In this phase we seek to improve the kinetic values of our enzymes by assaying previously described and new mutants of the respective enzymes. We hope to enhance the degradation rates of our biofilm to more effectively degrade micropollutants in wastewater by employing these optimized enzymes.

There are several publications describing improved mutants of our targeted laccases which were generated by site saturation mutagenesis or directed evolution approaches.^[1] Using these methods, improvements for example in kcat of 1.21-fold or in the redox potential of 100mV could be achieved. Further information on specific mutants is given in our enzyme engineering part. ^[2][3]

Mutations can have a positive effect on enzyme activity for multiple reasons, for example by improving the stability of the active site or by improving ligand binding. As an example, the CotA double-mutant T232P/Q367R described by Ouyang et al. possesses 4.45-fold higher activity over the wildtype.^[2] In this specific case a glutamate (G367) is exchanged into arginine, which forms stronger hydrogen bonds within the active site and can also interact with K402. Thus, formation and reinforcement of bonds can lead to increased stability and a potentially more favourable active site.

We created a library of possible point mutations in both laccase genes which can be realized by performing side directed mutagenesis on our laccase expression plasmids for E. coli (pET24). During this process multiple mutations can be introduced simultaneously to check for their compatibility and possible advantages in combined mutations.1 [Hyperlink Quick-Change Assays] Afterwards, the mutants can be checked for increased catalytic activity using the same assays previously described for laccase characterization. Examples for laccase assays are the degradation of the model substrate 2,2′-Azino-bis(3-ethylbenzothiazoline-6-sulfonic acid (ABTS) with a defined unit of enzymes or the degradation of our target molecule Diclofenac coupled with HPLC analysis. ^[3][4]

Since there are no publications on directed mutagenesis of EreB, we consider a site saturation mutagenesis approach. This way we hope to improve the binding pocket of the enzyme to bind and degrade azithromycin. Wild type EreB shows promiscuous activity to azithromycin but no antibiotic resistance is achieved yet.^[5] As shown before, enzymes with promiscuous activities towards other substrates are a great starting point for the directed evolution of enzymes for new substrate specificities.^[6] Positive mutants can be identified by Kirby Bauer assay, that measures cell toxicity of azithromycin and its degradation products after incubation with a defined unit of EreB enzyme.

Since our lab access this year is very limited due to the current COVID-19 pandemic, we attempt an in silico approach for EreB optimization. Therefore, we use the Rosetta Design application to improve the ligand binding for azithromycin in EreBs active site. The algorithm mutates specified amino acids and carries out molecular docking experiments with the protein variant and compares the variant to the wild type protein.^[7] This way we hope to find variants with improved catalytic activity. The structure for EreB was obtained via a comparative modelling run. Comparative modelling is a homology modelling approach that uses structures with high sequence homologies to determine the enzymes 3D structure. Mostly amino acids close to the catalytic site will be targeted for mutations. Rosetta Design was also used on both laccases CotA and CueO, introducing both random and previously described mutations. This way we hope to validate our researched mutation sites or find new ones that possess improved catalytic values especially for our target molecules. Thus, we can apply the mutated enzymes specifically on our problem granting higher accuracy of the predicted effects of the mutations.

Site directed mutagenesis

Mutants were obtained by site directed mutagenesis (Quik-Change™ polymerase chain reaction (PCR)) using the original gene fragments cloned into our expression vectors as templates. Therefore, primers were designed to contain the desired mutation within their hybridization sequence and an additional overlapping region. This way the site directed mutagenesis primers are aimed to amplify the whole plasmids. Consequently, the PCR product is the circular expression vector containing our gene of interest with the introduced point mutation. PCR does not include ligation steps therefore the product plasmids contain one nicked site on each DNA strand. The nicked plasmids are repaired by bacterial repair mechanisms after transformation.^[8]

PCR amplification mainly follows the standard reaction conditions according to the used polymerase, only the number of reaction cycles is lowered and template concentration raised [Hyperlink PCR amplification + polymerase]. The annealing temperature is chosen according to the primers melting temperature and the used DNA polymerase. After PCR amplification the product is digested with DnpI nuclease, which is specific for methylated DNA. This way the template DNA, that was obtained from E. coli and thus is methylated, is degraded, leaving only the modified circular PCR products.^[9]

Enzyme Assay

The assays that were briefly described above can be used to determine the activity of the enzyme mutants. In the ABTS assay the model substrate ABTS is oxidized by laccases to its diazonium salt cation that shows an absorbance peak at 420 nm light. Consequently, the enzyme kinetics can be measured by spectrophotometric analysis.^[10] Yet, ABTS is not the substrate we actually want to improve the activity for. Therefore, we aim to measure the reaction kinetics of laccase mutants towards Diclofenac, our main target molecule. Diclofenac shows no absorption of visible light so photometric analysis cannot be used. Instead the reaction is tracked by performing HPLC analysis of samples taken at certain time points. Diclofenac shows UV absorbance at 220 nm light but background absorbance makes separation steps such as HPLC necessary for precise measurement. The reaction is therefore stopped by precipitation of the enzymes with ethanol.^[11] [Hyperlink HPLC assay] Performing identical assays on both wild type enzymes and mutants guarantees comparability of our results. This way the best variant of our enzyme for application in the biofilm can be found.

We hope to achieve an activity increase of our targeted laccases by site-directed mutagenesis of residues in the active site. The targeted mutations refer to publications which used site-saturation mutagenesis on the respective enzymes.^[12]

Additional experiments following our in vitro laccase activity assays are necessary to quantify the increase or decrease in activity in the biofilm. Only mutants showing activity increase should be used in further experiments, others can be discarded. For EreB no mutations and their effects were previously determined, therefore we considered an in silico site saturation mutagenesis approach. Promising mutants can then be tested in the lab. For further information on EreB mutagenesis, please see our text on enzyme design and directed evolution.

For enzyme purification by affinity chromatography we designed EreB, CotA and CueO with an attached Strep-Tag, that can be used for chromatography with either Streptavidin columns or Streptavidin variants.^[1] By purification we are able to determine the enzymes reaction kinetics. (Hyperlink Laccase + EreB Assays)

CotA: The cotA sequence was taken from the uniport entry for CotA (H8WGE7) and codon optimized for E. coli using the GenSmart™ Codon Optimization tool. Sequence for Strep-Tag II was taken from the iGEM parts collection with the amino acid sequence “WSHPQFEK” and fused to the C-terminus (3’-end ) of CotA. A flexible linker peptide GGS was added between the two parts to guarantee the correct structure of the Strep-Tag, which is crucial for its functionality. The affinity tag was added to the C-terminal end of the laccase since the crystal structure suggests accessibility of the terminus (PDB: 1GSK).

CueO: cueO gene sequence was taken from its uniprot entry ((P36649)). It was fused to Strep-Tag II with a GGS linker similar to CotA. Strep-Tag II was added to its 3’-end C-terminally since this terminus is accessible, as identifiable in its PDB entry 1KV7.

EreB: ereB gene sequence was taken from its uniprot entry (P05789). Strep-Tag II was fused to its 3’-end (C terminus) without a linker peptide, since the C-terminus appears to be free in our modelled structure (Hyperlink modelling).

All plasmids were planned using SnapGene®, of which licenses were kindly provided by GSL Biotech LLC for us to use in our project. BioBrick® restriction enzyme sites were removed by adding different codons for the same amino acids.

For all plasmids a restriction site for restriction enzyme NdeI was added to the 5’ untranslated region (UTR) to allow modifications of the construct. For production and enzyme purification in E. coli we planned to use the pET24 plasmid with a strong T7 promotor. Cloning was planned with Gibson Assembly overhangs of the plasmid linearized via PCR. For application in our biofilm we planned using fusion proteins with TasA (EPS) expressed in B. subtilis.