Protocol

Competent Cell Check

The purpose of the competent cell check is to ensure that our E.coli bacterium cells are functioning properly before they are altered in our experiment.

1. Spin down the DNA tubes from the Competent Cell Test Kit/Transformation Efficiency Kit to collect all of the DNA into the bottom of each tube prior to use. A quick spin of 20-30 seconds at 8,000-10,000 rpm will be sufficient.
1. Note: There should be 50 µL of DNA in each tube sent in the Kit.
2. Thaw competent cells on ice. Label one 2.0ml microcentrifuge tube for each concentration and then pre-chill by placing the tubes on ice.
3. Pipet 1 µL of DNA into each microcentrifuge tube. For each concentration, use a separate tube.
4. Pipet 50 µL of competent cells into each tube. Flick the tube gently with your finger to mix.
5. Incubate on ice for 30 minutes.
6. Pre-heat waterbath now to 42°C.
7. Heat-shock the cells by placing them into the waterbath for one minute. Be careful to keep the lids of the tubes above the water level, and keep the ice close by.
8. Immediately transfer the tubes back to ice, and incubate on ice for 5 minutes. This helps the cells recover.
9. Add 200 µL of SOC media per tube, and incubate at 37°C for 2 hours. Prepare the agar plates during this time: label them, and add sterile glass beads if using beads to spread the mixture.
10. Pipet 20 µL from each tube onto the appropriate plate, and spread the mixture evenly across the plate. Do triplicates (3 each) of each tube if possible, so you can calculate an average colony yield.
11. Incubate at 37°C overnight or approximately 16 hours. Position the plates so the agar side is facing up, and the lid is facing down.
12. Count the number of colonies on a light field or a dark background, such as a lab bench. Use the following equation to calculate your competent cell efficiency. If you've done triplicates of each sample, use the average cell colony count in the calculation. Make sure to measure the area of each colony to see how effective our cells are.

Gibson Assembly

The purpose of the Gibson assembly was to synthesize DNA fragments of the E.Coli bacterium with three different Andersen promoters, labelled weak, moderate, and strong. Each of these promoter samples were tagged with OSMy on the N-terminus end.

1. Design primers to amplify fragments (and/or vector) with appropriate overlaps
2. PCR amplify fragments using a high-fidelity DNA polymerase.
3. Prepare linearized vector by PCR amplification using a high-fidelity DNA polymerase or by restriction digestion.
4. Confirm and determine concentration of fragments and linearized vector using agarose gel electrophoresis, a NanoDrop™ instrument or other method.
5. Add fragments and linearized vector to Gibson Assembly Master Mix and incubate at 50°C for 15 minutes to 1 hour, depending on number of fragments being assembled.
6. Transform into NEB 5-alpha Competent E. coli (provided) or use directly in other applications.

PCR Protocol

The purpose of the PCR was to prepare the E.Coli bacterium DNA fragments for the Gibson assembly procedure.

1. Combine 10 μl of dnTP, 5 μl of PCR buffer, 2 μl of DNA polymerase, 10 μl of DNA template, 5 μl of forward primer, and 5 μl of reverse primer into one tube.
2. Mix tube by tapping it on the table.
3. Put it into the thermocycler and run it for 30 cycles.

Cellular Transformation

The purpose of the cellular transformation was to transform samples of the E.Coli bacterium so that it incorporated the plasmid we constructed in the Gibson Assembly procedure.

1. Thaw New England Biolab (NEB) chemically competent cells on ice.
2. Transfer 50 μl of competent cells to a 1.5 ml microcentrifuge tube.
3. Add 2 μl of the assembly made during the Gibson assembly protocol to the NEB competent cells and mix the microcentrifuge tube by flicking the tube.
4. Place mixture on ice for 30 minutes.
5. Heat shock at 42 °C for 30 seconds.
6. Transfer tubes onto ice for 2 minutes.
7. Add 950 μl of room temperature SOC media to tubes.
8. Place tubes at 37 °C for 60 minutes.
9. Shake them them at 250 rpm.
10. Spread 100 μl of cells onto the agar plates.
11. Incubate overnight at 37 °C.

E. Coli PET Plastic Degradation Protocol

The purpose of the E.Coli PET Plastic Degradation procedure was to provide an accurate, reliable method by which we could track the changes in the mass of PET plastic film. Ideally, the mass of the PET plastic film would decrease for every sample tested, with the strongest promoter producing the greatest degree of degradation.

1. Weigh out the same mass of PET plastic to put into each e. Coli colony.
1. We had one control and one colony for each of our promoter strengths (0.33, 0.58, and 1).
2. The PET plastic film had to be weigh the same amount for each colony so that we would be able to accurately tell which promoter works the best.
3. Had 20 colonies in total (5 for each promoter and control)
2. Wait 5-6 days to give the e. Coli cells time to begin degrading the PET plastic film.
3. Weigh out the PET plastic film again to check for progress on the PET plastic film degradation.
4. After that put each PET plastic film back into each e. Coli colony tube and wait for another 5-6 before checking up on them again.
1. That was our second time checking up on the e. Coli, meaning that they have been degrading the PET plastic for about a week now.
2. However due to our time limit, we could only really give the e. Coli two weeks to degrade the plastic as our poster people needed our results then.
5. When the experiment was over, each tube containing an e. Coli colony had to be thrown out and the equipment washed with HCl (hydroclauric acid) and then rinsed with d. H2O.