Revision as of 14:12, 18 October 2020

Introduction Challenge and Idea Degradation of Substances Experimental Approach Safety Outlook

Introduction to Pharmaceutical Degradation

Wastewater toxicity - a topic that affects each and every one of us in our daily lives. If wastewatr is not treated properly, it can pollute our entire environment and affect the lives of people, animals and plants severely^[1]. But even if proper filtration and purification were achieved, some problematic substances such as diclofenac, carbamazepine or azithromycin would still not be degraded efficiently in wastewater treatment plants. This can even lead to the extinction of species^[2,^3,^4,^5,^6]. Since sustainability is an issue of significant importance to us, we have decided to tackle some substances that seem to be remarkably resistant and incredibly difficult to degrade in wastewater treatment plants.

GraphicalAbstract — Figure 1: Graphical Abstract.

By using a B. subtilis biofilm with the capability of degrading said substances, we are able to exceedingly reduce the toxicity of wastewater and work towards a more sustainable use of active ingredients. Our goal is to prove that the enzymes we have selected are able to efficiently degrade substances that cause immense problems in wastewater. By taking advantage of comparative modeling and Molecular Dynamics Simulation, we want to find out how we can modify these enzymes to increase substrate specificity and ensure that we are able to work with them on a larger scale. We also aim to prove that the products of our degradation are rendered less toxic to the environment in an efficient way.

Here is more information about how we have decided to study and modify our enzymes.

Challenge and Idea

What is the Problem?

Unfortunately, the pollution of wastewater caused by various drugs, antibiotics and other human influences is no longer a novelty. Several pharmaceuticals pollute our environment because they are not properly degraded in wastewater treatment plants^[3,^7]. Despite frequent analyses of water bodies in Europe, only indicative limit values for various drugs or painkillers such as diclofenac have been established so far^[8]. The effects on aquatic fauna^[1] and flora^[9] remain not merely enough emphasized. However, this is precisely where the problem lies.
Because of this, we kept coming back to issues of wastewater toxicity, while researching for this year’s iGEM project. As our team mostly consists of young scientists, it is our duty and will to work towards a more sustainable future. Purity of water is something every single person in the world has a right on, which motivated us even more to come up with a system to make sure, this right is being fulfilled. If we don’t act now, who knows what will happen to our environment in the near future.

Project Idea

This is where B-TOX, the reduction of wastewater toxicity using a B. subtilis biofilm, comes in. The ability of B. subtilis to form a natural biofilm is what this year’s iGEM team of TU Darmstadt means to utilize. With the combination of our bacterial strain and genetically modified laccases, we are eager to lower the toxicity of wastewater in order to preserve and protect our environment. A modification will enable the laccases, one extracted from E. coli and one from B. subtilis itself, to oxidize diclofenac to hydroxydiclofenac – making it less toxic. We also did our research on other substances than diclofenac, since we want to provide a modular system, suitable for other areas as well. For that you can find out more here. (Link zum Text weitere Substanzen)

Why Diclofenac?

Some substances appear more prevalent in this regard than others, dominating statistics of wastewater concentration and general toxicity. The painkiller diclofenac has been known to be potentially harmful since 1990, as it was the cause of an alarming mass extinction of three vulture species in Pakistan. Residues of diclofenac have also been found in many other species such as otters and trout^[2,^4]. Additionally it has been a burden on aquatic fauna: Fish that absorb micropollutants such as diclofenac via the water surface are exposed to damage to internal organs, bearing insufficiency of the kidney or liver. ^{(Sofija, was ist die quelle :o)}
In 2015, diclofenac has been placed on the EU watch list of priority substances^[2]. We decided to focus on this substance and wanted to develop a system that enables the degradation of diclofenac and thus has a positive impact on the environment. We found oxidoreductases, so-called laccases, which already show a degradation of diclofenac and a variety of other phenolic substances^[7,^10,^11]. Many experts such as Patrick Schröder, scientific administrator of the German Environment Agency, confirmed the danger that pharmaceuticals pose to the environment. (Hyperlink zu patrick schröder)
As only one enzyme was not sufficient for us, we also looked for a solution to degrade azithromycin. Azithromycin was also placed on the EU watch list with high priority, as it repeatedly exceeds its PNEC (Predicted No Effect Concentration, 0.019 ug/L) in various areas as you can see on the figure below (figure xy)^[4]. The horizontal line on each figure resembles the PNEC and strikingly, most measured concentrations are above that line. That also accounts for Diclofenac. During our research we came across an esterase called erythromycin esterase type II (EreB) which is able to efficiently degrade erythromycin, a substance with structural similarity to its semisynthetic successor azithromycin^[12].

Elephant at sunset — Figure 1: Measured Concentration of Diclofenac in open waters^[4].

Degradation of Substances

What Do Our Enzymes Do?

Modular Biofilm

Including Other Toxic Substances Into Our Project

Pharmaceuticals

Experimental Approach

Experimental Design

Cloning

Analytics: HPLC und ABTS

Kirby-Bauer-Assay and LCMS

Toxicity Assay

Enzyme Engineering

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]

Figure 1: Side directed mutagenesis. The plasmid containing the GOI is replicated using a primer pair with a built in mutation to replicate nicked plasmids containging the desired mutations. The plasmids are then transformed into competent cells for ligation and expression of the engineered protein.

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.

What Parts did we Design?

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.

Directed Evolution

Directed Evolution describes the process of creating a library of nucleic acids or proteins and their selection towards a specific ability. Consequently, this method mimics a normal evolution process in an accelerated form.^[1] Different approaches vary in their method of library creation and screening. Three of the most common possibilities for library construction are:
- DNA shuffling: DNase I digestion of DNA fragments with subsequent PCR reaction without additional primers. DNA fragments need to show partial sequence homology.^[1]
- Error prone PCR: PCR reaction introducing random copying errors through mutagenic reaction conditions such as Mn2+ ions instead of Mg2+ ions (cofactors + providing DNA binding) or polymerases without proof reading result in higher error rates.^[1]
- Site saturation mutagenesis: Targeted mutation of amino acids e. g. for structural observations of proteins.^[1]

The screening highly depends on the target for directed evolution. A few examples for screening techniques are phage displays, yeast displays or in vitro compartmentalization. In our case catalytic assays can be used to quantify the enzymes degradation rate towards a specific substrate, such as 2,2′-Azino-bis(3-ethylbenzothiazoline-6-sulfonic acid (ABTS) that’s oxidized to a solvable cation showing absorption of visible light. (Suitable for photometric analysis) A dramatic increase in effectiveness could be achieved by exchanging the wild type enzymes with mutants exhibiting increased catalytic activity towards our target molecules or increased stability. Therefore, we will select the best performing laccase for application in our biofilm to receive the highest possible transformation rate for pharmaceuticals in water, leading to decreased toxicity and cleaner water.

Table 1: Mutations on CotA Laccase from B.subtilis (Ouyang et al. Enhanced catalytic efficiency of CotA-laccase by DNA shuffling ^[1], Melo et al. Insight into stability of CotA laccase from the spore coat of Bacillus subtilis^[3])
Mutation	Publication	Effect	Kinetic values
Wild type	Enhanced catalytic efficiency of CotA-laccase by DNA shuffling	pH_Opt = 4.4 (ABTS) / 6.8 (SGZ) pH is strongly substrate-dependant Higly stable at pH ranging from 7-9 T_Opt = 60°C Hydrophobic interactions around active-site determine thermo-stability. Substitutions can lead to stability increase (f.e. T232F)	k_cat = 6.3 s^-1 K_M = 24.8 µM (SGZ)
T232P/Q367R	Enhanced catalytic efficiency of CotA-laccase by DNA shuffling	Gln-367 might be involved in hydrogen bonds with Arg-365, Asn-368, and Thr-406 Arg substitution allows new hydrogen bond between Arg-367 and Lys-402 Q367R might have changed the binding pocket into a potentially more favourable binding site	k_cat/K_M = 0.374 s⁻¹ µM⁻¹ 18.15% decrease in K_M and 1.21-fold increases in k_cat
M502L/I	Insight into stability of CotA laccase from the spore coat of Bacillus subtilis	Met⁵⁰² (weakly co-ordinating to the T1 copper) exchanged to non-coordinating residues stores geometry of copper-binding sites	Increases redox potential by approx. 100 mV

Table 2: Overview CotA mutations (Mate et al. Laccase engineering: From rational design to directed evolution^[2])
Laccase Host Organism	Approach	Property in study	Charakterization methods	Main results	Reference
CotA drom Bacillus subtilis	SM	Mechanism of reduction of O₂ to H₂O	Redox titration, EPR, CD spectroscopy, CAA^b	Small changes in the geometry of the Cu sites. Turnover rates highly reduced and optimal pH downshifted 1–2 unit	Brissos et al. 2012
	SM	Mechanism of reduction of O₂ to H₂O	Simulated pH titrations	Asp116 appears to be crucial in modulating Glu498 protonation	Silva et al. 2012
	CSM	Subststrate specifity	CAA^b	The CotA-ABTS-10 mutant was 132-fold more specific for ABTS	Gupta et al. 2010

Table 3: Mutations on CueO laccase from E. coli (Zhang et al. Directed Evolution of a Bacterial Laccase (CueO) for Enzymatic Biofuel Cells^[4], Wang et al. Crystal structures of multicopper oxidase CueO G304K mutant: structural basis of the increased laccase activity^[5])
Mutation	Publication	Effect	Kinetic values
Wild type	Directed Evolution of a Bacterial Laccase (CueO) for Enzymatic Biofuel Cells	The T1 Cu active site accepts four electrons of substrate oxidation and passes them to T2/T3 Cu cluster, where molecular oxygen is fully reduced to two water molecules by accepting four electrons CueO possesses a labile 5th copper binding residue in 7,5 A distance from T1 Cu active site Coordinated through 2 Met (M355 and M441), 2 Asp (D360 and D439) and a water molecule in triangular bipyramidal geometry.	E = 0.36 V
D439T/L502K	Directed Evolution of a Bacterial Laccase (CueO) for Enzymatic Biofuel Cells	D439 and L502 are located in the second coordination sphere of the T1Cu and form hydrogen bonds with coordinated ligands H443and C500.	E = 0.56 V (ABTS assay) 1.72-fold enhanced power output of the Microbial fuel cell.
G304K	Crystal structures of multicopper oxidase CueO G304K mutant: structural basis of the increased laccase activity	Residues D439 and L502 are adjacent to the T1-Cu coordinating ligands and targets for improvements of its onset potential	presence of excess Cu (II) was 2.7-folds higher

Safety

SafetyTextsPlaceholder

Outlook: What would we have done with more time?

Unfortunately, it was not possible for us to come to the lab and conduct actual experiments this year. Nevertheless, we came up with a lab plan with all the experiments and assays we would have done to get our project up and running.
At the beginning of our lab time we would have transformed our two laccases, CueO and CotA, into E. coli. After a successful expression of the enzymes, we would have used a 2,2’-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid (ABTS) assay to check the laccase activity in vitro^[101]. The kinetic activity or the determination of an effective laccase concentration would have been detected by HPLC. We were also planning a toxicity assay. Since our plan is to render diclofenac and other toxic substances less harmful to the environment by oxidation via our laccases, the detection of the substrate concentration with the lowest toxicity is a crucial step. Hence, we were planning an assay with zebrafish embryos, considering they are not classified as animal testing and commonly used in this respect^[102]. (see Tox-Assay, Hyperlink).
After the detection of the activity of laccase in vitro, the affinity of the substrate conversion would be increased using a quick-change PCR.
We planned to perform the analytical assays with our own expressed laccases as well as with laccase from Trametes versicolor in order to generate comparable values.
As we try to degrade a wide range of substances in wastewater with our laccase (see modular biofilm, hyperlink), we also wanted to measure the kinetic degradation values for different substances. Since it was very difficult to find comparable degradation values for different substances during our research, we would have also done this with the laccase from T. versicolor, so our work would have been useful for future projects.
In addition to our laccases, we were also focusing on the degradation of azithromycin via the enzyme EreB. Here too, we would have transformed and expressed the enzyme into E. coli at the beginning of our laboratory time. To detect the degradation of azithromycin, we were planning a Kirby-Bauer assay^[103]. Just as with the laccases, we would have performed HPLC to measure kinetic activity, as well as a LC-MS. To increase the substrate affinity of the enzyme, we would have performed site saturation mutagenesis. Our modeling team is also working on a rational design of the enzyme. (see modeling, hyperlink)
All assays would have been performed with both the naturally occurring enzyme variant and the optimized variant to generate comparability.
The next step would have been an implementation of our selected enzymes in a B. subtilis biofilm. We planned to realize this via a tasA fusion protein. To prove our concept, our biofilm sub-group has worked on an assay used to immobilize a fusion protein from tasA and sfGFP in the biofilm matrix (see Filmies, Hyperlink). The assay has already been performed with larger proteins, so we are positively encouraged to have it performed successfully with our laccases or EreB. A laccase-tasA and EreB-tasA fusion protein has already been designed for this purpose and would have been transformed and expressed in B. subtilis if it had been possible to go into the laboratory. This way, we would have managed to immobilize the active enzyme in the outer biofilm matrix, which harmful drugs and pharmaceuticals in wastewater reach without complications.

References

@@ Line 505: / Line 505: @@
       Unfortunately, it was not possible for us to come to the lab and conduct actual experiments this year. Nevertheless, we came up with a lab plan with all the experiments and assays we would have done to get our project up and running.  <br>
-  At the beginning of our lab time we would have transformed our two laccases, CueO and CotA, into <I>E.&nbsp;coli</i>. After a successful expression of the enzymes, we would have used a 2,2’-azino-bis(3-ethylbenzothiazoline-6-sulfonic&nbsp;acid&nbsp;(ABTS) assay to check the laccase activity <I>in vitro</i>.<sup>1</sup> The kinetic activity or the determination of an effective laccase concentration would have been detected by HPLC. We were also planning a toxicity assay. Since our plan is to render diclofenac and other toxic substances less harmful to the environment by oxidation via our laccases, the detection of the substrate concentration with the lowest toxicity is a crucial step. Hence, we were planning an assay with zebrafish embryos, considering they are not classified as animal testing and commonly used in this respect.<sup>2</sup> (see Tox-Assay, Hyperlink). <br>
+  At the beginning of our lab time we would have transformed our two laccases, CueO and CotA, into <I>E.&nbsp;coli</i>. After a successful expression of the enzymes, we would have used a 2,2’-azino-bis(3-ethylbenzothiazoline-6-sulfonic&nbsp;acid&nbsp;(ABTS) assay to check the laccase activity <I>in vitro</i><sup id="cite_ref-101"><a href="#cite_note-101">[101]</a></sup>. The kinetic activity or the determination of an effective laccase concentration would have been detected by HPLC. We were also planning a toxicity assay. Since our plan is to render diclofenac and other toxic substances less harmful to the environment by oxidation via our laccases, the detection of the substrate concentration with the lowest toxicity is a crucial step. Hence, we were planning an assay with zebrafish embryos, considering they are not classified as animal testing and commonly used in this respect<sup id="cite_ref-102"><a href="#cite_note-102">[102]</a></sup>. (see Tox-Assay, Hyperlink). <br>
   After the detection of the activity of laccase <I>in vitro</i>, the affinity of the substrate conversion would be increased using a quick-change PCR. <br>
@@ Line 513: / Line 513: @@
   As we try to degrade a wide range of substances in wastewater with our laccase (see modular biofilm, hyperlink), we also wanted to measure the kinetic degradation values for different substances. Since it was very difficult to find comparable degradation values for different substances during our research, we would have also done this with the laccase from <I>T.&nbsp;versicolor</i>, so our work would have been useful for future projects.  <br>
-  In addition to our laccases, we were also focusing on the degradation of azithromycin via the enzyme EreB. Here too, we would have transformed and expressed the enzyme into <I>E.&nbsp;coli</i> at the beginning of our laboratory time. To detect the degradation of azithromycin, we were planning a Kirby-Bauer assay.<sup>3</sup> Just as with the laccases, we would have performed HPLC to measure kinetic activity, as well as a LC-MS. To increase the substrate affinity of the enzyme, we would have performed site saturation mutagenesis. Our modeling team is also working on a rational design of the enzyme. (see modeling, hyperlink) <br>
+  In addition to our laccases, we were also focusing on the degradation of azithromycin via the enzyme EreB. Here too, we would have transformed and expressed the enzyme into <I>E.&nbsp;coli</i> at the beginning of our laboratory time. To detect the degradation of azithromycin, we were planning a Kirby-Bauer assay<sup id="cite_ref-103"><a href="#cite_note-103">[103]</a></sup>. Just as with the laccases, we would have performed HPLC to measure kinetic activity, as well as a LC-MS. To increase the substrate affinity of the enzyme, we would have performed site saturation mutagenesis. Our modeling team is also working on a rational design of the enzyme. (see modeling, hyperlink) <br>
   All assays would have been performed with both the naturally occurring enzyme variant and the optimized variant to generate comparability. <br>
@@ Line 570: / Line 570: @@
              <a  class="anchor" id="cite_note-428"></a>
              <a class="referencestd" href="https://doi.org/10.1038/s41598-018-32446-7" target="_blank">5. H. Whang et al. Crystal structures of multicopper oxidase CueO G304K mutant: structural basis of the increased laccase activity. Scientific Reports 2018, 8, https://doi.org/10.1038/s41598-018-32446-7</a>
+<br> <br>
+<a class="anchor" id="cite_note-101"></a>
+            <a class="referencestd" href="https://www.sciencedirect.com/science/article/abs/pii/S0045653512003335?via%3Dihub">101. I. Eichlerová, J. Šnajdr, P. Baldrian, Laccase activity in soils: consideration for the measurement of enzyme activity, Chemosphere, 2012, 88: 1154-1160, https://doi.org/10.1016/j.chemosphere.2012.03.019 </a>
+<a class="anchor" id="cite_note-102"></a>
+            <a class="referencestd" href="https://www.sciencedirect.com/science/article/abs/pii/S0045653504002759">102. A. V. Hallare, H.-R. Köhler, R. Triebskorn, Developmental toxicity and stress protein responses in zebrafish embryos after exposure to diclofenac and its solvent, DMSO, Chemosphere, 2004, 56: 659-666, https://doi.org/10.1016/j.chemosphere.2004.04.007 </a>
+<a class="anchor" id="cite_note-103"></a>
+            <a class="referencestd" href="https://asm.org/getattachment/2594ce26-bd44-47f6-8287-0657aa9185ad/Kirby-Bauer-Disk-Diffusion-Susceptibility-Test-Protocol-pdf.pdf">103. J. Hudzicki, Kirby-Bauer Disk Diffusion Susceptibility Test Protocol, American Society for Microbiology, 2009 </a>
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