Results

Short Summary

Detailed Results

One goal of our project is to compare different laccases. We have selected six laccases that looked promising for our planned application (Table 1). The genes were optimized for expression by the yeast Pichia pastoris. The genes were synthesized by the company IDT. Using PCR the TEV site and His-tag were removed and the corresponding fusion site was added to the 3’ end of the laccase genes. This left us with 12 different laccase genes.

Registry Number	Laccase	Origin Species	GenBank Entry Number
BBa_K5329009	CVL3	T. versicolor	D13372
BBa_K5329010	LAC1	T. hirsuta	EU492907
BBa_K5329011	LCC1	C. trogii	KU055621
BBa_K5329012	LCC1	T. versicolor	AY693776
BBa_K5329013	LCC2	T. versicolor	Y18012

Using golden gate assembly, we successfully cloned the laccases into different plasmid backbones (BB1). The BB1 assemblies were inserted into Escherichia coli using heat-shock transformation. We confirmed the insertion of the genes into BB1 by colony PCR and Sanger sequencing.

The expression cassettes consisting of the constitutive pGAP promoter, the different laccase genes, and the rps25aTT terminator were assembled using Golden Gate cloning into the BB3 (100_BB3rN). The assembled BB3s were again transformed into E.coli (NEB10beta) using heat shock transformation. The presence of the laccase genes in the plasmids was confirmed through restriction digest.

After confirming that the assembly was successful the BB3s were linearized and transformed into Pichia pastoris using electroporation. The transformation was confirmed by colony PCR.

The first attempts to express the laccases were made by inoculating 10 mL of YPD and growing the cells at 30°C for two days. To check for expression an ABTS Assay of the supernatant was made. No laccase activity could be detected in the media when performing an ABTS Assay. A Western Blot did show that no laccases were secreted into the media.

To improve the expression of laccases we switched to YNB media. We made an overnight preculture and inoculated the YNB Media at an OD₆₀₀ 8. After incubating for 48 hours laccase activity could be detected using the ABTS assay after extended incubation in two cultures. The culture expressing the Trametes hirsuta laccase and the culture expression the Trametes versicolor LCC2 laccase. Both of the expressed laccases do not contain the HIS-tag.

SDS-PAGE and staining (Coomassie and Silver) were performed on concentrated supernatants, to confirm that the laccases were indeed present in the supernatant. We expected a signal at the size of about 55 kDa but no band can be observed.

Because we could detect laccase activity with the ABTS Assay we decided to test if the culture supernatant can degrade Diclofenac. For this experiment, we mixed 1mL of culture supernatant with 1mL of Diclofenac stock solution (100µg/mL). The same reaction mix was also performed with the supernatant of a yeast culture which did not express the laccases (empty control – EC).

After incubation of the mix over the weekend the concentration of the pharmaceuticals was analyzed using HPLC. The results of the HPLC showed a 51,9 % reduction of diclofenac in the samples where laccases are produced compared to the sample containing the supernatant of yeasts not producing laccases (EC).

Table 2: Results of the HPLC analysis

Sample	concentration of diclofenac [µg/mL]	Reduction relative to empty control [%]
Empty control	45,21	-
LCC2 T. versicolor 10C	27,36	–39,5
Laccase T. hirsuta 4D	21,73	–51,9

An attempt was made to further improve the protein expression by switching to an inducible AOX1 promoter. The golden gate assembly was made, and the insertion of the expression cassette into the BB3 was confirmed by restriction digest but the transformation into P. pastoris was unsuccessful. Due to time constraints, no second attempt was made.

In summary, the team successfully performed cloning steps and managed, through optimization of cultivation conditions to successfully produce two different laccases. Although we could not detect our laccases with the SDS-PAGE and silver staining we were able to detect the enzyme activity and prove the ability of the laccases to degrade diclofenac.

Short Summary

Detailed Results

Short Summary

Degradation of Diclofenac with immobilized laccase

The HPLC results of our first immobilization approach reflect the degradation of Diclofenac and Ibuprofen after being treated with agar-agar immobilized laccase. The initial concentrations of the drugs were set at 100 µg/mL for Diclofenac and 500 µg/mL for Ibuprofen. The results show a varying degree of degradation across different samples, indicating the enzymatic activity of laccase.

This diagram illustrates the degradation process of Diclofenac using two different concentrations of immobilized laccase: 0.15 mg/mL and 0.60 mg/mL. The results indicate that the degradation efficiency is markedly higher with 0.60 mg/mL of laccase, demonstrating that increased concentrations of immobilized laccase lead to a more effective degradation of Diclofenac. Consequently, this higher concentration results in a faster and greater reduction in Diclofenac levels compared to the 0.15 mg/mL concentration.

The degradation process is more effective with the higher laccase concentration, and after 2 hours, both concentrations reach a plateau, indicating that no further degradation occurs beyond this point. Additionally, since no further degradation was observed after 24 hours, we excluded day 4 from the plot.

Additionally, during the reaction, the solution changes from transparent to yellow after 2 hours, showing that the immobilized laccase is active.

Degradation of Ibuprofen with Immobilized Laccase

The diagram below shows the degradation process of Ibuprofen using two different concentrations of immobilized laccase: 0.15 mg/mL and 0.60 mg/mL. Like Diclofenac, the degradation efficiency is higher with increasing concentration. However, the degradation of Ibuprofen was less effective than Diclofenac. Same as for Diclofenac after 2 hours of incubation no further degradation occurs.

Summary

Overall, the laccase enzyme showed higher degradation for Diclofenac compared to Ibuprofen, with degradation percentages for Diclofenac reaching as high as 57.3%, whereas the highest reduction for Ibuprofen was 25.9%. The immobilization of laccase in agar-agar was effective in breaking down both pharmaceuticals, but it appears to work more efficiently on Diclofenac. This experiment demonstrates the potential of laccase to mitigate pharmaceutical pollution, with varying effectiveness depending on the compound.

To further compare degradation results a new batch of agar-agar cubes was prepared. The subsequent experiment, which examined the degradation of Ibuprofen and Diclofenac using reused immobilized laccase and free laccase. Additionally, the experiment aimed to investigate whether the immobilized laccase retains its activity after being heat inactivated at 95°C post-immobilization and how its performance compares to free laccase. To assess this, we incubated the free laccase with the drug solution, taking two aliquots at specific time points during the incubation. One aliquot was then heat inactivated, allowing us to compare it with the untreated sample and determine if the laccase remained effectively immobilized within the agar cubes and was not leaking into the solution.

New cubes vs. reused cubes

There was no difference in Ibuprofen degradation between the reused and new immobilized laccase cubes, as both demonstrated the same degradation efficiency. In contrast, for Diclofenac, the reused cubes showed improved efficiency compared to the new ones, potentially due to differences in storage conditions.

Table 3: Degradation of Ibuprofen with Immobilized Laccase vs. post-incubation inactivated laccase

Time [h]	Ibuprofen [µg/mL]	Ibuprofen [µg/mL] heat treated
0	500.00	500.00
1	336.68	338.82
2	335.76	338.37
24	337.05	337.06
96	336.49	336.54

The results show that both the immobilized laccase and the heat-inactivated laccase after incubation achieved the same 32% degradation of Ibuprofen. This confirms that the laccase remained effectively immobilized within the cubes, as the degradation was not impacted by heat inactivation, indicating no enzyme leakage from the cubes.

Promoter isolation

We first designed primers to isolate promoters from S. cerevisiae genomic DNA (yeast strain S288c). These primers include homology overhangs that allow annealing during the assembly step. The melting temperatures we set to amplify our sequences from genomic DNA are initially calculated by NEB website (Tm calculator). Since primers are designed with homologous overhangs, we used a two-step PCR strategy to amplify the sequences; namely, the first 5 thermocycles are done with the primer melting temperature without the homology part, and the rest (about 30 thermocycles) are done with 72 degrees Celsius (maximal) annealing temperature. However, we have observed that our promoters are amplified with various efficiencies (see Fig.1a and b). At first, we hypothesised that this may be due to the bias that our target promoters might not be evenly distributed across the templates we took each time for our PCR reactions. Next, we used the amplicons that we purified from gels as templates to amplify more PCR products with maximal annealing temperature, that is, 72 degrees Celsius. However, we were not able to identify any amplification at the expecting electromobility (Fig.1c lane pHIS4 and opPDR5, Fig.1d lane pCWP2 and pSHM1). Furthermore, we repeatedly observed seemingly consistently low amplification efficiency from certain promoters (exemplified in Fig.1a, lane pHIS4, pPYK1; 1b, opPDR5; 1c, pPYK1; 1d, pPYK1, pHIS4, and opPRDR5). We thus postulated that primers might not have bound to our gel-purified templates.

We hypothesised that the estimated annealing temperature might not be accurate in this particular instance, and we sought an optimal melting temperature for each promoter by setting up thermogradient PCR reactions with melting temperatures ranging from 62 to 72 Celsius to amplify target sequences from gDNA. We assumed annealing temperature “sweet spots” for each corresponding promoter if the intensity of that band is the highest and that such band is surrounded by lower intensity bands, resembling a gaussian distribution, even though fluctuations in band intensity across temperatures due to sampling bias (that the amount of target is not always the same when the template is taken from the gDNA) is frequently observed (Fig.2). Next, we used the annealing temperature at which the band intensities are brightest for amplification from gDNA, and we were able to amplify a decent amount of product at expected electrophoretic mobility (Fig.3).

Extended primer PCR to amplify reporter gene, terminator and the backbone

We used similar approaches to amplify our fragments with overhang primer PCR. The primers have been designed such that each fragment will have overlapping sequences with the upstream and downstream elements in our goal plasmid (see Engineering Success - Biosensor – Introduction and Background, Fig.2). In general, we observed our target amplicon at the expected electrophoretic mobility for both our reporter gene (Fig.4a and b), terminator (Fig.4c), and BB3(2μ) backbone (Fig.4d and e). In addition to the 2μ plasmid, which is a high copy number plasmid for S. cerevisiae, we also amplified a low copy number plasmid with a centromere binding region (BB3_ARS/CEN) which integrates to the yeast genome and should contain the same sequence near multicloning site as in BB3_2μ plasmid. However, our initial attempt to use the same overhang primers to amplify BB3_ARS/CEN was unsuccessful (Fig.4f). We reasoned that the smear might be due to either one or more of at least three following reasons: a), artefacts from unspecific primer binding, b), amplification artefacts due to secondary structures of the plasmid and/or primers, and c), nuclease contamination. After using BamI (which cleaves once between the designed primer binding sites) linearised and PCR cleaned up plasmid as template, we still overserved an unspecific smear (Fig.4g). This excludes nuclease contamination as PCR clean-up kit (NEB #T1030S) should have removed nucleases. We then sought an optimal melting temperature for our primers to bind to the template, as at this point it appeared to us that unspecific binding might be the main cause of the smear (i.e., there are multiple similar elements in the template that could bind to our primers; thus, the amplicons will have various lengths). We observed a specific band at 62.9 degrees Celsius when we did a thermogradient PCR (Fig.4h). However, we were not able to reproduce this result when we attempted to amplify the target again (Fig.4i and j). From this point, we went on to our assembly step with BB3_2μ plasmid. Though, copy number can have a modulating effect on the sensitivity of our biosensor, it is more pressing to build a construct for the proof of concept.

Gibson Assembly and E. coli transformation

After we have amplified and purified all of our parts, we went on to assemble our target plasmids. We first set out a 0.1 pmol reaction, i.e., that all of our parts are at concentration of 0.1 pmol. Our initial transformation strategy was conventional heat shock transformation. However, we did not observe any colonies after our first transformation (data not shown). We confirmed by positive control that the assembly reaction indeed worked as cells transformed with assembled positive control DNA sample provided by the manufacturer showed colonies that are larger in size and higher in density (Fig.5b and c) than cells transformed with the same DNA sample without assembly reaction (Fig.5a). We hypothesized that this is due to poor assembly efficiency since our transformation positive control, in which cells were transformed by empty backbone BB3_2μ (Fig.5d and e) and BB3_ARS/CEN (not shown), showed decent number of colonies, implying that the transformation is not likely to be the cause. We thus increased the concentration of fragments to 0.25 pmol. After heat shock transformation of the assembled products, we still did not observe any colonies. We occasionally observed a few very tiny colonies, further colony PCR analysis showed negative results. We questioned whether our fragments indeed contained homologous sites and whether these sites are flanking the target sequence with designated order. To test this, we sent our fragments for Sanger Sequencing (Microsynth). All our BB3 plasmids, the 5’ of reporter genes, and 3’ of terminator sequence have shown desired homology sequence (data not shown); forward sequencing of pPYK1 and pCWP2 showed ambiguous results suggesting impurity (Fig.5f and h). However, this is not seen in the reverse sequencing data of the same primers (Fig.5g and i). We believed that these impurities may result from unspecific forward primer binding at the very 5’ sequence of these promoters as promoters frequently contain multiple copies of very similar sequences. We nonetheless assumed that these PCR products do contain desired promoter sequences, albeit only in a fraction. Taken together, these data suggest that we have to a large extent overestimated the actual amount of target plasmid within the Gibson Assembly mix. Since we do not have much time left before Wiki freeze, instead of using a higher concentration of fragments for the assembly reaction, we decided to transform E.coli cells via electroporation as electroporation has much higher transformation efficiency than heat shock1.

References

1. Lessard, J. C. Transformation of E. coli Via Electroporation. in 321–327 (2013). doi:10.1016/B978-0-12-418687-3.00027-6.