Wastewater toxicity - a topic that affects each and every one of us in our daily lives. If wastewater is not treated properly, it can pollute the 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 (WWTP). 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.
Figure 1: Graphical abstract of pollution sources The figure shows possible sources of micropollutants such as diclofenac, erythromycin and azithromycin. These substances can be broken down or transformed by enzymes such as laccase or EreB and converted into less toxic substances. The environmentally harmful consequences are thus significantly minimized[7].
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 future. 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.
Why Diclofenac and Azithromycin?
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[2].
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. For more information, click here.
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 the German Environment Agency repeatedly measured it exceeding it PNEC (Predicted No Effect Concentration, 0.019 ug/L) in various areas as you can see on the figures below (figure 2 and 3) (published data from the German Environment Agency)[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].
Figure 2: Diclofenac pollution The graph was taken from the German Environment Agency and shows the measuring program of 2016 at the measuring sites of the observation list (EU-Watch-List). The height of the columns shows the annual mean value of diclofenac at the respective measuring points in μg/L. The horizontal line indicates the predicted no effect concentration (PNEC). The abbreviation BG stands for determination limits. For diclofenac, every single measured concentration at least meets the PNEC line (published data from the German Environment Agency)[4].
Figure 3: Azithromycin pollution The graph was taken from the German Environment Agency and shows the measuring program of 2016 at the measuring sites of the observation list (EU-Watch-List). The height of the columns shows the annual mean value of azithromycin at the respective measuring points in μg/L. The horizontal line indicates the predicted no effect concentration (PNEC). The abbreviation BG stands for determination limits. It can be seen that either the PNEC or the BG is exceeded at each measuring site, except for one, where the BG is exceeded. On most measuring sites, the PNEC is exceeded (published data from the German Environment Agency)[4].
Degradation of Substances
What Do Our Enzymes Do?
For pharmaceutical degradation , we use laccases such as CotA from Bacillus subtilis, as well as CueO Escherichia coli[13]. Laccases show a natural capability of degrading phenolic substances through copper-catalyzed oxidation [13] and are therefore categorized as phenol oxidases. The oxidation of phenolic educts is coupled to the reduction of oxygen to water. It is our goal to engineer the selected laccases CotA and CueO for optimized diclofenac degradation. In a process of oxidization, laccases transform diclofenac to hydroxydiclofenac, which has been shown to be less toxic [19]. The laccase of the fungi Trametesvesicolor is well-studied to degrade a variety of substrates , such as trimethoprim, carbamazepine and sulfamethoxanzole [11].
Laccases possess a T1 copper site in close proximity to the substrate binding pocket and a coupled T2/T3 trinuclear copper cluster (TNC) , that gives them their ability to mediate electron transfers . T1 copper is ligated by two His and a Cys residue and additionally Ile and Phe/Leu/Met, both non- or weakly coordinating. T2 copper is ligated by two His and T3 by three His residues as well as an additional oxygen-bridged second copper . A substrate molecule, either phenolic derivates or metal ion substrates , binds to the T1 copper site. An electron is transported to the TNC by either extension of the coppers’ π-system over the ligating Cys residue, or via a different electron pathway involving an H-bond through a carbonyl oxygen[18]. O2 is ligated by the TNC and is reduced to H20 without the formation of active peroxide intermediates[14].
Bacterial laccases differ from laccases obtained from fungi regarding the axial complexation of the copper ion involved in catalytic activity[14]. Whilst bacterial laccases contain axial methionine ligands , fungi laccases such as the laccase found in Trametisvesiculor achieve a higher redox potential in the absence of complexing axial ligands[14].
Figure 4: Laccase CotA
Figure 5: Laccase CueO
Figure 6: Laccase from Trametisvesiculor
CotA is a laccase originally found in Bacillus subtilis, which is also our microorganism of choice for biofilm implementation. CotA has shown to oxidize polycyclic aromatic hydrocarbons such as anthracene and benzol. CueO derives from Escherichia coli and is specialized in degradation of synthetic dyes such as congo red and malachite green[16]. This blue copper oxidase contains a fifth, unstably coordinated copper ion, through which it differs greatly from Cu(II) to Cu(I), functioning as a redox mediator. Our enzyme optimization strategies target not only the copper complexation.
In literature, for both our utilized laccases the catalytic activity was improved by site saturation mutagenesis approaches [13].
Inappropriately disposed antibiotics are a risk factor for the development of antibiotic resistances. We therefore made it our goal to reduce antibiotic concentration in wastewater using suitable degradation enzymes. On the EU watch-list of most relevant antibiotics in wastewater pollution, azithromycin which is a semisynthetic derivate of erythromycin, ranks high, as it exceeds the PNEC (predicted no effect concentration) at multiple locations of measurement[14]. It is amongst the few antibiotics in the effluents of European wastewater treatment plants that pose a significant risk to the aquatic environment in several European countries[15]. As azithromycin is particularly relevant in clinical application, avoiding development of resistances in the environment is essential. To approach the issue of high azithromycin concentration in wastewater, we equip our biofilm with a variant of an erythromycin esterase, namely erythromycin esterase type II (EreB) , to degrade azithromycin amongst other antibiotics such as erythromycin itself. EreB is an esterase capable of hydrolyzation of a specific group of cyclic esters called macrolides [13]. EreB differs from other representative enzymes of the esterase group in its substrate promiscuity for azithromycin [13], which we intend to optimize by site saturation mutagenesis of its active site. The hydrolyzation product of both erythromycin and azithromycin is a cleaved ester forming a full acetal as shown in figure 5 [13].
Figure 7: Mechanism of Erythromycin degradation with EreB[13]
The reduction of azithromycin concentration in wastewater aids avoidance of antibiotic resistance and prevents the loss of clinical applicability for an important antibiotic. However, using a genetically modified organism to achieve this goal implies the important responsibility of containment .The B. subtilis strain used in our biofilm provides an artificial pathway for antibiotic degradation, which could be transferred to potentially pathogenic bacteria by horizontal gene transfer of the heterologous esterase gene. This may result in the clinical inapplicability of azithromycin that we are intending to counteract. Due to this issue, we made it our highest priority to guarantee the death of any bacteria leaving the biofilm using a quorum sensing based kill switch . For aspects related to biosafety please see our quorum sensing kill switch and our text on safety .
Modular Biofilm
Nowadays, pharmaceuticals are available everywhere on the market and in all possible shapes and forms. Most of them, from foot care products to antibiotics, eventually end up in wastewater in more or less large quantities, but all at the expense of the environment.
We have chosen diclofenac as our main target because of its current relevance[4][21], but there are many more problematic substances. Most of them are not yet fully handled by modern wastewater treatment plants[22]. Since specific enzymes are best suited for the detoxification of certain pharmaceuticals and we have the possibility to express multiple enzymes at once with our biofilm, we can therefore fight the pollution of water on a much broader angle. For this purpose, we plan to incorporate as many enzymes as possible that enable the detoxification of the most problematic substances.An advantageous property of our laccase is the ability to transform other relevant pharmaceuticals in addition to our main target diclofenac and even non-pharmaceutical substances that enter the environment via our wastewater, of which the most relevant are listed here:
Ibuprofen is a widely used pharmaceutical for the treatment of pain, inflammation and fever. Since it is a drug not subject to prescription, it is part of every modern medicine chest and its consumption is accordingly frequent. In 2018, more than 500 million daily doses of ibuprofen were consumed in Germany alone.[33] Worldwide, it occupies third place among the world's most widely used pharmaceuticals.[34] These enormous quantities are of course also reflected in the concentrations of ibuprofen in wastewater where it has been detected at concentrations of up to 3micrograms per litre.[34] Since most wastewater treatment plants are not yet equipped to handle that much pollution of the hard to remove pharmaceutical, ibuprofen often finds its way into the open waters, where it causes harm to both organisms and sub-organisms.[35]
Carbamazepine is an established anticonvulsant drug that is used the treatment of grand mal and psychomotor epilepsy as well as other non-epileptic seizures i.e. from multiple sklerosis or alcohol withdrawal.[36][25] In addition, it has proven to be effective against trigeminal neuralgia and is used for the prevention of manic phases of bipolar disorder.[36] It is mostly degraded in the liver and only 2-3% leave the human body unchanged.[25] These residuals, however, are problematic in wastewater treatment plants, since they are only able to remove up to 62% of the remaining Carbamazepine [25][37] and the leftover drug is toxic to lower life forms in the aquatic environment.[25][38]
Chloramphenicol is a broad-spectrum antibiotic which was first produced in 1947 by the chemists Paul R. Burkholder and David Gottlieb. Due to frequent serious side effects, it is now only used as a reserve antibiotic for very heavy diseases such as typhoid, paratyphoid, dysentery, diphtheria, malaria, Rocky Mountain spotted fever or rickettsialpox. In Germany, no chloramphenicol-containing preparations for human use have been on the market since 2019. In other countries, however, it is still widely used in animal production, especially for aquatic life.[39]
Bisphenol A is used in the manufacturing of plastics such as polycarbonates and epoxy paints and is one of the most widely used synthetic chemicals in the world. Independent studies have detected it in air, dust, surface water and seawater, but also in fruits from greenhouses and drinking water from plastic tanks and even in human blood and urine. It is released into the environment in such large quantities because packaging materials containing bisphenol A release it when heated or through passive extraction over a long period of time. This is particularly dangerous when food or water is packaged with such plastics and it absorbs the chemical directly, thus leading to increased human consumption. Bisphenol A acts as an endocrine disruptor, i.e. it interferes with the hormonal balance of living organisms in contact with it.[40] According to the WHO, frequent contact can contribute to diseases such as diabetes mellitus, obesity, thyroid dysfunction, developmental disorders (especially in children) and infertility.[41]
Nonylphenols are used in the production of NPEs (synthetic surfactants/detergents) and are also an intermediate in many other chemical processes. During the biological or thermal degradation of NPEs, nonylphenols are released into the environment. They accumulate in river sediment and aquatic organisms and exhibit estrogenic activity. The various nonylphenol isomers are degraded at different rates depending on their degree of branching. 4-Nonylphenol is the most unbranched and therefore most problematic, as it degrades slower than other nonylphenols.[42]
17β-Estradiol is a sex hormone and the most potent of the natural estrogens. It is formed in the follicles of the ovaries as well as in the testicles and ends up in wastewater through urine and faeces.[43] The substance interferes with the normal physiological process of water animals and can therefore create many deleterious effects [45] It constitutes a danger to the environment due to its low degradability in wastewater treatment plants and persistency in the aquatic environment, where it accumulates through the food chain.[44][45]
In order to estimate the effect of our laccase CotA on the various substances, we have also conducted Rosetta Docking simulations and compared the results with the widely used fungal laccase from Trametes Versicolor.
Through methods like directed evolution or rational design, we are also able to modify the binding sites of our enzymes CotA and CueO to specifically target other substrates.
Due to the many possibilities of enzyme combinations that we are able to express with our biofilm, we can address many major problems regarding water pollution that diverge from solely removing pharmaceuticals in municipal wastewater.The exchange of enzymes within the biofilm can be done on DNA level by the fusion of different enzymes to the extracellular protein TasA. On an early stage of our project we want to use plasmids with the TasA-enzyme fusion protein, later they could be genome integrated. Depending on the development stage of our project the enzymes could be exchanged by simple cloning methods like Gibson Assembly or with genome editing methods like CRISPR-Cas9. To immobilize more than one enzyme in the biofilm matrix, there could be used more than one plasmid in one bacterium or the biofilm could contain bacteria with DNA for different fusion proteins.
With these options it is possible to adapt the biofilm on many different conditions and therefore enables a wide range of applications.
. We discussed multiple different scenarios for the implication of our Biofilm, such as the detoxification of dyes that are a problem in the wastewater of textile factories[32], the detoxification of overused pesticides in agriculture or the recycling of water in closed systems such as the International Space Station
.
In conclusion, there are three levels of additional applicability of our biofilm:
Our laccase is able to transform a wide range of substrates and even without switching out the enzymes, our system can be used in many different situations, e. g. when different substances are present in the wastewater.
The enzymes inside of the biofilm matrix can easily be switched and so even a broader range of potential targets is reachable with our system.
Lastly, our system can be utilized for different places and various problems, as mentioned before: As a filter for dyes in river water or for water recycling in circular systems e.g. inside the International Space Station.
With these methods of adaptation and wide use of our biofilm, we can ensure clean water in many regions. Thereby our project contributes to two sustainable development goals of the United Nations: 6 (clean water and sanitation) as well as 14 (life below water).
Figure 8: Degradable pharmaceuticals
Pesticides for our Application
According to the world health organization “Pesticides are chemical compounds that are used to kill pests, including insects, rodents, fungi and unwanted plants (weeds).
Pesticides are used in public health to kill vectors of disease, such as mosquitoes, and in agriculture, to kill pests that damage crops.
By their nature, pesticides are potentially toxic to other organisms, including humans, and need to be used safely and disposed of properly”.[46]
Even with regulations, the pesticide limit is passed in some regions.[47]
Authorized pesticides in Germany for example are acetamiprid and chlortuloron.[48]
At first sight pesticides are an ideal second target.
They are present in the water and cause damage to aquatic wildlife.
There are some enzymes capable of degrading or rendering pesticides less toxic.
An example is the nitrile hydratase from Aminobacter.[49]
This nitrile hydratase is capable of degrading sulfoxaflor.
The nitrile hydratase from Ensifera meliloti is able to convert thiacloprid and acetamiprid into their corresponding amids.[50]
Another example is the degradation of diuron.
There are bacteria able to degrade diuron.[51]
Other iGEM projects are using enzymes to degrade pesticides, for example for liuron degradation.[52]
So why did we not use pesticide degradation in our final project?
The answer is quite simple.
Since our project focuses on wastewater treatment plants, there is no real pesticides concentration in wastewater treatment plants.[53]
The only hope left for expanding our degradation substances was pesticides used in cosmetics like malathion.
We even found enzymes able to degenerate malathion like the fungal cutinase from Fusarium oxysporum.[54]
But we could not find sources reporting sufficient malathion concentrations in wastewater.
Therefore, pesticides are a great target for enzymatic degradation, but they do not fit our application and were thus not included.
In view of this it should be noted that the laccase is able to render some pesticides like triclosan less toxic.[55]
Enzyme Immobilization
By immobilization of our transforming enzymes we aim to achieve an increase in stability, transformation efficiency and lifespan of our micropollutant-transforming biofilm.
Immobilization is enabled by fusing our enzymes to the extracellular matrix protein TasA (PDB entry 5OF2) of our biofilm former Bacillus subtilis.
Huang et al. (2018) have described this method for other fusion partners such as mCherry or MHETase and have shown that the proteins remain functional[56].
Figure 9: Enzyme immobilization
By growing the biofilm on so-called floating-bodies we are able to generate a large surface that is in contact with wastewater.
Immobilization of our enzymes in the biofilm’s extracellular matrix therefore drastically increases the processed volume of wastewater
and thus enhances the amount of pollutants that get in contact with the enzymes presented in high density on the biofilm’s extracellular polymeric matrix.
In theory if the enzymes are secreted, they would get washed away by the constant flow of wastewater. Consequently, most enzymes would collide with a very limited number of their target substrates resulting in a very low efficiency.
To compensate this effect a very high expression rate of the enzymes would be required, which would be energetically unfavourable and the bacteria are exposed to high stress leading to reduced growth rates.
Thus, this method would be unfavourable and inefficient compared to the immobilization of the enzymes.
Also, enzyme immobilization can have positive effects on half-life and thermal stability of the proteins[57].
This is especially interesting for EreB since CueO and CotA already possess an extremely high thermal stability more than sufficient for wastewater treatment[58][59].
Additionally, all enzymes profit from increased half-life leading towards a higher number of active enzymes immobilized in our biofilm and consequently higher transformation rates. Cooperative effects in pharmaceutical-transformation can cause an advantage in the catalytic values of a reaction[60].
Also, enzyme immobilization can enhance substrate binding by modifying the enzyme’s orientation, for instance by varying its anchoring point.
In further improvements of our project enzyme immobilization would also allow us to enable multi-step reactions to provide the full degradation of the target molecules through enzyme cascades.
At the moment we are focusing on one-step reactions to transform the target-molecules into less-toxic products. Additional studies on intermediate degradation products and possible enzymes for further processing would be needed.
To validate the most suitable method for enzyme immobilization with TasA, analysis can be carried out equal to the enzyme classification assays described in the pharmaceutical degradation part.
For both laccases either ABTS assays with photometric analysis or diclofenac transformation with HPLC analysis. For EreB Kirby-Bauer assay or HPLC analysis of azithromycin degradation products could be used.
Possible approaches would be TasA fusion at both termini to observe the effect of orientation on the transformation efficiency or variation of the used linker peptide from Huang et al. to other sequences.
Selection of the “right” linker peptide can have many effects like faster folding, increased fusion protein stability, increased activity of enzyme domains and higher expression rates[61].
Therefore, variation of the fusion protein domains holds multiple possible advantages relevant for functionality of our biofilm.
The fusion of our degrading enzymes to extracellular matrix protein TasA offers a great possibility to connect the immobilization with the correct formation of the B. subtilis biofilm.
Also, there should be no negative aspects on biofilm formation since a native protein can be used.
Immobilization offers many advantages like expanded bioactive surface area or the fact that water can flow past the active biofilm while being cleaned up, meaning that the additional layer in wastewater treatment plants can easily be added to the already existing parts.
Experimental Approach
Cloning
The foundation to most promising projects in synthetic biology is the successful implementation of genetic information into the host organism. For our purpose, this covers the optimization of cloning and protein production for our designed fusion proteins.
The microorganism we chose as a host for our project is Bacillus subtilis. For early studies however, we chose to work with Escherichia coli, as it is not only well-researched and exceptionally simple to handle , but it also simplifies high level protein production and characterization . E. coli and B. subtilis differ substantially in that they are gram-negative and gram-positive bacteria respectively. The enzymes utilized in our project are reported in literature to be functional in both types of bacteria [13][62]. For protein production, we opt for the E. coli strain BL21 (DE3), capable of high-level gene expression due to the T7 RNA-Polymerase and the IPTG induction system
As for B. subtilis, we considered the strain GP1672 due to the absence of the tasA and sinR genes or more generic strains for a proof of concept on the expression of TasA fusion proteins.
This approach implies that the plasmid designed for early expression studies and for tests of enzyme functionality must either be compatible with both bacteria, or redesigned respectively. It is of highest importance to check compatibility regarding the use of T7 RNA-Polymerases, the induction system relying on IPTG, antibiotic resistance genes and, to a certain degree, the codon optimization of the genes intended to be implemented. Because of this we codon optimized each gene separately for E. coli and B. subtilis. In addition to that, we checked each gene for unwanted restriction sites within the sequence, removing them, in order to conform to the BioBrick Assembly Standard (RFC10). On the same note we would have added the XbaI, EcoRI, SpeI and PstI to the flanks of our insert to provide compatibility. We decided on the kanR selection marker as it is both universally functional and conveniently available on our plasmid of choice.
Any in silico cloning and its planning was performed using SnapGene®, of which licenses were kindly provided by GSL Biotech LLC for us to use in our project. The plasmids we use rely on the established pET24 vector, into which we planned on cloning enzyme genes or enzyme fusion sequences of either CueO, CotA or EreB via Gibson-Assembly. For the eventual application of CotA in the B. subtilis biofilm as a TasA fusion protein, we would use the plasmid pSEVAeb67rb. For a more detailed explanation on how the plasmids were constructed in silico, please see our lab work masterplan.
The early expression plasmids would carry additional C-terminal Strep- or His-Tags to allow for protein purification and subsequent characterization. This is important to estimate the reliability of diclofenac or azithromycin turnover through our laccases or the erythromycin esterase II.
Analytical Toolbox: ABTS, HPLC, Kirby-Bauer-Assay and LC-MS Analytics
In this section we describe our toolbox of analytical methods we aim to use in practice to characterize our enzymes and to pave the way for targeted enzyme optimization.
ABTS Enzyme Activity Assay
Aims of the experiment
One goal of our project was the recombinant production of the laccase CotA from Bacillus subtilis and CueO from Escherichia coli, as well as the optimization of both enzymes via targeted mutation, to degrade water polluting drugs like diclofenac. The Laccase from T. Versicolor serves us for comparison and as standard enzyme. Essentially, the produced enzyme mutants need to be verified in their enzymatic activity. A widely used assay to determine laccases activity is the ABTS assay, which is described below.
How the ABTS assay work
Laccases are multicopper enzymes belonging to the family of oxidases that catalyse the oxidation of phenolic units in lignin as well as a wide variety of organic compounds, including various types of phenols and aromatic amines. A widely used model substrate for laccases is 2,2′-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid) (ABTS)[63]. The ABTS assay is based on the formation of a stable radical cation by enzymatic oxidation. This reaction can be photometrically measured by its absorbance at 420 nm to determine the enzyme’s activity (see Figure 10).
In our project, we aim to use ABTS in a spectrophotometric assay to determine the activity of our laccases. We therefore, according to A. A. Dias et al. and M. Wu et al., mix the enzyme with 1 mM ABTS in a reaction buffer (100 mM phosphate-citrate-buffer, pH 4.0), incubate at 30°C and photometrically measure the formation of the ABTS radical cation[63,64]. The enzyme activity is defined as the transformation of 1 µmol substrate to the corresponding product per minute (1 unit = 1 μmol/min)[63,64,65].
Figure 10: Enzymatic oxidation of ABTS. A stable radical cation is formed, which can be measured photometrically by the absorption at 420 nm.
What do we get out of it?
The assay serves as a quality control and to define a benchmark of enzymatic activity of our targeted laccases. Consequently, the ABTS assay allows us to compare the activities of production batches and mutated enzymes. The activity is essential for further experiments to optimize the degradation ability of diclofenac. Non-active enzyme most likely also prevents the substrate oxidation. However, we cannot conclude from a mutant’s activity in the ABTS assay on its activity towards diclofenac. Both compounds are structurally very different and thus a mutation benefitting the oxidation of ABTS may demolish the activity on diclofenac. In this case another activity assay is necessary for the optimization of laccases for diclofenac oxidation . For further details on how we would evaluate and handle unexpected results see our engineering success texts. In the next paragraph, we introduce how we aim to establish a high-performance liquid chromatography (HPLC) assay for measuring diclofenac oxidation by laccases.
HPLC Analysis
Aims of the experiment
We aim to use HPLC analysis to determine the enzymatic oxidation and turnover rate of our target substrates e.g. diclofenac to hydroxydiclofenac[66]. In addition, HPLC allows us to detect and verify further emerging transformation products.
What we wanted to achieve & why using HPLC
One of the main topics of our project is to enable and improve the oxidation of diclofenac by the bacterial laccases CueO and CotA. Therefore, we need appropriate analytical methods to evaluate and compare laccase activities.
Our target substance diclofenac is a nonsteroidal anti-inflammatory, small molecule drug. Its lipophilicity (logarithmized n-octanol-water partition coefficient, log Kow = 4.51[66]) makes diclofenac a well-suited analyte for reversed phase high-performance liquid chromatography (RP-HPLC)[66,67]. In RP HPLC the analyte diluted in a solvent (the commonly called mobile phase), in our case diclofenac, runs over a column with a nonpolar surface (the commonly called stationary phase), the typical octadecyl (C18) phase. Consequently, lipophilic substances are interacting more with the stationary phase than hydrophilic substances, and thus each molecule is specifically retarded depending on its lipophilicity. In addition, using solvents of different polarity, respectively, allows better separation of target molecules. For diclofenac, we aim to start with a polar solvent, e.g. water, to run our analyte over the column. In the presence of a polar solvent, lipophilic analytes bind particular well the nonpolar stationary phase. By gradually increasing the ratio of an organic solvent like acetonitrile, polarity of the solvent mixture is decreased. Accordingly, the hydrophobic interaction between substance and stationary phase is also decreased, resulting in the rapid elution of lipophilic substances. Eventually, each substance elutes at a specific time and acetonitrile concentration, depending on its lipophilicity[67]. Pure samples (the commonly called standards) of the expected products are used to determine the specific elution time of the target substance.
After the separation on the column, the substances are detected by an UV-VIS detector, either with fixed(DAD) or variable wavelengths (VWD)[67]. Detection is based on the Lambert–Beer law and results from the different light absorption of the analytes. In our case, the wavelength of maximum absorbance (λmax) for diclofenac is at 340 nm and the measured absorbance correlates directly with the amount of substance[68]. Consequently, with this method and a standard curve we are able to identify our target substance and determined its quantity, and consequently also its turnover by a laccase.
How we want to achieve it – RP HPLC
We developed the following experimental setup to achieve this goal. First, we incubate diclofenac in the presence of our laccases. The reaction conditions are chosen according to the optimal conditions of our laccases described in respective literature (CotA at 60 °C, pH 4.0; CueO at 55 °C, pH 5.5)[7,69]. Controls without enzymes and without substances are performed to ensure that only enzymatic effects are interpreted. After defined time steps, samples are taken and the reaction is quenched by precipitation of the enzyme with methanol. After centrifugation of the quenched samples, the supernatants are transferred into the initial solvent condition used for RP-HPLC analysis.
Prior to that workflow a RP-HPLC method for the target substance diclofenac needs to be established and validated by using bought substrate and product standards. At this step, we also select the appropriate conditions for UV-Vis detection and optimal chromatographic separation. Afterwards, we measured a serial dilution of the bought standards. Thus, we are able to measure the absorbance of known specific concentrations and can create a calibration curve. The calibration curve allows us to calculate the absolute concentrations of diclofenac in our samples. By analysing the amount of oxidized substance at specific time points, it is possible to calculate the enzymatic turnover rate. This rate is then used to identify superior laccase mutants and to rank them by comparing their degradation abilities to a well-known laccase from T. versicolor, which we used as a reference standard.
HPLC analysis allows us to monitor enzymatic degradation of the target substances and their rate. However, with this RP-HPLC method, it is only possible to observe the decrease of the respective substrate and to detect newly emerging compounds. Without known standards, no statement could be made about the chemical composition of these new emerging peaks. We aim to perform mass spectrometry analysis to obtain more information about the resulting transformation products and to compare them with already published structures (see chapter LC MS)[66].
Kirby-Bauer Assay
One of our subprojects was the characterization of an azithromycin-degrading enzyme as well as its optimization by protein engineering. For this purpose, we considered the enzymes erythromycin esterase type I & II (EreA & EreB), which catalyze the hydrolysis of the lactone ring of erythromycin and are known for their promiscuous activity on the erythromycin derivate azithromycin[1,70].
We aim to use the Kirby-Bauer assay, also known as disk or agar diffusion test, for the identification of azithromycin degrading enzymes. For this purpose, the respective enzyme of interest is incubated with a solution containing the macrolide antibiotics erythromycin or azithromycin. The reaction is then quenched by the addition of 50% (v/v) methanol. Due to the relatively high methanol concentration, the enzymes are immediately denatured and are then removed from the solution by centrifugation. The obtained supernatants are then further used for the Kirby-Bauer assay. These reaction mixtures were spotted on sterile paper platelets and incubated on agar plates with fresh cultures of a reporter organism which is sensitive to the antibiotics[71,72]. Consequently, remaining antibiotics from our platelets will diffuse into the agar plate and thereby create a decreasing concentration gradient in the media. The size of the affected area depends on the starting concentration of the antibiotic on the platelet. By the incubation with a microorganism sensitive to the antibiotics, the zone of effective concentrations becomes visible as the growth of microorganisms are inhibited in the presence of high antibiotic concentrations. Large zones of inhibition correspond to low enzyme activity, small up to no zones correspond to high enzyme activity. Since we orientated ourselves on the project of the iGEM team of Munich 2013, we aim to use the same reporter organism Micrococcus luteus. After incubation for 24 h at 24°C, we analyzed the zones of inhibition within the M. luteus culture (see Figure 11)[70]. As positive controls we use platelets soaked with fresh prepared antibiotic, depending on the experiment with azithromycin or erythromycin (platelet C), and platelets soaked with a buffer/methanol (50/50, v/v) mixture as negative control (platelet A)[71,72]. Enzymes with esterase activity on the used antibiotics will partially degrade the antibiotics and the zone of inhibition shrinks compared to the positive control (platelet B).
Figure 11: Schematic illustration of the Kirby-Bauer assay. Macrolide antibiotics containing solutions are incubated with the respective enzyme. Reaction is quenched with 50% (v/v) methanol and the enzyme is removed by centrifugation. Supernatant (platelet B), buffer/methanol (50/50, v/v) as positive control (platelet A) and solution without enzyme (platelet C) are spotted on sterile paper platelets. Platelets are incubated on agar-plates with the reporter organism M. luteus. Large zones of inhibition correspond to low enzyme activity, small up to no zones correspond to high enzyme activity.
The Kirby-Bauer assay allows us to qualitatively test EreA and EreB for their ability to degrade azithromycin and erythromycin. Furthermore, the assay can also be used for screening mutants of our protein engineering endeavor. We therefore start with EreA or EreB and further optimize the enzyme by means of modelling-assisted engineering and by site saturation mutagenesis. In each design-build-test-learn cycle one mutation is introduced, and by iterating the process multiple-point mutants are created.
LC-MS Analysis
When testing the degradation of our targeted pharmaceutical substances, diclofenac and the macrolide antibiotics, we will likely received several unidentified peaks in RP-HPLC analysis. Exemplary, the enzymatic oxidation of diclofenac by laccases can yield various products with the hydroxy diclofenac being the main product[11,66]. These products need to be identified to confirm the correct enzymatic reaction. For this purpose, liquid chromatography-mass spectrometry (LC-MS) is a powerful analytical method. LC-MS allows the precise measurement of the mass-to-charge ratio (m/z) of the occurring products. Based on this mass-to-charge ratio, we can determine the actual molecular masses of the compounds and thus their possible structures. Furthermore, fragmentation experiments can be performed to verify the assumed structures. For this purpose, the measured molecules are transformed into molecule fragments by the energetic impact with inert gas molecules, usually nitrogen. The breaking points of these fragments are usually located at more fragile bonds, e.g. bonds with double bond character, and rarely located at conjugated π-bonds/systems. This method also allows the identification of the initial structure of the measured substance, since the fragments that arise are usually specific to the molecule[73[74].
In addition to the data derived from our methods described above, LC-MS allows us to confirm the actual product formation of antibiotic degradation and diclofenac oxidation and confirms our proof of concept. This would also prove that our methods work and can be further developed for subsequent applications.
Toxicity Assay - Zebrafish Embryo Toxicity Test
One of the main aims of our project is the improvement of the enzymatic degradation abilities of laccases and the implementation of the improved enzyme into a working bacterial biofilm. By this we want to achieve an efficient degradation of problematic and persistent pharmaceutical compounds.
Pharmaceutical drugs such as diclofenac and the antibiotic azithromycin are emerging water contaminants due to the increasing availability and consumption. Those pollutants, which cause harmful impacts on the environment, are hardly biodegradable in sewage treatment plants and therefore represent a special problem of modern society. With our project we aim to relief the aquatic environment by reducing the amount of those substances or by improving their bioremediation using the enzymatic degradation capabilities of laccases[7].
Figure 12: Zebrafish assay By using a Zebrafish Embryo Acute Toxicity Assay, we efficiently prove the decrease of toxicity for aquatic environment. The assay allows measurement of toxicity to a tailored model-organism, without doing actual animal testing
For a target-oriented optimization of our laccases we had to know whether they were actually able to degrade diclofenac and to what extent. Further we needed to ensure, that the occurring transformation products have less or even nontoxic effects at all. Enzymatic transformation products have already been described in the literature. These products were generated by the oxidation of a laccase from Trametes versicolor (Figure 13)[75]. In addition, a lower toxicity of these transformation products has already been described[76]. Figure 13: Diclofenac transformation products The confirmed enzymatic transformation products of diclofenac by the laccase from T. Versicolor are shown
It has been shown that diclofenac can be oxidized by the laccase of the fungus T. versicolor. However, we want to achieve this with a bacterial laccase such as CotA from B. Subtilis and CueO from E. Coli. The use of bacterial enzymes has several advantages, such as better accessibility for optimization and are easier to produce in our host organism. We also want to optimize the whole system by integrating it into a biofilm and make it easier for technical applications.
Relief by Oxidation
We would have proved by previous HPLC
and LC-MS
analysis that our laccases are able to oxidize diclofenac. By these methods we determined the actual degradation and degradation rate of diclofenac by our laccases and identified the emerging transformation products. Now we needed to ensure, that the occurring transformation products have less or even nontoxic effects at all. To achieve this goal, we decided to use a toxicity assay that is directly related to the actual problem, water pollution. The first idea was to use animals like fish as a representative aquatic organism. However, we wanted to use an alternative to animal tests with adult or juvenile fish species. And so, we came up with the zebrafish embryo toxicity test as compromise between animal welfare and scientific benefit[77,78]. Therefore, we wanted to use embryos of the zebrafish (Danio rerio), which are used as an established general model in ecotoxicology and toxicology. Since 2007, this assay is a standardized method for risk assessment of ecotoxic chemicals in Germany (DIN 38 415-6, ISO 15088)[79].
The Assay – to fish or not to fish
The zebrafish embryo toxicity test is designed to determine acute toxicity and teratogenicity of chemicals on embryonic stages of fish.
For this purpose, only fertilized eggs are used for treatment. Twenty embryos are used per concentration of the test substance. Five concentrations are used in each case with a constant distance by a factor of two. An untreated control, a positive control with a fixed concentration of 4 mg/L 3,4 dichloroaniline and a solvent treated control are included. The reference substance used for the positive control, dichloroaniline, is a well-described toxic substance which should result in a minimum mortality of 30 % at the end of the exposure time. These experiments are performed with the original substance and the obtained enzymatic degradation products during the same experimental run[78].
The duration of treatment is 96 hours. Up to four apical observations are recorded every 24 hours as indicators of lethality:
Coagulation of fertilized eggs,
Lack of somite formation,
Lack of detachment of the tail-bud from the yolk sac,
Lack of heartbeat.
For the evaluation of mutagenicity, phenotypic changes such as a crooked tail are also recorded. The embryos are transparent and can therefore be classified well phenotypically.
After the finished treatment, the lethal concentration LC50 is calculated based on a positive outcome in any of the four apical observations recorded. To determine potential mutagenicity, the frequency of phenotypic changes will be considered.
This assay is based on the OECD guidelines for the “Fish Embryo Acute Toxicity Test” as well as the “Fish Embryo Toxicity Assays” of the German Federal Environment Agency[78,80].
The aim of this method was to determine the acute fish toxicity as well as the developmental and reproductive toxicity (DART) of the enzymatic transformation products from wastewater polluting drugs such as diclofenac. The comparison of the product with the substrate provides an evaluation of whether the laccase reduces the toxic impact of these drugs on the aquatic environment.
To guarantee animal welfare, we would not have performed these experiments on our own. We would have been aided by a team of experts from Prof. Boris Schmidt, who would have provided us with the necessary expertise and equipment. The correct performance of the experiments can thus be ensured under the instructions of the supporting team.
The fish tanks used for spawning have been approved by the regional council under examination and compliance with the requirements of the Animal Protection Act, the Hessian regulations and EU directives.
We are aware, that this test still uses animals and have discussed this at length. Nevertheless, we based our decision on the OECD guideline 236 as well as the background paper on fish embryo toxicity assays from the German Federal Environment Agency, which declares that the embryos do not fall under animal welfare regulation up to five days after fertilization, until the onset of independent feeding. Due to their still developing nervous system, it is suspected that the embryos do not suffer any pain. For this reason, the harm is reduced to a minimum[79,80,81].
Thanks to the combination of these analytical and toxicological methods, we would have been able to design the enzyme engineering in a targeted manner.
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[83].
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[84][85].
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.[84]
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 site 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. Also primary lower activity mutants can be considered for combination: In an effect called epistasis mutations can influence the phenotypic effect of other mutations. This way even unfavorable seeming mutants can cause an increase in enzymatic activity by changing the impact of other mutations by specific epistasis.[85]
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. [85][86]
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.[87]
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.[88]
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.[89]
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.[90]
Figure 13: Site directed mutagenesis The plasmid containing the GOI is replicated using a primer pair with a built in mutation to replicate nicked plasmids containing the desired mutations. The plasmids are transformed into competent cells for ligation and expression of the engineered protein. (Click to enlarge)
PCR amplification mainly follows the standard reaction conditions according to the used polymerase, only the number of reaction cycles is lowered and the template concentration is raised.
The annealing temperature is chosen according to the primers' melting temperature and the used DNA polymerase. After PCR amplification the product is digested with DpnI 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.[91]
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.[92] 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[93]. 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.[93]
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[106].
By purification we are able to determine the enzymes reaction kinetics.
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.
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].
Figure 14: Directed Evolution As we approach directed evolution, we're following a repetitive cycle of design-build-test-learn. After E. coli expression and generation of gene library, there would be an assessment of activity which allows us to screen through the generated enzyme variants. If there is a promising candidate, the desired variant would be isolated and the according gene would be amplificated via a mutagenic PCR reaction. The new variants created this way would undergo the same cycle, resulting in a variant with improved properties.
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.
pHOpt = 4.4 (ABTS) / 6.8 (SGZ)
pH is strongly substrate-dependant
Higly stable at pH ranging from 7-9
TOpt = 60°C
Hydrophobic interactions around active-site determine thermo-stability. Substitutions can lead to stability increase (f.e. T232F)
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
kcat/KM = 0.374 s−1 µM−1
18.15% decrease in KM and 1.21-fold increases in kcat
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 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])
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, CAAb
Small changes in the geometry of the Cu sites. Turnover rates highly reduced and optimal pH downshifted 1–2 unit
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.
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
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])
Safety
The application of genetically modified organisms (GMO)
requires particular caution in regard to safety and biocontainment.
For our strategies to guarantee the secure implementation of our
GMO in wastewater treatment plants, please see our quorum sensing dependent
kill switch approach and our text on biosafety.
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[103]. 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[104].
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, 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[105]. 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.
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. 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.
