Our protocols are based on standard lab protocols used in the lab of our advisers unless otherwise stated. These protocols were subsequently modified if necessary based on our experience. A knowledge of basic lab techniques such as pippeting, plating, etc. are assumed and not covered as there are pretty comprehensive tutorials on the web for those.
A list of materials mentioned in the protocols which we used is also included at the end of the page.

Materials Used

1. Cell strains. Cloning strains: DH5-alpha Expression cell strains: BL21(DE3), BL21(DE3)-pLys
2. Cell culturing: Luria Broth media, M9 media , Ampicillin, Chloramphenicol.
3. Dyes: Malachite Green ester [from MBIC], 3,5-difluoro-4-hydroxybenzylidene imidazolinone (DFHBI)

Kit Protocols

1. Z-Competent E. Coli Transformation Kit for making competent cells for easy transformation. Modified to incubate cells on ice for 1 hour during transformation
2. Phusion High Fidelity PCR Kit for amplifying our inserts and cassettes.
3. TAQ DNA Polymerase for verifying our cloned constructs
4. Qiagen Mini-prep Kit for isolating our plasmids from the cells, to be transformed into expression strains.
5. Zymo Clean and Concentrator kit for cleaning up our digestion products before ligation.

Cloning Protocol

1. If necessary, first prepare the insert using PCR amplification. This was done to add the necessary restriction sites for cloning. The PCR protocol follows the kit protocol outlined above, except with our specific range of melting and annealing temperatures calculated using NEB's Tm calculator
2. Start digestion of vector and insert DNA using desired restriction enzyme manufacturer protocol. [NEB]
3. After 2 hours, add Phosphatase [CIP] according to manufacturer protocol to insert to prevent self-ligation
4. Purify and clean DNA using kit protocol. [Zymo Research]
5. Measure vector/insert concentration. [Nanodrop]
6. Divide the concentration by the length of the sequence and calculate ligation ratios of 1 vector to 3 insert. Mix the ratios according to the calculations, including T4 buffer and ligase.
7. Leave ligation products at room temperature for 1 hour.
8. Transform ligation products into competent cells using appropriate protocol. Incubate on ice for 1 hour if using Zymo competent cells
9. Plate cells and incubate at 37 degrees overnight. Check for colonies the next morning. If there are colonies present, inoculate 1-3 colonies into (selective) L.B. in the evening and culture overnight.

Gel Protocol

1. For 7x7 cm, 0.5cm thick gel, with 1.75% agarose Concentration.
2. Add 0.35 grams of agarose to 20ml of 1x buffer (TBE) Note: Place mark on container to keep track of liquid level using marker, so water can be added to bring the liquid back to original level
3. Place gel solution in microwave. Use low/medium, set timer for 5 minutes. Stop the oven every 30 seconds and swirl gently to suspend undissolved agarose.
4. >Once dissolved, set aside to cool (~60 degrees). Once solution is warm to touch, add 0.5ug/ml ethidium bromide. [1µl of 10mg/ml]
5. Pour into casting tray, remember to add in comb at the cathode side (black). Gel will solidify in ~10 mins
6. Divide the concentration by the length of the sequence and calculate ligation ratios of 1 vector to 3 insert. Mix the ratios according to the calculations, including T4 buffer and ligase.
7. Remove comb for solidified gel, remove casting gates and submerge gel beneath 2 to 6mm of 1x buffer.
8. Mix loading dye with PCR/digested products, load mixture into wells, together with 10µl of ladder
9. Run gel at 75V for ~1 hour. Adjust voltage accordingly if require faster or more distinct gels.

Dosage Curve

1. The night before the experiment, start an overnight culture by seeding 1:100 of expression cell stock to fresh selection media [Typically 30µl stock to 3ml LB+Amp]. Place tube in a shaking water bath at 37 degrees celsius overnight.
2. Culture a fresh batch of cells using 1:100 of overnight culture prepared to selection media. Culture in a shaking 37 degrees celsius water bath for around 2 hours or until the solution is lightly turbid.
3. Induce cells by adding 1µl of 1M IPTG. Incubate cells in a shaking water bath at 37 degrees celsius for 2 hours.
4. Aliquot 1ml of induced cells into an 1.5ml Eppendorf tube and spin down the cells at 10,000rpm for 1min and discard the supernatant. Wash the cells with 1ml of PBS, repeat spinning down the cells at 10,000rpm and discard the supernatant. Resuspend the washed cells in 1mL M9 media.
5. To increase DFHBI fluorescence, add 1µL of 50nM Mg2+ to the tubes (from MgCl2 from PCR kit)
6. Aliquot (100µl/200µl) of cells into desired wells of a 96-well plate.
7. To each of the 96-wells, add 1µl of 1mM IPTG to maintain the production of FAP and Spinach. Add various doses of DFHBI to the wells intended for testing DFHBI dosage. To a separate set of wells, add the desired doses of MG.
8. Incubate for 45mins in 37 degrees celsius, as recommended by Paige et al. Plate read with plate reader (Tecan Safire II). First find cell density using OD600. Then use Excitation/Emission of 469/501 for Spinach and 635/660 for Malachite-green.

Time Lapse Protocol

1. For the temporal analysis, we will be characterizing different promoters. Culture fresh batches of cells using 1:100 of overnight culture of the different promoter constructs. Do include the wild type promoter to act a a form of comparison. Culture for around 2 hours or until the solution is lightly turbid.
2. Culture a fresh batch of cells using 1:100 of overnight culture prepared to selection media. Culture in a shaking 37 degrees celsius water bath for around 2 hours or until the solution is lightly turbid.
3. For each ml of cells required for the assay: Aliquot 1ml of cells into 1.5ml Eppendorf tubes and spin down the cells at 10,000rpm for 1min and discard the supernatant. Wash the cells with 1ml of PBS, repeat spinning down the cells at 10,000rpm and discard the supernatant. Resuspend the washed cells in 1mL M9+3µl 50nM Mg media.
4. Reserve part of the cells for testing the un-induced construct for leaky expression. Add 1µl of 1M IPTG per ml of remaining cells
5. Aliquot 100µl of induced and uninduced cells per well into a 96-well plate
6. Add the maximum dosage of DFHBI and MG as determined by the dosage curve into separate wells for each of the clones.
7. To each of the 96-wells, add 1µl of 1mM IPTG to maintain the production of FAP and Spinach. Add various doses of DFHBI to the wells intended for testing DFHBI dosage. To a separate set of wells, add the desired doses of MG.
8. As soon as the dyes are added, plate read with the plate reader (Tecan Safire II). First find cell density using OD600. Then use Excitation/Emission of 469/501 for Spinach and 635/660 for Malachite-green. Set the plate reader temperature to 37 degrees and to shake the plate for 10 seconds every 10mins. Repeat the OD and fluorescence measurements every 30minutes until the fluorescence output begins to decrease (~3 hours).