tetracycline (not that stable, use like Amp), but they seem to be quite stable as well. Tetracycline is light sensitive.

Liquid Media

Supplies Needed:

• diH2O
• Autoclavable container (500mL Glass Bottles)
• Antibiotics (Stock solutions: Amp - 100mg/mL in 50% ethanol, Kan - 50mg/mL in H2O, Chl - 35mg/mL in 100%? ethanol, Tet - 15 mg/ml in 50% ethanol - all at -20°C)
• Tinfoil (if using Tet)
• LB broth powder
• Colour scheme: Amp - red, Kan - blue, Chl - black, Tet - green

Steps:

1. Pour appropriate amount of diH2O (400mL) into autoclavable container.
2. Add LB broth powder (Check amount on stock container).
3. Mix as well as you can by swirling and gentle shaking.
4. Autoclave.
• *Mark bottle with autoclave tape, make sure the lid is loose*
• *Follow directions on autoclave*
• *Set to 30 min, turn timer counterclockwise* (CHECK LATER)
• *Before removing from autoclave make sure the pressure is at 0*
• *Use heat-resistant gloves to remove from Autoclave*
5. Cool until you can hold your wrist against the bottle for 5 seconds without pain.
6. Add antibiotics (final concentrations: A-100-150μg/mL, K-50μg/mL, C-25μg/mL, Tet-10-15μg/mL); if using recommended stock solution concentrations, add 1μL of antibiotic stock per 1mL of media.
7. Swirl gently to mix antibiotics.
8. For storage, seal with parafilm and label according to the colour scheme. Keep at 4°C. If using a light-sensitive antibiotic, wrap in foil as well.

Notes: Ampicillin degrades quickly, so don't make plates more than 2-3 weeks in advance and never leave them at 37°C for more than 16 hours. Kanamycin can last 2-3 months. I don't know about chloramphenicol and tetracycline (not that stable, use like Amp), but they seem to be quite stable as well. Tetracycline is light sensitive.

Competent Cell Preparation (Small Scale)

Protocol generously donated by Jeanette (Beatty lab manager).

Supplies needed:

• DH5(alpha)
• LB broth--about 50mL per culture (each culture makes about 25-30 * 100 uL aliquots); about 5 mL for the initial overnight culture (inoculates multiple cultures)
• 0.1M CaCl2 (filter sterilized and chilled)--about 40-50 mL per culture
• Ice water bath
• 60% glycerol--about 1 mL per culture
• Spectrophotometer
• 100-250 mL flasks, centrifuge tubes (50 mL Falcon or equivalent), microcentrifuge tubes

Note: When working with cultures (that you will use later), always work aseptically. That is, work near a flame and be sure to turn off the flame at the end and follow aseptic procedures.

Steps:

1. Inoculate 5mL overnight culture (see overnight culture protocol) and grow at 30°C. You can use AMBL lab rotating incubator (ask Lijuan if you need help). Approximately 30°C (can be a little higher or lower) seems to be fine. Otherwise, shaking speed ~200 rpm.
Note:A possible source of DH5(alpha) are frozen aliquots of DH5(alpha) competent cells. You can use the whole aliquot.
2. Dilute 0.5mL overnight culture into 50mL LB and incubate at 30°C, shaking vigorously. Label with name, date, strain, flask # (when making more than 1 culture), etc. You can use Finlay Lab common incubator for this (the one that says to not change the settings). Shaking speed ~200 rpm.
Note: You can inoculate and use several 50 mL LB cultures at the same time. A 125 mL or 250 mL Erlenmeyer flask works well as a culturing container. Make sure to not rip or contaminate the aluminum foil cover. When you’re done using the flasks, rinse with 10% bleach, then with water and put into dirty basket (ask Tony Lam for help if needed).
3. Harvest at Abs600nm = 0.401. Higher than this seems to work well anyway. Pipet the cultures into 50 mL Falcon tubes (don’t overfill).
Using a spectrophotometer: Take 1 mL from the culture and 1 mL from a LB stock and put into cuvettes. Turn on the spectrophotometer (on Antonio’s bench; make sure to ask if he needs to use it and ask him if you need help). Wipe the cuvette with Kimwipe (to clean the part where light passes through). Press the OD600 button. Put the 1 mL LB cuvette into the slot and press the Blank button. Replace the 1 mL LB cuvette with 1 mL culture cuvette and press sample. Rinse the cuvettes twice with 70% ethanol and twice with sdH20. Turn off spectrophotometer. Note: You don’t have to work aseptically when using the spectrophotometer. However, as always, transferring cultures that you want to use later on means you should work aseptically.
4. Centrifuge at 1600g for 7 minutes (4°C). You can use Kronstad lab common centrifuge (3rd floor) for this.
Kronstad lab centrifuge: Set the rotor type (e.g. 5.3), RCM (e.g. 1600 g), temperature =4°C, and time. Then press start (if machine is off, turn on). In any case, follow the instructions on the centrifuge. Make sure the samples are balanced. You may leave the temperature at 4°C (the machine will keep it nice for you). If someone else is going to use it, return the temperature to about 21°C. Turn off the centrifuge when you won’t use it again. Sign your name on the notepad. Emergency contact details are near the centrifuge.
5. Pour out the supernatant aseptically (also flame mouth and cap before and after). Wash gently in ~20mL COLD filter sterilized 0.1M CaCl2 (i.e. put the tube on ice or put in cooler). Swirl the tube using your wrist until the pellet disappears.
Filter sterilization: Use a syringe to suck up some CaCl2 (from a beaker probably). Attach a 0.4 micron filter unit to the tip. Press the plunger. Whether any part above the filter is sterile is unimportant; anything that passes through the filter is sterile. Make sure the filtrate enters a sterile container (e.g. Falcon tube). You can re-use the filter: remove the filter, use the syringe to suck up more CaCl2 and repeat. The original packaging material can hold the filter while you do this. Make sure the first few drops of filtrate does not go into the container with your previously sterilized CaCl2. You can drop it back into the unsterilized CaCl2. Label finished product.
6. Spin down gently: 1100g, 5 minutes, 4°C. See 4 about using the centrifuge.
7. Pour out the supernatant aseptically (flame mouth and cap before and after too). Resuspend in 12.5mL cold 0.1M CaCl2. Wrist.
8. Keep on ice 40 minutes.
9. Spin down: 1100g, 5 minutes, 4°C. See 4 about using the centrifuge.
10. Pour out the supernatant asepticall (flame mouth and cap before and after too). Resuspend in 2-2.5mL cold 0.1M CaCl2. Wrist.
11. Store over night at 4°C
12. Add glycerol to 15% and divide into 100uL aliquots, store at -80°C in Lagally Lab freezer; label with name, date, strain, flask where it came from, etc. Adding about 1 mL 60% glycerol to 2.5 mL of the final resuspended cells yields a ~15% glycerol solution.
13. Throw away your used tubes, pipets, filters, syringes, etc.

Three-Antibiotic (3A) Assembly (modified from OWW protocol)

Steps:

1. Perform a restriction digest (see Common Protocols) on all the parts you wish to assemble: Construction plasmid (EcoRI, PstI), Prefix part (EcoRI, SpeI), Suffix Part (XbaI, PstI).
Note: page 86 of the NEB catalog explains the 3A method well.
2. Purify restriction digests (Ethanol precipitation for <200bp, otherwise Qiagen PCR Purification Kit). This step is optional. You can just heat-inactivate your digests by raising the temperature of the reaction tube to 80ºC for 20 min. and then cooling it down again. You can use the PCR machine for this as long as nobody else is using it—Eric Ma. A water bath works too; make sure to turn it off at the end.
3. Ligate all parts together (Ligation protocol).
4. Transform ligation reaction into DH5α competent cells (Transformation protocol).
5. Perform cPCR on colonies to confirm insert length/presence (Colony screening).
6. Streak out correct colony (optional), inoculate overnight culture, miniprep and store (glycerol stock).

Transformation (adapted from OWW )

Supplies Needed:

• Competent cells in 100μL aliquots
• Water bath/heat block at 42°C
• Ice
• LB Broth
• LB agar plates (with appropriate selection)

Steps:

1. Remove competent cells (100uL aliquots) from -80°C and thaw on ice.
*Note, aliquots can be thawed GENTLY by hand if time is an issue*
2. Add 1µL ligation mix to thawed cells and incubate for 30 minutes on ice.
3. Heat shock in water bath/heat block for 60 seconds and put back on ice for two minutes.
• *Remember to turn off the water bath*
• *The following steps must be done near a flame*
• *Don’t forget a growth control, especially if you aren’t screening using antibiotics*
4. Add 400uL LB Broth and incubate at 37C for 2 hours.
*Note, 1 hour is also sufficient*
5. Spread plate entire 500µL.
*Turn off the flame*
6. Incubate overnight (<18hrs) at 37°C.

Rafael's Alkaline Lysis (for a miniprep)

1. 3-5mL of an overnight broth culture is spun down to a pellet in a microfuge tube (1mL of culture has also yielded DNA, if you don't have much to spare).
2. Decant the broth and pipet out the remaining small amount of broth; minimize broth as much as possible!
3. Add 200uL of buffer P1 (50mM Tris-Cl pH 8.0, 10mM EDTA, 100ug/mL RNAse A), resuspend pellet in this buffer.
4. Add 200uL of buffer P2 (200mM NaOH, 1%SDS), invert the tube 6-10 times and wait for 5 minutes at room temperature.
5. Add 300uL of 3M sodium/potassium acetate (pH 5.5), immediately invert the tube 6-10 times, and centrifuge for 10 minutes at 13 000 RPM.
6. Pour or pipette the supernatant into a microcentrifuge tube with 600uL isopropanol, make sure to avoid the white precipitate as much as possible!
7. Vortex the tube for 20 seconds and centrifuge for one minute at 13 000 RPM
8. Discard the supernatant and add 750uL of 70% ethanol to the tube, vortex briefly, and centrifuge for another 1 minute at 13 000 RPM
9. Discard the supernatant again and centrifuge for 10 seconds at 13 000 RPM to collect the remaining ethanol.
10. If you can see a pellet, pipette out the remaining ethanol, if not, heat the tube over a hot water bath (50C or more) for as long as it takes to evaporate the ethanol. A P10 tip works best because it won't freak out the ethanol as much when it comes in contact with the tip.
11. Resuspend the plasmid DNA in the desired amount of water or buffer of choice.

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