iGEM 2024 — Team tudarmstadt

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Notebook 📄

iGEM overview

Read our logbook, depicting our whole iGEM journey.

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Wetlab notebooks

Wetlab overview

Read the overview about all our wetlab activities.

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Domestication of gBlocks

Domestication of gBlocks: Learn about how we cloned the gBlocks in plasmid as a preparation for the operon assembly and enzyme production for characterization.

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Pyrene-Operon-Assembly

Pyrene-Operon-Assembly: See how we assembled the operon BBa\_K5043008 with all genes needed for the pyrene degradation.

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Enzyme characterization

Enzyme characterization: Read about our process of characterizing the pdoA2B2-phtAcAd enzyme complex.

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*Pseudomonas* cultivation with PAHs

*Pseudomonas* cultivation with PAHs: See how we produced data showing how *Pseudomonas putida* and *Pseudomonas vancouverensis* deal with exposure to PAHs.

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Immobilization

Immobilization: Read to understand how we got to our preferred immobilization methods.

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Improving PAH-degradation by using the DBTL-Cycle 🧬

To ensure engineering success we followed the Design-Build-Test-Learn Cycle (DBTL-Cycle). This iterative process allows us to improve our genetic and hardware design step by step to enhance degradation of polycyclic aromatic hydrocarbons (PAHs). First, we designed a strategy for a biotechnological cleanup of PAHs in rivers by reading and discussing existing literature. After genetically modifying our chassis to degrade PAHs and modeling the technical components of our project, we precisely analyzed them to learn about their performance and potential issues.

![Figure 1: The Design-Build-Test-Learn (DBTL) Cycle](https://static.igem.wiki/teams/5043/engineering/engineering-fig1-improv2.png)

Figure 1: The Design-Build-Test-Learn (DBTL) Cycle.

Overview

* [Design: Crafting the blueprint](#section1)
* [Build: Putting the device into practice](#section2)
* [Test: Analyzing pyrene degradation plasmid](#section3)
* [Learn: Drawing Conclusions](#section4)
* [Drylab engineering cycle](#section5)
* [References](#section6)

Design: Crafting the blueprint

*Pseudomonas vancouverensis DSM 8368* as chassis

Researching about microbial bioremediation, we quickly came across the *Pseudomonas* genus. Multiple *Pseudomonas* species show very promising abilities as chassis for biodegradation of pollutants [1]. They are known to be stress resistant, providing us with a resilient chassis to modify. Additionally, they show great metabolic diversity and promising abilities to degrade PAHs [2]. Naphthalene degradation, for instance, is studied best in **Pseudomonas** [3]. On top of that, they populate marine and freshwater environments ubiquitously [2, 4], meaning our chassis will be comfortable living in riverine conditions.

A number of *Pseudomonas* species show impressive PAH degradation abilities, like *Pseudomonas fluorescens, putida* or *stuzeri* [5]. We decided to work with *Pseudomonas vancouverensis DSM 8368* (aka. NCIB 9816, formally listed as *Pseudomonas putida*, from now on referred to as *P. vancouverensis*) [6, 7]. *P. vancouverensis* degrades naphthalene so efficiently, that it can even feed on it as sole carbon source. On top of that, it completely metabolizes fluorene and phenanthrene [8]. This makes it attractive as a degrader for a large variety of PAHs, effectively cleaning rivers from multiple pollutants.

As *P. vancouverensis* is a little studied, non-model organism, all experiments were carried out with *Pseudomonas putida KT2440* [9] as well. As *P. putida* is intensively studied as chassis for bioremediation [10], it served to us as a reference organism. We would like to thank Prof. Victor de Lorenzo for kindly providing us this strain.

Introduction of pyrene degradation

As PAHs are highly stable and poorly soluble in water, biodegradation is difficult. The degradation pathway’s initial step incorporates an oxygen molecule into the compound, breaking aromatic resonance and increasing solubility in water. This reaction is catalyzed by a dioxygenase enzyme and subsequently *cis*-dihydro-diol intermediates are created (see Figure 2) [5]. This first step is crucial for biodegradation, as aromatic resonance gets broken, and a more reactive compound is formed. In addition, the product’s higher water-solubility facilitates further degradation [11].

Regarding naphthalene degradation, in this way naphthalene is converted to *cis*-1,2-dihydro-naphthalene-1,2-diol. Next a dehydrogenase creates 1,2-dihydroxynaphthalene. Reaction cascade leads further via salicylate, which is subsequently fully degraded to tricarboxylic acid cycle (TCA) intermediates [5] (see Figure 2).

![Figure 2: naphthalene biodegradation in Pseudomonads](https://static.igem.wiki/teams/5043/engineering/engineering-fig2-2.png)

Figure 2: Naphthalene biodegradation in *Pseudomonads*.

Analogous phenanthrene gets oxidized to *cis*-3,4-dihydro-phenanthrene-3,4-diol first, which is then converted into 3,4-dihydroxy-phenanthrene. Most likely in *Pseudomonas* further oxidation and ring cleavage create 1,2-dihydroxy-naphthalene thus leading into naphthalene degradation pathway. [12, 13] (see Figure 3)

![Figure 3: phenanthrene biodegradation in Pseudomonads](https://static.igem.wiki/teams/5043/engineering/engineering-fig3-2.png)

Figure 3: Phenanthrene biodegradation in *Pseudomonads*.

In this project we introduced a pyrene degradation pathway in *P. vancouverensis* and linked it to its native phenanthrene pathway [14]. Similar to naphthalene- and phenanthrene-pathway, in pyrene-degradation the important initial step creates *cis*-4,5-dihydro-pyrene-4,5-diol [15]. This step is catalyzed by *nidA* and *nidB*, a rieske non-hemen iron ring hydroxylating oxygenase [16]. Though for its activity *phtAc* and *phtAd* from phthalate degradation are needed as extern electron carrier [15]. The pathway proposed in [15] shows further degradation involving one ring-cleavage leading to 3,4-dihydroxy-phenanthrene (see Figure 4).

![Figure 4: Pyrene biodegradation in <i>Mycobacterium vanbaalenii Pyr-1</i>](https://static.igem.wiki/teams/5043/engineering/engineering-fig4-1.png)

Figure 4: Pyrene biodegradation in *Mycobacterium vanbaalenii Pyr-1*.

3,4-dihydroxyphenanthrene is an intermediate of *P. vancouverensis’* native phenanthrene pathway (see Figure 3). In this way pyrene degradation gets channeled into our chassis’ native metabolic pathways. Thereby we only need to genetically introduce nine protein coding sequences forming five enzymes/enzyme complexes. As complete pyrene degradation includes around twenty enzymes [15], this is crucial to make our project feasible and prevent genetic overload.

Genetic design

To express genes necessary for pyrene degradation, we decided to arrange all nine protein coding sequences in one operon. In this way sequence lengths can be shortened, as transcriptional regulators are only needed once. All proteins will be translated from one mRNA-molecule resulting in spatial proximity to each other. This might increase the degradation pathway’s efficiency, as intermediate’s diffusion paths from one enzyme to the other are shrunk [17]. A scheme of the operon is displayed in figure 5.

![Figure 5: Scheme of cloned pyrene degradation operon](https://static.igem.wiki/teams/5043/engineering/engineering-fig5-1.png)

Figure 5: Scheme of cloned pyrene degradation operon.

Protein coding sequences were mainly taken from *Mycobacterium vanbaalenii Pyr-1*, the first bacterium reported to completely break down pyrene [18]. As shown in [15], for some proteins already characterized homologous proteins in other bacteria exist. Therefore some proteins in our pathway were derived from different organisms. We decided to add a hexahistidine-tag to every protein, to enable easy protein production verification via western-blot using just one primary antibody [19].

Initially we planned to use a strong promoter, hoping to thereby get high protein concentrations and hence good degradation rates. Yet Professor Victor de Lorenzo from CSIC advised us to use a low or medium constitutive promoter as PAH concentrations will be low and protein production costs lots of resources for our chassis. Therefore we decided to use Anderson promoter J23110 which shows medium expression in *Escherichia Coli* and *Pseudomonas* [20]. In this way the promoter shall be the rate limiting part for protein production. Hence, we decided to use an RBS showing good translation efficiency, namely BBa\_B0034 [21]. As a terminator we decided to use phage derived Luz7-T50, which shows good efficiency in *E. Coli* and *Pseudomonas* as well [22]. We used pSEVA231 from SEVA-collection as plasmid backbone [23], providing broad-host-range origin pBBR1 and kanamycin resistance marker, as advised by Prof. de Lorenzo. The backbone was generously provided to us by Prof. de Lorenzo. (See our [parts page](https://2024.igem.wiki/tudarmstadt/parts))

Build: Putting the device into practice

Cloning procedure

Genes were ordered from [IDT](https://2024.igem.wiki/tudarmstadt/sponsors). RBS and his-tags were added via overhang-PCR. Iterative assembly was carried out using GoldenBraid method [24]. We used Colabfold as protein structure prediction tool, to evaluate if the his-tag would be best placed at N- or C-terminus, in order to not interfere with protein’s native structure [25].

To make the pSEVA-backbone GoldenBraid-compatible, a DNA-fragment was ordered from IDT carrying promoter, lacZ-cassette, terminator as well as GoldenBraid-BsaI-restriction sides. By cloning of this fragment into the backbone, pSEVA231 served as a pseudo-GB-alpha-vector. A scheme of our cloning procedure is displayed in figure 6.

![Figure 6: Cloning procedure scheme](https://static.igem.wiki/teams/5043/engineering/engineering-fig6.png)

Figure 6: Cloning procedure scheme.

We would like to thank M. Sc. Jacob Mejlsted for helping us planning the cloning procedure.

Test: Analyzing pyrene degradation plasmid

Transformation

Utilizing electroporation, we successfully introduced the ligated SEVA plasmid into *Pseudomonas vancouverensis.* After carefully evaluating various protocols, we selected the most [effective](https://2024.igem.wiki/tudarmstadt/experiments). The transformation was confirmed when the modified bacteria grew on LB medium containing 50μg/ml Kanamycin. As negative control wildtype cells without plasmid were plated out (see Figure 7). In addition, plasmid miniprep and control digest using BamHI was carried out to verify correct plasmid uptake.

![Figure 7: Cells plated out on LB agar plates, 50mg/l Kanamycin](https://static.igem.wiki/teams/5043/engineering/engineering-fig7-2.png)

Figure 7: Cells plated out on LB agar plates, 50μg/ml Kanamycin.

At first, we tested our expression system consisting of promoter J23110, RBS B0034 and terminator Luz7-T50. For this purpose, amilGFP as reporter protein was used (see [BBa\_K5043011](https://parts.igem.org/Part%3ABBa_K5043011)). Protein production could be verified by observing green fluorescence of transformed cell pellets (see Figure 8).

![Figure 8: Cell pellets of bacteria carrying amilGFP expression system under blue light](https://static.igem.wiki/teams/5043/engineering/engineering-fig8-1.png)

Figure 8: Cell pellets of bacteria carrying amilGFP expression system under blue light.

Plasmid characterization

The transformed *Pseudomonads* strains were then characterized regarding their growth, pyrene minimal bactericidal concentration (MBC) and pyrene degradation using HPLC analysis.
 However the HPLC-assay of liquid cultures does not indicate successful pyrene degradation (see Figure 9). For details visit our [results page](https://2024.igem.wiki/tudarmstadt/results).

![Figure 9: Pyrene peak area over incubation time in HPLC assay of liquid cultures](https://static.igem.wiki/teams/5043/engineering/engineering-fig9.png)

Figure 9: Pyrene peak area over incubation time in M9-medium in HPLC assay of liquid cultures.

In addition, growth curves and MBCs for *P. vancouverensis* do not indicate pyrene degradation. In contrast, *P. putida* possessing pyrene degradation plasmid shows significantly higher pyrene and phenanthrene tolerance and grows faster than unmodified bacterium, when pyrene is present (see Figure 10). This could be an indication for pyrene degradation into less toxic intermediates.

![Figure 10: MBC-values and growth rates for <i>P. putida</i> unmodified and transformant](https://static.igem.wiki/teams/5043/engineering/engineering-fig10-1.png)

Figure 10: MBC-values and growth rates for *P. putida* unmodified and transformant.

Pyrene degradation could not be verified in our engineered strains. However, *P. putida* shows increased tolerance to pyrene and phenanthrene. This could indicate, our plasmid’s enzymes degrade phenanthrene and pyrene into less toxic intermediates in *P. putida.* Some of our chosen proteins can use a variety of PAHs as substrate [16], which could explain the increased phenanthrene tolerance.

Learn: Drawing conclusions

We clearly need to characterize our engineered strains in more detail. At first a reliable verification of the production of all nine proteins is necessary, for example using western blot. Second, more reliable pyrene degradation assays are needed, as our method showed great fluctuations in data. Cultures are also required to be incubated longer, to be sure whether pyrene is degraded or not. In addition, characterization of all our used enzymes in vitro is necessary to verify the integrity of the pyrene pathway.

Drylab engineering cycle

Introduction

At the heart of our project lies the genetic engineering of *P. vancouverensis* to successfully degrade PAHs. However, for our advanced bacteria to work effectively against PAH pollution in rivers, it needs to be brought in close contact with as many PAHs as possible. To achieve this, the drylab team worked alongside the wetlab team throughout the project to develop an effective platform for a successful *P. vancouverensis* deployment in polluted areas.
One of the main concerns when working on the deployment of GMOs is to comply with all regulations at hand and avoid an uncontrolled release of the GMOs. To achieve a strong degree of containment of the GMOs combined with a high level of exposure to the contaminated water, we came up with the idea to immobilize *P. vancouverensis* on the inner surface of a specialized device, which could be deployed at suitable sites in rivers, such as bridges, sluices or buoys.

From the beginning, it was clear to us that the interior design of the device, especially the interaction area of our chassis, would play a major role in the effectiveness of our solution.

First DBTL-cycle iteration

### Design and conceptualize: The membrane concept - a platform combining filtering and degradation of PAHs

The first idea that we got to work featured a membrane within the device. On this membrane, *P. vancouverensis* would be immobilized. It would also be able to filter the oil-micelles out, letting them accumulate on the same side of the membrane where *P. vancouverensis* would be placed, while letting other particles in the water pass through the entire system. This way, we could both separate the PAHs from the water and degrade them at the same place and time.

### Present and learn: Receiving input from experts at BFH Meetup and scrapping the membrane concept

We presented this design at the BFH Meetup in Bielefeld, where the jury and other experts were not convinced that this system could work. They saw the main problems in finding or developing a suitable membrane as well as in not clogging our system with too many PAHs. They instead encouraged us to look for a solution without a membrane, focusing more on water flow and the maximization of surface area inside the device. After further research into this and the uncertainty of how the immobilization could work on a membrane in the end, we decided to scrap the membrane and work on new concepts.

Second DBTL-cycle iteration

### Design: Labyrinths, Pillars and Balls - maximizing contact efficiency

We came up with three base configurations for an updated interior design of the device. One featured a labyrinth (see Figure 11). The focus of this design was really to get the maximum surface area out of the small inside of our device. If all sides of such a labyrinth were covered in silica, *P. vancouverensis* would be immobilized by the non-polar interactions of its membranes with the surface. Using the labyrinth structure we could reduce the probability of PAHs flowing through the entire system without ever encountering a bacterium. This would make for a very efficient design.

A second design featured large pillars filling the inside of the device (see Figure 12). *P. vancouverensis* would in this case be immobilized on the surface of these pillars. We planned on finding the best configuration of pillars through simulation, to make sure that the maximum amount of water would come in contact with one of the pillars at some point. An advantage of this system over the labyrinth would be the much smaller amount of forces the water would exert onto the interior of the device, since the water from the river would not be slowed as much through the pillars as through the harsh turns of a labyrinth.
The third design was based on small beads/balls containing *P. vancouverensis* (see Figure 13). When researching methods for immobilization, we discovered that many common methods result in permeable beads filled with bacteria. These beads would be free to move around inside the device. The creation of such beads is relatively simple and cost efficient. The materials used to create the alginate beads that turned out to be sufficient for this use case, are stable and non-toxic (learn more in [results](https://2024.igem.wiki/tudarmstadt/results)). A big advantage of this design would be the modularity of it. One could simply open the box and replace broken balls. However, it was unclear how we could stop the balls from simply getting pushed to the end of the box and being stuck there.

![](https://static.igem.wiki/teams/5043/model/model-fig1-1.png)

Figure 11: Labyrinth structure.

![](https://static.igem.wiki/teams/5043/model/model-fig2-1.png)

Figure 12: Pillar structure.

![](https://static.igem.wiki/teams/5043/model/model-fig3-1.png)

Figure 13: Ball structure.

### Build: Creating 3D-models and printing a prototype

Before we could start our simulations to test the viability of our ideas, we designed them using common computer aided design (CAD) software. The resulting 3D-models were not only the basis for the following simulations, but also gave us the opportunity to 3D-print our first prototype models to learn more about the sizing of our device.

Test and Learn: Simulating the flow and gaining valuable insight

After designing all three structures virtually, we proceeded with a [flow analysis simulation](https://2024.igem.wiki/tudarmstadt/model). In the simulations we focused mainly on the velocity and pressure of the fluid traversing the device. A low velocity at the contact points with the surface would be ideal for *P. vancouverensis* to work most efficiently. A constant decrease in pressure would then secure an optimal continuous flow throughout the device.
While the simulations were running, the different methods for immobilization were tested in the laboratory. As the method of immobilizing bacteria on a silica surface was verified, as well as the method of enclosing the bacteria in alginate beads, all three models would work in terms of biological feasibility (see our [results page](https://2024.igem.wiki/tudarmstadt/results)). Therefore, we could rely fully on flow analysis to show us which design would turn out optimal for our device.
Through the simulations we found that the ball design works best for our application. It also features to easy accessibility and replaceability of the beads containing the GMOs. This means that applications for other bacteria, targeting other riverine contaminations, could easily be installed as well. Through these advantages we concluded that the balls make for the best interior design of our device.

Our approach to a real-life flow simulation with a simple 3D-printed model of the pillar design. To make the path the water takes through the box visible, we used fluorescein.

References

[1] E. Martínez-García and V. de Lorenzo, "Pseudomonas putida as a synthetic biology chassis and a metabolic engineering platform," *Current opinion in biotechnology*, vol. 85, p. 103025, 2024, doi: 10.1016/j.copbio.2023.103025.

[2] D. C. Volke, P. Calero, and P. I. Nikel, "Pseudomonas putida," *Trends in microbiology*, vol. 28, no. 6, pp. 512–513, 2020, doi: 10.1016/j.tim.2020.02.015.

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[4] C. Valenzuela, V. L. Campos, J. Yañez, C. A. Zaror, and M. A. Mondaca, "Isolation of arsenite-oxidizing bacteria from arsenic-enriched sediments from Camarones river, Northern Chile," *Bulletin of environmental contamination and toxicology*, vol. 82, no. 5, pp. 593–596, 2009, doi: 10.1007/s00128-009-9659-y.

[5] B. Mohapatra and P. S. Phale, "Microbial Degradation of Naphthalene and Substituted Naphthalenes: Metabolic Diversity and Genomic Insight for Bioremediation," *Frontiers in bioengineering and biotechnology*, vol. 9, p. 602445, 2021, doi: 10.3389/fbioe.2021.602445.

[6] DSMZ, *Pseudomonas vancouverensis.* [Online]. Available: https://www.dsmz.de/collection/catalogue/details/culture/DSM-8368 (accessed: Aug. 20 2024).

[7] W. W. Mohn, A. E. Wilson, P. Bicho, and E. R. Moore, "Physiological and phylogenetic diversity of bacteria growing on resin acids," *Systematic and applied microbiology*, vol. 22, no. 1, pp. 68–78, 1999, doi: 10.1016/S0723-2020(99)80029-0.

[8] Y. Yang, R. F. Chen, and M. P. Shiaris, "Metabolism of naphthalene, fluorene, and phenanthrene: preliminary characterization of a cloned gene cluster from Pseudomonas putida NCIB 9816," *Journal of bacteriology*, vol. 176, no. 8, pp. 2158–2164, 1994, doi: 10.1128/jb.176.8.2158-2164.1994.

[9] M. Bagdasarian *et al.,* "Specific-purpose plasmid cloning vectors. II. Broad host range, high copy number, RSF1010-derived vectors, and a host-vector system for gene cloning in Pseudomonas," *Gene*, vol. 16, 1-3, pp. 237–247, 1981, doi: 10.1016/0378-1119(81)90080-9.

[10] Z. Zuo *et al.,* "Engineering Pseudomonas putida KT2440 for simultaneous degradation of organophosphates and pyrethroids and its application in bioremediation of soil," *Biodegradation*, vol. 26, no. 3, pp. 223–233, 2015, doi: 10.1007/s10532-015-9729-2.

[11] T. Pantsyrnaya *et al.,* "Biodegradation of Phenanthrene by Pseudomonas putida and a Bacterial Consortium in the Presence and in the Absence of a Surfactant," *Indian journal of microbiology*, vol. 52, no. 3, pp. 420–426, 2012, doi: 10.1007/s12088-012-0265-z.

[12] W. C. EVANS, H. N. FERNLEY, and E. GRIFFITHS, "OXIDATIVE METABOLISM OF PHENANTHRENE AND ANTHRACENE BY SOIL PSEUDOMONADS. THE RING-FISSION MECHANISM," *The Biochemical journal*, vol. 95, no. 3, pp. 819–831, 1965, doi: 10.1042/bj0950819.

[13] S. Gao, J.-S. Seo, J. Wang, Y.-S. Keum, J. Li, and Q. X. Li, "Multiple degradation pathways of phenanthrene by Stenotrophomonas maltophilia C6," *International biodeterioration & biodegradation*, vol. 79, pp. 98–104, 2013, doi: 10.1016/j.ibiod.2013.01.012.

[14] EPA, *Priority Pollutant List.* [Online]. Available: https://www.epa.gov/sites/default/files/2015-09/documents/priority-pollutant-list-epa.pdf (accessed: Aug. 21 2024).

[15] S.-J. Kim, O. Kweon, R. C. Jones, J. P. Freeman, R. D. Edmondson, and C. E. Cerniglia, "Complete and integrated pyrene degradation pathway in Mycobacterium vanbaalenii PYR-1 based on systems biology," *Journal of bacteriology*, vol. 189, no. 2, pp. 464–472, 2007, doi: 10.1128/JB.01310-06.

[16] O. Kweon, S.-J. Kim, J. P. Freeman, J. Song, S. Baek, and C. E. Cerniglia, "Substrate specificity and structural characteristics of the novel Rieske nonheme iron aromatic ring-hydroxylating oxygenases NidAB and NidA3B3 from Mycobacterium vanbaalenii PYR-1," *mBio*, vol. 1, no. 2, 2010, doi: 10.1128/mBio.00135-10.

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[18] A. A. Khan, S.-J. Kim, D. D. Paine, and C. E. Cerniglia, "Classification of a polycyclic aromatic hydrocarbon-metabolizing bacterium, Mycobacterium sp. strain PYR-1, as Mycobacterium vanbaalenii sp. nov," *International journal of systematic and evolutionary microbiology*, vol. 52, Pt 6, pp. 1997–2002, 2002, doi: 10.1099/00207713-52-6-1997.

[19] W. N. Burnette, ""Western blotting": electrophoretic transfer of proteins from sodium dodecyl sulfate--polyacrylamide gels to unmodified nitrocellulose and radiographic detection with antibody and radioiodinated protein A," *Analytical biochemistry*, vol. 112, no. 2, pp. 195–203, 1981, doi: 10.1016/0003-2697(81)90281-5.

[20] A. N. Pearson *et al.,* "The pGinger Family of Expression Plasmids," *Microbiology spectrum*, vol. 11, no. 3, e0037323, 2023, doi: 10.1128/spectrum.00373-23.

[21] S. G. Damalas, C. Batianis, M. Martin-Pascual, V. de Lorenzo, and V. A. P. Martins Dos Santos, "SEVA 3.1: enabling interoperability of DNA assembly among the SEVA, BioBricks and Type IIS restriction enzyme standards," *Microbial biotechnology*, vol. 13, no. 6, pp. 1793–1806, 2020, doi: 10.1111/1751-7915.13609.

[22] E.-M. Lammens, L. Putzeys, M. Boon, and R. Lavigne, "Sourcing Phage-Encoded Terminators Using ONT-cappable-seq for SynBio Applications in Pseudomonas," *ACS synthetic biology*, vol. 12, no. 5, pp. 1415–1423, 2023, doi: 10.1021/acssynbio.3c00101.

[23] E. Martínez-García *et al.,* "SEVA 4.0: an update of the Standard European Vector Architecture database for advanced analysis and programming of bacterial phenotypes," *Nucleic acids research*, vol. 51, D1, D1558-D1567, 2023, doi: 10.1093/nar/gkac1059.

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[25] M. Mirdita, K. Schütze, Y. Moriwaki, L. Heo, S. Ovchinnikov, and M. Steinegger, "ColabFold: making protein folding accessible to all," *Nature methods*, vol. 19, no. 6, pp. 679–682, 2022, doi: 10.1038/s41592-022-01488-1.

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Experiments🧪

Experiments

See wetlab protocols we used during our iGEM project

Read protocols

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Achievements 🎯

In our project we aimed to engineer *Pseudomonas vancouverensis* to fully biodegrade pyrene. This is, what we
achieved:

**- Cloning, transformation and characterization of pyrene degradation plasmid
- Characterization of important pyrene degradation enzymes
- Test and comparison of different immobilization methods
- Flow analysis simulation to optimize our device**

Read our results in detail:

Overview

* [Implementation of pyrene degradation](#section1)
* [*Pseudomonas* profiling](#section2)
* [Pyrene degradation HPLC assay](#section3)
* [*phtAcAd* characterization](#section4)
* [*pdoA2B2, phtAcAd* HPLC analysis](#section5)
* [Immobilization](#section6)
* [Future plans](#section7)
* [References](#section8)

Implementation of pyrene degradation

We plan to enable our chassis *Pseudomonas vancouverensis DSM8368* to degrade pyrene. For this purpose,
we put the nine proteins necessary to channel pyrene into its native phenanthrene pathway, under medium level
constitutive expression. To achieve this, at first, we needed to verify our expression system. We did so by
introducing amilGFP as reporter protein, regulated by our chosen promoter J23110, RBS B0034 and terminator
Luz7-, forming [BBa\_K5043011](https://2024.igem.wiki/tudarmstadt/parts), into our chassis. For test
amilGFP coding plasmid was also introduced into *P. putida KT2440*. Cell pellets of transformant were
analyzed under blue light (see Figure 1).

![](https://static.igem.wiki/teams/5043/engineering/engineering-fig8-1.png)

Figure 1: Transformed bacteria under blue light.

As clearly visible in the pictures, green fluorescence can be observed for cells carrying
amilGFP-coding-plasmid. This indicates that our chosen expression system works both in *P. putida*
as well as *P. vancouverensis*These results also prove our transformation method succesfull.

We then cloned an operon coding for all nine necessary enzymes into backbone pSEVA231 [1] using GoldenBraid
[2] method and verified it by sequencing. It was transformed into *P. putida*and *P.
vancouverensis* by electroporation. Correct plasmid uptake was in addition verified by plasmid
miniprep and control digest using BamHI (see Figure 2).

![](https://static.igem.wiki/teams/5043/results/results-fig2-1.png)

Figure 2: Control-digest of pyrene-degradation plasmid, miniprepped from *E. Coli DH5α* and
our two chassis.

*Pseudomonas* profiling

Growth curves

### Description

To better understand the growth dynamics of *P. putida* and *P. vancouverensis*, OD600 measurements were
plotted against the time. During the growth curve three key points were plated out to obtain viable colony
counts, as seen in the standard curve in Figure 4 and 5. This data helps to establish a more accurate
correlation between OD600 and colony forming unit (CFU), allowing for better prediction of cell density
during future experiments. For *P. vancouverensis*, a non-model organism with limited available data,
these results are particularly interesting and valuable as they provide crucial insights into the growth
characteristics of the strain.

### Results

![](https://static.igem.wiki/teams/5043/results/results-fig3.png)

Figure 3: Growth rate comparison of *P. putida* and *P. vancouverensis* in LB
medium. Incubation at 28°C and 180 rpm.

![](https://static.igem.wiki/teams/5043/results/results-fig4.png)

Figure 4: Growth and standard curve for *P. vancouverensis* unmodified. Incubation
at 28°C and 180 rpm.

![](https://static.igem.wiki/teams/5043/results/results-fig5.png)

Figure 5: Growth and standard curve for *P. putida.* Incubation at 28°C and 180 rpm.

The growth dynamics in Figure 4 and 5 of *P. putida* and *P. vancouverensis* were analyzed using
regression formulas to estimate colony-forming units (CFU) per milliliter based on optical density (OD600). For
*P. putida*, the regression formula of 4×1011 x indicates that at an OD of 0.1, the CFU count is
approximately 4×1010 CFU/ml. Similarly, for *P. vancouverensis*, the regression formula of
2×107 x results in a CFU count of 2×106 CFU/mL at an OD of 0.1.
In terms of growth rates, *P. putida* exhibited a growth rate (µ) of 0.63 1/h, with a generation time of
1.10h hours (66 minutes) in exponential phase. Comparatively, *P. vancouverensis* shows a slightly lower
growth rate of 0.59 1/h and a longer generation time of 1.17 hours (70 minutes). Additionally, *P.
vancouverensis* showed an extended lag phase.

### Discussion

While *P. putida* exhibits a slightly faster growth rate, the differences to *P. vancouverensis* do
not affect experimental outcomes in most cases. Both strains could be used effectively for various
biotechnological and environmental applications, with strain choice depending more on specific needs rather than
a significant difference in growth performance.
*P. vancouverensis* performs comparably well after a longer lag phase. However, *P. putida* has an
advantage in processes requiring quicker biomass accumulation with a CFU/ml count that is 1000-fold higher than
for *P. vancouverensis.* While cell size differences could contribute to variations in CFU counts, since
smaller cells would result in more cells per unit volume at the same OD600 [3], this is less likely because both
species belong to the same genus (*Pseudomonas*), suggesting comparable cell sizes which would not result
in such a high difference. It seems more likely that *P. vancouverensis* experiences a higher rate of cell
death compared to *P. putida*, resulting in lower CFU/ml count.
Nonetheless, the measurement of CFU/mL is cumbersome, and various factors contribute to variability in CFU
counts. This is reflected in the correlation coefficients, which is 0.916 for *P. vancouverensis* and 0.726
for *P. putida*. The greater variability is likely due to pipetting errors during the serial dilution,
particularly for *P. putida* a plating between the dilutions 10-8 and 10-12 was required, meaning more
dilution steps. Each additional step increases the chance for errors. As seen in the colony counts for both
strain, these were not consistent with the expected values based on the dilution factor. Furthermore, at higher
CFU the software OpenCFU [4] was used which introduces additional variability, because the threshold
for identifying a colony introduced another error factor.
Overall, despite having similar growth rates, *P. putida* exhibited a significantly higher CFU count than
*P. vancouverensis*. This difference is due to greater cell viability in *P. putida*, whereas *P.
vancouverensis* experienced higher cell death, contributing to optical density without forming viable
colonies. Despite variability introduced by factors like pipetting errors withing the dilution step an overall
picture for both bacteria, especially for *P. vancouverensis* could be taken.

Pyrene tolerance

The OD600 measurements with different pyrene concentration were plotted over incubation time for *P. vancouverensis* and
*P.
putida* including the unmodified and the engineered strain bearing the pyrene degradation plasmid. The
primary objective was to compare the growth rates in the presence of pyrene, determining if the plasmid enhances
the strain's tolerance to pyrene. Furthermore, the minimal bactericidal concentration (MBC) was determined.

[ ]
*P. vancouverensis DSM8368*

![](https://static.igem.wiki/teams/5043/results/results-fig6.png)

Figure 6: Growth rate comparison of *P. vancouverensis* unmodified and possesing pyrene
degradation plasmid. Incubation at 180 rpm, 28°C in LB media with different pyrene concentrations.

![](https://static.igem.wiki/teams/5043/results/results-fig7.png)

Figure 7: Growth curve of *P. vancouverensis* unmodified at 3 and 4g/L pyrene in LB
Medium. Incubation at 180 rpm, 28°C.

![](https://static.igem.wiki/teams/5043/results/results-fig8.png)

Figure 8: Growth curve of *P. vancouverensis* with pyrene degradation plasmid at 0, 3
and
4g/L Pyrene in LB Medium. Incubation at 180 rpm, 28°C.

### Result:

Growth curves of *P. vancouverensis* in LB medium containing pyrene are displayed in Figure 7 and 8. In
Figure 6 growth rates of *P. vancouverensis* unmodified and transformant are compared. Time error of 3min
was
assumed. For OD600 error of 0.05 was assumed, corresponding to standard deviation observed in sterile
control.

### Discussion

Due to the high variability observed between the growth rates, no definitive conclusion can be drawn regarding
the
pyrene degradation capability of the transformed strain. This variability suggests that the observed increase in
growth between 0 g/L and 3 g/L pyrene for the *P. vancouverensis* transformant may not be reliable or
conclusive.
However, some interesting assumptions for *P. vancouverensis* can be made: The unmodified strain exhibited
a
significantly higher growth rate at 3 g/L compared to 0 g/L, which could suggest that its native metabolic
pathways might be involved in the degradation of pyrene. Previous studies have shown that enzymes responsible
for
the breakdown of certain PAHs may also be capable of degrading structurally similar compounds, [5, 6]
Additionally
this would also indicate potential co-metabolization of pyrene, where pyrene is used additional to the nutrients
provided by the LB medium [7], which may resulted in a higher biomass and thus growth rate, as we [simulated](https://2024.igem.wiki/tudarmstadt/model). Nonetheless, these theories remain speculative,
and given the lack of direct
evidence for pyrene degradation, the higher growth rate is more likely due to pipetting errors or inconsistencies
in the experimental setup
At a pyrene concentration of 4 g/L, the growth rate of both strains declined compared to 3g/L, indicating a
potential inhibitory effect of pyrene at higher concentrations. To further investigate this decline, the minimal
bactericidal concentration (MBC) of pyrene was determined to show the concentration threshold where pyrene
inhibits the growth.

Table 1

Compound

MBC
*P. vancouverensis* unmodified [g/L]

MBC
*P. vancouverensis* transformed [g/L]

Phenanthrene

5

5

Pyrene

3

3

*Table 1: Comparison of MBC values of *P. vancouverensis* unmodified and transformed for
phenanthrene and pyrene after 16h incubation.*

Different pyrene concentrations in LB medium, derived from a pyrene-isopropanol stock solution, were incubated for approximately
16
hours, followed by plating on LB Agar to assess viable cell counts. At a concentration of 3 g/L pyrene, no
viable
cells were observed. This supports the observed decline in growth rate at 4 g/L, as pyrene concentrations at or
above the minimal bactericidal concentration of 3 g/L leads to toxic effects that impair the metabolic activity
of
both strains, resulting in complete cell death under our experimental conditions (after 16 hours). Prior to cell
death, a reduced growth rate is observed due to increased cell death rate and/or reduced reproductive capacity. That the strain growed at 4 g/l pyrene despite determined MBC of 3g/l could be due to toxic effects of isopropanol which could have resulted in lower MBC
values,
whereas in LB Medium pyrene was added directly as a solid. Additionally, no defined CFU was inoculated for the
determination of the MBC values. A higher CFU density could have contributed to increased pyrene tolerance. All
in
all, no statement about the degradational capabiltiy of *P. vancouverensis* transformant could be made. Thus a
comprehensive analysis using high-performance liquid chromatography (HPLC) will be conducted. This analysis will
allow for the precise quantification of pyrene and its degradation over time.

[ ]
*P. putida* KT2440

![](https://static.igem.wiki/teams/5043/results/results-fig9.png)

Figure 9 growth curve of *P. putida* unmodified and with the pyrene-degradation plasmid,
1mg/L
pyrene in LB Medium. Incubation at 28°C and 180 rpm.

Table 2

Compound

MBC
*P. putida* unmodified [g/L]

MBC
*P. putida* transformed [g/L]

Phenanthrene

2

5

Pyrene

<0.0001

2

*Table 2: Comparison of MBC values of P. putida modified and transformed for phenanthrene and
pyrene after 16h incubation*.

In Figure 9 the growth behavior of unmodified and transformed *P. putida* strain in 1mg/L pyrene is
plotted.
Errors were assumed as above. The growth rate for the unmodified *P. putida* is at 0.25±0.02 1/h and for
the
transformant at 0.36 ±0.05 1/h.

### Discussion

The growth rate of the engineered *P. putida* strain is 28% higher compared to the unmodified *P.
putida* in 1mg/L pyrene, suggesting a higher tolerance to pyrene. This could also be seen at the MBC value
for pyrene where initially even at pyrene concentration of 0.1mg/L no cell viability after an incubation of 16h
could be seen. In contrast, the engineered strain tolerates concentrations up to 1g/L of pyren. The growth observed
in
the unmodified strain at 1 mg/L pyrene, despite its lower tolerance indicated by the MBC test, migth have the same
reasoning as above (variable CFU and the isopropanol solution). Interestingly, the engineered strain also
demonstrated an increased MBC value for phenanthrene from 2g/L (unmodified) to 5g/L. This could be due to pyrene degradation enzymes also breaking down or detoxifying phenanthrene [16].
Overall, the higher growth rate and MBC values, indicate a greater tolerance to pyrene in the engineered strain.
This enhanced tolerance could be attributed to the transformant's ability to degrade pyrene, due to the
introduction
of pyrene degradation pathway indicating a sucessful pathway design.

Pyrene degradation HPLC assay

### Description

To test pyrene degradation abilities of transformed *P. vancouverensis* and *P. putida*, HPLC-analysis
of bacterial cultures determining pyrene concentration was carried out.

### Method

Liquid cultures of *P. vancouverensis* and *P. putida* in LB and M9 media with different pyrene
concentrations were incubated at 28°C, 180 rpm. Probes were taken regularly and centrifuged. As pyrene is poorly
water-soluble [8], most of it should be found in pellets after centrifuging. Cell pellets were resuspended in
dimethyl sulfoxide (DMSO) and cells were lysed using an ultra sonic device. Lysed cells were centrifuged again, and
supernatant was applied to Zorbax SB-C18 HPLC column.

### Results

Pyrene showed 7.8 to 8.1 minutes retention time on HPLC column. Pyrene peak was best visible at 270nm
absorbance. In Figure 10 and 11 results of HPLC-analysis are shown. Pyrene A270 peak area correlating to pyrene
concentration in probe is plotted against incubation time. A time error of 1h is assumed. For pyrene peak area
an error of 10% as result of pipetting errors is assumed.

![](https://static.igem.wiki/teams/5043/results/results-fig10.png)

Figure 10: Pyrene peak area over incubation time is shown for liquid cultures in LB-medium. *P.
putida* was incubated with 1mg/l pyrene, *P. vancouverensis* with 3g/l pyrene.

![](https://static.igem.wiki/teams/5043/engineering/engineering-fig9.png)

Figure 11: Pyrene peak area over incubation time is shown for M9-medium cultures with 100mg/l
pyrene.

HPLC-analysis for *P. putida* in LB-medium indicates constant pyrene concentration in media. *P.
vancouverensis* analysis shows also constant pyrene concentration for transformed bacteria but great
fluctuations for unmodified control. Both exhibit unusual high peak area (>3500mAU\*s) after 21h of incubation
(not shown in diagram).
All probes in M9-medium show initial pyrene decrease which is followed by constant pyrene amounts. Data for
unmodified *P. putida* strain shows great fluctuations.

### Discussion

Data cannot provide proof of pyrene degradation for any of the examined bacterial cultures. Pyrene decrease in
all M9-probes including sterile-control could indicate spontaneous decomposition of pyrene. Though this seems
unlikely as pyrene is thermodynamically stable [9] and decline could not be observed in LB-cultures.
It stands out, that especially graphs for *P. vancouverensis* in LB medium and unmodified *P. putida*
in M9 medium show great fluctuations, beyond assumed error tolerances. This most likely indicates that the
chosen extraction method does not work quantitatively reliable. This could also explain, why pyrene peak area
does not correlate well with pyrene concentration in medium.
In summary, data suggests that no bacterial strain, neither unmodified nor bearing pyrene degradation plasmid,
is capable of degrading pyrene. Therefore graphs in all cultures, except *P. vancouverensis* in LB-medium,
show approximately constant pyrene concentrations. This could be due to several reasons.
At first, due to time constraints, we were not able to prove protein expression by transformed strains. As
amilGFP was successfully produced using the same expression system, most likely at least some of pyrene
pathway’s proteins get produced. Yet there is no proof, the whole operon gets expressed.
In addition, as mentioned earlier, pyrene is poorly water soluble (135μg/l at 25°C) [8]. This means only few
supstrate will be available for degradation resulting in overall slow pyrene degradation, so that no pyrene
decrease could be observed during our chosen incubation times. In this case, production of biosurfactants
enhancing PAH-solubility like shown by 2015 iGEM-team from Uppsala, could be helpful [10].
Furthermore, cellular uptake of pyrene could also be a problem. As all expressed degradation enzymes are
cytosolic, pyrene is required to pass bacterial cell membrane. As pyrene shows high octanol-water partition
coefficient [8], it can be suspected to diffuse passively through membranes. However for naphthalene active
uptake by *Pseudomonas fluorescence* has been reported [11], indicating an active PAH import system could
be necessary for efficient degradation.

In conclusion, most data shows that the transformed bacteria are not capable of degrading pyrene. Only growth curves and MBCs of transformed *P. putida* indicate a higher pyrene tolerance. This could be due to pyrene degradation or at least detoxification. In summary further analysis, improvements and iterations of the DBTL-Cycle are necessary to implement successful pyrene degradation.

*phtAcAd* characterization

To get more insight into pyrene degradation, we decided to characterize some of our pathway’s enzymes *in vitro*. It is established that [*phtAc*](https://parts.igem.org/Part%3ABBa_K5043009)  and [*phtAd*](https://parts.igem.org/Part%3ABBa_K5043010), as electron transfer components, along with a ring-hydroxylating dioxygenase system formed by [*pdoA2*](https://parts.igem.org/Part%3ABBa_K5043006)  and [*pdoB2*](https://parts.igem.org/Part%3ABBa_K5043007), aggregate to create a complex exhibiting dioxygenase activity. [13–15] This complex is capable of converting phenanthrene-4-carboxylate, an intermediate product of pyrene degradation, into *cis*-3,4-dihydroxy-phenanthrene-4-carboxylate (see Figure 12) [14, 15]. Given this reaction’s importance in pyrene degradation, we opted to conduct a more detailed characterization. The components of the electron transport chain, ferredoxin *phtAc* and ferredoxin reductase *phtAd*, were produced and purified to elucidate their kinetic parameters. Subsequently, we aimed to analyze the complex formation with *pdoA2B2* through HPLC analysis.

![](https://static.igem.wiki/teams/5043/registry/bba-k5043010-fig1.png)

Figure 12: Enzymatic reaction of *phtAcAd-pdoA2B2* complex [16].
pdoA2B2 dimer exhibits catalytic side, which attacks substrate. phtAcAd transport two electrons from NADH to active side.

Enzyme production and purification

Enzyme coding sequences were cloned into pQE bacterial expression vector with a N-terminal, hexahistidine tag. Proteins were expressed in *E. coli* BL21 (DE3). Main cultures were incubated at 37°C until an OD600 of 0.5 was reached. Subsequently, cultures were induced with a final concentration of 0.5 mM IPTG and incubated overnight at 30°C. Purification was performed using Immobilized Metal Affinity Chromatography (IMAC). Both production and purification samples were analyzed via SDS-PAGE and Coomassie staining (data not shown). Enzyme concentration was determined using the Bradford Assay [17].

Kinetic characterization of *phtAd*

The characterization of *phtAd* activity was performed by evaluating its ability to reduce 2,6-dichlorophenolindophenol (DCPIP), which serves as an electron acceptor [16]. In its oxidized form, DCPIP displays a blue color, which transitions to colorless upon reduction, facilitating the evaluation of *phtAd* activity (see Figure 13). The decrease in absorbance was monitored at 600nm [16].

![](https://static.igem.wiki/teams/5043/registry/bba-k5043010-dcpip-2.png)

Figure 13: Reduction of DCPIP catalyzed by *phtAd*

The activity of *phtAc* was assessed using a coupled assay. The increase in absorbance resulting from an electron transfer to cytochrome c signifies an interaction between ferredoxin *phtAc* and ferredoxin reductase *phtAd*. The increase in absorbance was monitored at 550nm. Both reactions needed NADH and FADH as cofactors [16].

### Method

The optimal reaction time and enzyme concentration for the assay were determined by measuring the decrease in absorbance at 2-minute intervals over a 30-minute period. This approach was necessary due to the uncertainty regarding the concentration at which the reaction would proceed and the duration for which the enzyme could effectively catalyze the reaction before reaching substrate saturation. Following the determination of these parameters, the reaction was assessed using varying concentrations of NADH while maintaining a constant enzyme concentration to determine the kinetic parameters. The reaction was initiated by the addition of a specified amount of *phtAd* and monitored at 600nm using a UV/Vis spectrometer.

### Results

**Determination of the optimal enzyme concentration**
Initial optical analysis indicates that the assay conducted with the highest concentration of *phtAd*, 1.25 µM, resulted in the complete reduction of DCPIP, as evidenced by the solution's transition to a fully colorless state (see Figure 14). On the other hand, Figure 15 shows that there is no significant difference in the absorbance measurements between the assays conducted with 1 µM and 1.25 µM *phtAd*. However, the reaction conducted with 0.5 µM *phtAd* exhibited a lesser decrease in absorbance over the 30-minute period, indicating that the reaction proceeded at a relatively slow rate.

![](https://static.igem.wiki/teams/5043/results/results-fig12.png)

Figure 14: Reduction of DCPIP with different *phtAd* concentration [μM].

Regarding the optimal reaction time, Figure 15 illustrates that within the initial 10 minutes, absorbance decreases rapidly, indicating that the reduction of DCPIP occurs predominantly during this period. Beyond the 10-minute mark, absorbance measurements stabilize, suggesting that the reaction has reached a saturation point. This phenomenon may be attributed to the complete reduction of DCPIP or the inability of the reaction to proceed further due to the lack of NADH regeneration.

![](https://static.igem.wiki/teams/5043/results/results-fig13.png)

Figure 15: Reduction of DCPIP performed with different *phtAd* concentrations.
The
reduction of DCPIP by *phtAd* was monitored for 30 min at a constant substrate quantity and varying
amount of enzyme. The reaction mixture contained 100 mM Tris/HCl (pH 7.5), 100 µM NADH, 30 µM DCPIP, 0.5 µM
FAD and 0.5, 1, 1.25 µM *phtAd*. Reaction was performed at room temperature.

Based on the results obtained, we selected a concentration of 1.25 µM *phtAd* for the subsequent assay, as this concentration demonstrated a complete reduction of DCPIP.

### Kinetic parameters of *phtAd*

For the determination of kinetic parameters the assay was run with different NADH concentrations ranging from 40 µM to 250 µM. Data obtained from these assays was then analyzed using Michaelis-Menten kinetics (see Figure 16) and a Lineweaver-Burk plot (see Figure 17) was constructed to provide a linear representation of enzyme kinetics to determine key parameters such as Vmax and Km.

![](https://static.igem.wiki/teams/5043/results/results-fig14.png)

Figure 16: Michaelis-Menten Curve for *phtAd*
The velocity was calculated utilizing the
Lambert-Beer Law with the extinction coefficient of Cytochrome C (ε600 23 mM-1 x cm–1) [16]. A logarithmic
trend line was applied as a preliminary approximation.

![](https://static.igem.wiki/teams/5043/results/results-fig15.png)

Figure 17: Lineweaver-Burk plot
Graphs show the fitted linear function and its
correlation coefficient.

The Vmax, Km, kcat, kcat/Km values of the ferredoxin reductase *phtAd* were 0.0011 mM/min, 0.028 mM, 1.12 min-1 and 39.46 mM-1 x min-1, respectively. Notable discrepancies from the values reported in the literature [16] were identified, which can be attributed to variations in enzyme concentration. The reaction was performed using a higher enzyme concentration due to the observed diminished activity of the produced enzymes.
Furthermore, the R2 value of 0.4668 indicates that the Lineweaver-Burk plot does not perfectly describe the relationship between 1/[NADH] and 1/V. While the fit is moderate, it suggests that factors like experimental variability, biological complexity, or non-linearity in enzyme behavior might be contributing to deviations from the ideal linear relationship.

Kinetic characterization of *phtAc*

### Method

Initial measurements were conducted in accordance with the methodology outlined by Wu et al. (2020) [16]. However, the assay produced inconclusive results, as no variation in absorption was observed over time. This lack of change was attributed to the high concentration of cytochrome c (600 µM) utilized in the initial trials. Consequently, we opted to conduct further assays employing varying concentrations of cytochrome c and NADH to identify the optimal concentrations that would facilitate a measurable increase in absorption. In alignment with the approach taken by Wu et al. (2020) [16], subsequent assays were performed using a *phtAd* to *phtAc* ratio of 1:3. The increase in absorbance was measured over a 30-minute period at 2-minute intervals at 550 nm. After the optimal cytochrome c concentration was established, further assays were performed varying the NADH concentration to determine the kinetic parameters.

### Results

**Determination of the optimal substrate concentration**
In the initial assay conducted with 250 µM cytochrome c, no change in absorbance was observed (see Figure 18A). Considering this outcome, the subsequent assay was performed using 100 µM cytochrome c in conjunction with a higher concentration of NADH (200 µM). This combination resulted in a measurable increase in absorbance, demonstrating a linear progression over a duration of 26 minutes (see Figure 18C) indicating the oxidation of cytochrome c. To validate this outcome, additional assays were conducted utilizing 100 µM cytochrome c alongside varying concentrations of NADH, both higher and lower. The assay employing 250 µM NADH also demonstrated a linear increase in absorption (Figure 18B) in the first 26 minutes. Conversely, the assay with 100 µM NADH (See Figure 18D) also exhibited a linear increase in absorption, albeit at a reduced rate. Notably, no saturation point was observed at this concentration, in contrast to the higher concentrations, which indicated saturation occurring approximately 30 minutes into the assay.

![](https://static.igem.wiki/teams/5043/results/results-fig16.png)

Figure 18: Oxidation of cytochrome c catalyzed by *phtAc*, in conjunction with *phtAd*,
under varying concentrations of cytochrome c and NADH.
The oxidation of cytochrome c by *phtAc*
coupled with *phtAd* was monitored for 30 min at a constant enzyme concentration and varying amount of
substrate. The reaction mixture contained 100 mM Tris/HCl (pH 7.5), 100 – 250 µM NADH, 100 and 250 µM Cyt.
C, 0.5 µM FAD, 1 µM *phtAd* and 3.5 µM *phtAc*. Reaction was performed at room temperature.

Based on the results obtained, we chose a concentration of 100 µM cytochrome c for the subsequent tests, as this concentration showed a measurable increase in absorbance. For the reaction time, we decided to run further reactions for 34 minutes to confirm the saturation point.

### Kinetic parameters of *phtAc*

For the determination of kinetic parameters the assay was run with different NADH concentrations ranging from 40 µM to 400 µM. Data obtained from these assays was then analyzed using Michaelis-Menten kinetics (see Figure 19) and a Lineweaver-Burk plot (see Figure 20) was constructed to provide a linear representation of the enzyme kinetics to determine key parameters such as Vmax and Km.

![](https://static.igem.wiki/teams/5043/results/results-fig17.png)

Figure 19: Michaelis-Menten curve for *phtAc*
The velocity was calculated utilizing the
Lambert-Beer Law with the extinction coefficient of cytochrome c (ε550 21 mM-1 x cm–1) [16]. A logarithmic
trend line was applied as a preliminary approximation.

![](https://static.igem.wiki/teams/5043/results/results-fig18.png)

Figure 20: Lineweaver-Burk plot
Graphs show the fitted linear function and its correlation
coefficient.

The Vmax, Km, kcat, kcat/Km values of the ferredoxin *phtAc* were 0.00043 mM/min, 0.31 mM, 0.12 1/min and 3.9 1/mM x min, respectively. Significant deviations from the literature values [16] were observed; however, the reaction was conducted with a substantially lower concentration of cytochrome c and elevated concentrations of enzymes, as the produced enzymes exhibited reduced activity.
Moreover, the R2 value of 0.5378 derived from the kinetic analysis of *phtAc* is marginally higher than that of *phtAd*. This marginally elevated value may indicate a greater reliability of the data or a more consistent enzymatic response across varying NADH concentrations. Nevertheless, the Lineweaver-Burk plot does not accurately characterize the relationship between 1/[NADH] and 1/V. Although the fit is moderate, it implies that factors such as experimental variability, biological complexity, or non-linear enzyme behavior could be responsible for the observed deviations from the expected linear relationship.

*pdoA2B2, phtAcAd* HPLC analysis

Method

We successfully characterized and demonstrated the activity of *phtAc* and *phtAd* both individually and as a complex. This complex functions as an electron carrier for *pdoA2B2* [13–15]. The subsequent step involved conducting an activity assay in conjunction with HPLC analysis with the aim to evaluate the function of *pdoA2B2* both independently and in complex with *phtAcAd*, forming a tetramer.

Method

Initially, the activity assay was conducted, wherein the enzymes were incubated at 30°C for approximately 15 to 20 minutes, both in the presence and absence of NADH to examine the effect of NADH on enzymatic aggregation. Subsequently, phenanthrene-4-carboxylate (P4C) was added to initiate the reaction. The reaction mixture was then incubated for 30 minutes at 30°C, after which the reaction was terminated by the addition of 100% methanol. Following this, HPLC analysis was performed using a Zorbax SB-C18 column, employing an acetonitrile/water gradient at a flow rate of 0.4 ml/min, with the column oven maintained at room temperature. The gradient elution program commenced with an initial mobile phase of 60:40 (v/v) acetonitrile to water, transitioning linearly to 100% acetonitrile over 14 minutes, followed by a return to the initial phase (60:40) after 5 minutes. The total duration for each analysis was 35 minutes. For the negative control, an assay was conducted using denatured enzymes, achieved by heating the enzymes at 95°C for 10 minutes. Peaks were analyzed at 270nm absorbance.

Results

Initial activity assays were performed solely with *pdoA2* and *pdoB2* to form a dimer, thereby validating their enzymatic activity and evaluating their aggregation characteristics. Monomers were incubated together for aggregation in the presence and absence of NADH to determine whether NADH, as a cofactor, influences the aggregation process.
The efficacy of the reaction was assessed by measuring the absorbance of phenanthrene-4-carboxylate (P4C) and NADH using HPLC. The area under the resulting peaks corresponding to the levels of P4C and NADH was determined, enabling a comparative analysis of the consumption of P4C and NADH against established standards and between the samples. In this context, phenanthrene-4-carboxylate consumption refers to the extent of P4C degradation, as indicated by a reduction in peak area compared to the standard. In the absence of additional samples with different concentrations, it is not possible to quantify the remaining P4C in the samples. The only conclusion that can be drawn is whether P4C has been degraded, as evidenced by the decrease in the area of the corresponding peak; this applies also to NADH.
Furthermore, to eliminate potential statistical errors in the assessment of our results, we employed triplicate measurements for each sample and calculated their averages. The results are presented in Figure 21. Results indicated that the assays in which NADH was added prior to aggregation exhibited enhanced P4C degradation and, correspondingly, increased NADH consumption compared to the assays where NADH was added after aggregation. However, the observed NADH consumption was relatively low, which may be attributed to the absence of the electron carrier dimer (*phtAcAd*) in the sample, thereby limiting NADH consumption.

![](https://static.igem.wiki/teams/5043/registry/bba-k5043010-fig10.png)

Figure 21: Comparative Analysis of phenanthrene-4-carboxylate (P4C) and NADH Consumption in
*pdoA2*:*pdoB2* activity assays.
The degradation of P4C by *pdoA2*:*pdoB2* was
monitored for 30 min at a constant substrate and enzyme concentration. The reaction mixture contained 100 mM
Tris/HCl (pH 7.5), 25 µM P4C, 1 µM *pdoA2* and 1 µM *pdoB2*. The 100 µM NADH were either added
after or before aggregation.

Considering the initial results, the analysis of the tetramer aggregation process and the degradation of phenanthrene-4-carboxylate was conducted by introducing NADH prior to the aggregation process.

**Degradation of phenanthrene-4-carboxylate by enzyme complex with dioxygenase activity using NADH as a
cofactor**
The presence of the functional enzyme complex with dioxygenase activity is anticipated to result in a reduction in the quantity of phenanthrene-4-carboxylate. This reduction can be quantified by determining the decrease in the area of the corresponding peak in comparison to the standard. Given the unknown stoichiometry of this enzyme complex, we conducted assays utilizing two distinct stoichiometric ratios. The first assay employed a 1:1 stoichiometry of all monomers, while the second utilized a ratio of *pdoA21*:*pdoB21*:*phtAc*3:*phtAd1*. This latter stoichiometry was selected based on previous kinetic analyses of *phtAc*, which were performed with a ratio of three equivalents *phtAc* to one *phtAd* for determining kinetic parameters [16]. This ratio was employed in the present assays to assess its influence on the degradation of phenanthrene-4-carboxylate and to determine whether the presence of three equivalents *phtAc* enhanced the efficiency of the electron transfer process, thereby accelerating the breakdown of phenanthrene-4-carboxylate. However, results showed (see Figure 22) that the breakdown of phenanthrene-4-carboxylate was slightly more efficient with the 1:1 ratio of the monomers; this could suggest that a higher ratio of *phtAc* does not enhance the electron transfer process. And that the complex with 1:1 ratio of all enzymes aggregates better.
To confirm the aggregation of the enzyme complex and the successful degradation of P4C by this complex, negative controls were prepared. For this, assays were prepared with denatured enzymes with the stoichiometries described above. Results showed (see Figure 22) that the peak area of P4C within these assays was higher in comparison with the assays with the active enzyme complex. The comparison of these results indicates that the enzyme complex was successfully formed and demonstrated the capacity to degrade P4C.

![](https://static.igem.wiki/teams/5043/registry/bba-k5043010-fig11-1.png)

Figure 22: Comparative Analysis of phenanthrene-4-carboxylate (P4C) and NADH Consumption in
*pdoA2*:*pdoB2*:*phtAc*:*phtAd* activity assays.
The degradation of P4C by the enzyme
complex was monitored for 30 min at a constant substrate and enzyme concentration. The reaction mixture
contained 100 mM Tris/HCl (pH 7.5), 200 µM NADH, 40 µM P4C, 1 µM *pdoA2*, 1 µM *pdoB2*, 1 or 3 µM
*phtAc* and 1 µM *phtAd*.

Another negative control was conducted by excluding any enzymes from the assay. In comparison to samples containing denatured enzymes, the results showed a reduced peak area in the peak corresponding to phenanthrene-4-carboxylate, suggesting either a diminished quantity of P4C in the sample or its degradation. However, the possibility of cross-reactivity with other components in the assay can be discounted, as such interactions would also manifest in the other negative controls. This observation indicates the presence of an independent source of error. Potential issues may involve the unintentional introduction of enzymes or inadequate substrate concentration. The presence of enzymes in the assay can be corroborated by taking into consideration the amount of NADH; notably, the peak area of NADH in this sample was diminished suggesting the possibility that a reaction has taken place, and the results are more consistent with those obtained from the assay conducted with a 1:3 enzyme ratio (see Figure 22). While numerous errors could have arisen, a systematic issue such as cross-reactivity, which would render the enzyme ineffective, can be dismissed by comparing the sample to those with non-functional enzymes.
Moreover, as anticipated, the samples containing the active enzyme complex demonstrated a decrease in the peak area associated with NADH relative to the standard (see Figure 22). This finding corroborates the functionality of the electron carrier dimer, which collaborates with *pdoA2B2* in the degradation of P4C. In contrast, the assay involving denatured enzymes at a 1:1 ratio revealed no consumption of NADH, indicating the absence of any reaction.
However, the results of NADH consumption from the assay involving denatured enzymes, utilizing three equivalents of *phtAc* in conjunction with a 1:1 ratio of the other enzymes, demonstrate a reduction in NADH consumption, as evidenced by the diminished area of the corresponding peak relative to the standard. This observation indicates the presence of an independent source of error, as the other assay performed with denatured enzymes shows no NADH consumption. Potential issues may include insufficient NADH concentration. If the NADH concentration was low, degradation could have occurred over time, as the activity assays were conducted days prior to the HPLC analysis. The thawing process and exposure of the sample to temperature fluctuations may have contributed to NADH degradation due to its inherent instability. Additionally, the possibility of incomplete denaturation should be considered. If the enzymes were not fully denatured during the assay, residual active enzyme could remain, resulting in NADH consumption. The denaturation conditions may not have been adequate to completely inactivate the enzymes. This is further supported by the observation that P4C consumption was slightly lower in comparison to the sample with denatured enzymes with the 1:1 ratio.

Conclusion

In conclusion, the enzyme complex with dioxygenase activity demonstrated its ability to degrade phenanthrene-4-carboxylate (P4C), with slightly greater efficiency observed in the assay utilizing a 1:1 ratio of monomers compared to the 1:3 ratio of *phtAc* to *pdoA2*:*pdoB2*:*phtAd*. This suggests that a higher ratio of *phtAc* does not enhance electron transfer efficiency. Negative controls, including assays with denatured enzymes and no enzymes, confirmed that the active enzyme complex was responsible for P4C degradation, while the reduction in NADH in some denatured enzyme assays points to possible errors, such as NADH degradation or incomplete enzyme denaturation. Further assays with stricter controls are needed to confirm these findings and minimize potential sources of error.

Immobilization

Description

Immobilization of our chassis is important, to make a self-maintaining device of riverine deployment possible. For this we chose five evidence-based methods by the criteria feasibility and sustainability. We included silica beads and a larger silica surface as *P. vancouverensis* was immobilized on silica in preceding research [28]. The other three materials (modified alginate, carboxy methyl cellulose and rice bran) were chosen due to them having proven to work in the related model organism *P. putida*. The physical properties were tested, as well as viability and immobilization efficiency. All tests were done with the *P. vancouverensis* transformant containing the pSEVA231-plasmid.

Results

### Physical properties

1. Alginate
   beads do not melt at temperatures up to 37°C so they would work
   even in very hot rivers. They are difficult to destroy with physical
   force because they withstand being squished to some degree and then
   dodge the force exerted by the object by slipping away.
2. Silicabeads are almost indestructible by the physical force a human can
   induce. They do not melt or disintegrate with high temperatures (in
   our experiments they had no problem with 120°C and we could have
   gone much higher).
3. Silica pieces with larger surface than the beads, are slightly easier to
   crush but still a very stable material.
4. Carboxy methyl cellulose (CMC)
   beads are easy to crush, even a water flow can crush them, and they
   melt at 28 °C once damaged. They are fully solved in the watery
   solution after 48 hours. Therefore, they are not suitable for our
   application, but we only figured that out later while doing
   biological tests.
5. Rice
   bran turned out to be impractical for our use case because the
   formed beads disintegrate in water, especially fast when there is
   water flow. Therefore, the idea was discarded after testing the
   physical properties.

### Viability tests

1. The
   creation of alginate beads does not kill the bacteria. The method is
   established and often used with a variety of bacteria [25]. For
   our viability tests, the beads were destroyed by physical force and
   put in liquid culture as well as being plated out over 48 h. The
   tests proved that P. vancouverensis is no different and survives the
   procedure.
2. Immobilizing
   bacteria on silica beads is also known to work [26-28]. The
   nonpolar surface of silicon dioxide
   leads to *P.
   vancouverensis* immobilizing itself. The bacteria are just cultivated in liquid
   culture with the silica beads, making it the gentlest treatment.
   Therefore, the viability of the cells is expected to remain fully
   intact. Our results prove these expectations, showing that
   immobilized bacteria are still viable.
3. The
   immobilization on the larger silica surface is done in the same way
   as on the beads. Therefore, our expectations were the same and the
   results turned out the same as well.
4. The
   bacteria in CMC beads showed viability as well, even though the
   beads were not physically stable.

![](https://static.igem.wiki/teams/5043/results/resultsimmoviability.png)

Figure 23: Cell viability assay.
Viability tests were carried out by propagating the immobilized cells into liquid cultures. The materials were either left intact or their integrity was destroyed by physical force.

### Immobilization efficiency

To determine immobilization efficiency, meaning how difficult it is to wash bacteria off, the materials with immobilized cells were first washed multiple times in water and then inverted in LB medium, simulating water flow. The LB medium was subsequently plated out. The less viable cells could be observed, the more effective immobilization worked.

1. As the bacteria are enclosed in the alginate beads, its not possible to wash off all the bacteria. After washing there were still some bacteria coming of. In the first test there were around 900 colonies, in the second test 16 colonies could be counted.
2. Silica beads showed a lot of bacteria remaining on the surface after washing five times, but there were still bacteria that came off when simulating water flow. When plated out the washing medium, around 10.000 colonies could be counted after several days. There were no second tests of this method due to a lack of material.
3. The Silica surface also showed a lot of bacteria remaining on the surface after washing, but there were still bacteria that came off when simulating water flow as well. When plated out the washing medium, around 2.500 colonies could be counted after several days. There were no second tests of this method due to a lack of material.
4. In the CMC beads, the bacteria are enclosed as well. When simulating water flow and plating the medium in the first test, only 15 colonies could be observed. In the second test there were no colonies visible.

![](https://static.igem.wiki/teams/5043/results/resultsimmoefficency.png)

Figure 24: Immobilization efficiency assay.
 The immobilization efficiency was tested by washing immobilized cells and subsequently inverting them in LB medium kanamycin. LB medium was then plated out.

### Conclusion

Our tests show that rice bran and carboxy methyl cellulose are insufficient for our application, due to their physical properties. On the other hand, using alginate or silica is suitable for our purposes. Both methods have good physical properties and immobilize the bacteria without killing them. When testing the efficiency of the immobilization, it turned out that after several washes, most of the bacteria remained immobilized, but many were still washed off.

Discussion

It is not our desired outcome, that a lot of the bacteria were still washed off after several washes. But when talking to Julia Polat, whose research is centered around immobilization with alginate, she reassured us there is no way to prevent bacteria from leaving their immobilized state, especially as time progresses. For us, this means we need a very reliable kill switch to prevent biocontamination. We can be sure that the only bacteria that grew is *P. vancouverensis* because a transformed variant was used that contained a plasmid with kanamycin resistance, and all tests were done in LB medium with 50μg/ml kanamycin. The creation of the beads and further tests were also carried out in a sterile environment so the chances of another kanamycin resistant bacterium entering the experiment are very low. Nevertheless, we additionally analyzed the visual properties of the colonies, and they all looked like the typical *P. vancouverensis* colonies. To conclude, our tests of the two methods using alginate and silica showed promising results. They should undergo more testing in the future, especially simulations of a river with a typical current, to see which one is the best method for our use case.

Future plans

Genome Editing

As in our application in rivers no antibiotic resistance pressure can be applied and our chassis would lose
pyrene degradation plasmid, it is necessary to integrate the pyrene pathway into the host’s genome.
Furthermore, our advisor M. Sc. Christian Hake pointed out, that PAHs are very stable molecules and therefore
represent unattractive nutrition sources. Hence our chassis might not produce naphthalene and phenanthrene
degradation enzymes, as more nutritious carbon sources are around in rivers. Therefore we plan to put naphthalene
and phenanthrene degradation genes under low constitutive expression as well. This could be done for example
using CRISPR systems [18].

Implementing an appropriate kill switch

In order to avoid contamination or an unwanted upsurge of GMOs in the environment implementing a kill switch is
crucial. An light activated kill switch could be promising, like shown by [iGEM Team Ashesi 2022](https://2022.igem.wiki/ashesighana/design) [19], which
kills the engineered bacteria upon leaving our device. Though the light in the depth of a riverbed could not be
enough to activate the kill switch. Another possibility could be implementing an auxotrophic marker to create a
dependency on a substance only supplied by the surface inside the container on which the bacteria grow, but that
requires attendance and care for the system which is supposed to function autonomously.
Therefore, the best cut out would be one designed especially for this purpose. A kill switch activated by not
being immobilized on a surface like a RNAse or toxin bound to an extracellular receptor. Even here might be a
downside, as not every receptor in the membrane of a cell could be attached to the surface and would cause an
RNAse or toxin to be moving freely inside the cell. A promising possibility could be a toxin-antitoxin system,
where the bacteria express a toxin constitutively and selectively express an antitoxin or inhibitor based on the
containment in our device. In the project of [TU Darmstadt 2020](https://2020.igem.org/Team%3ATU_Darmstadt/Project/Kill_Switch) [20] they thought of a kill switch connecting their
bacteria to a biofilm. They did so by giving the gene for the toxin to one colony and the gene for the antitoxin
to another. When they coexisted in a biofilm they survived and otherwise were killed by an excess amount of the
toxin. The idea would need to be tested long term on *P. vancouverensis* colonies to see if it works for
us. This complex and difficult aspect of our project might inspire future generations of iGEMers to find a
solution to our kill switch problem.

Increasing PAH’s solubility using biosurfactants

PAHs are poorly water soluble [8], which limits substrate availability for degradation and probably thereby
shrinks bioremediation’s efficiency [21]. This is why Prof. Victor de Lorenzo from CSIC Madrid advised us to use
biosurfactants. Biosurfactants are amphipathic, surface-active molecules which can increase solubility of
hydrophobic substances [21]. Though recent research shows doubts, it is suspected that in this way PAH
degradation could be enhanced [22]. Therefore we plan to produce biosurfactants *in situ* by introducing its
biosynthesis pathway genetically into our chassis.
iGEM teams [Sorbonne 2023](https://2023.igem.wiki/sorbonne-u-paris/description) and [Columbia-University 2016](https://2016.igem.org/Team%3AColumbiaU_NYC/Project) already used *Pseudomonas putida* to produce
sophorolipids [23] and rhamnolipids [24]. As we would be happy to build on previous iGEM team’s success, we are
eager to try out biosurfactant production in *P. vancouverensis* and study its effects on PAH degradation.

Drylab

Since the engineering cycle is an iterative process, continuous improvements can be made to the design of the device. Additional simulations could help us to further optimize the internal fluid dynamics of the device, maximizing the contact between PAHs in the water and *Pseudomonas* bacteria in the immobilization material. The results of these simulations would then be validated using 3D-printed prototypes.
The next phase would involve constructing full-scale prototypes with the material that is already used for buoys, which would need to be tested under real-world conditions, along with the engineered bacteria.
Several potential issues need to be identified and addressed through testing, including the risk of clogging due to floating debris, the stability of the device under adverse weather conditions and animal interactions, the ability of *Pseudomonads* to adhere to the interior surfaces as intended, and whether they can effectively degrade PAHs in this scenario. These tests would show us potential for improvement, which can be used to develop a market-ready product.
When the fundamental functionality of the device has been demonstrated, the focus would shift towards its longevity and ease of maintenance. This would require long-term testing, which must still be planned and conducted.

Once the device has been fully tested and its functionality confirmed, the next step would involve transitioning it into a complete, market-ready product. This process would begin with sourcing a supplier for the necessary materials and identifying a manufacturer capable of producing the device on a large scale. With these partnerships established, we could accurately calculate production costs and use this information to determine an appropriate price point for the product. Mostly practical would be the use of the material of already used buoys, which are proved already as a durational, cheap and water friendly option.
Subsequently, efforts would be directed towards customer acquisition. Potential customers include companies seeking to remediate polluted waterways they are responsible for, as well as municipalities or local governments aiming to clean their rivers. Given the versatility of our design, the device can be deployed globally, provided the water body does not have excessively strong currents and there is a suitable method for securing the device in place.
To anchor the device, we propose using existing buoys, which are already widely available in many river systems. Buoys offer a practical solution for maintaining the device’s position and are typically well-distributed, making it feasible to deploy multiple units if required for larger-scale water treatment efforts. To use an already existing infrastructure would be wasteless and a cheap opportunity.

Our hope is for future iGEMers to build upon our project, expand and improve it, as it represents an innovative and low-effort method for cleaning rivers from dangerous pollutants. We wish to inspire many new
projects no matter if it’s in bioremediation, upscaling, foundational advance or anything else. Either way, one
thing’s for sure. There’s much to do for generations of iGEMers to come.

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