Experiments

Procedure

1. Pellet culture by centrifugation for 30 seconds and remove supernatant.
2. Use 200 µL of Monarch Buffer B1 to resuspend pellet.
3. Add 200 µL of Monarch Buffer B2, and invert 5 times. Incubate at room temperature for one minute.
4. Add 400 µL of Monarch Buffer B3, and invert the tube to neutralise.
5. Centrifuge for 5 minutes and transfer the supernatant to spin column. Centrifuge for one minute and remove the flow-through.
6. Re-insert the spin column to the collection tube. Add 100 µL of Monarch Buffer BZ and centrifuge for one minute.
7. Add 400 µL of Monarch Buffer WZ and centrifuge for one minute.
8. Transfer the column to a new microfuge tube.
9. Add 30 µL of Monarch Buffer EY. Wait for one minute and spin for one minute to elute the DNA.

Protocol: Plasmid Miniprep Protocol

High Efficiency Transformation Protocols

• Protocol (C2987H/C2987I)
• Protocol (C3010)

Cell Transformation Procedure

1. Thaw competent cells on ice. Gently mix and add 50 µL of cells into a new tube on ice.
2. Add 10 µL of DNA to the cell mixture. Gently flick the tube to mix the cells and DNA.
3. Place the tube on ice for 30 minutes without mixing.
4. Heat shock at 42 °C (time varies depending on competent cell type).
5. Place on ice for 5 minutes.
6. Add 950 µL of appropriate media into the mixture.
7. Incubate at 37 °C for 60 minutes at 250 rpm.
8. Warm plates at 37 °C.
9. Mix the cells by gently flicking the tube. Perform serial dilution in appropriate media.
10. Spread the dilution onto the selection plate and incubate overnight at 37 °C.

Procedure

1. Prepare solution to be tested.
2. Add 10 drops of Fo-1 to the solution.
3. Dip the test strip for 5 seconds.
4. Wait for 1 minute.
5. Match the color of the test strip with the results table.

Protocol: Supelco Formaldehyde Test Strips Protocol

Gotaq Hot Start Green 2× Mastermix was used according to manufacturer’s protocol.

PCR Setup:
For 25× cycles

Notes:

• Annealing temperature (Tm) is based on primers used.
• Extension time depends on length of expected amplicon.

Procedure

1. Set up gel casting tray.
2. Make a 1% (w/v) agarose gel in 50 mL 1×TAE buffer. Add 1 µL of dye per 10 mL of agarose gel. Pour into the gel casting tray and allow the gel to set.
3. When the gel is set, transfer the gel into a gel tank. Pour enough 1×TAE into the tank so the gel is submerged. Pull out the gel comb.
4. Mix the QuickLoad ladder and PCR products with 6× loading dye and load into the gel.
5. Electrophorese at 120 V for 45–60 minutes. Check the dye front at 45 minutes, continue to electrophorese if necessary.
6. When finished, visualise PCR products using the transilluminator.

Procedure

1. Prepare media and drug stocks:
   • Make 2× media (LB or YPD depending on organism).
   • Prepare 2× drug stock at the highest concentration for column 1.
2. Plate setup:
   • Add 100 µL 2× drug stock to column 1.
   • Add 50 µL 2× media to columns 2–10.
   • Serially transfer 50 µL from column 1 → column 10, mixing each time.
   • Column 11 = growth control (50 µL 2× media).
   • Column 12 = blank (100 µL media only).
3. Add inoculum:
   • Prepare 2× inoculum in 1× media.
   • Add 50 µL inoculum to wells in columns 1–11.
   • Add 50 µL media to column 12.
   • Final volume = 100 µL per well.
4. Incubation:
   • E. coli & B. subtilis: 37 °C overnight.
   • S. cerevisiae: 30 °C overnight.

Inoculation & Growth

1. Inoculate 25 mL SOB in a 150 mL flask with 300 µL of overnight culture (LB @ 37 °C).
2. Incubate at 18 °C (preferred) or 30 °C if necessary.
3. Monitor until OD₆₀₀ = 0.5–0.6 (approx. 4 h at 30 °C, 3 h at 37 °C).

Harvesting Cells

4. Place culture on ice for 10 minutes and transfer to 50 mL falcon tube on ice.
5. Centrifuge at 2500 × g for 10 minutes at 4 °C.
6. Discard supernatant.

Washing

7. Resuspend pellet in 10 mL ice-cold TB buffer.
8. Keep on ice for 10 minutes.
9. Centrifuge again at 2500 × g for 10 minutes at 4 °C.
10. Discard supernatant.

Final Resuspension

11. Gently resuspend pellet in 930 µL ice-cold TB buffer and 70 µL DMSO.

• This mix is for −80 °C storage.
• DMSO is unnecessary for immediate use.
• Ensure 7% DMSO is measured accurately.

Aliquoting

12. Prepare 50 µL aliquots into 20 eppendorf tubes.
13. Store aliquots at −80 °C.

Materials

• E. coli cultures: 250 mL induced culture and 250 mL uninduced culture
• 50 mL Falcon tubes ×10
• 1 mL Eppendorfs ×4
• 2 mL Eppendorfs ×24
• TE buffer (for protein extraction reserves) 4 mL
• PBS (for resting cell assay) ~26 mL
• Methanol (for treatment at different concentrations) ~200 µL
• Formaldehyde test strips – 25

Procedure

1. Culture Harvesting

1. Split each culture (induced and uninduced) into 50 mL aliquots in Falcon tubes (5 × 50 mL for each condition). Reserve any extra in suitable tubes.
2. Centrifuge to pellet the cells and discard all supernatant (10 min at highest rcf). Use a pipette if needed to remove residual liquid, being careful not to lose any pellet.

2. Protein Extraction Reserves

3. Take one pellet each – Induced “I” and Uninduced “−ve”.
4. Resuspend each pellet in 2 mL TE buffer.
5. Split into 2 × 1 mL aliquots (4 tubes).
6. Label clearly:
   • “E. coli T7X Induced pMoxF1 pMxa TE” (with date) Lid label: “pMDH I”
   • “E. coli T7X −ve pMoxF1 pMxa TE” (with date) Lid label: “pMDH −ve”
7. Store in −80 °C. Record location in lab book.

3. Prepare Resting Cells for Assay

7. Resuspend remaining 4 pellets each (“I” and “−ve”) with 12.6 mL PBS.
8. Transfer pellets sequentially from one Falcon tube to the next (within each condition) until all material is pooled. Keep “I” and “−ve” separate.

4. Methanol Treatment Setup

9. Set up 12 × 2 mL Eppendorfs for induced and 12 × 2 mL for uninduced samples.
10. Label: “I” (induced) or “−ve” (uninduced). For each, prepare triplicates labelled “0”, “0.1%”, “2%” methanol concentrations.
11. Also label time points: “t = 0”, “t <1 min”, “t = 10 min”, “t = 1 h”.
12. Aliquot 4.2 mL of the 12.6 mL into three Falcon tubes labelled “I” or “−ve” and “0”, “0.1%”, “2%”.

Protocol: Resting Cell Methanol Assay

Conduct Methanol Assay

11. Pre-label 24 test strips to match tubes. You will measure at 4 time points: t = 0, t <1 min, t = 10 min, t = 1 h.
12. Add methanol to cultures: 3.2 µL for 0.1% or 64 µL for 2%. Add simultaneously across samples. Take t = 0 samples before addition.
13. t = 0: Add 1 mL sample to labelled tubes. Add 2 drops Fo1, shake, insert strip. Photograph strips.
14. t <1 min: Immediately add methanol, then 1 mL sample to labelled tubes. Add 2 drops Fo1, shake, insert strip. Photograph strips.
15. t = 10 min: Take 1 mL sample, add 2 drops Fo1, shake, insert strip. Photograph strips.
16. t = 1 h: Repeat sampling and strip assay as above. Photograph strips.

Protocol Reference: ThermoFisher Dynabeads™ His-tag Isolation & Pull-down Protocol

We recommend preparing your sample containing the histidine-tagged protein in a total volume of 700 µL 1× Binding/Wash Buffer.

Procedure

1. Thoroughly resuspend the Dynabeads™ magnetic beads in the vial (vortex >30 s or tilt and rotate for 5 min).
2. Transfer 50 µL (2 mg) Dynabeads™ magnetic beads to a microcentrifuge tube. Place the tube on a magnet for 2 min. Aspirate and discard the supernatant. Add your sample (in 1× Binding/Wash Buffer) to beads. Mix well.
3. Incubate on a roller for 5 min at room temperature (or colder if protein is unstable). Incubation may be extended up to 10 min.
4. Place the tube on the magnet for 2 min, then discard the supernatant.
5. Wash the beads 4 times with 300 µL 1× Binding/Wash Buffer. For each wash: place the tube on the magnet for 2 min, discard the supernatant, and resuspend thoroughly.
6. If the protein is to be eluted:
   • Resuspend bead/protein complexes in 1× Pull-down Buffer (or compatible buffer) for downstream applications.
   • For protein pull-down, continue with the protocol on page 2 of the reference manual.
7. Add 100 µL His-Elution Buffer. Incubate on a roller for 5 min at room temperature (or colder if protein is unstable).
8. Place on the magnet for 2 min and transfer the supernatant containing the eluted histidine-tagged protein to a clean tube.

Protein samples were separated using (SDS-PAGE) following the procedures described in the protocol by the Coleman Lab. The specific sections used included Loading Samples, Running the Gel, Staining and Visualising (Coomassie Blue Staining), as well as the preparation and use of Coomassie Blue Staining Solution, High Destain Solution, and Low Destain Solution.
The gel was run under denaturing conditions, stained with Coomassie Brilliant Blue, and destained to visualise protein bands.

Protocol Reference: Coleman Lab: Protein Analysis by SDS-PAGE