iGEM 2025 — Team macquarie-australia

{% extends "base.html" %}
{% block hero\_title %}Notebook{% endblock %}
{% set sections = [
{
"id":"apr-2025",
"title":"April 2025",
"open": true,
"body": '

#### Week 1 · 14 Apr – 20 Apr

* **IGEM Bootcamp**
  While still finalising the project and plasmid design, the team decided to do a bootcamp for members to practice and be familiar with the laboratory techniques that will be used throughout the project. Preparations include making LB media, plates (LB, LB Cam and LB Spec), doing a plasmid mini prep, and making glycerol stock.
* Separately, our team set up overnight cultures for *E.coli* amilcp and *E.coli* rfp to decide on which chromoprotein would be best used for the project. After this, a plasmid mini prep was done and were later sent for sequencing.

#### Week 3 · 28 Apr – 4 May

* **Minimum Inhibitory Concentration (MIC) for Bacillus, *E.coli* and Yeast**
  **28/04/25**
  Three treatments per organism: MIC Methanol; MIC Ethanol; and pH test
  Two organisms: Escherichia coli DH5α and Saccharomyces cerevisiae.
  E. coli prefers LB media and 37℃. S. cerevisiae prefers YPD media and 30℃.
  Overnight cultures were set up for the following organisms.
* **29/04/25**
  OD measurements were taken using 1:10 culture.
  MIC experiments using different media were done in 96 well plates for the two organisms above:
  Ethanol 80% media
  Methanol 20% media
  PH media
  A serial dilution was done for both ethanol and methanol MIC experiments. Plate setups are as shown below. Growth curve measurements were taken using the Agilent Biotek LogPhase 600 Microbiology Microplate Reader. Results were analysed using GraphPad Prism.

---

#### MIC Plate Setups

**Methanol concentration**
Start with *20% v/v (2×) methanol*    note: methanol is 32.04 g per mol

MIC Plate Set-UP — % Final concentrations (methanol)

| Row | 1 | 2 | 3 | 4 | 5 | 6 | 7 | 8 | 9 | 10 | 11 Neg ctrl | 12 Pos ctrl |
| --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- |
| a | 10 | 5 | 2.5 | 1.25 | 0.625 | 0.3125 | 0.15625 | 0.078125 | 0.039063 | 0.019531 | Neg ctrl | Pos ctrl |
| b | 10 | 5 | 2.5 | 1.25 | 0.625 | 0.3125 | 0.15625 | 0.078125 | 0.039063 | 0.019531 | Neg ctrl | Pos ctrl |
| c | 10 | 5 | 2.5 | 1.25 | 0.625 | 0.3125 | 0.15625 | 0.078125 | 0.039063 | 0.019531 | Neg ctrl | Pos ctrl |
| d | 10 | 5 | 2.5 | 1.25 | 0.625 | 0.3125 | 0.15625 | 0.078125 | 0.039063 | 0.019531 | Neg ctrl | Pos ctrl |
| e | 10 | 5 | 2.5 | 1.25 | 0.625 | 0.3125 | 0.15625 | 0.078125 | 0.039063 | 0.019531 | Neg ctrl | Pos ctrl |
| f | 10 | 5 | 2.5 | 1.25 | 0.625 | 0.3125 | 0.15625 | 0.078125 | 0.039063 | 0.019531 | Neg ctrl | Pos ctrl |
| g | 10 | 5 | 2.5 | 1.25 | 0.625 | 0.3125 | 0.15625 | 0.078125 | 0.039063 | 0.019531 | Neg ctrl | Pos ctrl |
| h | 10 | 5 | 2.5 | 1.25 | 0.625 | 0.3125 | 0.15625 | 0.078125 | 0.039063 | 0.019531 | Neg ctrl | Pos ctrl |

**Initial load**: 100 µL 20% methanol (col 1); 50 µL media in cols 2–10; 100 µL media in col 11; 50 µL media in col 12.
**Next**: Transfer 50 µL serial dilution till column 10.
**Finally**: Add 50 µL bacterial culture. Start loading culture from column 12, skip 11, then 10 → 1 so the same tips can be used.

**Ethanol concentration**
Start with *80% v/v (2×) ethanol*

MIC Plate Set-UP — % Final concentrations (ethanol)

| Row | 2 | 3 | 4 | 5 | 6 | 7 | 8 | 9 | 10 | 11 Neg ctrl | 12 Pos ctrl |
| --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- |
| a | 40 | 20 | 10 | 5 | 2.5 | 1.25 | 0.625 | 0.3125 | 0.15625 | Neg ctrl | Pos ctrl |
| b | 40 | 20 | 10 | 5 | 2.5 | 1.25 | 0.625 | 0.3125 | 0.15625 | Neg ctrl | Pos ctrl |
| c | 40 | 20 | 10 | 5 | 2.5 | 1.25 | 0.625 | 0.3125 | 0.15625 | Neg ctrl | Pos ctrl |
| d | 40 | 20 | 10 | 5 | 2.5 | 1.25 | 0.625 | 0.3125 | 0.15625 | Neg ctrl | Pos ctrl |
| e | 40 | 20 | 10 | 5 | 2.5 | 1.25 | 0.625 | 0.3125 | 0.15625 | Neg ctrl | Pos ctrl |
| f | 40 | 20 | 10 | 5 | 2.5 | 1.25 | 0.625 | 0.3125 | 0.15625 | Neg ctrl | Pos ctrl |
| g | 40 | 20 | 10 | 5 | 2.5 | 1.25 | 0.625 | 0.3125 | 0.15625 | Neg ctrl | Pos ctrl |
| h | 40 | 20 | 10 | 5 | 2.5 | 1.25 | 0.625 | 0.3125 | 0.15625 | Neg ctrl | Pos ctrl |

**Initial load**: 100 µL 80% ethanol (col 1); 50 µL media in cols 2–10; 100 µL media in col 11; 50 µL media in col 12.
**Next**: Transfer 50 µL serial dilution till column 10.
**Finally**: Add 50 µL bacterial culture. Start loading culture from column 12, skip 11, then 10 → 1 so the same tips can be used.

**pH test**
Start with media of 10 different pH measures

MIC Plate Set-UP — pH measures

| Row | 1 | 2 | 3 | 4 | 5 | 6 | 7 | 8 | 9 | 10 | 11 Neg ctrl | 12 Pos ctrl |
| --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- |
| a | 2 | 2.5 | 3 | 3.5 | 4 | 4.5 | 5 | 5.5 | 6 | 6.5 | Neg ctrl | Pos ctrl |
| b | 2 | 2.5 | 3 | 3.5 | 4 | 4.5 | 5 | 5.5 | 6 | 6.5 | Neg ctrl | Pos ctrl |
| c | 2 | 2.5 | 3 | 3.5 | 4 | 4.5 | 5 | 5.5 | 6 | 6.5 | Neg ctrl | Pos ctrl |
| d | 2 | 2.5 | 3 | 3.5 | 4 | 4.5 | 5 | 5.5 | 6 | 6.5 | Neg ctrl | Pos ctrl |
| e | 2 | 2.5 | 3 | 3.5 | 4 | 4.5 | 5 | 5.5 | 6 | 6.5 | Neg ctrl | Pos ctrl |
| f | 2 | 2.5 | 3 | 3.5 | 4 | 4.5 | 5 | 5.5 | 6 | 6.5 | Neg ctrl | Pos ctrl |
| g | 2 | 2.5 | 3 | 3.5 | 4 | 4.5 | 5 | 5.5 | 6 | 6.5 | Neg ctrl | Pos ctrl |
| h | 2 | 2.5 | 3 | 3.5 | 4 | 4.5 | 5 | 5.5 | 6 | 6.5 | Neg ctrl | Pos ctrl |

**Initial load**: 100 µL media into cols 1–12.
**Finally**: Add bacterial culture.

'
},
{
"id": "jul-2025",
"title": "July 2025",
"open": true,
"body": "

#### Week 14 · 14 Jul – 20 Jul

* **14/07/25**
  Supelco Formaldehyde Test

  A formaldehyde test was done using Supelco Formaldehyde Test. To test for the first time, we created different formaldehyde solutions to match those in the kit’s protocol, and also included 5mg/L and 200mg/L to test for concentrations outside its detection range. We confirmed that the color matching was consistent with the protocol.

* **16/07/25**
  Bromothymol Blue (BTB) Formaldehyde Test

  A BTB Experiment was done according to ThepChuay Experiment.

#### Week 15 · 21 Jul – 27 Jul

* **29/07/25**
  In addition to the BTB Experiment, the same experiment was done using 1mM Iron.

"
},
{ "id":"aug-2025",
"title":"August 2025",
"open": true,
"body": "

#### Week 17 · 4 Aug – 10 Aug

* **08/08/25**
  Supelco Formaldehyde Test Strips With Ethanol

  This experiment was done using a range of 5mL ethanol concentrations (0%, 5%, 10%, 20%, and 40%) with a range of formaldehyde added (0mg, 40mg, and 100mg).

  Supelco Formaldehyde Test Strips With Ethanol

  This experiment was repeated without ethanol. However, 1mL of each of the concentrations were used instead of 5mL which was sufficient enough to submerge the test strips. Results show that there is a clear gradient but the colors were all darker than expected. It also showed that the formaldehyde test works in strong 40% ethanol background.

* **11/08/25**
  Cell Transformation of DH5a for plasmid amilCP with copper promoter

  DH5a was transformed with the amilCP plasmid containing the copper promoter. This was done using the NEB High Efficient Transformation Protocol. Transformants were plated in different dilutions (1x, 10x, and 100x). pUC19 in Amp plate was used as a positive control, while pUC19 in Kan plate as a negative control.
* **12/08/25**
  Rususpension of Primers

  The following primers were resuspended using TE buffer to a final concentration of 0.1nM.
  mxaA\_F
  mxaK\_R
  amilCP\_R
  amilCP\_F
  MDH\_R
  MDH\_F
  Chim\_R
  Chim\_R

  Primers were diluted in 1:10 to have 10uM as aliquots to be easily used.

* **12/08/25**
  PCR for amilCP with copper promoter transformants
  A PCR was done to the amilCP transformants. A mastermix was prepared containing GoTaq Hot Start Green 2x MM buffer, amilCP\_F, amilCP\_R, and MilliQ. 5 colonies from the 10^-3 diluted transformant plates were picked, along with an amilCP plasmid as a control.

* **14/08/25**
  A gel electrophoresis was performed to the PCR products. Results show that bands are visible for all the 6 colonies and the positive control.
  Colonies with correctly assembled plasmid contructs that were verified using PCR were inoculated into LB media with KMSO and incubated at 37C with 200 rpm shaking.

* **14/08/25**
  Induction of amilCP transformants with copper promoter using copper chloride
  Cultures were inoculated in LB with 50mg/mL KMSO antibiotic to OD600 = 0.1. These were growth to OD600 = 0.5-0.7 at 37C with 200rpm shaking. The cultures were split into 3mL aliquots. Copper chloride was added to 1uM, 10uM, and 20uM, and then the cultures were placed into shaking incubator at room temperature with 200rpm shaking for 5 hours and checked for color change.

* **15/08/25**
  Plasmid Mini Prep of plasmid with amilcp was done according to protocol except for the final step. Once wash 2 was discarded, the samples were centrifuged for another 2 minites to remove residual wash.
  Following the induction of the amilcp gene by cusC, we spun down the samples to check if there was color. After 25hours induction, no color has been produced.

#### Week 18 · 11 Aug – 17 Aug

* **18/08/25**
  Mxa support protein plasmid transformation in 5-alpha and BL21 cells
  Transformation was done according to experimental protocol.
  Competent cells used: C2987 NEB 5-alpha competent *E.coli* cells & C3010 T7 express lysY competent *E.coli* cells
  Plasmid: MDH support protein plasmid DNA
  Media: SOC recover media
  After transformation, a serial dilution was done for 10x and 1000x for samples and plated into LB Cam plates and incubated in 37C.

* **19/08/25**
  PCR was done according to protocol.
  Primers used: mxaA\_F & mxaK\_R
  Colonies used: 8 colonies from 18/08/25
  8 reactions were done, including one control (mxA plasmid)
  mxaA\_F: Tm = 55.8C
  mxaK\_R: Tm = 55.8C
  For 25 cycles

| Temperature | Time |
| --- | --- |
| 95°C | 2 min |
| 95°C | 15 sec |
| 50°C | 30 sec |
| 72°C | 45 sec |
| 72°C | 5 min |
| 4°C | HOLD |

* After PCR, an agarose gel electrophoresis was done. 5uL of samples were loaded onto the wells, and a quickload ladder was used.
* Overnight cultures were also set up for the following samples
  Mxa plasmid *E.coli* BL21
  Mxa plasmid *E.coli* Dh5a
  AmilCP plasmid *E.coli* Dh5a

#### Week 19 · 18 Aug – 24 Aug

* **21/08/25**
  From previous overnight cultures, glycerol stocks were made, followed by a plasmid mini prep.
  Overnight cultures were done for the following:
  Mxa *E.coli* Dh5a cam
  Mxa *E.coli* BL21 cam

* **22/08/25**
  A plasmid mini prep was done for the MDH accessory proteins (DH5a and BL21).
  SOB and TB mdia were also prepared.

**Transformation Buffer (TB) Recipe**
Final composition: 10 mM HEPES, 15 mM CaCl₂, 250 mM KCl, 55 mM MnCl₂. Adjust pH to 6.7 with KOH before adding MnCl₂. Do not adjust pH after MnCl₂ is added. Filter sterilise and store at 4 °C.

*100 mL Transformation Buffer*

| Reagent | Amount | Notes |
| --- | --- | --- |
| HEPES | 0.238 g 1.0 mL (1 M stock) | Choose one option |
| KCl | 1.864 g | — |
| CaCl₂ | 0.167 g (anhydrous) 0.221 g (dihydrate) | — |
| MnCl₂ | 0.692 g (anhydrous) 1.089 g (tetrahydrate) | Dissolve separately and add last |

*250 mL Transformation Buffer*

| Reagent | Amount | Notes |
| --- | --- | --- |
| HEPES | 0.596 g (free acid) 0.651 g (sodium salt) 2.5 mL (1 M stock) | Choose one option |
| KCl | 4.660 g | — |
| CaCl₂ | 0.416 g (anhydrous) 0.551 g (dihydrate) | — |
| MnCl₂ | 1.730 g (anhydrous) 2.721 g (tetrahydrate) | Dissolve separately and add last |

*Preparation Protocol*

1. In beaker, add ~80% of the final volume of milliQ water.
2. Add HEPES (powder or 1 M stock), dissolve.
3. Add KCl, dissolve.
4. Add CaCl₂, dissolve.
5. Adjust pH to 6.7 with KOH.
6. Dissolve MnCl₂ in a small volume of milliQ water separately (~10% of the final volume).
7. Add MnCl₂ solution gradually to the base solution with stirring.
8. Do not adjust pH after adding MnCl₂ (a drop in pH is expected).
9. Bring to final volume with milliQ water.
10. Filter sterilise.
11. Store at 4 °C (indefinitely stable).

#### Week 20 · 25 Aug – 31 Aug

* **25/08/25**
  Formaldehyde Experiment
  For this experiment, 0.04% of BTB was used which is 0.004g of BTB in 10mL H2O.
  2 concentrations of Hydrogen Peroxide were used (0.1M and 0.005M) alongf with 0.01M of Sodium Hydroxide.
  Iron chloride was also used for this experiment alongside iron sulfate.
  FeSO4 (0.4% and 0.8% final concentration)
  FeCl3 (0.4% and 0.8% final concentration)
  These were done on a 96-well plate. Images of the plate can be found in the result section of the wiki.

* **26/08/25**
  Transformation of plasmids

  Plasmids used: MDH mxaAK (KanR), MDH (MoxF1), amilCP MxaY, and MDH MoxF1
  Cells used: Dh5a, 10b, abnd T7lysY
  Dh5a was transformed with MDH mxaAK (KanR). 10b was transformed with MDH MoxF1 and amilCP MxaY. T7lysY was transformed with MDH MoxF1
  Transformation protocol was used using SOC media for DH5a and T7lysY cells and using specific 10b media for 10b cells.
  2 LB plates were done for each (1/100 or 1x10^-2 dilution) and 100uL of culture.

* **28/08/25**
  Colony PCR was done for plasmids NEB 10b PMDH MoxF1, Dh5a pMDH MxaAK, and NEB 10b pAmilCPMxaycuss.
  PCR protocol was used.
  PCR setup:

| Primers | Annealing Temperature (°C) | Extension Time (sec) |
| --- | --- | --- |
| MDH | 46 | 45 |
| Mxa | 50 | 45 |
| amilCP | 50 | 45 |

* A gel electrophoresis was also done according to protocol.
  Colony 1 from the PCR plasmid came out to be the best and was inoculated into 10mL LB broths with Km antibiotic(10uL).
  Strains were inoculated at 37C.

* **29/08/25**
  Glycerol stocks were done in 2 replicates to dh5a pMDHMxaAK, NEB 10b pMDH MoxF1, and NEB 10b pAmilCPMxaYCuss. Following this, a plasmid miniprep was done according to protocol.
  Overnight cultures were set up for NEBa pAmilcp and no plasmid, as well as NEB10beta pamilcpMxaYCusS and no plasmid for induction.

" },
{ "id":"sep-2025",
"title":"September 2025",
"open": true,
"body": "

#### Week 21 · 1 Sep – 7 Sep

* **02/09/25**
  Transformation
  NEB T7 express competent cells were transformed with pMDH MoxF1, pMDH MoxF1 + pMca, or PUC19. Transformation was done according to protocol except was heatshocked for only 10 seconds, put on ice for 5 minutes. SOC media was used.
  Streak plates were also done for the transformants.

* **03/09/25**
  Induction of chimera sensor
  5mL cultures were set up
  1. NEBa no plasmid
  2. NEB10beta no plasmid
  3. NEBa amilcp
  4. NEB10beta amilcp chimera
  Cultures were incubated at 37C

* **04/09/25**
  The OD600 for the overnight cultures were measured. 0.1% (w/v) glucose cultures were inoculated to OD600 and grown at 37C to mid exponential phase.
  Copper chloride induction
  3mL cultures were set up. 3 concentrations were done (1mM, 2mM, and 500uM)
  Methanol induction
  3mL cultures were set up. 4 concentrations were done (0%, 1%, 2.5%, and 5%)
  Cultures used were NEBa amilcp, NEB10beta amilcp chimera, NEBa no plasmid, and NEB1b no plasmid

* **05/09/25**
  Transformation
  T7 express cells were transformed accoriding to transformation protocol.
  4 plasmids were used: moxF1, MxaAK, MxaAK + MoxF1 and Puc19.
  These were plated. MoxF1 and MxaAK were diluted in 1x, 1/10x and 1/100x, double trasnformation with 1x and PUc19 in 1x and 1/10x.
  One extra plate was done for each transformation. Concentrated cells were spin down and the pellet was plated labelled as concentrated cells.
  For double plasmids, these were recivered for 60 minutes at 37C. Recovered cells were added to 2mL of LB with Kan+Cam. Additional recovery was done for 60 minutyes at 37C and plated into LB Kan+cam plates.

* **06/09/25**
  Cells were spun down. Results are photograpged. Amilcp cells were then resuspended and transferred to 15mL tubes

#### Week 22 · 8 Sep – 14 Sep

* **08/09/25**
  Cultures were inoculated at 10am. OD600 measurements were taken at different time points until it reached 0.6. Glycerol stocks were made after according to protocol and were stored in –80C.
  Control AmilCP Copper were induced using 10uL of 20mM Cu2Cl. No immediate color response during the day and will be further observed the following day.
  Colony from pMDH MoxF1 KanR (05/09/25) was inoculated in 3mL LB +Kan in 15mL falcon tube and incubated in 37C.

* **09/09/25**
  Cells were inoculated in SOB 25mL(25uL Kan) with 500uL of overnight culture and incubated at 30C.
  T7x MDH MoxF1: KanR cells were made chemically comptent according to protocol.

* **10/09/25**
  Transformation was done to pMDH MoxF1 + pMxa KanR/CamR and Puc19 according to protocol.
  Puc19 was plated with 1/10x dilution. MDH MoxF1 + pMxa KanR/CamR was plated in 1x, 1/10x, and 1/100x dilution. Plates were stored in 37C.
  Overnight cultures for pamilCP and pchimera were done for copper inductions the following day. LB kan was used for cells with plasmid, and LB only for controls (NEB alpha and NEB 10b cells)

* **11/09/25**
  Expression of methanol dehydrogenase was done using transformants plates from 10/09/25. Colonies from the 1:10 dilution plate was suspended into 5mL LB.
  Copper induction of amilcp and chimera plasmids:
  1. *E.coli* 10b pamilcp chimera LB Kan
  2. *E.coli* NEBa pamilcp LB Kan
  3. *E.coli* 10b no plasmid
  4. *E.coli* NEBa no plasmid
  Cells were spun down to pellet and supernatant was discarded. Pellet was resuspended woth 1xTBS (500uL). 1mM CuCl2 was added to *E.coli* NEBa pamilcp and NEB10alpha. 5mM CuCl2 was added to *E.coli* 10b and *E.coli* 10b pamilcp chimera.

  Preparation of PQQ (330g/mole)
  In a falcon tube, add 1 tablet of PQQ and add 12mL of MilliQ. Each tablet had 40mg of PQQ [10mM]. Due to starch present, it does not dissolve properly, spin the solution for 10 minutes. Make a sterilised PQQ solution by transferring it using a syringe and 0.2um filter into a new falcon tube.

  Protein expression using T7 express
  OD of *E.coli* T7x pMDHMoxF1 KanR + pMxa was checked and grown to OD=0.4. 3uM of PQQ was added. The cell culture was divided into two conical flask. One is induced with0.4mM IPTG and other is uninduced. Flasks were incubated at 16C overnight.

  The following buffers were prepared: 2x binding/wash buffer, His elution buffer, and Buffer modifiers.

  |  | Sodium Phosphate | NaCl | Imidazole | Tween–20 |
  | --- | --- | --- | --- | --- |
  | 2× Binding/Wash Buffer | 0.69 g | 1.75 g | — | 10 μL |
  | His Elution Buffer | 0.345 g | 0.877 g | 1.03 g | 5 μL |
  | 1 M NaCl | — | 2.92 g | — | — |
  | 0.1 M Imidazole | — | — | 0.345 g | — |
* **12/09/25**
  Methanol Dehydrogenase was extracted and purified from the induced and uninduced cells with 2 replicates each. Cells were spun down and resuspended in 2.5mL of TE buffer, spund down and discreded the TE buffer. It was then resuspended in 2.5mL 1x binding buffer, sonicated for 3 mins on 10 sec pulse to lyse. Protein Isolation and purification were done according to protocol. This was then stored in the –80C.

#### Week 23 · 15 Sep – 21 Sep

* **15/09/25**
  Protein gel was run using cell pellet, supernatant containing lysed cells, and purified protein (induced vs. Uninduced) according to protocol.

  Gel running buffer preparation was done. 100mL of 4x Tris/Glycine/SDS and 900mL of RO water were used to make 1L of 1x Tris/Glycine/SDS Buffer.
  Protein concentration for supernatant and purified protein were measured using the nanodrop.
  Loading samples were prepared in PCR tubes for the following (1 set for 10ul and 1 set for 15uL of protein sample)
  Samples:
  Purified protein (induced) 2 replicates
  Purified protein (uninduced) 2 replicates
  Lysates (induced) 1 replicate
  Lysates (uninduced) 1 replicate

  For 10uL protein samples, 10uL of 2x loading dye was added. For 15uL protein samples, 15uL of 2x loading dye was added.
  SDS-PAGE was done according to protocol.
  Overnight staining and destraining were done according to Coleman protocol.
  Overnight cultures were done for amilcp, amilcp chimera, and pMDH MoxF1 + Mxa and their controls.

* **16/09/25**
  Cells were grown to OD = 0.4
  1. T7x MDH
  2. NEBa amilcp
  3. NEB10beta chimera
  4. NEBa
  5. NEB10beta
  Cells were induced with either (0, 1, 2.5 and 5%) methanol and (0, 0.5, 1mM) copper
  For T7x pMDH MoxF1 + pMxA induced as per 11/09/25. 6uM of PQQ was added to the 50mL culture then split into two flasks. One was for inducing with 0.8mM IPTG.

* **17/09/25**
  Purification of protein was done according to protocol. Samples were placed in –80C freezer.
  Samples from 16/09/25 were spun down and photographed results.
  Overnight cultures for pMDH MoxF1 + pMxa t7x was done.

* **18/09/25**
  pMDH MoxF1 + pMxa t7x was induced at OD=0.65. 300uL of the 10mM PQQ stock was added to 500mL culture then split culture into 2 lots of 250mL both in 2L flasks, added IPTG 250uL of 1M stock to 250mL culture to induce 1 flask. PQQ = 6uM and IPTG=1mM.

  SDS Page was done according to protocol for the following samples. 15uL and 10uL protein samples were loaded.
  1. MDH induced
  2. MDH uninduced
  3. Contaminated MDH induced
  4. Contaminated MDH uninduced
  5. Supernatant induced
  6. Supernatant uninduced
  7. *E.coli* pellet induced – only for 10uL
  8. *E.coli* pellet uninduced – only for 10uL

  The following were prepared: 2x xT sample buffer, 10% v/v beta-merck to make up the gel running buffer.
  For 5uL protein, 9.5uL of samples were loaded while 20uL was loaded for 10uL protein.

  Destaining solution was also made using 10% (v/v) methanol and 10% (v/v) acetic acid.
  Gel was stained in Coomassie blue staining solution for 30 minutes. Stain was removed and added high destain for 30 minutes. After removing high destain solution, pour low destain solution and keep overnight.

* **19/09/25**
  1. Testing of PBS affects the formaldehyde test strips, testing the strips at 40mg/L
  Control: 900uL water + 100uL (0.04%) formaldehyde
  Test: 900uL PBS + 100uL (0.04%) formaldehyde
  Added two drops of Fo1
  Result – PBS does affect formaldehyde strips. Color change was inconsistent with reference concentration

  2. Repeat of experiment, did one formaldehyde strip just with PBS and water, no formaldehyde added but 2 drops Fo1
  Result – no color change for PBS or water

  3. Repeat of experiment
  Control: 900uL water + 100uL formaldehyde (0.04%) + 2 drops pf Fo1
  Test: 900uL + 100uL formaldehyde (0.04%) + 4 drops pf Fo1
  Result – correct color change observed

  4. Repeat experiment
  Control: 900uL NaCl (0.9%) + 100uL formaldehyde
  Test: 900uL of sodium chloride (0.9%) + 100uL formaldehyde + 2 drops of Fo1

  0.15M =0.9% NaCl used filter sterilised sodium chloride for the experiment “resting cell assay” rather than LBS as PBS interferes with the ph
  Results – 0 formaldehyde for all

#### Week 24 · 22 Sep – 28 Sep

* **22/09/25**
  Uninduced and induced pmdh moxf1 mxa thawed at room temperature, pellet cells and resuspended in 1x binding buffer. These were sonicated for 3 minutes. Dynabeads protocol was followed with 700uL samples to compensate, insteadd of 4 washes, doing 5 wases with 1x binding buffer, 500uL. The resulting supernatant was transferred to fresh centrifuge tubes and stored in –80C freezer.
  Samples in the end: 2x supernatant, 2x pellet and 2x pellet with lower concentration, and 2x his tagged purified.

* **23/09/25**
  SDS- gel run for protein (similar procedure from 18/09/25)
  The following samples were used and run in the gel. 20uL was loaded into each well
  1. MDH (in)
  2. MDH (un)
  3. Supernatant (in)
  4. Supernatant (un)
  5. Pellet (in)
  6. Pellet (un)
  Pellet was resuspending in 1x loading dye.
  Gel stain and destain was also done.

"}] %}
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{
"id": "intro-dbtl",
"title": "Introduction",
"body": "

The Design-Build-Test-Learn (DBTL) cycle is a core engineering framework in synthetic biology, enabling systematic iteration and optimization of biological systems. In our project, we applied DBTL across four interconnected experimental lines: chassis selection, formaldehyde detection assay, whole-cell biosensor design, and enzyme-based biosensor design. Rather than treating these components as isolated modules, we leveraged insights from the Learn phase of each to inform and enhance the others. This integrative approach fostered cross-component refinement and accelerated development. The interplay between these DBTL cycles is illustrated in Figure 1.

![Overview of DBTL cycle integration in MethaNO project](https://static.igem.wiki/teams/5787/engineering/1.avif)

**Figure 1:** Overview of the DBTL integration across chassis selection, formaldehyde detection, whole-cell, and enzyme-based biosensor design pathways.

",
"open": true
},
{
"id": "chassis-selection",
"title": "Chassis Selection",
"body": "

For both our whole-cell two-component system and methanol dehydrogenase enzyme designs, we needed to select one chassis organism. Correct chassis selection is vital when working in backgrounds with varying conditions. For a methanol biosensor that works in cocktails, it is crucial our chassis microbe can tolerate different levels of methanol, ethanol, and varying pH. Our lab commonly sees synthetic biology work conducted in *E. coli* and *Saccharomyces cerevisiae*. These two model organisms were thus subject to the DBTL cycle to assess their tolerance of our required conditions.

##### CYCLE 1

###### Design

To mimic the conditions present in a methanol-tainted alcoholic beverage, we designed a maximum tolerance test for *E. coli* DH5α and *S. cerevisiae* BY4742. The range of ethanol and methanol concentrations were designed to match those potentially found in cocktails, while the pH levels ranged from neutral pH to 2, as cocktails can vary from neutral to acidic.

###### Build

A set of serial dilutions were prepared in 96-well plates. Methanol and ethanol concentrations ranged from low to high, and pH values ranged from neutral to acidic. Tolerance testing conditions are listed in Table 1. A constant amount of cell culture — either LB or YPD for the respective microbes — was added to all wells.

**Table 1.** We chose to subject the microbial candidates to the ranges listed below. Varying conditions of Methanol, Ethanol, and pH levels.

| Conditions | Varying Concentrations | | | | | | | | | |
| --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- |
| Methanol | 10% | 5% | 2.5% | 1.25% | 0.63% | 0.31% | 0.16% | 0.08% | 0.04% | 0.02% |
| Ethanol | 40% | 20% | 10% | 5% | 2.5% | 1.25% | 0.63% | 0.31% | 0.16% | 0.08% |
| pH | 2 | 2.5 | 3 | 3.5 | 4 | 4.5 | 5 | 5.5 | 6 | 6.5 |

###### Test

Plates were incubated overnight in a Log Phase 600 plate reader, with optical density measurements taken to assess growth. Growth curves were visualized using the Log Phase 600 App software. To quantify overall growth inhibition across the tested concentration ranges, the area under the curve (AUC) was calculated for each tolerance assay. AUC values enabled a direct comparison of tolerance profiles between *E. coli* and *S. cerevisiae*.

###### Learn

Both chassis options showed progressive decline as the methanol and ethanol concentrations increased and pH decreased. AUC results showed that *E. coli* performed better than yeast across methanol and ethanol concentrations and comparably in lower pH levels. The results guided the choice of *E. coli* as the optimal chassis for future biosensor research. The findings underscore the significance of empirical chassis selection in biosensor design and provide a basis for developing resilient, field-deployable synthetic biology instruments to combat methanol adulteration in alcoholic drinks.

",
"open": true
},
{
"id": "formaldehyde-detection",
"title": "Formaldehyde Detection",
"body": "

For our biosensor's enzyme design, we planned to employ a colorimetric assay to indicate the presence of methanol. As the enzyme design involves the reaction of methanol dehydrogenase oxidising the methanol to formaldehyde, a colorimetric test for detecting formaldehyde was required.

##### CYCLE 1

###### Design

We began by reviewing literature on colorimetric assays for formaldehyde detection, prioritising methods that were simple to perform, cost-effective, used readily available reagents and equipment, and produced a rapid and distinct colour change. The assay described by Thepchuay et al. [1] met these criteria and was selected for further development. In this method, formaldehyde is oxidized to formic acid using hydrogen peroxide under alkaline conditions. Bromothymol blue (BTB) serves as the pH indicator, shifting from blue (alkaline) to yellow (acidic) as formic acid is generated.

**Methanol to formaldehyde reaction**
`2 CH₃OH + O₂ → 2 CH₂O + 2 H₂O`

**Formaldehyde to formic acid reaction**
`2 CH₂O + H₂O₂ + 2 NaOH → 2 HCOONa + 2 H₂O + H₂`

The stoichiometric conversion of one mole to one mole provides a simple proxy for methanol detection via formaldehyde.

###### Build

Assay reagents were prepared according to the optimized concentrations reported in the paper Thepchuay et al. [1]. Unfortunately, we found the assay could not be replicated as described in the paper and as such required further refinement (see [Notebooks](https://2025.igem.wiki/macquarie-australia/notebook) and [Results](https://2025.igem.wiki/macquarie-australia/results) pages).

###### Test

Multiple reagent combinations with varying component volumes were tested to identify the most effective setup for formaldehyde detection. A range of concentrations of formaldehyde were used to evaluate the performance of the reagent. The goal was to determine which formulation produced the clearest and fastest colour change.

###### Learn

Thepchuay et al. [1] reported that the colour change should occur within 5 minutes before formic acid begins to evaporate. However, in our trials, no colour change was observed within this timeframe across all tested formaldehyde concentrations.

##### CYCLE 2

###### Design

To address the lack of colour change observed in Cycle 1, we conducted a further literature review and modified the assay design by incorporating iron salts into the reagent solution. Hydrogen peroxide alone is a relatively weak oxidising agent; however, through the Fenton reaction, iron ions catalyse its activity, enhancing the oxidation of formaldehyde to formic acid [2]. This adjustment aimed to improve the sensitivity and reliability of the colorimetric response.

###### Build

We continued to revise the assay using a 96-well microplate format to allow for replicates and visual comparison across a gradient of formaldehyde concentrations. The plate was configured to test two concentrations of hydrogen peroxide and two types of iron salts — iron(II) sulphate and iron(III) chloride — to evaluate their catalytic efficiency.

###### Test

We tested a range of formaldehyde concentrations aligned with the detection range of Supelco® formaldehyde test strips, enabling direct benchmarking against a commercial standard. Additionally, the observation window was extended beyond 5 minutes to assess whether delayed colour change could occur under the modified conditions.

###### Learn

Our finding was that it took one week to achieve the most discerning colour range indicative of the formaldehyde concentration range tested (see [Results](https://2025.igem.wiki/macquarie-australia/results) page). We would like to further explore measures to accelerate the speed of detection since it has been reported possible within 5 minutes [1]. Despite having access to the commercially available Supelco® formaldehyde test strips, the 96-well format chemical assay represents an opportunity for high throughput assessment of both methanol dehydrogenases (MDH), and promoter–MDH combinations, in future experiments.

",
"open": true
},
{
"id": "tcs-chimera-biosensor",
"title": "TCS Chimera Biosensor",
"body": "

For our whole-cell biosensor, we chose to utilise a two-component system (TCS), as it enables bacteria to sense and respond to specific environmental stimuli. These systems offer modularity and tunability, two important attributes for synthetic biology biosensors [3]. We began with a literature review to identify a well-characterised TCS responsive to methanol. Previous studies have demonstrated successful methanol detection using an engineered EnvZ/OmpR system, which initially captured our interest [4][5]. These described a chimeric protein that had the MxaY methanol sensing domain fused to the EnvZ histidine kinase, allowing the system to respond to methanol and signal a downstream response [4]. While this work inspired our approach, the papers did not present their methods in a reproducible manner. We were also wary that since EnvZ histidine response is associated with outer membrane porins, such a design may unintentionally allow excess toxic methanol and ethanol into the cells [6].

We next looked for a more well-characterised TCS. The CusRS TCS was recently optimised for biosensor applications by adjusting signal amplification and expression levels of CusR and CusS [7] for the purpose of copper detection. We decided to move forward by designing our TCS with the CusRS system fused with the MxaY domain from *Paracoccus denitrificans*.

##### Plasmid Synthesis

##### CYCLE 1

###### Design

The CusRS two-component system detects environmental stimuli through its membrane-bound histidine kinase, CusS, which undergoes autophosphorylation upon activation. The phosphate group is then transferred to the response regulator, CusR, enabling CusR to bind DNA and initiate transcription from the CusR-regulated promoter [7]. In our design we utilise a chimeric histidine kinase that responds to methanol, while the response regulator remains unchanged.

For reporter output, we placed the gene encoding amilCP, a bright blue chromoprotein, under control of the CusR-regulated promoter. The strong visual signal of amilCP may provide an easily detectable output upon methanol exposure; amilCP is a standard iGEM biological part and common in teaching labs [Part: BBa\_K592009](https://parts.igem.org/Part%3ABBa_K592009). Additionally, the bidirectional nature of the CusR promoter allows for placement of two copies of the amilCP gene, further enhancing signal output. Our design includes both the chimeric histidine kinase and response regulator expression under a constitutive promoter ensuring the biosensor components are continually available to sense and respond. To maximize expression, we selected the constitutive promoter for *gapA*, which includes multiple transcriptional start sites (P1, P2, and P3) and ensures strong, constitutive expression in *E. coli* throughout various growth phases. In designing the plasmid we used the IDT plasmid design tool for codon optimisation.

###### Build

Synthesis of the plasmid was initially outsourced to Twist Bioscience. Sequences were uploaded to their systems to check the complexity which came back as standard. However, the initial plasmid synthesis was unsuccessful. Twist was informative, suggesting potential problems including a hairpin formed by the bidirectional amilCP, and repetitive regions.

###### Re-design

We aimed to eliminate the problems in the first plasmid design that led to failed synthesis. The bidirectional arrangement of the amilCP reporter gene resulted in the formation of a large hairpin structure, which complicated DNA synthesis and stability. To resolve this, we used two different synonymous DNA sequences encoding the same amilCP protein, effectively removing the hairpin issue. To further address repetitive regions, we incorporated three different terminators (T7, T7hyb10, and rrnBT1), thus breaking up sequence repeats. IDT tools were used to monitor the sequence complexity, which initially scored in the unacceptable range due to both repeats and filler DNA. We corrected these problems by deleting unnecessary filler sections, and once the complexity score was lowered, we ordered the improved design again from Twist. Unfortunately, Twist could not synthesise this new design either. Thankfully, ANSA Biotechnology offered to sponsor the Macquarie University team and successfully synthesised our new plasmid design.

Additionally, to enable functional validation of the chromoprotein output, a simplified plasmid was designed consisting only of the CusR promoter and bidirectional amilCP gene arrangement. This was successfully synthesised by IDT. In this design, the absence of the chimeric methanol sensor allows for direct induction by copper, leveraging the natural E. coli CusRS system to drive amilCP expression. This provided an efficient platform to confirm chromoprotein production and assess promoter responsiveness independent of the synthetic methanol-sensing module.

![TCS Chimera plasmid design schematic](https://static.igem.wiki/teams/5787/engineering/2.avif)

**Figure 2:** Plasmid for the whole-cell biosensor before redesign, highlighting the issue of identical terminators and potential hairpin formation with bidirectional amilCP.

###### Re-build

Synthesis of the main chimera plasmid was successful with ANSA, a pivotal step that allowed us to progress with our project. The resulting plasmid included the codon-optimised chimeric methanol-responsive histidine kinase, the CusR response regulator, the bidirectional amilCP reporter arrangement, and kanamycin resistance. The simplified plasmid ordered from IDT was successfully synthesised and will be used to test our chromoprotein.

###### Test

After receiving the synthesised plasmids, they were transformed into DH5α *E. coli* cells. A plasmid miniprep was performed to purify the plasmid DNA from the bacterial cultures. The purified plasmid samples were then sent for sequencing. Whole plasmid sequencing was performed by Plasmidsaurus using Oxford Nanopore Technology with custom analysis and annotation, which confirmed the sequences were as designed.

###### Learn

This whole process proved to be an invaluable lesson in plasmid construction, highlighting critical factors that must be considered at each stage. The challenges we faced, particularly regarding repetitive regions and hairpins, underscore the importance of rigorous sequence design. These insights will guide all subsequent plasmid projects. Prioritising sequence uniqueness and minimising repetitive elements will be essential for improving synthesis reliability and expediting successful construct assembly.

##### Functional Validation of Methanol and Copper-Responsive Reporter Systems

##### CYCLE 1

###### Design

The plasmids designed and sequence confirmed in the plasmid design section were transformed into *E. coli*. Experiments were designed to test the ability of our engineered cells to respond to methanol or copper respectively by producing amilCP. Both NEB 5-alpha and NEB 10-beta strains were chosen. Inductions with copper and methanol from low to high at multiple concentrations were planned to compare the sensitivity and specificity of each system.

###### Build

Overnight starter cultures were grown to mid-exponential phase, providing optimal conditions for induction. For each experimental condition, negative controls without plasmid were included to establish a baseline for non-specific colour development. All technical controls and biological replicates were run in parallel to ensure reproducibility and reliability. All induction concentrations were set up at increasing concentrations from minimal detection ranges to those becoming inhibitory for growth.

###### Test

Induction assays were carried out across a range of copper concentrations as well as varying levels of methanol. Post-induction, cultures were incubated, pelleted and then assessed for visible colour development. Negatives were included to monitor baseline colour and confirm specificity of the response, especially since copper chloride has a blue tinge. No visible colour development was detected in the chimera-containing cells under any induction condition, indicating a lack of functional reporter expression or activation. Visible blue colour development was observed in the amilCP-only cells following copper induction, confirming that the reporter system and induction method were functioning as expected.

###### Learn

Given the absence of colour development in chimera-containing cells, the next steps would involve troubleshooting the functionality, such as confirming protein expression and optimising induction protocols. The successful production of colour from the simplified amilCP design led us to register the bidirectional CusR promoter as a composite part [BBa\_25zau49i](https://registry.igem.org/parts/bba-25zau49i).

However, after seeking input from academic advisors and stakeholders, the consensus is to streamline efforts by focusing on a single approach. While building the chimera has been instructive, the enzyme-based design now presents a less risky, less regulated, and more scalable pathway. Although neither strategy has yet yielded complete success, the enzyme system is considered more promising for future development.

",
"open": true
},
{
"id": "mdh-enzyme-biosensor",
"title": "MDH Enzyme Biosensor",
"body": "

Methanol dehydrogenases (MDH) are essential enzymes that catalyse the oxidation of methanol to formaldehyde—the first and critical step in methanol metabolism. There are three main types of MDHs, classified by their electron acceptors: NAD-dependent, pyrroloquinoline quinone (PQQ)-dependent, and O₂-dependent [8]. NAD and PQQ-dependent MDHs are typically found in methylotrophic bacteria, while O₂-dependent variants are derived from yeast [8]. Based on learnings from our Chassis Selection DBTL cycle, we opted for a bacterial system over yeast. Although NAD-dependent MDHs are often favoured in synthetic methylotrophy due to their simpler genetic requirements, they exhibit lower substrate affinity for methanol compared to PQQ-dependent MDHs. Given that high substrate specificity is vital for MethaNO’s performance, we chose to pursue PQQ-dependent MDHs despite the added engineering complexity.

##### Plasmid Synthesis

##### CYCLE 1

###### Design

We selected the MDH from *Methylorubrum extorquens* AM1 (strain ATCC 14718), as it is one of the most extensively studied PQQ-dependent MDHs. Key literature, including gene identification [9], metabolic mechanism [10], and enzyme kinetics and purification technique [11], provided foundational insights for our construct. Based on this, we identified a set of essential genes required for functional MDH expression and sourced their protein sequences from UniProt. A His-tag was added to *mxaF* (Table 2) to facilitate downstream protein purification.

**Table 2.** Four genes identified as essential for a functional MDH synthesis – *mxaF* and *mxaI* code for the enzyme subunits, whilst *mxaA* and *mxaK* code for supporting proteins to facilitate co-factor Ca²⁺ incorporation into the enzyme.

| Gene | Function | Notes |
| --- | --- | --- |
| *mxaF* | Large (α) subunit of PQQ-MDH catalytic core | His-tag added for purification |
| *mxaI* | Small (β) subunit; completes MDH holoenzyme | Pairs with *mxaF* for activity |
| *mxaA* | PQQ/Ca²⁺ insertion/assembly factor | Supports cofactor loading |
| *mxaK* | Accessory protein for MDH maturation | Facilitates Ca²⁺ incorporation |

To streamline plasmid construction, we used the Integrated DNA Technologies (IDT) plasmid design tool to codon-optimise the sequences for *E. coli*, remove restriction sites incompatible with iGEM assembly standards, and reduce the overall complexity score of the design. The T7 promoter was chosen for its high transcriptional efficiency, enabling strong protein expression. Since MDH enzymes require an oxidising environment to function properly, signal peptides were incorporated to direct recombinant proteins from the cytoplasm to the periplasmic space. We adopted signal sequences with enhanced performance including optimised translation initiation regions (TIRs) as described in Merzadeh et al. [12].

###### Build

Synthesis of the initial plasmid design (Figure 3) was outsourced to Twist Bioscience. However, the synthesis attempt was unsuccessful. Twist kindly shared informative information regarding potential repetitive regions and hairpins which may have thwarted these initial synthesis attempts.

![Formaldehyde detection assay DBTL cycles](https://static.igem.wiki/teams/5787/engineering/3.avif)

**Figure 3:** Both mxaFI and mxaAK operons in one plasmid design for illustrative purpose.

###### Re-design

The redesigned plasmids minimised repetitive regions by utilising a greater variety of signal peptides. The tools from IDT proved particularly helpful in identifying areas of complexity. To further favour successful synthesis the new design was ordered with genes distributed across two distinct vectors (Figure 4 and Figure 5). Care was taken to ensure these included different antibiotic resistance markers and origins of replication for co-transformation compatibility:

* The *mxaFI* operon was cloned into a kanamycin-resistant pET vector.
* The *mxaAK* operon was cloned into a chloramphenicol-resistant pTwist vector.

![Formaldehyde detection assay DBTL cycles](https://static.igem.wiki/teams/5787/engineering/4.avif)

**Figure 4:** mxaFI operon only in one plasmid design for illustrative purpose.

![Formaldehyde detection assay DBTL cycles](https://static.igem.wiki/teams/5787/engineering/5.avif)

**Figure 5:** mxaAK operon only in one plasmid design for illustrative purpose.

Additionally, both plasmids were ordered from ANSA Biotechnology, who only offered kanamycin-resistant vectors.

###### Re-build & Test

Synthesis was successful for both plasmids via ANSA, whereas Twist successfully synthesised only the *mxaAK* plasmid. We greatly appreciate ANSA’s support in providing free synthesis services for complex DNA design and it was their successful synthesis of the *mxaFI* plasmid that allowed us to progress with our experiments.

###### Test

Whole Plasmid Sequencing was performed by Plasmidsaurus using Oxford Nanopore Technology with custom analysis and annotation. This confirmed the plasmids we cloned in C3019I NEB 10-Beta competent *E. coli* cells and C2987I NEB 5-Alpha competent *E. coli* cells were the sequence as designed.

###### Learn

Avoiding repetitive regions is critical for successful plasmid synthesis. Future designs should prioritise sequence uniqueness to enhance synthesis reliability.

##### MDH Enzyme Expression

##### CYCLE 1

###### Design

Plasmid expression was induced using 0.4 mM isopropyl β-D-1-thiogalactopyranoside (IPTG), as per documentation in [Notebook](https://2025.igem.wiki/macquarie-australia/notebook) page. To support cofactor-dependent enzyme activity, pyrroloquinoline quinone (PQQ) was supplemented at an initial concentration of 3 µM. Although *E. coli* do not synthesize PQQ endogenously, they possess high-affinity PQQ transporters that function effectively in environments with elevated PQQ concentrations (0.1–3 µM) [13].

###### Build

The double plasmid transformants were prepared on LB agar plates containing kanamycin and chloramphenicol (KanR and CamR), then overnight cultures inoculated. Three sample types were prepared: pellet resuspension, cell lysates, and purified protein. Detailed protocols are available in the Experiment and Notebook sections.

###### Test

Lysate and purified protein samples were analysed via SDS-PAGE to separate proteins by molecular weight and identify MDH. The MDH, assembled by two subunits, were expected at band ~70 kDa for *mxaF* and band ~10 kDa for *mxaI*. No clear bands of these sizes were observed in the purified protein samples.

###### Learn

Key areas for optimisation in Cycle 2 include:

* Induction efficiency: IPTG concentration or timing may require adjustment
* Cofactor loading: PQQ concentration may be suboptimal
* Cell density: Achieving OD600 ≥ 0.6 is critical for sufficient biomass
* Purification yield: Current protocol may result in low recovery of target protein

##### CYCLE 2

###### Design

To enhance protein expression, the IPTG induction concentration was increased to 0.8–1 mM, based on recommendations from other researchers in our lab who reported successful yields at 1 mM. The concentration of PQQ was also doubled to 6 µM to ensure sufficient cofactor availability, given the unconventional source used. Instead of research-grade PQQ from laboratory suppliers, we opted for a commercially available dietary supplement due to cost constraints. A bottle of 30 capsules (40 mg each) costs approximately US$23, whereas Sigma-Aldrich’s reagent-grade PQQ is priced at US$238 per milligram. Notably, the supplement contains PQQ in its disodium salt form, which is reported to be more stable and water-soluble than the free acid form [14].

###### Build

Cultures were induced with the revised concentrations of IPTG and PQQ, and growth was monitored to ensure the target OD600 of 0.6 was achieved. Sample preparation followed the same protocol as in Cycle 1. To address previously observed low protein recovery, the purification step was modified by increasing the quantity of magnetic beads used for His-tag capture and performing additional wash steps to improve purity and reduce background protein contamination.

###### Test

Purified protein, crude lysate, and resuspended cell pellet samples were analysed via SDS-PAGE using the same protocol as Cycle 1, but with double the loading volume to enhance protein detectability. Despite this adjustment, no visible bands corresponding to our target protein were observed in the purified sample. In contrast, the lysate and cell pellet lanes displayed broad, high-intensity bands, likely due to overloading, which compromised resolution.

###### Learn

Across both cycles, we found that optimising IPTG induction, cell biomass, and purification yield did not result in successful protein expression. Further investigation, including direct communication with ANSA, revealed that the vector used to synthesise our plasmid was a single-copy backbone, which only transitions to medium-copy upon arabinose induction—a detail we had not anticipated. This likely meant our expression relied on just one plasmid copy per cell, explaining the absence of detectable protein on SDS-PAGE.

In future work (beyond our current timeline), we plan to repeat the expression protocol with arabinose induction to activate higher plasmid copy number. Additionally, we aim to reconstruct the ANSA-synthesised insert into a more conventional protein expression backbone using Gibson Assembly. If SDS-PAGE confirms successful expression in purified samples, we will proceed with periplasmic extraction to evaluate the functionality of our signal peptides.

##### MDH Functional Characterisation

##### Cycle 1

###### Design

Following advice from our mentor, Prof. Rob Willows, we adopted a top-down experimental strategy—starting with resting cells, then progressing to crude lysate and finally purified protein. The rationale was that if methanol oxidation could be demonstrated at the whole-cell level, downstream characterisation would be more straightforward. In this cycle, we focused on implementing the resting cell methanol assay. This assay was also appropriate as we did not successfully obtain gel bands for lysate nor purified protein expression.

###### Build

The resting cell assay was designed to evaluate methanol oxidation activity in MDH-expressing cells without cell lysis. Induced cell pellets were resuspended and exposed to methanol at two concentrations: 0.1% and 2.0%. Detailed protocols are available on the [Experiments](https://2025.igem.wiki/macquarie-australia/experiments) page.

###### Test

To detect formaldehyde production as a proxy for methanol oxidation, we used Supelco formaldehyde test strips. Measurements were taken at multiple time points post methanol addition, ranging from immediate exposure to 1 hour. However, no formaldehyde was detected across conditions. Full results are documented on the [Results](https://2025.igem.wiki/macquarie-australia/results) page.

###### Learn

Recent literature highlights the essential role of the *mxaJ* gene in MDH enzyme maturation [15]. Without *mxaJ*, the enzyme remains non-functional even if *mxaF* and *mxaI* are expressed [15]. *mxaJ* encodes a chaperone protein that facilitates PQQ incorporation into the catalytic site of the *mxaF*-encoded alpha subunit, enabling proper assembly with the *mxaI*-encoded beta subunit [15]. Moving forward, we plan to include *mxaJ* in our plasmid design and switch to research-grade PQQ to ensure cofactor purity and functional enzyme assembly.

",
"open": true
},
{
"id": "references",
"title": "References",
"body": "

1. Y. Thepchuay, W. Chairit, N. Saengsane, P. Porrawatkul, and R. Pimsen, “Simple and green colorimetric method for the detection of formaldehyde in vegetable samples,” *Journal of Food Composition and Analysis*, vol. 111, p. 104623, 2022.
2. P. Wang et al., “A pH-responsive production of hydroxyl radical in Fenton process,” *Environmental Science and Ecotechnology*, p. 100566, 2025.
3. G. Özer Bergman, S. Mecacci, V. A. P. Martins dos Santos, and E. Asin-Garcia, “Engineering chimeric signaling proteins for microbial whole-cell biosensors: from design to deployment,” *Trends in Biotechnology*, 2025, doi: 10.1016/j.tibtech.2025.08.002.
4. I. Ganesh, S. Vidhya, G. T. Eom, and S. H. Hong, “Construction of Methanol-Sensing *Escherichia coli* by the Introduction of a *Paracoccus denitrificans* MxaY-Based Chimeric Two-Component System,” *Journal of Microbiology and Biotechnology*, vol. 27, no. 6, pp. 1106–1111, 2017, doi: 10.4014/jmb.1611.11070.
5. V. Selvamani, M. K. Maruthamuthu, K. Arulsamy, G. T. Eom, and S. H. Hong, “Construction of methanol sensing *Escherichia coli* by the introduction of novel chimeric MxcQZ/OmpR two-component system from *Methylobacterium organophilum* XX,” *The Korean Journal of Chemical Engineering*, vol. 34, no. 6, pp. 1734–1739, 2017, doi: 10.1007/s11814-017-0063-8.
6. L. J. Kenney and G. S. Anand, “EnvZ/OmpR Two-Component Signaling: An Archetype System That Can Function Noncanonically,” *Ecosal Plus*, vol. 9, no. 1, 2020, doi: 10.1128/ecosalplus.esp-0001-2019.
7. Y. Fu, J. Li, J. Wang, E. Wang, and X. Fang, “Development of a two component system based biosensor with high sensitivity for the detection of copper ions,” *Communications Biology*, vol. 7, no. 1, Art. no. 1407, 2024, doi: 10.1038/s42003-024-07112-6.
8. T.-K. Le, Y.-J. Lee, G. H. Han, and S.-J. Yeom, “Methanol dehydrogenases as key biocatalysts for synthetic methylotrophy,” *Frontiers in Bioengineering and Biotechnology*, vol. 9, p. 787791, 2021.
9. K. Mirzadeh et al., “Increased production of periplasmic proteins in *Escherichia coli* by directed evolution of the translation initiation region,” *Microbial Cell Factories*, vol. 19, no. 1, p. 85, 2020.
10. C. J. Morris, Y. M. Kim, K. E. Perkins, and M. E. Lidstrom, “Identification and nucleotide sequences of mxaA, mxaC, mxaK, mxaL, and mxaD genes from *Methylobacterium extorquens* AM1,” *Journal of Bacteriology*, vol. 177, no. 23, pp. 6825–6831, 1995.
11. C. Anthony and P. Williams, “The structure and mechanism of methanol dehydrogenase,” *Biochimica et Biophysica Acta (BBA) - Proteins and Proteomics*, vol. 1647, no. 1–2, pp. 18–23, 2003.
12. T. Karaseva et al., “Isolation and characterization of homologically expressed methanol dehydrogenase from *Methylorubrum extorquens* AM1 for the development of bioelectrocatalytical systems,” *International Journal of Molecular Sciences*, vol. 23, no. 18, p. 10337, 2022.
13. F. Munder, M. Voutsinos, K. Hantke, H. Venugopal, and R. Grinter, “High-affinity PQQ import is widespread in Gram-negative bacteria,” *Science Advances*, vol. 11, no. 22, p. eadr2753, 2025.
14. “Is Pyrroloquinoline Quinone the Same as PQQ Disodium Salt?,” Santa Biotech. [Online]. Available: https://santabiotech.com/ (accessed Sep. 26, 2025).
15. H. Zhou et al., “Deciphering the assembly process of PQQ dependent methanol dehydrogenase,” *Nature Communications*, vol. 16, no. 1, p. 6672, 2025.

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{% block hero\_title %}Experiments{% endblock %}
{% set sections = [
{"id":"miniprep", "title":"Plasmid Miniprep", "body":"

**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](https://www.neb.com/en-au/products/t1110-monarch-spin-plasmid-miniprep-kit)

", "open":true},
{"id":"transformation", "title":"Transformation", "body":"

**High Efficiency Transformation Protocols**

* [Protocol (C2987H/C2987I)](https://www.neb.com/en-au/protocols/0001/01/01/high-efficiency-transformation-protocol-c2987?pdf=true)
* [Protocol (C3010)](https://www.neb.com/en-au/protocols/0001/01/01/high-efficiency-transformation-protocol-c3010?pdf=true)

**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.

", "open":true},
{"id":"formaldehyde", "title":"Supelco Formaldehyde Test Strips", "body":"

**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](https://www.sigmaaldrich.com/AU/en/product/mm/110036?srsltid=AfmBOoqK6L9uX2KshRrh-nN1_v9e46P0sZbIb7JMtGusl9XQ_lpNZwGH)

", "open":true},
{"id":"pcr", "title":"PCR", "body":"

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

**PCR Setup:**
For 25× cycles

| Step | Temperature | Time |
| --- | --- | --- |
| Denaturation | 95 °C | 15 s |
| Annealing | Adjusted based on primer | Adjusted based on primer |
| Extension | 72 °C | Adjusted based on primer |
| Final Hold | 4 °C | Hold |

*Notes:*

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

", "open":true},
{"id":"gel-electrophoresis", "title":"Gel Electrophoresis", "body":"

**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.

", "open":true},
{"id":"maximum-tolerance", "title":"Maximum Tolerance", "body":"

**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.

", "open":true},
{"id":"competent-cells", "title":"Making Cells Chemically Competent", "body":"

**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.

", "open":true},
{"id":"ecoli-methanol", "title":"Resting Cell Methanol Assay and Protein Extraction", "body":"

**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.

", "open":true},
{"id":"dynabeads", "title":"Dynabeads His-tag Isolation and Purification", "body":"

**Protocol Reference:**
[ThermoFisher Dynabeads™ His-tag Isolation & Pull-down Protocol](https://documents.thermofisher.com/TFS-Assets/LSG/manuals/DynabeadsHisTagIsolationPulldown_man.pdf)

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.

", "open":true},
{"id":"sds-page", "title":"SDS-PAGE", "body":"

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](https://coleman-lab.org/wp-content/uploads/2022/07/Protein-analysis-by-SDS-PAGE.pdf)

", "open":true}
] %}
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{"href": url\_for('pages', page='engineering'), "title": "Engineering", "img": "svg/suit.svg"},
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{"href": url\_for('pages', page='members'), "title": "Members", "img": "svg/team.svg"}
] %}

{% extends "base.html" %}
{% block hero\_title %}Results{% endblock %}
{% set sections = [
{
"id":"pcr-agarose-gel-electrophoresis",
"title":"PCR and Agarose Gel Electrophoresis",
"open": true,
"body": "

### 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.

![Colony PCR confirmation (AmilCP)](https://static.igem.wiki/teams/5787/results/1.webp)

#### 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.

![Agarose gel electrophoresis of colony PCR (mxa)](https://static.igem.wiki/teams/5787/results/2.webp)

#### 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).

![28/08/25 Agarose gel electrophoresis of plasmid constructs](https://static.igem.wiki/teams/5787/results/3.webp)

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.

"
},
{
"id":"choosing-a-reporter-protein",
"title":"Choosing a Reporter Protein​",
"open": true,
"body": "

### 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.

![Choosing a reporter protein – amilCP vs RFP](https://static.igem.wiki/teams/5787/results/5.webp)

![Choosing a reporter protein – amilCP vs RFP (additional)](https://static.igem.wiki/teams/5787/results/6.webp)
"
},
{
"id":"testing-commercial-formaldehyde-test-strips",
"title":"Testing Commercial Formaldehyde Test Strips​",
"open": true,
"body": "

### 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.

![Supelco formaldehyde test strips 5–200 mg/L](https://static.igem.wiki/teams/5787/results/7.webp)
"
},
{
"id":"methanol-dehydrogenase-enzyme-design",
"title":"Methanol Dehydrogenase Enzyme Design",
"open": true,
"body": "

### 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

![Adding Formaldehyde t=0 4/8/25](https://static.igem.wiki/teams/5787/results/8.webp)

**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

![Adding Formaldehyde t=1 week](https://static.igem.wiki/teams/5787/results/9.webp)

**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](https://static.igem.wiki/teams/5787/results/10.avif)

**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

![Addition of formaldehyde t=0 25/08/25](https://static.igem.wiki/teams/5787/results/11.avif)

**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

![Addition of formaldehyde t=12 days](https://static.igem.wiki/teams/5787/results/12.avif)

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

![Resting cell assay 1](https://static.igem.wiki/teams/5787/results/13.webp)

![Resting cell assay 2](https://static.igem.wiki/teams/5787/results/14.webp)

![Resting cell assay 3](https://static.igem.wiki/teams/5787/results/15.webp)

![Resting cell assay 4](https://static.igem.wiki/teams/5787/results/16.webp)

![Resting cell assay 5](https://static.igem.wiki/teams/5787/results/17.webp)

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

![SDS-PAGE Analysis Showing No Detectable MDH Protein Bands](https://static.igem.wiki/teams/5787/results/18.webp)

**Figure 18:** SDS-PAGE Analysis Showing No Detectable MDH Protein Bands

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

![SDS-PAGE Shows Unexpected High Molecular Weight Band Instead of Predicted MDH Subunits](https://static.igem.wiki/teams/5787/results/19.webp)

**Figure 19:** SDS-PAGE Shows Unexpected High Molecular Weight Band Instead of Predicted MDH Subunits

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

![SDS-PAGE analysis of MxaF and MxaI protein expression following IPTG induction](https://static.igem.wiki/teams/5787/results/20.webp)

**Figure 20:** SDS-PAGE analysis of MxaF and MxaI protein expression following IPTG induction

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.

"
},
{"id":"t1", "title":"Whole Cell Model", "body":"

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

![Ethanol MIC results](https://static.igem.wiki/teams/5787/results/21.webp)

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

![Methanol MIC results](https://static.igem.wiki/teams/5787/results/22.webp)

*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.

![Copper induction Chimera 0.5–2 mM](https://static.igem.wiki/teams/5787/results/23.webp)

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

![Chimera non-induced vs 2 mM](https://static.igem.wiki/teams/5787/results/24.webp)

![Chimera additional replicate](https://static.igem.wiki/teams/5787/results/25.webp)

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

![Non-transformed 2 mM comparison 1](https://static.igem.wiki/teams/5787/results/26.webp)

![Non-transformed 2 mM comparison 2](https://static.igem.wiki/teams/5787/results/27.webp)

![Copper induction replicates](https://static.igem.wiki/teams/5787/results/28.webp)

![Copper induction replicate 2](https://static.igem.wiki/teams/5787/results/29.webp)

![Copper induction replicate 3](https://static.igem.wiki/teams/5787/results/30.webp)

![Copper induction replicate 4](https://static.igem.wiki/teams/5787/results/31.webp)

![Copper induction replicate 5](https://static.igem.wiki/teams/5787/results/32.webp)

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.

![AmilCP copper induction 0.5–2 mM](https://static.igem.wiki/teams/5787/results/33.webp)

![AmilCP copper induction additional](https://static.igem.wiki/teams/5787/results/34.webp)

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

![No copper induction comparison](https://static.igem.wiki/teams/5787/results/35.webp)

![Copper induction 2 mM comparison](https://static.igem.wiki/teams/5787/results/36.webp)

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

![Copper induction replicate 6](https://static.igem.wiki/teams/5787/results/37.webp)

![Copper induction replicate 7](https://static.igem.wiki/teams/5787/results/38.webp)

![Copper induction replicate 8](https://static.igem.wiki/teams/5787/results/39.webp)

![Copper induction replicate 9](https://static.igem.wiki/teams/5787/results/40.webp)

![Copper induction replicate 10](https://static.igem.wiki/teams/5787/results/41.webp)

![Copper induction replicate 11](https://static.igem.wiki/teams/5787/results/42.webp)

![Copper induction replicate 12](https://static.igem.wiki/teams/5787/results/43.webp)

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%.

![Methanol induction Chimera 0.5–10%](https://static.igem.wiki/teams/5787/results/45.webp)

![Non transformed cells 0–10%](https://static.igem.wiki/teams/5787/results/46.webp)

![Methanol induction replicates](https://static.igem.wiki/teams/5787/results/47.webp)

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.

", "open":true}
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{"href": url\_for('pages', page='notebook'), "title": "Notebook", "img": "svg/notebook.svg"},
{"href": url\_for('pages', page='members'), "title": "Members", "img": "svg/team.svg"}
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