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From 2012.igem.org

LB Agar

We used a premade Becton, Dickinson and Comapany mix to make our LB agar.

Adding Antibiotics to LB

How to make LB liquid plus antibiotics:

• Ampicillin – The frozen stock solutions of ampicillin are at 50mg/ml and 100mg/ml in H2O, and are marked with a red sticker. The final concentration for LB liquid culture is 50ul/ml. To obtain this in 100ml (the amount in each LB bottle), add 100ul stock solution.
• Kanamycin – The frozen stock solutions of kanamycin are at 50mg/ml in H2O, and are marked with green. The final concentration for LB liquid culture for growing plasmids is 50ug/ml, and for cosmids is 20ug/ml. To obtain 50ng/ml in 100ml of LB, add 100ul stock solution, and to obtain 20ug/ml, add 40ul stock solution.
• Tetracycline – The frozen stock solutions of tetracycline are at 15mg/ml in methanol and are marked with black. The final concentration for LB liquid culture is 15ug/ml. To obtain this in 100ml of LB, add 100ul stock solution.
• Chloramphenicol – The frozen stock solutions of chloramphenicol are at 25mg/ml in 100% ethanol and are marked with purple. The final concentration for LB liquid culture is 25mg/ml. To obtain this in 100ml of LB, add 100ul stock solution.

How to make LB plates plus antibiotics:

• Follow the recipe card in box for making LB plates, being sure to add the agar. After autoclaving, and when the agar has cooled enough that it’s not too hot to touch (about 1 to 1.5hrs), add antibiotics as follows:
• Ampicillin – add 1ml ampicillin (at 100mg/ml) per liter of agar to obtain a final concentration of 100ug/ml. Mark the plate with a single red line on the side.
• Kanamycin – add 1ml kanamycin stock (at 50mg/ml) per liter of agar to obtain a final concentration of 50ug/ml. Mark the plates with a single green line on the side.
• Tetracycline – add 1ml tetracycline stock (at 15mg/ml) per liter of agar to obtain a final concentration of 15ug/ml. Mark the plates with a single black line on the side.
• Chloramphenicol – add 1ml chloramphenicol stock (at 25mg/ml) per liter of agar to obtain a final concentration of 100ug/ml. Mark the plates with a single purple line on the side.

SOB Media Recipe

• 2% w/v bacto-tryptone (20 g)
• 0.5% w/v Yeast extract (5 g)
• 8.56mM NaCl (0.5 g) or 10mM NaCl (0.584 g)
• 2.5mM KCl (0.186 g)
• ddH2O to 1000 mL[4]

For maximum effectiveness, SOB media should have its pH adjusted to 7.0 by adding concentrated sodium hydroxide. Autoclave media to ensure sterility

SOC Media Recipe

In addition to the contents of SOB media

• 10mM MgCl2 (0.952 g) or 20mM MgSO4 (2.408 g)[2]
• 20mM glucose (3.603 g)
• Alternatively, SOC can be made by adding small amounts of concentrated magnesium chloride and glucose solutions to pre-prepared SOB.

For maximum effectiveness, SOC media should have its pH adjusted to 7.0 by adding concentrated sodium hydroxide. Autoclave media to ensure sterility

Creating Chemically Competent Cells

The protocol for creating chemically competent cells is provided by the parts registry and can be found here.

Transforming Chemically Competent Cells

The protocol for transforming chemically competent cells is provided by Open Wet Ware and can be found here.

Updated Protocol for Transforming Chemically Competent Cells (as of 6/20)

Due to our low yield from our previous transformations and minipreps, we are now using the following protocol.

1. Thaw cells on ice ( the cells must be kept on ice as much as possible to work properly)
2. Gently mix approximately 25μL of cells into chilled polypropylene tube (if ligation then 50μL)
3. Add 1.25μL or less plasmid DNA
4. Swirl and flick with finger
5. Heat shock at 42°C for approximately 45 seconds (no shaking)
6. Put tubes back on ice for 2-5 minutes
7. Add 900μL of SOC
8. Gently shake and incubate at 200 rpm for 30-60 minutes at 37°C
9. Spread on LB plates
10. Place in 37°C for 14 hours

Tarun's Transformation Protocol (used 6/21)

1. Warm cells on ice and by holding
2. Shake cells for several seconds
3. Place 1μL of DNA on the side of the cell tube
4. Obtain 25μL of cells
5. Place cells on the side of the tube where the DNA is and mix by pipettor
When getting cells out of the container, pipette up and down to mix cells before adding to the tube
6. Flick the tube with a finger to mic and place the tube back on ice
7. Let the tube sit on ice for 20-30 minutes
8. Heat shock the cells at 42°C for 40 seconds
9. Put the cells on ice for 2 minutes
10. Shake the cells at 37°C for 8 hours

Miniprep Procedures

For this procedure, we used the QIAprep Spin Miniprep Kit by Qiagen.

Original Miniprep Procedure

1. Add 1.5mL of overnight culture to a micorcentrifuge tube
2. Spin for 3 minutes at 800rpm
3. Decant the supernatant so that the DNA pellet is all that remains
4. Repeat steps 1-3 three times
5. Add 250μL of buffer P2 and invert several times to mix (do not allow the solution to sit for more than 5 minutes)
6. Add 300μL of buffer N3 and immediately invert to mix
7. Centrifuge at 13,000rpm for 10 minutes
8. Apply the supernatant to the QIAprep spin column by decanting or pipetting
9. Centrifuge at 13,000rpm for 1 minute and discard flow through
10. Wash the QIAprep spin column by adding 500μL buffer PB
11. Centrifuge at 13,000rpm for 1 minute and discard flow through
12. Wash the QIAprep spin column by adding 750μL Buffer PE
13. Centrifuge at 13,000rpm for 1 minute and discard flow through
14. Centrifuge at 13,000rpm for 1 minute
15. Place the QIAprep column in a clean 1.5mL microcentrifuge tube
16. Elute the DNA by adding 50μL of Buffer EB to the center of the QIAprep spin column
17. Let stand for 1 minute
18. Quantify DNA concentration with nanodrop

Alterations to Miniprep Procedure (as of June 19)

1. The P2 and N3 steps (steps 5 and 6) were done twice
2. The final elution (step 16) was completed using 50°C nanopure water

PCR protocol

We used a PCR mastermix provided by 5 Prime . The components of the mastermix are as follows:

• Taq DNA Polymerase (62.5 U/ml)
• 2.5x Taq reaction buffer (with 125 mM KCl, 75 mM Tris-HCl pH 8.3, 4 mM Mg2+, 0.5% Igepal®-CA630)
• 500 µM each of dNTP
• stabilizers

The full manual for the mastermix can be found here . The procedures we used are described below.

1. Add 1ul of 100-200nM of the foward and reverse primer to a .2ml tube
2. Add 10-200ng of the template DNA to the .2ml tube
3. Adjust the final volume to 15μL (for a 25μL reaction) or 30μL (for a 50μL reaction) with nanopure water
4. Add 10μL (for a 25μL) reaction or 20μL (for a 50μL reaction) of mastermix to a PCR tube
5. Add the template DNA/primer mix to the PCR tube, close the tube, and mix well. Avoid foaming. If necessary, centrifuge briefly and place the tube on ice.
6. Cycle:
1. 95°C for 1-5minutes (usually 4min)
2. 95°C for 1min
3. 55°C for 1min. Cycle 30 times
4. 72°C for 1.5 to 2min (usually 2min)
5. 72°C for 10min
6. 4°C hold

A video showing the theory behind PCR can he found here

3A Assembly

The protocol for 3A assembly is provided by Open Wet Ware and can be found here

Gibson Assembly

Prepare a master mix, as detailed below.

• 320 μL 5X Isothermal Master Mix (recipe below)
• 0.64 μL 10 U/μL T5 exonuclease
• 20 μL 2 U/μL Phusion DNA Pol
• 0.16 μL 40 000 U/μL Taq DNA Ligase
• 860 μL ddH2O
• 1.2 ml Total

Prather Recipe 5x Isothermal Reaction Mix:

• 3 ml 1 M Tris-Hcl (pH 7.5)
• 300 μL 1 M MgCl2
• 60 μL 100 mM dGTP
• 60 μL 100 mM dATP
• 60 μL 100 mM dTTP
• 60 μL 100 mM dCTP
• 300 μL 1 M DTT
• 1.5 g PEG-8000
• 300 μL 100 mM NAD
• balance ddH2O
• 6 ml Total

Store the master mix in 15 ul aliquots at -20 °C. Then:

1. PCR up your fragments of choice and gel purify
2. Thaw a 15 μl assembly mixture aliquot and keep on ice until ready to be used.
3. Add 5 μl of DNA to be assembled to the master mixture. The DNA should be in equimolar amounts. Use 10-100 ng of each ~6 kb DNA fragment. For larger DNA segments, increasingly proportionate amounts of DNA should be added (e.g. 250 ng of each 150 kb DNA segment).
4. Incubate at 50 °C for 15 to 60 min (60 min is optimal).
5. Transform as usual

The protocol for Gibson assembly is provided by Open Wet Ware The original paper for the Gibson assembly protocol can be found here

Silica Creation

1. Obtain 14.4g of nanopure water
2. Obtain 50µL of .04M HCL and combine it with the water. Put this solution in an ice bath
3. Obtain 7.6g of tetramethyl orthosilicate (TMOS) in a separate container
4. Combine the TMOS and HCL/water
5. Immediately begin the vortex the solution. The solution will be cloudy while the reaction is taking place. Once the solution is clear again, the reaction is finished. This will take approximately 5 minutes
6. When the reaction is finished, let the solution set for 1-2 minutes
7. Use a rotary evaporator at 47ºC to remove the methane from the solution. This should take 2-3 minutes. The final solution volume should be approximately 13mL
8. If the silica is to be used immediately, put it on ice. It can be stored for up to a week if it is refrigerated.
9. Filter the silica before use

The procedures are provided by Rickus lab group.

Growth Rate Assay

1. Measure out 5mL of SOB media into 7 culture tubes
2. Add 5uL of NEB to 3 of the tubes and label
3. Add 5uL of DH5 to 3 of the tubes and label
4. Do not add anything to the last tube to keep as a blank
5. Place the culture tubes in an incubator for 1 hr
6. Record the OD after an hour
7. Check and record the OD for every hour until the OD starts to double, then check ever 30 minutes
8. Record all data

Static Biofilm Assay

1. Grow the E.coli to stationary phase
2. Dilute bacteria to 1:100 with SOB media
3. Place 200uL of the culture samples in each well making sure to do triplicates of each dilution and a control well
4. Set up duplicats of the plates for each time interval needed
5. Set up trays:
   • 1st tray - No water, used to collect waste
   • 2nd tray - Fill 1 to 2 inches of tap water, used to wash out bacteria
   • 3rd and 4th tray - Also filled with 1 to 2 inches of water, used to wash off the trays when the wells have been stained
6. After growing for the given time intervals, remove platonic cells by shaking the plate over the waste tray and submerging it and shaking it over the 2nd tray.
7. Add125uL of 0.1% crystal violet solution to each well and allow to sit for 10 minutes
8. Wash out the stain with trays 3 and 4
9. Add 200uL of 80% ethanol/20% acetone to each well and let sit at room temperature for 10-15 minutes
10. Mix the wells by pipetting
11. Transfer 125 uL to a container that can be used to measure OD
12. Measure the OD and record results