Scientific background

Peroxisomal matrix proteins are imported post-translationally and in their folded state (Lazarow and Fujiki 1985). The peroxisomal protein import depends on two pathways, both involving a different Peroxisomal targeting signal (PTS) and respective receptor (PEX5 and PEX7). The import cycle can be divided into five conceptual steps: (i) the cytosolic receptors bind their cargo proteins and guide them to a docking site at the peroxisomal membrane, (ii) the receptor-cargo complex translocates to the peroxisomal matrix, (iii) the complex is disassembled, (iv) the receptor is returned to the cytosol (v).

The vast majority of peroxisomal matrix proteins are imported by the PEX5 importer. PEX5 recognizes the C-terminal PTS1 peptide whose evolutionarily conserved Sequence is (S/A/C)-(K/R/H)-(L/M) (Gould et al. 1989). Upon recognition of the PTS1 in the cytosol, PEX5 binds its cargo (i). It docks to the peroxisomal membrane complex, consisting of PEX13, PEX14 and PEX17 (ii). This docking complex is connected to the RING-finger complex, consisting of PEX2, PEX10 and PEX12, via PEX8. This multi-protein complex is known as the importomer. PEX5 and PEX14 form a pore in the membrane, through which the cargo is translocated (iii). The receptor–cargo complex dissociates at the matrix site of the membrane (iv). The integral PTS1-receptor is either monoubiquitinated by the E2-enzyme PEX4 or polyubiquitinated by Ubc4 or Ubc5. The AAA peroxins PEX1 and PEX6, which are anchored to the peroxisomal membrane by PEX15, dislocate the ubiquitinated PEX5 from the membrane back to the cytosol (v). The polyubiquitinated PTS1-receptors are degraded by the proteasome, whereas the monoubiquitinated receptors are recycled for further rounds of import.

Some proteins are instead imported by the PEX7 importer, together with the co-receptors PEX18 and PEX21. The targeting signal (PTS2) for this pathway is localized near the N-terminus of the cargo-protein and is comprised of nine different amino acids with a highly variable five amino acid core region and the consensus sequence (R/K)/(L/V/I)X5(H(Q))(L/A). In contrast to the pore formation by PEX5, the pore for import of PTS2 proteins is formed by the co-receptor PEX18 and the docking complex.

Engineering of PEX5 and PTS1

Designing our receptors

To achieve the engineering of an orthogonal import pathway, we followed two approaches regarding PEX5. The first is based on targeted mutagenesis based on educated guesses which is first verified by molecular dynamics and later experimentally in the laboratory. The second approach is based on a recently published paper: We searched for literature dealing with the modification of the peroxisomal import machinery. During our research we came across a paper of Alison Baker et al., published in 2017, in which they present a synthetic construct of the PEX5 protein, partly Arabidopsis thailana and partly Physcomitrella patens. Compared to the wild type PEX5, this one shows different binding affinities since it interacts with a PTS1* variant that does not interact with the wild type PEX5. Since the protein sequences of yeast's and arabidopsis's PEX5 differ quite a lot, we aligned both sequences to understand where the mutations were set.

The alignment shows three red marked amino acids we changed in our receptor sequence. Interestingly, these mutations are located within the TPR motifs of our PEX5 protein and this persuaded us to try out this receptor, we call it R19. Due to lack of time we tested this PEX5 variant in silico and in vivo simultaneously. We started molecular dynamics simulations with a couple of PTS variants that we already tested with our previous designs − one of them was actually the variant they used in the paper (YQSYY). The details and results of our structural modeling can be found in the modeling section.
Furthermore we synthesized this variant and together with two receptors we designed based on educated guesses we got three receptors for our experimental work.

Experimental design

The easiest way to verify successful import is the localization of a fluorescent protein within the peroxisome. Due to that, we decided to tag mTurquoise with our PTS variants. Additionally, we wanted to mark the peroxisomal membrane, to be absolutely sure about the localization within the peroxisome. For that cause we chose the transmembrane domain of PEX13, tagged with the fluorescent protein mRuby.

Pex13−mRuby

We used the peroxisomal membrane protein PEX13 as a fluorescent marker − by just using the transmembrane domain of PEX13 with a short linker, we make sure that it has no influence on the peroxisomal features. To get a higher differentiation from mTurquoise, which we use for another construct, we chose to work with mRuby. Literature research revealed that such constructs have been tested before − Erdmann et al. (2004) described a construct containing only PEX13_200-310 with a C-terminal GFP.

PEX5 variant

In order to achieve an orthogonal import we used a PEX5 knockout strain and transformed it with a plasmid based PEX5 variant that is supposed to detect a non native PTS variant instead of the wild-type one. The construct contains a medium strength promotor, the PEX5 gene and a terminator. The remaining plasmid parts can be seen in the plasmid map below.

mTurquoise−PTS

Our approach for import verification is based on a fluorescent protein tagged with the PTS variants. After several promotor tests with different strength, we decided to express this construct only in low amounts. This is advantageous because we detect only a low signal if the import does not work, hence small amounts of the protein are distributed in the whole cytosol. In contrast we see a clear signal if the import does work due to the relative high concentration inside the peroxisome.
Our construct is depicted in the figure below.

Combination of our constructs

To combine our constructs, we cloned our PEX5 and mTurquoise constructs into a level 2 plasmid portrayed below.

We then did a co-transformation with the PEX13−mRuby plasmid and the level 2 plasmid to combine everything that is needed into our yeast.

PTS screening

Trusting on our targeted approach alone seemed risky − that is why we planned a PTS screening to find the most favorable PTS for our three receptors. Dueber et al. (2016) used the Violacein assay for a similar purpose. They screened for the best PTS for the wild type receptor and were successful. Hence another subproject of our team is the integration of the Violacein pathway into the peroxisome ( Violacein ), we were already supplied with all necessary enzymes − VioA, VioB and VioE.

The figure above shows the principles of the assay. VioA and VioB are localized in the cytosol and lead to the production of the IPA imine dimer while VioE is tagged with a PTS1 variant. Successful import leads to white colonies whereas missing import results in green colonies due to the cytosolic production of Prodeoxyviolacein.
Our rests upon the following two plasmids which are co-transformed into yeast.

As shown above, we created one plasmid containing VioA, VioB and one of our PEX5 variants while the other plasmid only contained VioE. We then designed primers which bind to the VioE plasmid to amplify the whole plasmid except the terminator − random PTS1 variants were attached to VioE with the help of a random primer library. Following up, we did the ligation with the corresponding terminator and obtained a mix of several different VioE-PTS1 plasmids.
After plasmid amplification in Escherichia coli we then co-transformed yeast with the two constructs and waited for the colonies to grow. With the yeast growing prodeoxyviolacein should be produced in yeast cells with absent import and the IPA imine dimer (white color) should be produced in those with functional import.

Mutagenesis of PTS2

To characterize the import efficiency for the site-directed PTS2 firefly luciferase was used. Luciferase is a luminescent protein which can be split in a C- and a N-terminal part. Only when combined, luminescence can be detected. To measure the import efficiency the two parts will be expressed and imported into the peroxisome in a separated way. The smaller part (Split2) of the split luciferase will be brought into the peroxisome first via the PTS1 dependent pathway. The other part is imported via the respective modified PTS2 sequence. The better this sequence is recognized by Pex7, the stronger the luminescence of the assembled luciferase can be detected in the peroxisome. There is a chance of split parts of the luciferase assembling in the cytosol if the import is too slow. To avoid wrong conclusions of the luciferase localisation, we designed a negative control experiment. It includes a split luciferase similar to the one used in the initial experiment, but without the peroxisomal targeting sequence. Consequently there will be no import into the peroxisome. If we subtract the luminescence of the negative control experiment from the luminescence of the main experiment we can define the degree of import.

In addition to a directed approach according to Kunze and colleagues we also want to perform a random mutagenesis experiment to alter the five variable amino acids of the core region of the PTS2 sequence in an unbiased manner. The aim is to generate a library of different peroxisomal PTS2. The 15 nucleotides are assembled by chance. In the DNA synthesis this sequence will either be described as [NNN]5 or [DNK]5. N stands for all four nucleotides mixed, K for either G or T and D for A,G or T. The “DNK” composition prohibits two out of three termination codons. Additionally with this library the amino acid frequency is improved towards a balanced ratio in between the different kinds. DeLoache, William C., Zachary N. Russ, and John E. Dueber (2016)

Each approach could generate up to 415 DNA sequences, which is roughly 1,07 billion. On the level of the amino acid sequence there are 3,2 million possibilities, since each residue can be taken by 20 different amino acids. For the assay we therefore need a high throughput method.

We adapted work of DeLoache, Russ and Dueber using the violacein pathway to measure the import effectiveness of tripeptides. The pathway consists of Violacein A (VioA), Violacein B (VioB) and Violacein E (VioE). It converts tryptophan into the green product prodeoxyviolacein (PDV). The first two enzymes, VioA and VioB, are expressed in the cytosol, and third one, VioE is targeted to the peroxisome with a PTS1 sequence [1]. The degree of import can be measured by the intensity of green colour of the colonies. An efficient import signal leads to a strong import of the VioE into the peroxisome and subsequently to white colonies, because the intermediates cannot diffuse into the peroxisome to its respective enzyme. DeLoache et al. showed that there is a proportional correlation between the concentration of the green product PDV and a red fluorescent substance. The concentration of this product displays the import efficiency of the respective sequence.

This assay has been used for the evaluation of the generated PTS2 sequences. The VioE-PTS2 plasmids are harvested and cotransformed with a VioA-VioB plasmid. Each plasmid contains a specific auxotrophy marker. Consequently every growing colony contains both plasmids. To evaluate the respective sequence the concentration of the red fluorescence is measured. The more fluorescence is detected the more VioE is in the cytosol. Therefore the respective PTS2 is not that efficient. The other way around a low concentration of the fluorescent substance correlates with an efficient import via the respective PTS2.

In order to test our hypothesis we fused the last 59 amino acids of the C-terminus of human PEX26 (AA 246-305) to a red fluorescent protein, called mRuby2, to further elucidate the Pex3/Pex19-dependent import. mRuby is generally used as a marker in combination with a fluorescent microscope to visualize the localization of the fusion protein. The C-terminus of PEX26 contains a helical signal-anchor, which serves as both, a mPTS and transmembrane domain. We designed our construct with mRuby2 fused to the N-terminal side of the PEX26-C-Terminus, this way the mRuby should face the cytosolic side of the peroxisomal membrane. Quite similar to our mRuby-PEX26 approach, we designed a construct for the ER-dependent import. Therefore, we fused the mRuby2 fluorescent protein to the N-terminus of Pex3 (AA 1-39). This construct should be N-terminally anchored in the peroxisomal membrane, with mRuby2 again facing the cellular lumen.

Our main goal is to introduce a rather complex membrane protein to the peroxisome that can alter specific traits. For that we fused the Pex3 N-terminus (AA 1-39) to a Halobacterium salinarum Bacteriorhodopsin protein (AA 16-262), replacing the first 16 amino acids ( Pex3-BacR). The original archaeal Bacteriorhodopsin acts as a proton pump by capturing light energy to move protons across the membrane out of the cell. The resulting proton gradient is subsequently converted into chemical energy. Our assumption is that the first transmembrane segment determines the orientation of the following protein and that therefore due to the N-terminal anchoring signal the Bacteriorhodopsin will be inserted in reverse orientation, pumping the protons into the peroxisome. This way the pH of the peroxisomal lumen could actively be controlled and adjusted.

Finally, our project involved combining the work of other subteams to verify the localization of our constructs in the peroxisome and analyze the effects they have on the import. Therefore, we are using the superfolder-gfp protein, another fluorescent marker, which is in our case fused to the peroxisomal import sequence pts1, and a version of Pex11 that is fused to the fluorescent marker Venus. Both markers emit light in the green light spectrum, were as mRuby2 emits light in the red part of the spectrum, giving us a strong contrast and an easy way of differentiating between the two under the fluorescent microscope.

To physically create our constructs, we researched the DNA sequences of Bacteriorhodopsin, Pex3 and PEX26 via UniProt and pre-designed our fusion constructs with the software tool „Geneious“. We ordered the synthesis of three separate parts ( Pex3, PEX26 and Pex3-BacR) from IDT. To ease out the process of assembling our plasmids, we used the „Dueber Toolbox“, containing various parts such as promoters and terminators, to tailor the plasmids specific to your needs. Finally, to combine all the selected parts, we used the „Golden Gate” assembly method.

Scientific background

Downstream processing is a very important part of industrial biological compound production. For most biotechnological produced compounds, it is the most expensive part of the production [7]. One step to decrease the costs is to secrete the products into the supernatant [1]. After secretion, it is possible to remove most cellular compounds from valuable products with one simple centrifugation step. Due to this, secretion is not only a great tool for a compartment toolbox but also has an economic value.
In regards to the whole project, this is an important part for making the compartment more applicable. Through it we go a step further by thinking about the extraction of products after production.
At the end of this sub project it should be possible to secrete every compound produced in the modified compartment to the supernatant. This is not trivial because peroxisomes, which are the basis of our compartment do not possess a known natural secretion mechanism.
We overcome this problem by using the "peroxicretion" concept of Sagt and colleagues[9]. They used a v-SNARE (vesicle- synaptosome-associated-Soluble N-ethylmaleimide-sensitive-factor Attachment REceptorprotein) fused to a peroxisomal membrane-protein to secrete the content of peroxisomes. V-SNAREs interact with the t-SNARE (target synaptosome-associated-SNARE) at the cell membrane, which leads to an fusion of the vesicle with the membrane [3]. Sagt and colleagues applied this system in the filamentous fungi Aspergillus niger.

Experimental Design

We will adapt the system of Sagt and colleagues *needs to be change* to secrete the content of our modified compartments [9].
For the application of this system in S. cerevisiae we use a truncated version of the v-SNARE Snc1 to decorate our compartments(Figure 1) [5].

To decorate the compartments with the SNARE we use a peroxisomal transmembrane protein . In our case we use the proteins Pex15 or PEX26, which were further investigated in another sub project, and fuse Snc1 to the N-terminus. We expressed these constructs of membrane anchor and Snc1 constitutively under control of the RPL18B promotor. In case of Pex15 we used a truncated version, lacking a large part of the N-terminus, only consisting of the transmembrane domain (315-383) (Figure 1). For PEX26 we use the truncated version published in Halbach et al. (2006). [6]

We verified our secretion using Beta-glucuronidase (GUS) as a reporter protein. In 2012 Stock and colleagues described the GUS reporter assay for unconventional secretion [10]. With it, it is possible to determine whether a protein is secreted conventional and is N-glycosylated or secreted unconventional and not N-glycosylated. GUS is a bacterial protein with an N-glycosylation-site, which is active only if the protein is not N-glycosylated. The GUS-activity can be measured with different reagents in plate or liquid assays. Liquid assays can be applied qualitatively as well as quantitatively to measure differences in activity. If GUS is secreted by the conventional pathway the N-glycosylation leads to inactivation of the enzyme [10] (Fig 2).

GUS will be imported to the peroxisome with the PTS1 sequence and measured quantitatively in the supernatant. We will use a coexpression of GUS-PTS1 and Snc1-Pex15 or Snc1-PEX26 to identify the secretion of the compounds. Furthermore, we will use GUS-PTS1 expressed in S. cerevisiae without Snc1 fused to a membrane anchor for a control. We will measure the active GUS in the supernatant with a liquid assay based on the turnover of 4-methylumbelliferyl-beta-D-glucuronide to 4-methyl umbelliferone (4-MU) [2]. Here we expect a higher activity of GUS in the supernatant of cultures with Snc1 decorated peroxisomes.
To increase the variability of our constructs we also designed vectors with and without a GS-Linker connecting the Snc1 with the Pex15. Additionally we tested our constructs in strains with a deletion of Pex11 . This deletion leads to formation of larger peroxisomes and may increase the efficiency of our secretion mechanism.

Heading

Sensors

To enrich our toolbox we decided to measure four essential physiological factors: ATP, NADPH, Glutathione and the pH.
ATP/ADP conversion is used as an energy currency in many cellular processes like translocation of proteins and metabolites or anabolic and catabolic turnovers. It is assumed that ANT1P an ATP/AMP antiporter is located in the peroxisomal membrane and that the re shuttle mechanism of the peroxisomal protein import machinery is ATP dependent(Palmieri L. et al. 2001). NADPH plays an important role in anabolic pathways and is also indispensable to our desired pathways of nootkatone and violacein. Glutathione is an antioxidant and redox buffer which is also found in yeast peroxisomes. It is used as cofactor by at least two types of peroxisomal proteins the glutathione peroxidases and glutathione transferases, which reduce lipid- and hydrogen peroxides or transfer glutathione to lipid peroxides for the purpose of detoxification (Horiguchi H. et al. 2001). Furthermore, it partly represents the redox state of the peroxisome. Knowing the pH of a compartment is important to predict the activities of almost all enzymatic processes inside of it and to follow up acidification and basification upon conversion of metabolites.

We finally chose two ratiometric sensors to perform measurements with. The pH sensitive and glutathione redox state reporting green fluorescent proteins pHLuorin2 (Mahon M. J. et al. 2011) and roGFP2 (Schwarzländer M. et al. 2016). We aim to target them either in the peroxisomal lumen or the cytosol. To achieve peroxisomal targeting we attach the peroxisomal targeting signal 1 with Golden Gate cloning Link zur toolbox beschreibung

Using promoters with differnt expression strength in order to find optimal measurement conditions is of high interest. There is a trade off between a high signal-to-noise ratio and self induced effects which are both dependent on expression levels (Schwarzländer M. et al. 2016). This cannot be generalized for each sensor. For example, pHlourin2 has sparse influence on the existing pH because of the buffer effect of proteins Quelle.
Validation of the peroxisomal localization can be achieved via fluorescence overlap of the sensor and a peroxisomal marker in our case pex13-mRuby(link to pex5 import).

It can also be validated by transforming the sensors attached to the PTS1 sequence into PEX5 knockout yeast strain. The sensor is expected to show no specific localisation, because of the missing import sequence. We calibrate the sensors in living yeast cells and physiological ranges so that we can not only perform relative but also quantitative measurements. We aim to confirm our hypothesis of an more oxidized redox state of roGFP2 in peroxisomes with violacein pathway violacein activity and want to measure differences in pH within yeast strains with peroxisomal membrane anchored pex3-bacteriorhodopsin protein.

Once expression and localization of the sensor is proven by microscopy, measurements with a plate reader or a fluorometer are acceptable. This allows a high number of replicates to be measured accurately in reasonable time. Microscopy is performed with a filter based Nikon Eclipse TI fluorescence microscope at 100-fold magnification and plate reader measurements are performed with a Tecan infinite plate reader(Mahon M. J. et al. 2011). For pHLuorin2 emission intensity is measured at 535 nm upon excitation at 405 nm and 485 nm. Same settings were used for roGFP2.(Schwarzländer M. et al. 2016) . Evaluation of the reported signals is done by the excitation ratio of the the corresponding excitation wavelength.

Nootkatone

In order to have a proof of concept for our compartmentation strategy we intend to establish the nootkatone pathway inside the peroxisome. Therefore we aim to target the three enzymes for the nootkatone production to the peroxisome. Before doing this there are several aspects that have to be considered.

The initial step is to find a reliable source to prove the abundance of our precursor FPP in yeast peroxisomes. In yeast there is no prove of existence of FPP in the peroxisome yet, but it is predicted to be present since it was detected in mammalian and plant peroxisomes. Olivier et al. (2000) .

The precursor FPP is converted into valencene by a valencene synthase (ValS). We chose the one from Callitropsis nootkatensis because of its comparably high efficiency in microorganisms. It achieves greater yields in yeast than the citrus valencene synthase. Furthermore, the product specificity is relatively high, while producing less byproducts Beekwilder et al. (2014). The valencene synthase was also chosen because of its robustness towards pH and temperature changes. Beekwilder et al. (2014) . Our modelling approach revealed that for optimal yields an overexpression of valencene synthase is necessary because of its slow conversion rate. This is why we chose the strongest promoter of the yeast toolbox (Dueber Toolbox) for this attempt [https://2017.igem.org/Team:Cologne-Duesseldorf/Model]

The intermediate valencene is then converted into nootkatol by a P450 monooxygenase as well as into small amounts of our desired product nootkatone. The P450 monooxygenase we chose for this project was taken from the bacterium Bacillus megaterium. In this case it is not a simple P450 monooxygenase, but an entire P450 system, consisting of a soluble P450 fused to a cytochrome P450 reductase (CPR) enzyme, making an additional reductase obsolete De Mot Ret al. (2002) . Unlike eukaryotic P450s, which are mostly membrane bound, this prokaryotic BM3-P450 is located in the cytosol facilitating an easier transport into the peroxisome, as membrane integration of proteins is a more difficult task to achieve than import of cytosolic proteins. Girvan et al. (2006) . BM3 normally catalyzes the hydroxylation of long chain fatty acids Narhi et al. (1986) . , which in our case could inhibit the conversion of valencene. For that reason we used a mutated version of BM3, which is called AIPLF. This variant is an enhanced version of the BM3 AIP version, which has a ten times better substrate oxidation rate for valencene than the wildtype BM3 and produces less byproduct when valencene concentration is saturated. Additionally to previous named benefits the AIPLF variant with 5 point mutations in the active side has a significantly lower binding affinity towards long chain fatty acids and therefore increase the transposition rate of valencene. Schulz et al. (2015) Lehmann (2016)

The alcohol dehydrogenase from Pichia pastoris subsequently converts nootkatol into nootkatone by oxidation. It uses NAD+ as a cofactor which is reduced in the reaction. The regeneration of this cofactor is facilitated by the BM3, which oxidizes NADH Schulz et al. (2015) .

The first milestone to achieve our goal is the separate integration of each of our three enzymes ValS, BM3 and ADH into level 1 vectors (Dueber Toolbox) in yeast and to verify their expression by Western Blot analysis. Therefore, a 3xFlag/6xHis-tag was added to the N-terminus of each of the proteins. It enables us to use an anti-His antibody followed by an anti-mouse-antibody to make protein abundances visible Subsequently, the two enzymes ADH and ValS were combined in a level 2 cassette plasmid. BM3 is designed as a level 1 plasmid and will be co-expressed with the level 2 plasmid to achieve a nootkatone production. The expression of the enzymes in the cytoplasm is again verified by Western Blot analysis. The further approach aims to provide the enzymes with C-terminal peroxisomal targeting signals type one, which finally converts the nootkatone pathway into our artificial compartment.

Since the production of nootkatone does not lead to a change of colour we need to apply different methods for verifying substrates. Once we succeed with the qualitative validation via Western Blot analysis, we can verify the presence of nootkatone by using high performance liquid chromatography and mass spectrometry, respectively.

Model influence on Nootkatone expression

We modeled the nootkatone biosynthesis pathway using ordinary differential equations in order to optimize nootkatone production. We found two hard and an easy problem, all of which we could find a solution for. The easy problem is optimization of the enzyme concentrations of the biosynthesis pathway. According to our model of the Nootkatone pathway we found that overexpression of Valencene Synthase is necessary to maximize the Nootkatone yield, while both alcohol dehydrogenase and p450-BM3 have only minor effects on the yield.

One of the hard problems, as shown in our penalty model is the toxicity of nootkatone and nootkatol. Since the toxicity most likely stems from both nootkatone and nootkatol clogging up the cell wall we present our peroxisome as a solution for this problem. When comparing the cytosolic model to our peroxisomal model we found that, if our assumption that neither Nootkatone nor Nootkatol are able to pass the peroxisomal membrane holds up, we can greatly increase Nootkatone production.

The last hard problem is the influx of the pathway precursor farnesyl pyrophosphate (FPP). We used OptKnock analysis to design yeast strains with optimized FPP production. With this analysis we got hints that growing the yeast cells on a fatty acid medium might be a simple alternative to knocking out the desired genes.

Violacein

Working on a project is a process of well planned steps. It starts with an idea and theoretical research to create a design. When finally demonstrating the mechanism of the project it is important to point out the benefits with a suitable application.
Synthetic biology offers countless numbers of new opportunities, especially in the field of metabolic engineering. To testify some main parts of our artico project, we decided to relocate a metabolic pathway into yeast peroxisomes. There are several reasons why this approach fits perfectly as a proof for our concept.
Based on described advantages of violacein this pathway was chosen. Violacein is naturally produced in numerous bacterial strains, most popular in the gram-negative Chromobacterium violaceum . It is related to biofilm production and shows typical activities of a secondary metabolite (Seong Yeol Choi et al., 2015).

relocating the pathway into the peroxisome enables proximity of the enzymes and substrates. Furthermore the yeast cell is protected from the toxic substance hydrogen peroxide. Yeast peroxisomes have no problem with this as their main function is the beta-oxidation of fatty acids and the detoxification of the thereby produced H₂O₂. (Erdmann R. et al., 2007) Because VioC and VioD are FAD-dependent, it is additionally an evidence for FAD location inside of the peroxisome, if the synthesis of Violacein works. Otherwise the two enzymes would not be able to catalyze the reaction.

The genes for VioA, VioB and VioE were amplificated via PCR with Golden Gate compatible overhangs from the biobrick VioABCE (Part:BBa_K274004).

By Golden Gate cloning the peroxisomal targeting sequence (PTS1) was attached to the C-terminus of every pathway protein. Combined with the other necessary parts of the toolbox they represent the level 1 plasmids.

The PTS-tag marks the proteins for the import into peroxisomes. This should first of all point out the functionality of the yeast’s natural import mechanism and also be the basis for demonstrating our own modeled PTS*, proving our designed orthogonal import mechanism. Furthermore we also aim to optimize the working conditions for the enzymes inside of the reaction room - the peroxisomes. For example to vary the pH with new membrane proteins such as bacteriorhodopsin. To secure this change, we can also check the current conditions by our designed sensors.

There are several methods to verify the pathway’s enzymes. First of all, violacein and several intermediates (prodeoxyviolacein, deoxyviolacein, proviolacein) are colorful and the production in yeast can be visualized easily. Furthermore we added a His-/Flag-tag to the N-terminus of every protein (see geneious plasmid cards) to confirm their expression via SDS page and Western Blot. After verifying the presence of the enzymes the next step is to test their functionality. Before performing in vivo experiments in yeast an in vitro assay was implemented. For this the three enzyme pathway leading to PDV was reconstructed, testing VioA, VioB and VioE. To enable the best conditions for the enzymes, the pathway was studied intensively and all needed cofactors were calculated and added to the in vitro reaction (see protocol prodeoxyviolacein in vitro assay). This included FAD, MgCl₂, catalase for decomposition of hydrogen peroxide, and the substrate L-tryptophan. The in vitro reaction was followed by qualitative analysis via HPLC and mass spectrometry.

Outlook

Our outlook is characterized by the vision to use our own modeled PTS* import sequence for an real world application. But first we want to further investigate our in vitro strategy including the missing enzymes VioC and VioD and move on towards already developed in vivo tests. It would be great to qualify a statement about the efficiency of different import combinations into the peroxisome. To do so, different level 2 plasmids were planned. On each plasmid the combination of enzymes being peroxisomal or cytosolic is different.

To assure the import of the enzymes and for further analysis it is indispensable to perform peroxisomal purification for an overall quanti- and qualification. Also measurements of the pathway intermediates and their fluxes across the peroxisomal membrane have to further be analyzed. The final step would be the comparison of the differences in the yield level, depending on the localization of the pathway enzymes (cytosolic or peroxisomal). With this the assumed better production in peroxisomes could be shown.