Difference between revisions of "Team:Marburg/Results"

Gibson cloning in Vibrio natriegens

The next step in our project is to test different cloning methods. A new and fast chassis needs a fast and clever cloning strategy. We wanted to show that the most commonly used cloning methods, Golden Gate cloning (Carola Engler et al. 2009)

and Gibson assembly (Daniel G Gibson et al. 2009),

work, using the toolbox we designed for V. natriegens. Demonstrating the functionality of Gibson cloning in V. natriegens, we successfully exchanged a GFP dropout from the pYTK plasmid from the Duber Toolbox (Michael E. Lee et al. 2015), with an RFP dropout. Alex Lepak advised us to try if it made a difference if we take prepared plasmids from E. coli or from Vibrio natriegens. So in total we prepared six Gibson assemblies, combining RFP 1 to 3 (V. natriegens) with pYTK 1 to 3 ( V. natriegens) and RFP 1 to 3(E. coli) with pYTK 1 to 3 (E. coli). After repeating one of the transformations we had colonies on every plate.

Consequently, we send sequenced samples of colonies showing red fluorescence and found all of them to harbor the correctly assembled plasmid without mutations. Cloning efficiency for Gibson assembly is usually higher than 90%, where very few cells still showed green fluorescence. ( Picture 4)

With this, we also prove that the efficiency of electroporation is high enough for Gibson cloning in V. natriegens. We found no significant difference when taking the plasmids from an E. coli preparation. After showing Gibson cloning with two fragments we wanted to check if we could do this method with up to five fragments. To test this we amplified pieces of the pYTK and RFP plasmids according to figure X and Y.

Using the same protocol as mentioned before, and also successfully demonstrated correct assembly, which we affirmed by sequencing. While we had the Echo in our Lab, a machine for “pipetting” very low volumes, down to 2.5 nL, together we also tried doing Gibson assembly with it. The total volume of the mix was exactly one µL. After assembling the samples in the 96 well plate and after transformation we got colonies for every cloning. Again, we counted the cells to calculate the efficiency and again reached over 90% of correct transformants.

Golden Gate cloning and cloning in one day

Our Toolbox, the Marburg Collection is build for Golden Gate cloning and intended to be used in V. natriegens. Implementing this method, we want to enable combinatorial building of plasmids in which you can decide on the position for every part. To demonstrate that this method works, we devised an experiment, of building a Lvl2 strep tag. To create the short tag we ordered two oligonucleotides for an annealing. For the backbone, we again took the pYTK plasmid to have a visual output to identify the incorrect colonies. Triplicates of the annealing and the backbone were combined to give 9 combinations. We performed 18 variants of the experiment in parallel, in V. natriegens and E. coli, to have a direct comparison. After the Golden Gate assembly with BsmB1 we transformed 5µL of each mix by electroporation. All clonings worked fine. Regrettably, the high efficiency resulted in a bacterial lawn, so we couldn’t count the colonies in total. The ratio between fluorescing and non-fluorescing colonies looked quite good. To give a better impression we made pictures under UV light to visualize the colonies producing GFP better.

We sequenced in six different plasmids the Strep-tag and his flanking regions. The sequencings where all correct. The next step for Golden Gate cloning after seeing that it worked, was to attempt if we could do the cloning from transforming the Golden Gate assembly to the miniprep in one day (12h) and build a level two plasmid in three days. For every day the workflow is nearly the same. Transformation, recovery, plating, picking of colonies and harvesting the plasmids from the cells. Overnight you do the Golden Gate assembly for the next day. So we made transformations of eleven level zero plasmids to assemble for the next day a LVL1 GFP LVL 1 RFP ) and a LVL 1 RFP plasmid. From these level one plasmids we wanted to build a level two plasmid with the RFP and the GFP cassette . The only varying factor things in the timeline for each day are the times to pick colonies. On the first day we needed 11.5h to get from the first transformed plasmids to the first finished minipreps. Proving that cloning in under 12h is definitely possible. In Weinstock et al. 2016, it is recommended that you can see the first colonies after four hours and that incubation takes 3h. Also, we could see the first colonies after 4h. Unfortunately, not on all plates. Still, except for one plasmid we could inoculate all after 6h. We then also incubated for 3h. Waiting for the last Plasmid to grow properly took two additional hours, so 13.5 in total. Golden Gate assembly takes about 8.5h

After transforming the LVL 1 and LVL 2 plasmids we got the impression that more complex plasmids take longer to grow. So the whole workflow took us 53h to get the LVL 2 plasmid, which we confirmed later on by sequencing. One major problem is that the visual output of GFP and RFP is that it sometimes takes a lot of time. Our hypothesis is, that because many fluorescent proteins need oxygen to function, and Vibrio natriegens uses a lot of oxygen for his growth, local oxygen levels are depleted so much that they can’t become functional. Therefore, it takes time until we see the visual output on agar plates. A possible solution could be using plates with vents. For picking the LVL1 cells we waited nine hours and incubated 4.5h to be sure to select the correct colonies. The LVL 2 cells took ten hours to grow properly. We think that Plasmid for complex proteins or over-expressing something, the metabolic burden could increase the time of waiting.

Cloning in Vibrio natriegens on the other hand is still much faster than cloning in E. coli, where you also have to struggle with the same problems. Weather you have a normal work day of eight hours or if you work like an iGEMer, you can save in each case a whole day by using Vibrio natriegens. That is immense. You can save third of the time by not waiting that long for cultures to grow. Additionally, there is still a lot of potential to build faster Vibrio natriegens strains. This could revolutionize cloning. All this combined with lab automatisation, clever machines doing the cloning steps, as we did with with the echo or the opentron ,performing heat shock transformations in 96 well plates and pick automatically after automatically plating the colonies. You could start your cloning on Friday, and as we are used to say in Marburg: “ Monday its finished”. This could be what accelerating synbio could look like.

Aquacloning

Aquacloning was first described by Beyer et al. (2015) and is a method for in vivo DNA assembly without the requirement of any additional enzymes. Similar to yeast recombination, linear PCR fragments are transformed into bacterial cells where the fragments are combined by the endogenous DNA repair machinery. The required fragments are PCR amplified from plasmid or genomic templates with primers that add homologous sequences (25 – 30 bp) to the neighboring fragments in a similar manner like Gibson Assembly. Aquacloning can be used for many different applications like site directed mutagenesis or assembling multiple fragments.

Initially, we tried to reproduce the experiment carried out by Beyer et al. (2015), to turn a sfGFP into a blue fluorescent protein by changing a single amino acid (Y66H). We could observe blue colonies and sequenced the plasmids to confirm the desired mutation. An alignment of the chromatogram is shown in figure 1.

After having shown the general applicability in V. natriegens, we tried to use Aquacloning to reproduce the results we gained with Gibson Assembly. We were able to observe correctly assembled plasmids with three but neither with four nor with five fragments.

The experiment was designed such that correctly assembled plasmids result in RFP expressing V. natriegens colonies. The sfGFP CDS of pYTK, the plasmid used in the previous experiment was replaced by a RFP CDS. The resulting agar plate is shown in figure 2.

We do not observe a single white or green colony. The phenotypic appearance of the colonies can be seen as a strong evidence for correctly assembled plasmids. Therefore we were able to demonstrate high efficiency of Aquacloning with three fragments and therefore we present an in vivo alternative for Gibson Assembly.

Due to the limited amount of time, we were not able to repeat the experiments with four and five fragments, but we are certain that optimizing transformation protocols, the amount and ratio of DNA fragments or the length of homologous flanks will result in successfully assembled plasmids even for this very challenging application.

Growth experiments

After reading papers about V. natriegens deciding to order the strain ATCC 14048, we were eager to make our first growth curves.

Because V. natriegens is a marine bacterium organism and from literature (et al), we knew that we had to supplement our standard LB media with an additional 1.5% NaCl.

Being inexperienced with the new organism, we only managed to get a doubling time slightly better than for E. coli. Not disheartened by this, we did some more research and found several points we wanted to try.

We tested the following media, LB 2.5 NaCl, LB v2 salts and Brainheart infusion v2 salts.

v2 salts are: 204 mM NaCl, 4.2 mM KCl, 23.14mM MgCl₂ Brainheart infusion can not be autoclaved together with the v2 salts. From this we learned that growth was most vigorous in BHIv2, closely followed by LBv2. So we decided to make LBv2 our standard media going forward. It is cheaper as well as just being standard LB media, supplemented after autoclaving with the three common salts NaCl, KCl and MgCl₂.

Additionally, some aspects of handling could be improved. For instance reducing the time outside of the incubator for measuring, thereby avoiding cooling of the media, and using baffled flasks to improve aeration. Applying these, we managed to get much better results, even beating the reported doubling time of 9.8 min

(Eagon et al. 1962). The graph shows a growth curve of V. natriegens in BHIv2; V.natriegens in LBv2 and the fastest E.coli stain (NEB Turbo) in LB media.

We achieved an incredible growth rate of 7 min. Also, using baffled flasks, our overnight cultures routinely reached OD's from 14 to over 16. By using enpresso capsules, which slowly release glucose into the media, we could even best that and reach staggering OD's of around 26.

pH- Tolerance of Vibrio natiregens

A defining trait of microorganisms is their pH tolerance. Since we want to produce 3-hydroxypropionate in V. natriegens it is important to know how much acidity our strain tolerates. Therefore, we analyzed its growth under different pH conditions. In order to do this, we prepared LBv2 medium and tested on pHs ranging from 2.5 to 12. Then we inoculated them from a preculture of V. natriegens. After incubating for 15 minutes or for 6 hours we plated the cultures on LBv2 plates with a physiological pH of 7. The exact protocol can be seen in the experiments section.

The experiment showed that V. natriegens tolerates a pH ranging from 6 to 9.5 and has its optimum at pH 7. It was observed that the growth capacity drops drastically when the pH drops under 6 or over 9.5. Nevertheless, in the span from 6 to 9.5 it shows sufficient growth for lab-work. From these results, we learned, that a growth medium optimized for 3HPA production would have to have a pH of 9.5. This would create a larger buffer towards lower pH values, enabling a higher 3HPA yield without killing the organism.

Furthermore, these results confirm that V. natriegens has no problems with pH values in physiological ranges. Since most growth media and buffers work with pH values inside this range, there shouldn’t be any problems related to the pH. Nevertheless, we have information on growth capacity, but we don’t know how protein expression or other cellular processes are influenced. More experiments should be made to further characterize the pH tolerance of V. natriegens, but our results delivered some basic first predictions.

Salt experiments

To find the perfect salt concentration for V. natriegens, we conducted new experiments using different salt concentrations ranging from standard LB media with no additional salt and then added NaCl in 1% steps, up to a concentration of 12 % in total.

We measured the OD600 undiluted too see to which concentration the cells grew in a certain period of time. In a salt concentration of 9%, V. natriegens reached an OD600 of 0.3 after 48h of incubation after slowly adapting from six hours of incubation to 7% NaCL and after 24 hours to 8% NaCl. ( Pictures 1 to 3 )

The pictures 11 to 13 show a gradient of the salt concentration from one to twelve % NaCl in total.

Afterwards we incubated the cultures that showed no growth on media with the ideal concentration of 2.5% and let them recover to check if they were hibernating, but still alive after being exposed to the high salt concentration. Until 10% of total salt in the media, V. natriegens did not recover from growing in higher salt concentrations.

Antibiotic tolerance of Vibrio natriegens

The required concentrations for selection in V. natriegens is still unknown for most standard antibiotics. Since these were a prerequisite for our project, we strove to find them early on.

An easy method to get a good idea of the effective concentrations is to make agar plates with a gradient , then set spots of your culture along that gradient. (Figure 1)

We found that the penicillins ampicillin and penicillin, the arguably most used antibiotics, seemed to be completely ineffective at inhibiting V. natriegens . Fortunately, V. natriegens susceptible for the penicillin derivative carbenicillin. Since the carbenicillin resistance is mediated through the same enzyme as ampicillin resistance, the ß-lactamase, all constructs carrying this resistance can still be selected for.

The concentrations in the following table are working fine in our lab for selecting transformants with the resistance cassettes from our Toolbox.

	WT ingibition [µg/mL]	WT counter-selection [µg/mL]	inhibition with resistance [µg/mL]
Chloramphenicol	2	2	<12
Carbenicillin	50	200	600
Kanamycin	65	200	400
Tetracycline*	7	11	<12
Streptomycin	80	/	/
Gentamycin	30	/	/
Glycloserine	<200	/	/
Penicillin	<100	/	/

(Figure 2 Scheme of the most common antibiotics in synthetic biology and their relevant concentration in correlation with selection V. natriegens with and without a resistance cassette )

The first four antibiotics, Chloramphenicol, Carbenicillin, Kanamycin and Tetracycline are the antibiotics we commonly used in our project. For Golden Gate cloning we routinely switch between Chloramphenicol and Kanamycin. Because, it is highly sensitive to light, and we are using inducible promoters that are activated by a derivative of tetracycline and also reacting to the antibiotic, tetracycline only is used as our emergency antibiotic.

The first transformations we tried were based on E. Coli standard protocols and did not work. In the paper of Weinstock et al. 2016 you can find a protocol for transformation with chemically competent cells and for electroporation. In the protocol for chemically competent cells it is stated that there must be a knockout of the DNAses for it to work. We tried it without the knockout with positive results, but for a re-transformation at least 550 ng of the desired DNA is needed.

We used the pYTK plasmid from the Dueber Toolbox pYTK which has a GFP dropout for visual differentiation, as well as a Chloramphenicol resistance.

We think the issue is that during the making and freezing of the competent cells, a lot of cells die, which releases the DNase from the wild type into the media. The DNases digest the DNA before it can get into the cells.

The reason why the transformation works without the knockout is that the DNases can't digest all 550 ng DNA and some is left over and can be taken up. You only get single colonies and this efficiency is too bad for cloning. The amount of DNA needed is too great for realistic applications and you would still need E. Coli for re-transformation. Since we think the DNase is the root of our problem, after we knock it out we want to give chemically competent cells and heat shock transformation another try. Next we tested the electroporation protocol, because it is reported to have a much higher efficiency. Akira Taket 1988 We made some minor adjustments to the protocol because we did not have the same equipment that was used in the paper and we wanted to use 3000xg instead of the rpm value for the centrifuge to make the protocol applicable to other labs.

We used an electroporator from Eppendorf whose only variable parameter was the voltage. This protocol worked fine with 100 ng of DNA. But the efficiency still left something to be desired. Especially when we think about cloning, so we optimized the protocol and our handling again. We made sure that all buffers and centrifugation tubes were cooled to 4 °C when making competent cells before they reached an OD_600nm of around 0.5 and that the samples were kept on ice to inhibit further growth.

We increased the centrifugation strength because after diluting the cells to a fix OD of 16, we produced 30 aliquots which isn't enough when you do triplicates. We also froze the cells in liquid nitrogen instead of in dry ice. For the electroporation we tested with different gap sizes, ranging from 1 to 2 mm. 1mm proved to work better for Vibrio natriegens. We preheat the recovery media to 50 °C because mixing the 4 °C cells in the chilled cuvette lowers the temperature of the media. Preheating the media to 37 °C would result in a temperature approximating 30 °C after pipetting and mixing with the cold cells. Afterwards we put the electroporated aliquots in a 37 °C water bath to help them recover while we are treating the other samples. More cells survive, therefore the efficiency increases.

In the paper of Weinstock et al. 2016, a voltage between 700 and 900V is recommended. One of our advisers said that a higher voltage increases efficiency so we tested voltages of 700V up to 1600V in steps of 100V.

The difference between 1600V and 900V was not noteworthy. It might be that with an increasing voltage (efficiency)also more cells die during the procedure, resulting in a consistent final efficiency. Electroporating 24 samples takes time and during the first and the last electroporation the first cells have about 15 min more time to recover, directly i a water bath. After applying these new changes the efficiency increased we are now able to successfully transform with concentrations as low as DNA of 0.1ng. Which is nearly what is achieved when working with E. Coli with heat shock. E. Coli still performs better in this, but we are working with the Vibrio natriegens wild type and it is already good enough for cloning.

Working with V. natriegens all the time, we also wanted to see it with our own eyes. To do that, we used several methods of transmission electron microscopy ( TEM ) and scanning electron microscopy (SEM).Our microtomes we made in two ways, by fixation in epoxy resin and in freezing to a grid. Thereby we can show you cuts through the cell (Picture 1-4 ) The method we also showed was platinum coal steaming the cells (Picture 5) and negative staining for visualising attachments like flagella.( Picture 6 and 7 )

On one picture we also have a three dimensional impression of the cells from scanning electron microscopy. (Picture 8 and 9 ) From the size of the pixels and the magnification you can calculate a scale bar, or directly measure the cells. What we can conclude from this is, Vibrio natriegens is about one to two µm long and has a diameter of around 1µm. It has a single polar flagellum and it looks like it is dividing along its equatorial axis. We could also make out something in the cell what could be storage compartments. ( Picture 10 )

At many different occasions, at meetups or conferences, we showed the growth curve of V. natriegens compared to E. coli. Other scientists were impressed by the extremely fast growth rate but even more by the high OD₆₀₀ that we could show. We were asked many times if the high OD₆₀₀ is really due to a high cell density or if it is rather caused by other components like cell debris or substances secreted to the medium which contribute to the measured absorbance.
To find out more about the growth dynamics, we decided to acquire a growth curve of V. natriegens in the most direct way, by counting cells in a flow cytometer. We inoculated three baffled flasks from stationary cultures and took samples in 15 minute intervals while the bacteria were incubated at 37 °C with shaking at 220 rpm. The OD₆₀₀ of these samples was measured in a normal photometer and the cultures were then immediately analyzed by flow cytometry. The flow cytometer directs the samples through a thin capillary so that the cells pass a laser bean one by one and thus, can be counted and analyzed independently. A constant flow rate and time for data acquisition was set, which results in measuring a defined sample volume. Together with the counted events, the cells per volume can be calculated.

A comparison of the OD₆₀₀ to events/µL values are shown in figure 1.
When we planned this experiment, we were most curious about the composition of the culture in the stationary phase to answer the question if the high _OD that V. natriegens can reach is the result of a high cell density or if it can be traced back to other substances. Interestingly, both values, OD₆₀₀ and events/µL start to stagnate at a similar time point (165 min). We interpret this result as a confirmation that the high _OD is indeed caused by bacterial cells.

By carefully comparing the shape of both growth curves, we realized that, in fact, the most striking data in this plot can be found at the beginning of the experiment. While exponentially increasing values can be seen right from the start for the curve created from the OD₆₀₀ data, a short lag phase is apparent when events/µL are plotted (figure 1). We tried to find an explanation for this observation and realized that the absorbance of a culture does not necessarily correlate with the concentration of cells but rather with the biomass inside the flask.

Fortunately, additional data can be obtained from the forward and side scatter of the laser beam in a flow cytometer which provide information about the size and inner complexity of the analyzed cells, respectively. Figure 2 exemplary shows one sample at t = 45 min. The side scatter (SSC-A) is plotted on the Y-Axis versus the side scatter (FSC-A) on the X-Axis. Each dot in this scatter plot represents one detected event and a heatmap can be used to visualize many events with the same properties (red = high event cont; blue = low event count). The population in the top right corner comprises roughly 98 % of all events and represents fully viable cells, since this is the population that increases in number throughout the growth experiment (peaking at >99% of all measured events at mid-log phase). The population in the bottom left corner most likely consists of sick or dormant cells and cell debris, since the number of events in this population remains low throughout the experiment.

We plotted the mean forward scatter values of all cells together with the events/µL. The mean side forward value dramatically increases during the first data points with a peak after 45 minutes. This is also the same time point which we identified as the beginning of the exponential growth phase. During the subsequent course of the experiment the forward scatter values decrease again, reaching a minimum when the culture goes into stationary phase.
Taking all three datasets into account, we suggest that the cells start to grow in size upon provision of fresh medium but initially without undergoing cell division. This results in an increase volume of individual cells and thus, an increase of the measured OD₆₀₀ but without significant changes to the cell concentration in the culture. After 45 minutes, when the forward scatter peaks, we assume that a majority of cells reach maximum cell volume initiate rapid division, quickly entering the exponential phase. During the ensuing time points, exponential growth can be observed and the decrease of the forward scatter is a hint for a reduction in mean cell size.

To additionally visualize the composition of the measured cells in regard to the forward scatter, we created figure 4 showing histograms of the forward scatter at different timepoints. It is apparent that the population is heterogeneous at the beginning of the experiment and at the end when the cultures again reaches stationary phase. During the period of exponential growth, the sample is more homogenous. The already discussed trend in the forward scatter curve can also be observed with these histograms which show a shift to the right when the forward scatter peaks and a shift to the left for the following time points.

We want to thank Dr. Max Mundt who carried out the experiments with us and who helped with analyzing the data.

The goal of synthetic biology is to modify biological organisms in an engineering manner using them as nano-factories for a cheap and environmentally friendly production of valuable resources. To reach this goal a detailed genotyping of the corresponding host is a crucial step. Concerning this we sequenced the genome of our host Vibrio natriegens for our synthetic biological purposes.
One of our goals was to identity and characterize chromosomal integration sites. This made the genotyping a critical step for the design of the correct homologous flanks. But the genome map we built became as well a valuable tool for our whole team for the planing and design of genetic constructs.

We got the organism Vibrio natriegens strain ATCC 14048 from the german collection of microorganisms and cell cultures and sequenced it by Illumina sequencing. Illumina sequencing is a next generation sequencing (NGS) method acquiring data by ‘sequencing by synthesis’ (SBS) chemistry. Since 2014 the short read, massively parallel sequencing technique lead to an explosion of sequencing data output and a drop in cost per base pair, launching the next generation of sequencing methods (Illumina). Using Illumina sequencing, we were able to sequence the whole genome in a manner of hours at an affordable cost.
The workflow of Illumina is based on three steps: Library preparation, short read sequencing and genome assembly. For library preparation several kits can be used. Firstly the template DNA needs to be isolated and purified. Then, the DNA fragmentation and tagging are done simultaneously by a method developed by Illumina called tegmentation. Here bead-linked transposomes carrying Illumina sequencing adapters perform DNA fragmentation, normalization and adapter ligation in one step LINK. After that the fragments are amplified and the fragments of suitable size purified. Then, the library can finally be loaded on a flow cell for sequencing. The flow cell is covered by embedded adapter sequences were the fragments can bind to. By bridge amplification the single fragments are amplified into clonal clusters. Finally the clusters are synthesized with fluorescent labeled nucleotides. A designed software records the light’s wavelength and intensity and saves them into a text file containing the different reads LINK. Finally the reads are assembled into bigger scaffolds by bioinformatic software tools like in this case SPAdes 3.12.0 (Bankevich et al., 2012).

We prepared the library using the Nextera DNA Flex Library Prep Kit LINK by Illumina and sequenced the genome with the MiSeq systemLINK with a coverage of 500-fold. The protocols for library preparation and sequencing were given by Illumina LINK. Subsequently we analyzed the data with SPAdes 3.12.0 (Bankevich et al., 2012), Prokka 1.12 (Seemann, 2014) and Geneious 11.1.5.
The Illumina sequencing yielded multiple DNA contigs – short snippets of the genome – that were assembled to bigger scaffolds using SPAdes 3.12.0 (Bankevich et al., 2012). SPAdes is an open source software developed for genome assembly of single-cell data as well as multi-cell data. Version 3.12.0 is available since May, 2018. Subsequently we looked for annotations with Prokka 1.12 (Seemann, 2014). Prokka is an open source software for prokaryotic genome annotations written in Perl. Finally we checked the genome for secondary metabolites using antiSMASH 3.0 (Weber et al., 2015). Here we found three secondary metabolite clusters for nonribosomal peptides (NRPs), bacteriocin and ectoine (see link below for the .gb file).
NRPs are biologically active compounds built by nonribosomal-peptide-synthetases in a ribosome independent manner. These compounds are interesting for clinical applications due to their use as last resort antibiotics, antitumor or antifungal drugs or as immunosuppressants (Walsh, 2008; Strieker et al., 2010). Bacteriocins are heat resistant antimicrobial peptides produced by bacteria to kill or inhibit the growth of other strains without harming the bacteria themselves. Due to their large diversity in structure and function they are interesting for the industry for food preservation or as new potential drugs in the future (Yang et al., 2014). The ectoine cluster may make this organism interesting for further research and for industry applications as well due to the use of ectoine in medicaments against cold and dermatitis or their use in eye drops and nasal sprays (Werkhäuser et al., 2014).

We checked for published genomes on the NCBI database and found two annotations for Vibrio natriegens ATCC 14048. Using Geneious we aligned our sequences against the database and got an 80.5% similarity against the genome with the accession number NZ_CP009977.1 and a 99.97% identity against the genome with the accession number NZ_CP016345.1. Our sequences can be downloaded from the links below.

Both sequences downloaded from the NCBI database have been sequenced with PacBio. The sequence NZ_CP009977.1 has a coverage of 96.48-fold and was assembled with RS HGAP v. 3 while the sequence NZ_CP016345.1 has a coverage of 178.8-fold and was assembled with HGAP v. 2.

Acknowledgements

We would like to thank Dr. Javier Serrania for the possibility and support sequencing the genome of our host Vibrio natriegens. This work became a useful and very valuable tool for our whole team and our whole project and could not have been done without his support.

Secondary metabolites
Vibrio natriegens Chromosome 1
Vibrio natriegens Chromosome 2



Bankevich A, Nurk S, Antipov D, Gurevich AA, Dvorkin M, Kulikov AS, Lesin VM, Nikolenko SI, Pham S, Prjibelski AD, et al. 2012. SPAdes: A New Genome Assembly Algorithm and Its Applications to Single-Cell Sequencing. Journal of Computational Biology 19: 455–477.
https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3342519/ (October 18, 2018)

Seemann T. 2014. Prokka: rapid prokaryotic genome annotation. Bioinformatics 30: 2068–2069.
https://academic.oup.com/bioinformatics/article/30/14/2068/2390517 (October 18, 2018)

Strieker M, Tanović A, Marahiel MA. 2010. Nonribosomal peptide synthetases: structures and dynamics. Current Opinion in Structural Biology 20: 234–240.
http://www.sciencedirect.com/science/article/pii/S0959440X10000126 (October 18, 2018)

Walsh CT. 2008. The Chemical Versatility of Natural-Product Assembly Lines. Accounts of Chemical Research 41: 4–10.
https://doi.org/10.1021/ar7000414 (October 18, 2018)

Weber T, Blin K, Duddela S, Krug D, Kim HU, Bruccoleri R, Lee SY, Fischbach MA, Müller R, Wohlleben W, et al. 2015. antiSMASH 3.0—a comprehensive resource for the genome mining of biosynthetic gene clusters. Nucleic Acids Research 43: W237–W243.
https://academic.oup.com/nar/article/43/W1/W237/2467910 (October 18, 2018)

Werkhäuser N, Bilstein A, Sonnemann U. 2014. Treatment of Allergic Rhinitis with Ectoine Containing Nasal Spray and Eye Drops in Comparison with Azelastine Containing Nasal Spray and Eye Drops or with Cromoglycic Acid Containing Nasal Spray. Journal of Allergy.
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Yang S-C, Lin C-H, Sung CT, Fang J-Y. 2014. Antibacterial activities of bacteriocins: application in foods and pharmaceuticals. Frontiers in Microbiology 5.
https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4033612/ (October 18, 2018)

When we first presented our project, many people asked about the mutation rate of Vibrio natriegens, worrying its high growth speed would be accompanied by a higher mutation rate. Some were concerned that random mutations could undo their work in the lab, or lead to a pathogenic Vibrio natriegens mutant, others were hoping this could speed up their mutation experiments.

The mutation rate is the frequency with which new mutations appear in an organism. A mutation in a gene can be a silent mutation, meaning it has no effect, or lead to gene function loss, or alter the genes function. These mutations can be the result of DNA-damage, caused for example by radiation. They can also happen spontaneously through mistakes the DNA-polymerase makes during DNA-replication. The spontaneous mutation rate of most organisms is known, such as Escherichia coli’s and Vibrio cholerae’s. It was determined by selecting for mutants which had changed phenotype, (S. E. Luria and M. Delbrück 1943) or in recent years, as it became cheaper, by analyzing the genome trough genome sequencing. This method is more precise than estimating the mutation rate based on a phenotypical change (Patricia L. Foster 2015) (Marcus M Dillon 2017). The mutation rate plays a major role in multiple arears of biology, influencing for example the rate with which pathogenic microorganisms gain antibiotic resistances or adapt to a new environment (Christopher B Ford 2013).

We conducted mutation experiments to estimate the mutation rate (probability of mutation per cell per division or generation) of Vibrio natriegens by using the number of mutation events and the final number of cells in a culture (Patricia L. Foster 2006). To determine the number of mutation events, mutants have to first be identified. This can be done with rifampicin, an antibiotic which inhibits mRNA transcription by obstructing its elongation path through binding to the β-chain. Some microorganisms can gain a rifampicin resistance through a specific point mutation or deletion in the β-chain of the RNA-Polymerase (Wu and Hilliker, 2017). By plating Vibrio natriegens out on rifampicin plates, thes Vibrio natriegens mutants can be selected.

By testing sister colonies of Vibrio natriegens in this manner and counting the resistant colonies we estimated the mutation rate with the Lea-Coulson median estimator (Lea and Coulson, 1949). For this we inoculated 14 sister cultures of Vibrio natriegens with the same OD_{600 (}About 0.0001) out of an exponential pre-culture. When the sister cultures almost reached the stationary phase, we platted them on rifampicin- and on plats without an antibiotic. After evaluating these plates, we could estimate the Median number of mutants in a culture, by counting all rifampicin resistant cultures and applying the Lea-Coulson median estimator. The final number of cells in the culture was estimated by counting the colony forming units on the plates without antibiotics.

The calculated number of mutations per culture with the Lea-Coulson median estimator was 5,7925 and the estimated final number of cells in the culture were 3165000000 cells. The number of mutations per culture is divided by the final number of cell (Patricia L. Foster 2006). The mutation rate we estimated was 1,83017E-09, and there for a bit lower than the mutation rate of Escherichia coli which was estimated by Luria and Delbrück in a comparable experiment for of about with was calculated to be 3,2E-09 (S. E. Luria and M. Delbrück 1943). Since our number of mutations per culture was estimated to be between 4 and 15 we determined the number of mutations per culture with the Lea-Coulson median estimator, unlike how the mutation rate of Escherichia coli was determined. For Escherichia coli the p⁰ method was used, because there were  between 0,3 and 2,3 mutations per culture (Patricia L. Foster 2006). Because Escherichia coli grows to a lesser OD₆₀₀, cultures do not undergo as many cell divisions, thus fewer cells have the opportunity to mutate and less mutation events take place. However, our experiment suggests, that the number of mutants per cells are higher in Escherichia coli, meaning that the possibility to choose a mutated colony is higher when working with Escherichia coli.

We are now able to answer the question of our fellow researchers regarding Vibrio natriegens’ mutation rate. After these experiments we know that concerns about unwanted spontaneous mutations when switching from Escherichia coli to Vibrio natriegens are unwarranted. It can be assumed that Vibrio natriegens can speed up mutation experiments anyway, since there are more mutations occurring in the culture, simply because there is a higher density, increasing genetic variability within it. Our data is not conclusive but enables an estimation of the mutation rate of Vibrio natriegens. A more precisely calculated mutation rate can be obtained through whole genome sequencing of multiple Vibrio natriegens cultures as has been done for Escherichia coli (Heewook Lee et al. 2012).

When we first presented our project, many people asked about the mutation rate of Vibrio natriegens, worrying its high growth speed would be accompanied by a higher mutation rate. Some were concerned that random mutations could undo their work in the lab, or lead to a pathogenic Vibrio natriegens mutant, others were hoping this could speed up their mutation experiments.

The mutation rate is the frequency with which new mutations appear in an organism. A mutation in a gene can be a silent mutation, meaning it has no effect, or lead to gene function loss, or alter the genes function. These mutations can be the result of DNA-damage, caused for example by radiation. They can also happen spontaneously through mistakes the DNA-polymerase makes during DNA-replication. The spontaneous mutation rate of most organisms is known, such as Escherichia coli’s and Vibrio cholerae’s. It was determined by selecting for mutants which had changed phenotype, (S. E. Luria and M. Delbrück 1943) or in recent years, as it became cheaper, by analyzing the genome trough genome sequencing. This method is more precise than estimating the mutation rate based on a phenotypical change (Patricia L. Foster 2015) (Marcus M Dillon 2017). The mutation rate plays a major role in multiple arears of biology, influencing for example the rate with which pathogenic microorganisms gain antibiotic resistances or adapt to a new environment (Christopher B Ford 2013).

We conducted mutation experiments to estimate the mutation rate (probability of mutation per cell per division or generation) of Vibrio natriegens by using the number of mutation events and the final number of cells in a culture (Patricia L. Foster 2006). To determine the number of mutation events, mutants have to first be identified. This can be done with rifampicin, an antibiotic which inhibits mRNA transcription by obstructing its elongation path through binding to the β-chain. Some microorganisms can gain a rifampicin resistance through a specific point mutation or deletion in the β-chain of the RNA-Polymerase (Wu and Hilliker, 2017). By plating Vibrio natriegens out on rifampicin plates, thes Vibrio natriegens mutants can be selected.

By testing sister colonies of Vibrio natriegens in this manner and counting the resistant colonies we estimated the mutation rate with the Lea-Coulson median estimator (Lea and Coulson, 1949). For this we inoculated 14 sister cultures of Vibrio natriegens with the same OD_{600 (}About 0.0001) out of an exponential pre-culture. When the sister cultures almost reached the stationary phase, we platted them on rifampicin- and on plats without an antibiotic. After evaluating these plates, we could estimate the Median number of mutants in a culture, by counting all rifampicin resistant cultures and applying the Lea-Coulson median estimator. The final number of cells in the culture was estimated by counting the colony forming units on the plates without antibiotics.

The calculated number of mutations per culture with the Lea-Coulson median estimator was 5,7925 and the estimated final number of cells in the culture were 3165000000 cells. The number of mutations per culture is divided by the final number of cell (Patricia L. Foster 2006). The mutation rate we estimated was 1,83017E-09, and there for a bit lower than the mutation rate of Escherichia coli which was estimated by Luria and Delbrück in a comparable experiment for of about with was calculated to be 3,2E-09 (S. E. Luria and M. Delbrück 1943). Since our number of mutations per culture was estimated to be between 4 and 15 we determined the number of mutations per culture with the Lea-Coulson median estimator, unlike how the mutation rate of Escherichia coli was determined. For Escherichia coli the p⁰ method was used, because there were  between 0,3 and 2,3 mutations per culture (Patricia L. Foster 2006). Because Escherichia coli grows to a lesser OD₆₀₀, cultures do not undergo as many cell divisions, thus fewer cells have the opportunity to mutate and less mutation events take place. However, our experiment suggests, that the number of mutants per cells are higher in Escherichia coli, meaning that the possibility to choose a mutated colony is higher when working with Escherichia coli.

We are now able to answer the question of our fellow researchers regarding Vibrio natriegens’ mutation rate. After these experiments we know that concerns about unwanted spontaneous mutations when switching from Escherichia coli to Vibrio natriegens are unwarranted. It can be assumed that Vibrio natriegens can speed up mutation experiments anyway, since there are more mutations occurring in the culture, simply because there is a higher density, increasing genetic variability within it. Our data is not conclusive but enables an estimation of the mutation rate of Vibrio natriegens. A more precisely calculated mutation rate can be obtained through whole genome sequencing of multiple Vibrio natriegens cultures as has been done for Escherichia coli (Heewook Lee et al. 2012).

After having established an experimental and data analysis workflow and after determining the optimal plasmidal context for reporter experiments, we started to apply our knowledge to characterize the parts in our Marburg Collection.

We started by measuring the promoter strength of the Anderson Promoter library in V. natriegens. Firstly, we assembled 19 test plasmids with golden-gate-assembly and measured their expression strength, following our selfmade workflows. The results are shown in figure 1. We observed an even distribution of the tested promoters throughout the dynamic range. The strongest promoter (J23100) yielded 40 fold stronger signal than the promoter dummy and was used as a reference to calculate relative promoter strengths. The test constructs were built with dummy connectors which did not possess insulator elements. We assume that this resulted in additional expression caused by transcription throughout the rest of the plasmid, e.g. ori and antibiotic resistance. This is thought to add the same extent of signal to all measured promoters thus reducing the overall dynamic range. To further evaluate this assumption, we could repeat this experiment with one of our insulators instead of the dummy connector.

Characterization of pTet

In addition to constitutive promoters, the Marburg Collection contains two inducible promoters, pTet and pTrc. For all experiments with inducible promoters, we added the respective inducer concentration to the preculture as well as to the main culture to ensure constant expression.The first experiments were performed with the pTet promoter that can be induced by the tetracycline derivative anhydrotetracycline (ATc). ATc is much less cytotoxic but still capable of binding and altering the structure of the repressor TetR, leading to release of the promoter and enabling transcription.

To measure the dose response behavior of the pTet, we made a dilution series of ATc. Following the recommendation of our advisors (Stefano Vecchione), we started with the concentration commonly used in E. coli, started with the concentration (100 ng/mL). The starting concentration was diluted twofold in 20 subsequent steps. Our results are shown in figure 2. The absence of bars for the four highest concentrations is due to the fact that the cultures did not reach an OD₆₀₀ of 0.2 in the six hours of the measurement.
Remarkably, we observed reasonable growth of those same cultures in the preculture already induced with the identical amount of ATc. Knowing that luminescence is produced at the end of an enzymatic cascade, starting with intermediates of the phospholipid metabolism ( Meighen 1991), we reckon that very strong induction could decrease the fitness of cells and that after dilution in room temperature medium, strained cells are not able to recover from the stationary phase.
However, we only observed this phenomenon in experiments with pTet, although we obtained higher signals for the strongest constitutive promoters as well as for the highly induced pTrc. We checked for toxicity of ATc but could not see a measurable effect. Another possibility is that TetR interacts with components inside the cell and that high ATc increases these interactions.
Blast searches of TetR against the genome of V. natriegens identified one protein that shares some homology with the N-terminal part of TetR which could result in cross talk between the host and the inducible promoter.

All measured data were normalized to the strongest constitutive promoter J23100. Saturation occurred at a dilution of 2^6 (~ 1.6 ng/mL) and an exponential reduction of luminescence signal can be observed for higher dilutions. In the absence of ATc, the signal is twelve fold lower compared to saturation.
pTet allows relatively tight control of gene expression and is therefore well suited for driving the expression of potentially toxic proteins. On the other hand, we were not able to induce strong expression that can compete with strong constitutive promoters or the fully induced pTrc.

Characterization of pTrc

pTrc is the second tested inducible promoter. It contains lac operator sites and is therefore regulated by the repressor LacI which is constitutively expressed from a downstream gene.
pTrc can be induced Isoopropyl-β-D-thiogalactopyranosid (IPTG), a chemical derivative of lactose ( Camsund et al.2014). Similar to our experiments with pTet, we made a dilution series starting with the commonly used IPTG concentration for E. coli 0.5 mM. We observed a five fold induction and a saturation that occurred at a dilution of 2^5 (~15 µM). The strongest expression is similar to the expression gained from the strongest constitutive promoter J23100 while the expression in the absence of inducer equals medium strong promoters.
As a consequence, we do not recommend using pTrc in constructs where a tight control of gene expression is desired. Instead, pTrc is well suited when strong expression is required.

Taking the results of both inducible promoters into account, we made two observation. In both cases, the dynamic range is smaller compared to E. coli and the inducer concentration that facilitates saturation is 32 and 64 fold lower for pTrc and pTet, respectively, than the concentration that is typically used for E. coli. A possible explanation could be found in the fast growth of V. natriegens which might result in a lower concentration of the repressor proteins in the cells, finally leading to a less restricted control of the negatively regulated promoters. However, we do not have experimental support for our idea.

One novel key feature of our toolbox are the connectors. They were designed in order to function as insulators to prevent crosstalk between neighboring transcription units (Link to Design). Therefore a perfectly insulating connector would prevent the readthrough from backbone sequences that most probably caused the notably high expression that was measured in the promoter experiment for the dummy promoter. In addition to blocking transcriptional readthrough, a good connector must not possess any cryptic promoter activity.

We focused on characterizing the 5’ Connector because we expect the stronger influence on signal strengths. For characterizing our connector parts, we created 20 test plasmids with the lux operon as the reporter.
In our toolbox we provide five short connectors, which solely possess the fusion sites for LVL2 cloning, and five long connectors which additionally harbor self-designed insulators (Link to Design).
Each of these ten connectors were cloned with the constitutive promoter J23100, to check for effects on an active promoter, and with the Promoter Dummy to quantify the extent of transcriptional activity that reaches the Promoter Dummy.

A

B

The acquired data are shown in figure 4. The data were normalized over the test construct J23100, that was used in the promoter experiment and constructed with the connector dummies. For the five constructs with the active promoter and the long connectors we observed extremely varying signals (figure 4, A). We measured a range from 0.2 to 2 fold change compared to the reference construct. It has been shown that the sequence directly upstream of small synthetic promoters can greatly impact the transcription efficiency ( Carr et al.2017). In case of the long connectors, the sequence upstream of the promoter forms the terminator and could affect the efficiency of RNA-polymerase binding to the -35 and -10 regions.
For the constructs built with small connectors, we also observed varying signals but to a lesser extent compared to the long connectors (figure 4, B).
For all ten connectors that are provided in our toolbox, we show a tenfold range in the measured luminescence/OD₆₀₀ signal. As a conclusion, we recommend to carefully consider the combination of promoter and 5’ Connector for rationally designing constructs.

Taking a look at the constructs that were built with the Promoter Dummy, we also see a huge difference in the expression signals. For the long connectors we expected a negligibly low reporter expression which we observed for two out of five long 5’ Connectors resulting in a 14 fold signal reduction compared to the “Promoter Dummy” reference. We interpret this result as the confirmation that those two connectors function as insluators!
The remarkably strong signal observed for the remaining three connectors could be due to inefficient terminators or cryptic promoters in the pretended “neutral sequence”.

For the remaining five constructs possessing the five short 5’ connectors we observed a range from 0.3 to 5.5 fold compared to the “Promoter Dummy” reference. We are not able to give an experimental explanation for this observation but we could imagine that the LVL2 fusion sites, the only four bases that differ in these constructs, could constitute a weak promoter together with surrounding sequences.
Summarizing the connector characterization, we found that sequences upstream of short synthetic promoters greatly affect reporter expression, which is in accordance with literature ( Carr et al.2017). Moreover, we demonstrated that two of our five self-designed connectors efficiently reduce the signal resulting from other sources than the actual promoter. We additionally conclude that algorithms that predict the “neutrality” of sequences alone are not sufficient to create well functioning insulators.

Natural Competence

In our project we wanted to build many different strains. For this we have to make many genomic changes of the wildtype of Vibrio natriegens. Natural transformation is a good method for multiplex genome editing in a short time (Figure 1).

To get in touch with multiplex genome editing by natural transformation, we constructed a linear DNA fragment for transformation. We have chosen an antibiotic resistance for selection, which has to be different to the resistance marker on the plasmid in the strain (Figure 2).

With a linear DNA fragment containing the chloramphenicol cassette with upstream and downstream regions of the dns gene, we try to delete this gene by natural transformation in a V. natriegens strain without genetic modifications (Figure 3A). To check if the deletion and the natural transformation was successfull, we did colony-PCRs with primers binding inside the inserted fragments (1. upstream-CmR -downstream: ~1940bp; 2. CmR -downstream: ~1440bp; 3. upstream-CmR: ~1440bp). We could prove that the deletion cassette was inserted into the genome and this shows that the natural transformation was successful (Figure 3b).

Using the natural competence regulator Tfox, we were able to transform pYTK. To our best knowledge we were able to show the first natural transformation with a plasmid. We could demonstrate natural transformation using our lab automation process by downscaling all reaction volumes, which has not been shown before. We were surprised since it has been reported that high amounts of DNA are needed for the uptake of the DNA (Zeaiteret al.2017). To make the tfox part accessible to the scientific society, we wanted this part to be integrated into the Marburg Collection. By utilizing our MoClo based cloning system for our tfox expression, the expression level can specifically tailored for the intended application like transforming linear DNA fragment, ssDNA oligos or plasmids.

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Lvl0

For implementing and domesticating the tfox gene into our Marburg collection, we had to built a Level 0 plasmid. At first we amplified the tfoX gene from the pMMB-tfox plasmid (Daliaet al.2017) and build a Lvl 0 plasmid to integrate this part in the Marburg Toolbox (if you want to see how to build a level 0 plasmid take a look at the Marburg Collection). After we were able to construct and build the plasmid, we first did a test digest with EcoRI and PstI to see if it was correct (Figure 4). Then we sent the plasmid for sequencing, we also received confirmation of the accuracy of our plasmid.

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Lvl1

In order to construct two plasmids with full transcription units, with two different promoters we assembled tfoX together with the constitutive promoters J23100 or J23110 into a transcriptional unit by golden gate assembly.

After transformation of this reactions into E. coli we were able to get colonies that seemed to be positive for the Lvl 1 plasmids encoding tfoX because they showed the right size of fragment after plasmid preparation and test digestion, whenever we sent them for sequencing, some bp of the tfox gene were always missing (Figure 6).
We suggest that these problems could be caused because tfox is maybe somehow toxic to E. coli cells.

tet promoter

In addition to constitutive promoters, the Marburg Collection contains two inducible promoters, pTet and pTrc. To build our level 1 plasmid we chose the pTet promoter. The pTet promoter can be induced by the tetracycline derivative anhydrotetracycline (ATc). ATc is much less cytotoxic than IPTG but still capable of binding and altering the structure of the repressor TetR, leading to release of the promoter and enabling transcription.

Using this suitable promoter for V. natriegens we successfully assembled a plasmid containing a full transcriptional unit with a pTet promoter and did not had any problems during the cloning process. By test digestion and sequencing we could prove, that now the sequence of tfox was correct (Figure 7 and 8).

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Establishing Flp/frt in V. natriegens

One of our goals was to create fast genomic modifications, establishing the Flp/frt system in V. natriegens. Therefore, we assembled a plasmid encoding the FLP recombinase, amplified from pBR-flp into the backbone K2560002, creating a Lvl 0 plasmid by Golden Gate assembly. Next, we used this created part K2560271 to assemble a Level 1 plasmid, encoding a full transcriptional unit under the control of a constitutive promoter. This plasmid was transformed into a V. natriegens strain with a successfully integrated chloramphenicol resistance cassette by electroporation. The resulting clones were tested by colony PCR and positive clones were cultivated in LBV2-Kanamycin. The three-phase streaking pattern on LBV2-Kanamycin was performed to get single colonies. These colonies were streaked on replica plates with LBV2-Kanamycin and LBV2-Kanamycin-Chloramphenicol to screen for chloramphenicol sensitive clones. Of 100 clones, only eleven clones were not chloramphenicol sensitive. Eight chloramphenicol sensitive clones were tested by four different colony PCRs: one to amplify the region of the gene of interest, a second PCR to amplify the upstream and chloramphenicol fragment, a third PCR would amplify the downstream-chloramphenicol fragment and the last PCR would amplify only the chloramphenicol cassette. The first PCR should result in a fragment size of 2400 bp in case of a successful excision mediated by the FLP recombinase. In the wildtype genotype, this PCR would result in a fragment size of 3900 bp. The second, third and fourth PCR would not show a fragment in case of the chloramphenicol excision.

In all eight tested clones, the excision of the chloramphenicol resistance cassette was proved.

Thus, we established a new method for creating genomic modifications in the world’s fastest growing organism V. natriegens. This is again one example, that commonly used methods are transferable to this amazing organism, without much effort.
Here we just show that Flp mediated excision is working. In future the functionality of Flp mediated insertion, translocation and inversion need to be demonstrated.

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Based on our self-developed methods for fast genomic modifications, we could establish three strains for accelerated everyday lab use. We established our first cloning strain VibriClone 1.0 and lay the foundation for VibrioClone 2.0. We got first results for VibriExpress on which we can do further work on in the future. With VibriInteract we successfully performed a Vibrio Two-Hybrid assay enabling fast protein-interaction studies.

In our project, we provided a first indication of a successful mutation via CRISPR/Cas9.

As outlined in the description of this experiment, we assembled a LVL2 plasmid possessing the Cas9 CDS as well as the gRNA with the dns spacer. In comparison with the cotransformation of the Cas9-Plasmid and the gRNA-plasmid, the transformation of one plasmid has a higher efficiency, especially when transformed in combination with the donor nucleooligotides.

For the knock-out of the dns in Vibrio natriegens, we cotransformed the LVL2 plasmid once with the suitable oligotides of the lagging strand, once with those of the leading strand and as a control once in the absence of the oligos. Figure 1 shows the plates of these transformations. The control lacking the donor DNA showed no bacterial growth. This indicates that Cas9 is able to cleave the DNA in V. natriegens when coexpressed with gRNA, targeting the chromosome. This leads to the double strand break (DSB) of the DNA and cell death (Jiang et al.,2013). The product of the transformation in which the donor nucleooligonucleotides were provided presented a strong bacterial growth on the plates. As was shown, the LVL2 plasmid is lethal to Vibrio. The addition of donor nucleooligotides facilitates the repair of the DSB through homologous recombination, thereby incorporating the desired mutation into the strand (Wright et al.,2013).

This first experiment suggests that our CRIPSPR/Cas9 is able to generate mutations.

Enzyme activities of Acc from different organisms

Our overall goal was to accelerate metabolic engineering by developing a workflow for high throughput pathway assembly and pathway screening. The first step for that is to find the best enzyme candidates. We tested the native acetyl-CoA carboxylase (Acc) from V. natriegens as well as the Acc from E. coli and S. elongatus together with their activating biotin ligase BirA. In the Choice of Enzymes section of Metabolic Engineering we explained why we chose these enzyme candidates ( link to ori characterization of the part collection). We cultivated V. natriegens expressing these enzyme variants, harvested them and used the French press to get cell extracts. The cell extracts were used for an enzyme activity assay to see if the Accs are functional and which of them is the most active one (Figure 1). For the native Acc we got nearly no measurable activity no matter which amounts of cell extract or acetyl-CoA we added. This result correlates with our expectation that the native Acc is less active than the other enzymes, because it is located on the chromosome and has therefore just one copy. Moreover, the Acc is strictly regulated and only active when fatty acid biosynthesis is required. Since we harvested the cells in the stationary phase, we assume that the enzyme wasn’t in its active state. The Acc from E. coli showed a higher activity but it was beaten by the Acc from S. elongatus, that has a specific activity of 164 mU/mg. We think this high activity is due to the phylogenetic distance, that makes different regulation mechanisms more likely. Therefore, the native regulation mechanisms don’t influence activity of Acc from S. elongatus. Furthermore, the need of high amounts of fatty acids for thylakoid formation makes an active Acc crucial.

The specific activity was calculated via determination of absorption slopes (Fig 2). To ensure that the decrease of absorption is definitely due to malonyl-CoA addition and not addition of cell extract, we first added cell extract and saw no change of absorption. However, after malonyl-CoA addition the absorption instantly dropped. Therefore, we suppose that the measured NADPH consumption correlates with Acc activity. Unfortunately, we haven’t measured Acc activity from C. glutamicum yet because we built the corresponding plasmid library just recently and had no time for biochemical characterization. However, we measured a rationally designed construct bearing accBC, accD and birA from C. glutamicum and mcr from C. aurantiacus and were able to see activity (Fig. 6).

Mcr is soluble and functional in V. natriegens

The second enzyme in our pathway is the malonyl-CoA reductase (Mcr). The enzyme comes from C. aurantiacus, a thermophilic organism, therefore we wanted to test, if the Mcr is correctly expressed and functional in V. natriegens. For that we overexpressed it, lysed the cells via French press and made an SDS-PAGE as well as an enzyme activity assay.

On the SDS-PAGE we compared cell extract from wild type cells with extract from Mcr producing cells. Figure 3 shows, that the band for Mcr with a mass of 131 kDa is only visible in Mcr producing cells. This result confirms, that the protein is soluble and that Mcr production works.

In the enzyme activity assay, we also used cell extract from wild type cells and Mcr producing cells. Figure 4 shows a significant activity with extract of Mcr producing cells. The specific activity is 48 mU/mg. Therefore, we know now, that acetyl-CoA carboxylase, as well as malonyl-CoA reductase are both functional in our strain. The model predicted a higher activity of Mcr compared with Acc, but our measurements suggest the opposite. However, it isn’t possible to compare these data directly, because the calculations of specific activities depend on the protein amount and we used extract instead of purified proteins. If in the Acc extract the percentage of Acc is higher than the Mcr percentage in the Mcr extract, we got incomparable results.