Results

PCR and Agarose Gel Electrophoresis

To determine whether the E. coli colonies had successfully taken up the plasmids, colony PCR was performed using specific primers. PCR products were analysed by agarose gel electrophoresis.

14/8/2025 Colony PCR confirmation (AmilCP)

14/8/2025 Colony PCR confirmation of E. coli NEB 5-alpha transformants carrying AmilCP plasmid from 11/08/25.

Lanes: L, NEB 1 kb DNA ladder; 1–6, colony PCRs from colonies 1–6; +ve, positive-control PCR using AmilCP plasmid DNA.

Primers: AmilCP-F / AmilCP-R (expected amplicon: 372 bp).

Colony PCR was carried out on E. coli NEB 5-alpha transformants to confirm uptake of the AmilCP plasmid. Amplification with AmilCP specific primers produced bands at the expected size of 372 bp in several colonies, consistent with successful transformation. The positive control generated a clear band at the same size, while the DNA ladder was used for fragment size estimation.

Agarose Gel Electrophoresis of Colony PCR (mxa)

Colony PCR was performed using primers mxaA_F and mxaK_R to confirm uptake of the mxa plasmid. Five colonies were screened and their PCR products were loaded into wells 4–8. The mxa plasmid was included as a positive control in well 2, while BL21 cells without plasmid served as a negative control in well 3. The expected amplicon size was 590 bp.

28/08/25 Agarose gel electrophoresis of plasmid constructs

28/08/25 Agarose gel electrophoresis of plasmid constructs pMDHMoxFI, pMDHMxaAK, and pAmilCPMxaycusS.

NEB 1 kb DNA Ladder was loaded in the first lane. Negative controls (1, 5, and 9) show no amplification.

Lanes 2–4: NEB 10β pMDHMoxFI (three biological replicates).
Lanes 6–8: E. coli DH5α pMDHMxaAK (three biological replicates).
Lanes 10–12: NEB 10β pAmilCPMxaycusS (three biological replicates).

Agarose gel electrophoresis was performed to confirm the presence of the plasmid constructs pMDH MoxF1, pMDH MxaAK, and pAmilCP MxaycusS in transformed E. coli and NEB 10β cells. The NEB 1 kb DNA ladder was loaded in lane 1 as a molecular weight reference.
As expected, no amplification was observed in the negative controls (lanes 1, 5, and 9). Clear DNA bands were visible in the replicate lanes corresponding to each construct:
Lanes 2–4: NEB 10β pMDH MoxF1 (three biological replicates)
Lanes 6–8: E. coli DH5α pMDH MxaAK (three biological replicates)
Lanes 10–12: NEB 10β pAmilCP MxaycusS (three biological replicates)
Distinct bands of expected size were observed in the corresponding lanes, confirming successful plasmid presence and transformation for all three constructs.

Choosing a Reporter Protein

Testing amilCP and RFP

After testing both amilCP and RFP, our team chose to proceed with amilCP for the project, as it provided easier colorimetric visualisation compared to RFP.

Testing Commercial Formaldehyde Test Strips

Testing Supelco Test Strips

We tested Supelco formaldehyde test strips across a concentration range of 5 mg/L to 200 mg/L. The strips showed clear, progressive colour changes with increasing concentration, ranging from pale peach at low levels (5–10 mg/L) to deep purple at high levels (100–200 mg/L). At 200 mg/L, the concentrated sample saturated the strip, while dilution to 100 mg/L produced a colour consistent with the expected scale. These results confirm the strips provide a reliable, semi-quantitative indication of formaldehyde concentration.

Methanol Dehydrogenase Enzyme Design

Formaldehyde Experiment

The formaldehyde assay relies on the pH indicator bromothymol blue (BTB), which shifts colour according to pH. In the presence of hydrogen peroxide, formaldehyde is oxidised to formic acid, lowering the pH and triggering a visible colour change in the BTB solution. At neutral pH, the dye appears green/blue, while acidification shifts the colour toward yellow.

After addition of Formaldehyde t=0

T=0 (After addition of formaldehyde)
Assay was expected to produce a colour gradient within 5 minutes but no clear response was observed.
No visible gradient corresponding to increasing formaldehyde concentration was detected.
Higher BTB concentration (0.6% BTB) shifted the reagent to yellow before any addition of formaldehyde sample.

After addition of formaldehyde t=7 days

T=7 (Seven days after addition of formaldehyde)
0.04% BTB produced a visible gradient across formaldehyde concentrations in both hydrogen peroxide concentrations, indicating this lower BTB concentration is more suitable.
Lower Fe concentration(4 µM) gave slightly more distinct than the higher (8 µM) though both conditions showed some activity.
The gradient at 0.04% BTB + 4 µM Fe demonstrates that the assay can distinguish formaldehyde concentrations after 7 days

The experiment was repeated using FeSO₄ and FeCl₃ as iron sources, under the same hydrogen peroxide and bromothymol blue (BTB) concentrations as before.

After addition of formaldehyde t= 0

T=0 (After addition of formaldehyde)
No visible gradient was observed within 5 minutes across any condition, consistent with earlier findings.

After addition of formaldehyde t=7 days

T=7 (Seven days after addition of formaldehyde)
A clear gradient became visible, indicating delayed but successful detection.
No gradient was observed under 0.01 M H₂O₂ regardless of BTB concentration.

Conclusion
3.8 mM H₂O₂ consistently produced the best results across both Fe salts.
0.04% BTB remains optimal, aligning with previous findings.
4 µM for either Iron(II) sulfate or Iron(III) chloride. The 8 µM is not as consistent.

After addition of formaldehyde t=7 days

When viewing our results for the formaldehyde assay optimisation it is important to understand we are generally referring to stock concentrations used. Labels indicating 0.04 or 0.6 % BTB are not referring to final concentrations.
For details of the final concentrations for the working reagent please see the table in our Contributions section.

Resting Cell Assay

A resting cell assay was carried out to test for formaldehyde production by MDH-expressing E. coli. Supelco formaldehyde test strips were used to compare induced and uninduced samples at three time intervals (<1 minute, 10 minutes, and 1 hour).

Across all conditions and timepoints, no colour change was observed on the strips. Both induced and uninduced samples consistently matched the baseline reference, corresponding to 0 mg/L formaldehyde. These results confirm that no detectable formaldehyde was produced under the assay conditions.

Protein Gels

Protein samples from E. coli cultures expressing the MDH constructs were analysed using SDS-PAGE to assess protein expression and purification outcomes. Samples included both induced (Indc) and uninduced (Unindc) cultures, with 10 µL and 15 µL of purified and lysate samples loaded per well.

The MDH enzyme was expected to produce two visible subunit bands corresponding to the MoxF + Histag (68 kDa) and MoxI (10 kDa) proteins. However, no distinct bands were observed at these expected molecular weights in any of the purified or lysate samples. The absence of visible protein bands suggests that either expression was unsuccessful or protein yield was below the detection limit of Coomassie staining.

MDH protein gel run 18 Sept 2025

Protein samples from E. coli expressing the MDH constructs were analysed by SDS-PAGE to assess protein expression and purification. Samples (10 µL and 15 µL) included induced (Indc) and uninduced (-ve) cultures, as well as lysate and pellet fractions. The MDH complex was expected to yield two subunit bands corresponding to MoxF + Histag (~68 kDa) and MoxI (~10 kDa). However, a single unexpected band was observed at approximately 80 kDa (indicated by red arrows).

MDH protein gel run 29 Sept 2025

Protein expression was induced with IPTG in the presence of PQQ and analysed via SDS-PAGE. Purified His-tagged samples and corresponding lysate and pellet fractions were loaded. The MxaF (~68 kDa) and MxaI (~11 kDa) subunits were expected to appear as distinct bands. However, no clear bands were detected at the predicted molecular weights, suggesting unsuccessful protein expression. This may be attributed to the use of a single-copy expression vector supplied by ANSA, resulting in low expression levels below detection limits.

To compare tolerance of potential host organisms, the minimum inhibitory concentrations (MICs) of ethanol and methanol were determined for E. coli and S. cerevisiae. Growth was measured across a dilution series of alcohol concentrations, and relative growth (AUC%) was plotted for each condition.

Minimum Inhibitory Concentrations

To compare tolerance of potential host organisms, the minimum inhibitory concentrations (MICs) of ethanol and methanol were determined for E. coli and S. cerevisiae. Growth was measured across a dilution series of alcohol concentrations, and relative growth (AUC%) was plotted for each condition.

Ethanol

Analysis of relative AUC% revealed that ethanol had a concentration-dependent inhibitory effect on both organisms. E. coli sustained relatively high growth up to 0.63% (v/v) ethanol, after which growth declined sharply. In contrast, S. cerevisiae was more sensitive, showing a gradual reduction in AUC% from the lowest concentrations tested. By 0.63% (v/v), S. cerevisiae growth had fallen below 50%, and was almost completely inhibited at 5% (v/v).

Methanol

E. coli maintained a stable growth profile across the methanol concentrations tested, with AUC% values remaining above 70% even at the highest levels. In contrast, S. cerevisiae showed increasing sensitivity as methanol concentration rose, with a decline in AUC% beginning at 1.25% (v/v) and complete inhibition by 10% (v/v). These findings demonstrate that E. coli has a considerably greater tolerance to methanol than S. cerevisiae.

These results indicate that E. coli is more tolerant than S. cerevisiae to both ethanol and methanol, supporting its use as the primary chassis for subsequent experiments.

Copper Induction of AmilCP Chimera

To assess copper responsiveness of the double-plasmid construct, E. coli transformants were induced with increasing copper concentrations (0.1, 0.2, 0.5, 1, and 2 mM). Cell pellets from induced and uninduced cultures were compared visually to non-transformed controls (10b).

Chimera transformant induced with copper 0.5 mM, 1 mM or 2 mM.

Chimera transformant: Three from the left have no copper and the fourth induced with 2 mM.

Non transformed cells: from left to right two have not been induced and the third induced with 2 mM (cells are NEB 10b).

Across all conditions tested, there was no visible colour change in the induced samples relative to uninduced controls or non-transformed cells. Even at the highest copper concentration (2 mM), chimera transformants showed no detectable amilCP expression. These results indicate that the double-plasmid system did not respond to copper induction under the tested conditions.

Copper Induction of AmilCP Plasmid

To assess copper-inducible expression of AmilCP, E. coli transformants carrying the AmilCP plasmid were induced with increasing copper concentrations (0.1–2 mM). Visual inspection of cell pellets revealed a detectable colour change in induced samples compared to uninduced controls. The colour change was most apparent at higher copper concentrations (≥1 mM), consistent with copper-responsive induction of AmilCP expression.

AmilCP transformant induced with copper 0.5 mM, 1 mM or 2 mM.

No copper induction. Left is the non-transformed cells and right is the AmilCP transformant.

Copper induction with 2 mM. Left is the non-transformed cells and right is the AmilCP transformant.

Colour change was observed in transformants induced with copper chloride at concentration from 0.5mM and higher. As shown in images above, tubes labelled with AmilCP showed cell pellets producing purple colour of AmilCP as compared to beige cell pellets in the negative control tubes labelled α. This is a positive result that confirmed the AmilCP plasmid had functioned as expected.

Methanol Induction of Chimera Plasmid

Chimera transformant induced with methanol 0.5%, 5% or 10%.

Non transformed cells on left and Chimera transformant are two sample on the right. Induced with 5% methanol (cells are NEB 10b).

To test whether the chimera construct was responsive to methanol, transformants were induced with 0.5%, 5%, and 10% methanol. Non-transformed E. coli (NEB 10β) cells were used as negative controls.

No visible colour change was observed in the chimera transformants at any methanol concentration when compared to the controls. These results indicate that the chimera plasmid did not respond to methanol induction under the tested conditions.