Design
We designed a novel toolbox for complete control over all major functions of the peroxisome. The toolbox is our solution to improve the engineering workflow and predictability of synthetic constructs. Interested? Find out how.
Scientific background
The root problem
Synthetic biology is an engineering discipline. And while we are able to plan our constructs with tools like biobricks, a major difference to e.g. electrical engineering is that our constructs are not nicely isolated on a chip, but surrounded by all types of interfering agents. One of the major issues regarding protein expression in a novel chassis is unwanted and unexpected crosstalk between engineered pathways and the native cellular processes of the production host. The other one is toxicity of the products or intermediates of the pathway. Both can greatly change our system’s behaviour which in some cases leads to us to having to trial-and-error find a solution, making our previous modeled optimization useless.
Our approach
The natural approach of organisms to deal with metabolic interference and toxic byproducts is subcellular compartmentalization. This has proven to be a functional solution in either naturally occurring pathways in eukaryotes or in new synthetic pathways for biotechnological application. Thus, the creation of a synthetic organelle presents a suitable strategy to increase the efficiency and yield of non native pathways. A common approach is to build up artificial compartments from scratch. Many breakthroughs have been achieved in the last decade, however the creation of a fully synthetic compartment is yet a milestone to reach for. We on the contrary want to start by engineering artificial compartments through orthogonalization.
The peroxisome is the ideal starting candidate as it has many advantages over other compartments, including it being able to import fully folded proteins and not being essential in yeast under optimal growth conditions. Our projects aim is to create a toolbox for manipulating and creating customizable peroxisomes as a first step towards synthetic organelles.
By modifying the import machinery of yeast peroxisomes only selected proteins will be imported into the peroxisomes leaving the researcher in full control over the content of the peroxisomal lumen. Furthermore our toolbox will include a secretion mechanism for the synthesized products, various intra-compartmental sensors, modules for the integration of proteins into the peroxisome membrane, as well as optogenetic control for some of these parts for a more precise spatiotemporal control.
As a proof of concept for the functionality of the toolbox and the customizable compartment two metabolic pathways will be integrated into the altered peroxisome: (i) Violacein biosynthesis and (ii) Nootkatone biosynthesis. Violacein, a bisindole formed by condensation of two tryptophan molecules, is a violet pigment and thus easy to quantify in the cell. Nootkatone on the other side is a natural compound found inside the peel of the grapefruit, which gives it its characteristic taste and smell. In addition, Nootkatone is a natural mosquito and tick repellent that is already being commercially used and industrially manufactured. Unfortunately, the production costs are extremely high. Furthermore, the production of Nootkatone inside yeast is challenging as it is toxic for yeast and thus, the production efficiency is rather low. A successful implementation of the synthetic compartment will show increased yields in the production of these compounds and showcase the potential of this approach for similar future applications.
Cloning strategies and the Yeast Toolbox for Multipart-Assembly
Describing our cloning strategies we mentioned several levels, which stand for different stages of our plasmids. They are further described in the work of J.M. Dueber and colleagues, who designed the well established yeast toolkit we used in this project Michael E. Lee (2015) . The toolkit offers the possibility to design plasmids with desired antibiotic resistances, promoters as well as terminators from standardized parts. It also provides fluorescence proteins, protein-tags and many more useful components as part plasmids. These part plasmids are distinguished in different part types due to their specific overhangs to ensure their combination in the correct order (e.g. promoter - gene of interest - terminator) all in a versatile one-pot Golden Gate reaction without time-consuming conventional cloning steps.
The cloning steps regarding the plasmid levels are implemented in E.coli in order to reduce the required time to generate the final plasmids. The different levels are therefore defined by their part content and their antibiotic resistances.
To generate a level 0 plasmid, the gene of interest is ligated into the provided level 0 backbone via golden gate assembly using the enzyme BsmBI. The backbone contains a resistance to Chloramphenicol, as well as an origin of replication, creating a very basic yet functional plasmid.
The level 1 plasmid contains a promoter and terminator suited for S. cerevisiae. There is the possibility of including a polyhistidine-tag if there is a need for Western blot analysis. The antibiotic resistance contained in the level 1 plasmid changes from chloramphenicol to ampicillin which enables filtering out residual level 0 plasmids contained in the Golden Gate product. Furthermore, the Dueber toolbox includes the possibility of designing GFP-Dropout cassettes. These are custom-built level 1 backbones whose inserts are sfGFP as well as promoter and terminator suited for E. coli. Upon a successful cloning step the GFP is replaced by the part(s) of interest, and a correct colony shows a white colour. In case of a wrong ligation event colonies show a green fluorescence. This provides a very useful tool to detect unsuccessfully cloned colonies. The enzyme used for level 1 changes from BsmBI to BsaI to avoid any interference between different steps.
The level 2 plasmid combines two or more genes of interest with their respective promoters, terminators and tags. The resistance changes from ampicillin to kanamycin. The enzyme of this step is BsmBI again. This level is useful, if the construct you are designing requires multiple genes to be transformed into one yeast strain.
Design of our sub-projects
Protein Import
The peroxisome has two pathways for importing proteins with the main transport proteins being PEX5 and PEX7. We created an orthogonal PEX5 binding pocket and corresponding recognition peptide (PTS1) by structural modeling. We also created a library of PEX7 recognition sequences for import of proteins incompatible with the PTS1 peptide.
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 developed at late point in our project − as it turned out the research group of Alison Baker et al. did similar work on the PEX5 receptor and published a paper in September 2017. They designed a receptor which is a combination of the PEX5 from Physcomitrella patens and Arabidopsis thaliana that did interact with a new PTS. Understandably, we were curious if we could use their results to get an orthogonal import pathway in yeast and did sequence alignments to figure out which amino acids have to be changed.
We then set point mutations at the corresponding positions in the yeast’s PEX5 and started molecular dynamics simulations with a couple of PTS variants − one of them was actually the variant they used in the paper (YQSYY). The details and results of our strucutral modeling can be found in the modeling section. Fortunately, evaluation revealed that this receptor variant could be a promising candidate and thus we synthesized it. Together with the 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 − we chose the transmembrane domain of PEX13 tagged with mRuby.
Once we figured out our experimental design, we thought about our constructs.
Pex13−mRuby
We wanted to use the peroxisomal membrane protein PEX13 as a fluorescent marker, but instead of using the whole protein which could severely influence the peroxisomal properties we just used the transmembrane domain with a short linker. Literature research revealed that such constructs have been tested before − Erdmann et al. (2004) described a construct containing only PEX13200-310 with a C-terminal GFP. Instead of GFP we used mRuby to get a higher differentiation from mTurquoise which we use for another construct described later.
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 binding 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
Membrane integration
PEX19-dependent
ER-dependent
Peroxicretion
Membrane permeability and size control
Sensors
Applications
Nootkatone
Model influence on Nootkatone expression
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. 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.
Violacein
Working on a project includes 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. Because of several reasons this approach fits perfectly as a proof for our concept.
On the basis of described advantages of violacein ( link to project description ) 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)
.
The synthesis of Violacein requires five enzymes encoded by the VioABCDE operon. VioA, a Flavin-dependent L-tryptophan oxidase and VioB, a heme protein, work in combination to oxidise and dimerize L-tryptophan to an IPA imine dimer. Hydrogen peroxide is released as a by-product of the VioA reaction. Next step by VioE is the rearrangement of the IPA imine dimer to prodeoxyviolacein acid, which can non-enzymatic oxidise to the prodeoxyviolacein or, by VioC via deoxyviolaceinic acid, oxidate to pink deoxyviolacein. VioC and VioD,flavin-dependent oxygenases, contain 19 nucleotide binding amino acids, which require interaction with the oxidised form FAD (flavin-adenine dinucleotide) (uniprot) (uniprot) . The two enzymes act sequentially: first, VioD hydroxylates protodeoxyviolaceinic acid, leading to protoviolaceinic acid. Second, VioC creates the oxindole at the 2-position of one indole ring, leading to violet violacein (Balibar CJ et al., 2006) .
By relocating the pathway into the peroxisome, the yeast cell is protected from the toxic substance. 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, we additionally have 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 GoldenGate compatible overhangs from the biobrick VioABCE
(Part:BBa_K274004).
By GoldenGate cloning the peroxisomal targeting sequence (PTS1) was attached to the C-terminus of every pathway protein. Combined with the other necessary parts ( toolbox explanation ) they represent the level1 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 (LINK import).
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 (LINK membrane proteins). To secure this change, we can also check the current conditions by our designed sensors (LINK 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 prodeoxyviolacein 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 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.
