During our time in the lab, we designed and conducted the experiments that would offer the most valuable insights at the outset of our project. Following thorough discussions, we pinpointed specific aspects of our project's design to concentrate on, aiming to provide a practical and realistic demonstration. For better comprehension, we encourage you to first read the ‘Cloning method’ section on our Engineering page.

One of the goals we set for our experiments was to shed light on a pivotal component of the malonyl-CoA-based negative feedback system we have implemented in our design, the PFR1 promoter. (for more details, refer to our Design page ). To comprehensively characterize this component, we devised two experiments:

1. Testing a level 1 construct featuring the PFR1 promoter controlling the syfp2 reporter (Fig. 1) to assess its relative strength.


2. Testing a level 2 construct that includes the aforementioned construct but also incorporates a transcriptional unit of the fapR , regulated by the AraC/PBAD system (illustrated in (Fig. 2).

For more details, refer to our New Basic Part page.

Fig. 1 .Schematic representation of the level 1 (alpha) construct for the characterization of the PFR1 (1st experiment).

Fig. 2.Schematic representation of the level 2 (omega) construct for the characterization of the PFR1 (2nd experiment).

Process

We obtained and domesticated the sequences of these three components using the [Domestication tool]. This process involved eliminating internal restriction sites incompatible with the GoldenBraid 2.0 grammar while also incorporating the appropriate 3' and 5' overhangs. To confirm the accuracy of this domestication, we imported the modified parts into Snapgene to simulate the restriction-insertion reaction and ensure that the correct overhangs were added to each part. After verifying all the parts, we were ready to begin the cloning process. The initial step involved the insertion of the fragments into a pUPD2 vector to generate level 0 constructs, following our standard digestion-ligation cloning protocol. We maintained the optimal 3:1 insert:vector molar ratio, as suggested by our protocol. Continuing, 10 ul of the digestion-ligation reaction was used to transform DH5α chemically competent cells. The following day, single colonies were picked (through blue-white screening), inoculated in 5 ml of LB medium, with the appropriate antibiotic, and finally the cultures were incubated overnight at 37°C and 210 rpm. These cultures were used to isolate the plasmids, which were subsequently subjected to restriction-digestion reactions to confirm the successful insertion of the fragment (Fig. 3, 4 and 5).

Fig. 3 . Diagnostic digestion of pUPD2_pFR1 with EcoRI and EcoRV, expected bands (bp): 1305 and 896. Lane 2: pUPD2 (no insert)

Fig. 4. Diagnostic digestion of pUPD2_araC/pBAD with EcoRI and EcoRV, expected bands (bp): 2107, 896 and 466. Lane 2: pUPD2 (no insert).

Fig. 5. Diagnostic digestion of pUPD2_fapR with EcoRV, expected bands (bp): 1477 and 1196. Lane 2: pUPD2 (no insert).

The resulting level 0 constructs, along with selected parts from the distribution kit, such as syfp2 (BBa_K864100), B0015 double terminator (BBa_J428092), and B0030 RBS (BBa_J428032), were combined to build 3 level 1 constructs : 1. a construct carrying the PFR1 with the syfp2 downstream, 2. the PJ23118 Anderson promoter, again regulating syfp2, and 3. the fapR gene under the control of AraC/PBAD .
After a few failed attempts, we tried to optimize our digestion-ligation protocol, adjusting the insert:vector ratio depending on a few parameters. Specifically, the length of the final construct was taken into account. In cases where the final construct surpassed 2.0 kb, higher quantities of both vector and insert were used (e.g. 40-120 fmol instead of 20-80 fmol respectively ). Secondly, the length of the part itself was also considered, where the ratio of smaller parts was increased.
With those adjustments to our process, we managed to successfully build the level 1 construct we required for the 1st part of this experiment (Fig. 1), the PFR1 strength evaluation.

PFR1 strength evaluation

Having the correct level 1 construct, we were ready to start the 1st part of this experiment. The verified plasmid was used to transform E. coli BL21 (DE3) chemically competent cells. Single colonies were picked and inoculated in 5 ml LB medium with 5 ul kanamycin and cultured O/N at 37οC and 210 rpm. The following day, the cultures were taken to the cold room and intermediate dilutions 1:10 were prepared with the addition of 500 ul of culture in 4,5 ml of LB broth + kanamycin. Then intermediate dilutions were measured for absorbance at 600 nm and the values were inserted in the culture dilution calculator, [https://static.igem.org/websites/technology/interlab/2023/protocolsculture-dilution-calculator-template.xlsx] provided by the InterLab committee. Using those measurements, we prepared final dilutions for all three cultures (positive, negative and test construct). Fluorescence measurements at wavelengths of 511 nm (excitation) and 529 nm (emission) were taken after 6 hours of incubation and the results we got are shown below (Fig. 6). For the interpretation of the results see New Basic Part page.

Fig. 6. Normalized fluorescence intensity of the pFR1-syfp-rrnB T1/T7TE construct, compared to a positive control (pJ23118-syfp-rrnB T1/T7TE) and a negative control (non-transformed E. coli cells) after 6h incubation.

Evaluation of FapR - mediated PFR1 repression

After testing the level 1 construct, we were ready to use it to build the level 2 construct, in order to perform the second part of the experiment, the evaluation of the repression FapR poses on the promoter. To achieve this, we had to combine the two level 1 constructs containing the PFR1-driven reporter module and the PBAD-driven FapR repressor module via a new digestion-ligation reaction. The standard protocol was followed with the appropriate optimizations. Eventually, we were able to build the construct (Fig. 2).
The verified plasmid was used to transform E. coli BL21 (DE3) chemically competent cells. The next day, single colonies were picked and inoculated in 5 ml LB + 5 ul spectinomycin and cultured O/N at 37οC and 210 rpm. The O/N cultures were retrieved the next day, intermediate 1:10 dilutions were prepared and their OD600 was measured. From those intermediate dilutions we prepared final dilutions for the level 2 construct, the positive and negative controls. The appropriate amounts of arabinose were added in each sample (+ the blanks) so in the end we were left with 5 repeats of each culture, each containing increasing concentrations of arabinose. The results we got are shown below (Fig. 7). For the interpretation of the results see New Basic Part page.

Fig. 7. Normalized fluorescence intensity for the level 2 construct (pDGB3ω1_pFR1-syfp-rrnB T1/T7TE + araC/pBAD-fapR-rrnB T1/T7TE) in different concentrations of L-arabinose, after 3h and 6h incubation.

The objective of this experiment was to create six constructs: three level 1 constructs aimed at assessing the relative strength of the PfadBA , PLR and PAR promoters, and three level 2 constructs that included the aforementioned level 1 constructs along with fadR to evaluate the level of promoter inhibition exerted by the regulator. This would help us get closer to selecting the optimal promoter, one that would offer the highest sensitivity and efficiency in regulation. However, we were only able to construct and test the level 1 constructs and determine the strengths of the PfadBA , PLR , and PAR promoters (Fig. 8) (see Engineering for more details on the results).

Fig. 8. Schematic representation of the standard construct for the characterization of PLR and PAR .

Process

The sequences for PfadBA , PLR and PAR were acquired and domesticated using the domestication tool, to be compatible with the GoldenBraid standards. The domesticated part sequences were then imported into Snapgene to simulate the restriction-insertion reaction procedure and verify the 3’ and 5’ overhang compatibility. Once all constructs were checked, all the fragments were ordered by IDT. The DNA fragments were then resuspended and the first digestion-reaction was performed for the insertion of the parts into a pUPD2 domestication vector. For the level 0 clonings, the insert:vector ratio was kept at a 3:1. Following the standard protocol, 5 ul of the digestion-ligation reactions were used to transform E. coli DH5α chemically competent cells. Plates were streaked and 10ul of IPTG and 40 ul of X-gal were added for the blue-white screening. The following day, white colonies were picked, inoculated in 5 ml LB broth + 5 ul chloramphenicol, and finally the cultures were incubated O/N at 37οC and 210 rpm. From the O/N cultures, we performed plasmid isolation and next restriction-digestion to confirm that we had indeed isolated the right plasmids. After a few rounds of this procedure, we had successfully acquired all level 0 constructs (Fig. 9 and 10) and we continued with the level 1 clonings.

Fig. 9. Diagnostic digestion of pUPD2_pfadBA with EcoRI and EcoRV, expected bands (bp): 1309 and 896. Lane 2: pUPD2 (no insert).

Fig. 10. Diagnostic digestion of (1) pUPD2_pLR and (2) pUPD2_pAR with EcoRI and EcoRV, expected bands (bp): (1) 1300 and 896, and (2) 1305 and 896. Lane 3: pUPD2 (no insert).

The same pattern as with the previous experiment was observed where the first attempts for the construction of the level 1 parts failed. Thus, the same optimization took place, adjusting the quantities of the smaller parts, and the constructs were eventually successful (Fig. 11 and 12). Once we had successfully built our constructs, we were ready to test the devices

Fig. 11. Diagnostic digestion of pDGB3α1_pfadBA-syfp-t1te with BsaHI, expected bands (bp): 2327, 2026, 1628 and 1291. Lane 2: pDGB3α1 (no insert).

Fig. 12. Diagnostic digestion of (1) pDGB3α1_pLR-syfp-t1te and (2) pDGB3α1_pAR-syfp-t1te with BsaHI, expected bands (bp): (1) 2327, 2017, 1628, 1291 and 58, and (2) 2327, 2022, 1628, 1291 and 58. Lane 3: pDGB3α1 (no insert).

To begin, once we had all the devices in our hands, we transformed E. coli BL21 (DE3) chemically competent cells. The next day, single colonies were picked for the preparation of liquid cultures, which were incubated O/N at 37οC and 210 rpm. The cultures were retrieved for the preparation of 3 final dilutions for each construct, including positive and negative controls. The constructs were tested in three different conditions: with the addition of non-dissolved oleic acid, addition of oleic acid dissolved in DMSO (final concentration 10 mM in each case) and without oleic acid. The plate was prepared with four repeats for each condition and placed into the plate reader. Measurements were taken at 0h and 6h timepoints. The final results are depicted in Fig. 13. For the interpretation of the results see Engineering page.

Fig. 13. Normalized fluorescence intensity for pfadBA-syfp-rrnB T1/T7TE, pLR-syfp-rrnB T1/T7TE and pAR-syfp-rrnB T1/T7TE constructs on three different conditions (No oleic acid addition, addition of non-dissolved oleic acid, addition of oleic acid dissolved in DMSO) after 6h incubation. E. coli BL21 (DE3) cells with pJ23118-syfp-rrnB T1/T7TE construct and non-transformed cells were used as (+) and (-) control respectively.

The secretion of a functional laccase from E. coli is essential for our system since it would mean that the poly-phenolic content of OMW would be degraded to the maximum. To establish that, we conducted two experiments: 1. an assay to determine the location of the enzyme inside or outside the cell and 2. an assay for the calculation of the enzyme activity, to prove functionality of the secreted laccase. Below, we describe the procedures for each of the two aspects and our results.

Process

To test the secretion of the enzyme, we decided that the best way for us would be to fuse the genetic signal peptide-laccase entity with the sequence of SYFP2 fluorescent protein, the reporter we had chosen for the rest of our experiments. However, to evaluate the enzyme activity, we deemed that the enzyme should be tested without being fused with the fluorescent protein, since that might interfere with its functionality. So, for each peptide, including the native, 2 constructs were built: one containing the signal peptide-laccase sequence in fusion with the syfp2 (Fig. 14) and one where the coding sequence of the mature laccase carries upstream only the appropriate signal peptide (Fig. 15), making a total of 8 level 1 constructs.

Fig. 14. Schematic representation of the standard level 1 (alpha) construct of the laccase fused with the syfp and carrying one of the signal peptides (Na.SP, NSP4, PelB and MalE).

Fig. 15. Schematic representation of the standard level 1 (alpha) construct of the laccase carrying one of the signal peptides (Na.SP, NSP4, PelB and MalE).

  The sequences for the signal peptides and the laccase were domesticated to the Golden Braid standards. However, because of the Golden Braid method's characteristics, specific 3' and 5' overhangs are limited to use in a single position within the transcription unit. So, to create constructs where the SP-laccase genetic entity is fused with the syfp2 and constructs where it is not, we needed to create two distinct sets of overhangs for the laccase. These sets were designated for the B4 and the B4+B5 position respectively within the transcriptional unit. For the addition of the overhangs, we designed primer pairs specifically for each of the 2 positions we wanted to assemble laccase in. The primers we designed for this procedure have been added to the iGEM registry and can be found on our Parts page.
  Once the compatibility of the parts was confirmed, the parts were ordered to be synthesized by IDT. The DNA fragments were resuspended and we performed 2 different PCR reactions, one for each position of the laccase in our transcriptional units (B4 and B4+B5). The samples were then prepared for gel electrophoresis, and once we confirmed that the main product was indeed in the appropriate height, we followed the standard protocol for Gel Extraction. Then the 2 laccase parts, along with the rest of the domesticated sequences were inserted into pUPD2 part domestication following standard protocol. A diagnostic restriction digestion reaction and gel electrophoresis followed for each of the level 0 constructs to verify their correct insertion (Fig. 16-19).
Finally, the resulting level 0 constructs were combined with the ones of AraC/PBAD and B0015 double terminator, for the creation of the complete level 1 (alpha) constructs (Fig. 15 and 16). The inserts were once again confirmed through restriction-digestion reactions and electrophoresis (Fig. 20 and 21).

Fig. 16.Diagnostic digestion of (1) pUPD2_NSP4, (2) pUPD2_PelB and (3) pUPD2_Na.SP with EcoRV and NotI, expected bands (bp): (1) 1143 903 and 122, (2) 1143, 903 and 128 and (3) 1143, 903 and 125. Lane 4: pUPD2 (no insert).

Fig. 17. Diagnostic digestion of pUPD2_MalE with EcoRI and EcoRV, expected bands (bp): 1290 and 896. Lane 2: pUPD2 (no insert).

Fig. 18. Diagnostic digestion of pUPD2_laccase(B4) with EcoRV and NotI, expected bands (bp): 1562 1143 and 903. Lane 2: pUPD2 (no insert).

Fig. 19. Diagnostic digestion of pUPD2_laccase(B4+B5) with EcoRV and NotI, expected bands (bp): 1563, 1143 and 903. Lane 2: pUPD2 (no insert).

Fig. 20.Diagnostic digestion of (1) pDGB3α1_pBAD-Na.SP-laccase-syfp-rrnB T1/T7TE (2) pDGB3α1_pBAD-NSP4-laccase-syfp-rrnB T1/T7TE, (3) pDGB3α1_pBAD-PelB-laccase-syfp-rrnB T1/T7TE and (3) pDGB3α1_pBAD-MalE-laccase-syfp-rrnB T1/T7TE with BsaHI, expected bands (bp): (1) 2327, 2049, 1628, 1361, 1291, 610, 423, 185, 163, 81 and 58, (2) 2327, 2049, 1628, 1361, 1291, 625, 423, 185, 163, 81 and 58, (3) 2327, 2049, 1628, 1361, 1291, 613, 423, 185, 163, 81 and 58 and (4) 2327, 2049, 1628, 1361, 1291, 625, 423, 185, 163, 81 and 58. Lane 5: pDGB3α1 (no insert).

Fig. 21. Diagnostic digestion of (1) pDGB3α1_pBAD-Na.SP-laccase-rrnB T1/T7TE, (2) pDGB3α1_pBAD-NSP4-laccase-rrnB T1/T7TE, (3) pDGB3α1_pBAD-PelB-laccase-rrnB T1/T7TE and (4) pDGB3α1_pBAD-MalE-laccase-rrnB T1/T7TE with EcoRV and BsaHI, expected bands (bp): (1) 2327, 1628, 1291, 1290, 807, 610, 554, 423, 185, 163, 81 and 58, (2) 2327, 1628, 1291, 1290, 807, 607, 554, 423, 185, 163, 81 and 58 (3) 2327, 1628, 1291, 1290, 807, 613, 554, 423, 185, 163, 81 and 58 and (4) 2327, 1628, 1291, 1290, 807, 625, 554, 423, 185, 163, 81 and 58. Lane 5: pDGB3α1 (no insert).

Having all 8 constructs ready, we were able to begin with the first experiment, the secretion evaluation assay.

Secretion evaluation

For the secretion evaluation, we decided to measure both the absorbance and fluorescence intensity of the supernatant fraction and pellet fraction after centrifugation. Single colonies of the 4 constructs were picked, inoculated in 5 ml of LB medium with the appropriate antibiotic and cultured O/N at 37oC and 210 rpm. The next day, the OD600 of the O/N cultures was measured and used to prepare dilutions with the same starting value. Arabinose was added to a final concentration of 100mM for the induction of the devices and the samples were placed for incubation at 30oC and 160 rpm. After 6 hours the cultures were retrieved and 2 ml of each was transferred in 2 ml eppendorf tubes and centrifuged at 11,000 x g for 3 minutes. The supernatant was collected and loaded onto a black plate with a transparent bottom. The cell pellets were resuspended in fresh LB broth with the appropriate antibiotic and then also loaded onto the plate. Measurements of the absorbance were taken at 600 nm and for the fluorescence at 511 nm for excitation and 529 nm for emission. Two negative controls were used, E.coli BL21 (DE3) cells with a construct containing the syfp2 under the control of the J23118 Anderson promoter and non-transformed cells. For the interpretation of our results we got (Fig. 22) please see our Engineering page.

Fig. 22. Normalized fluorescence intensity of the supernatant and pellet fraction for pBAD-Na.SP-laccase-syfp-rrnB T1/T7TE, pBAD-NSP4-laccase-syfp-rrnB T1/T7TE, pBAD-PelB-laccase-syfp-rrnB T1/T7TE and pBAD-MalE-laccase-syfp-rrnB T1/T7TE constructs after 6h incubation with 1,5% arabinose, (-) control 1: pJ23118-syfp-rrnB T1/T7TE, (-) control 2: non-transformed cells.

Laccase activity

  For this experiment, we followed the ABTS assay designed to assess laccase activity in the culture of the white-rot fungus Trametes Versicolor. The protocol was provided by Dr. Panagiotis Karas, a Post-Doctoral researcher in the Plant Biotechnology lab of the Department of Biochemistry and Biotechnology. The basis of this assay involves the oxidation of the non phenolic dye ABTS (2,2’-azino-bis(3-ethylbenzthiazoline-6-sulfonic acid) by laccase to the radical cation ABTS*+, a soluble chromogen that is green in color and can be determined spectrophotometrically at 425 nm. After discussing the protocol with Dr. Karas, he suggested that we should optimize it by adjusting the ratio of bacterial culture to the ABTS reagent. This adjustment is necessary since E. coli is unlikely to produce the same quantity of the enzyme as fungi naturally do. Having all the needed devices transformed into E. coli BL21 (DE3) cells, liquid cultures were prepared and incubated O/N at 37oC and 210 rpm. The next day, these cultures were retrieved and used in order to prepare the final cultures with the same starting OD600 values. Addition of L-arabinose to a final concentration of 100mM was followed and the cultures were incubated at 30oC and 160 rpm for 6h. So, after 6h incubation we began the procedure by adding 1,2 ml of sodium tartrate 0.1 M (pH of 4,5 adjusted with NaOH addition) in a cuvette. Next, 0,6 ml of the bacterial culture was added, the cuvette was placed in the spectrometer and the absorbance was set to zero. To initiate the reaction, 0,4 ml of ABTS solution 3 mM were added in the cuvette and pipetted three to four times to mix. The instrument was set back to zero again and the timer started. Absorbance measurements at 425 nm were taken every 20 seconds for 2 minutes 1. The above procedure was repeated 3 times to get as reliable results as possible. The results for each construct are summarized in Fig. 23. Non-transformed E. coli BL21 (DE3) cells were used as negative control.

Time (sec)	20			40	60	80	100	120
Na.SP	0,029			0,039	0,043	0,046	0,048	0,049
	0,026			0,033	0,037	0,041	0,044	0,047
	0,019			0,021	0,022	0,026	0,028	0,030
NSP4	0,067			0,078	0,082	0,084	0,085	0,086
	0,016			0,067	0,076	0,081	0,085	0,087
	0,02			0,025	0,075	0,083	0,088	0,09
PelB	0,043			0,049	0,052	0,056	0,058	0,06
	0,03			0,042	0,046	0,048	0,049	0,05
	0,038			0,047	0,051	0,054	0,056	0,056
MalE	0,012			0,045	0,051	0,056	0,059	0,062
	0,003			0,034	0,04	0,044	0,05	0,051
	0,007			0,037	0,043	0,047	0,051	0,054
(-) control	0,026			0,034	0,038	0,043	0,047	0,05
	0,003			0,013	0,014	0,025	0,027	0,028
	0,003			0,008	0,021	0,03	0,034	0,039

Fig. 23.Fig. 23. Absorbance measurements at 425 nm of the 4 constructs (Fig. 15) designed for laccase activity calculation. Measurements were taken every 20 sec after the addition of the ABTS reagent for 2 minutes. Non-transformed E. coli BL21 (DE3) cells were used as (-) control.

Once the measurements were collected, we calculated the enzyme activity based on the absorbance changes using the formula:
      U / L = (ΔΑ x Vt)/(Δt x ε x Vs)
U enzyme activity umol min^-1 L^-1, ΔΑ (final absorbance - initial absorbance) the maximum absorbance difference measured at 2 min, Vt total reaction volume (ml), Δt elapsed time between two measurements (min), ε molar extinction coefficient (L mol^-1 cm^-1) = 36 L mol^-1 cm^-1, Vs sample volume (ml). The enzyme activity values for each sample are depicted in Fig. 24.

Fig. 24. Laccase activity of the four different constructs estimated with ABTS assay, after 6h incubation with 1,5% arabinose. Non-transformed E. coli BL21 (DE3) cells were used as (-) control.

Comparing the above results with those of the laccase secretion, we notice that although there is no perfect measurements matching for the PelB and MalE signal peptides, the NSP4 signal peptide exhibited the best results in both aspects. Therefore, the results of both experiments indicate the selection of NSP4 as the ideal one for our system.

At the outset of our experiments, we recognized firsthand the critical importance of accurately and comprehensively documenting a scientist's daily work. This documentation is crucial for interpreting results and drawing credible, logical conclusions. Consequently, we promptly made the decision to maintain a digital lab notebook, which encompassed nearly allall of the experiments we conducted throughout the iGEM competition. Our goal was to present our work in as realistic a manner as possible, with the aim of providing future teams with a valuable introduction and insight into the role of a wet lab member in an iGEM team.

Here you can find all the protocols our team used during our time in the lab, in great detail.