iGEM 2022 — Team unilausanne

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Notebook

## Lab Notebook FitD

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## Lab Notebook Zosteric Acid

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## Lab Notebook Mussels

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Every notebook was made using Benchling and directly exported as pdf.

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Engineering success FitD

## Overview

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Through existing literature on combating invasive zebra and quagga mussels, we came across the bacterium *P. protegens*, which naturally produces an insecticidal toxin called FitD. FitD is encoded by the *fitD* gene and its transcription is activated by FitG. Both genes, *fitD* and *fitG*, are present in the genome of *P. protegens*. Scientific publications on a related *Pseudomonas* strain suggest that the FitD toxin is not only effective as an insecticide, but also toxic to *Dreissena* mussels (Daniel P.Molloy et al., 2013).

Our project consists in genetically modifying *P. protegens* to improve its production of the FitD toxin to kill the invasive quagga mussel effectively. When we genetically reprogrammed our bacteria, we applied engineering principles and underwent the following four stages: design, build, test, and learn.

## 1st cycle - Design

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In order to increase our chances of success, we designed different strategies which independently aimed at overexpressing the FitD toxin in the bacterium *P. protegens*. We, indeed, thought of overexpressing the toxin by either (a) introducing simply an extra copy of its gene (*fitD* ) on a plasmid, or (b) introducing an extra copy of its activator *(fitG* ). Considering that expressing extra copies of these genes might be toxic or represent a significant metabolic burden on our cells, we designed and built vectors bearing both genes under the regulation of a constitutive or inducible promoter.

## 1st cycle – Build

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To achieve the overexpression of the FitD toxin in *P. protegens*, we built four different plasmids containing either the *fitD* or *fitG* genes, which were amplified directly from the genome of the strain *P. protegens* CHA0 (see [Experiments](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27experiments%27%29%20%7D%7D)):

![E.coli growth curve](https://static.igem.wiki/teams/4209/wiki/t-engineering-fitd-bild1.png)

**Figure 1:** Plasmid maps of the vectors built for the overexpression of the FitD protein.

We registered the four plasmids with the numbers BBa\_K4209011, BBa\_K4209012, BBa\_K4209013 and BBa\_K4209014 in the registry for composite [Parts](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27parts%27%29%20%7D%7D).

We chose pSEVA plasmids as backbones for our cloning as they are designed to be compatible with *Pseudomonas* bacteria. Gibson assembly was used to clone either the *fitD* or *fitG* gene in our vector of choice (see Figure 1; see page [Design](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27design%27%29%20%7D%7D) , see page [Experiments](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27experiments%27%29%20%7D%7D) ). The inducible plasmids are based on the promoter lacIq-Ptrc, which IPTG can induce. The constitutive plasmids are based on the promoter EM7. All four constructs provide resistance to the antibiotic kanamycin.

The resulting plasmids were then introduced into competent *E. coli* DH5α cells by electroporation (see [Experiments](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27experiments%27%29%20%7D%7D) ) and cells were plated onto selective media.

## 1st cycle - Test

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Our transformations resulted in colonies growing on our selective media. To check if we had obtained the expected plasmids, we performed colony PCR and visualized our PCR products by gel electrophoresis.

Results from our colony PCR showed DNA fragments of the expected sizes, indicating that we successfully cloned our desired plasmids (Figure 2).

![E.coli growth curve](https://static.igem.wiki/teams/4209/wiki/t-engineering-fitd-bild2.png)

**Figure 2:** Electrophoresis result of colony-PCR products of transformed E.coli. Fragments are labeled and their sizes expected as follows: A is amplified from inducible *fitG* plasmid, 3356bp; B is amplified from constitutive *fitG* plasmid, 1968bp; C is amplified from inducible *fitD* plasmid, 3480bp; D is amplified from constitutive *fitD* plasmid, 3480bp. Ladder used: 1kb Plus DNA Ladder.

Additionally, the PCR products were sent for sequencing to check for any potential mutations and the results obtained confirmed that our plasmid assemblies were successful (see [Notebook](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27notebook%27%29%20%7D%7D)).

Having built our plasmids of interest, we introduced them to *P. protegens* by electroporation. Unfortunately, our first trial to electroporate DNA into our cells was unsuccessful, as we could not obtain transformants on our selective media. (see [Notebook](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27notebook%27%29%20%7D%7D))

## 1st cycle - Learn

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We hypothetised that the failure of our attempt to electroporate DNA into *P. protegens* was due to low concentration of our plasmids. To circumvent this issue, we followed the protocol kindly provided by a laboratory technician from our department working with *P. protegens* CHA0. He recommended using 100 ng plasmid DNA for 50 uL of competent cells. We, therefore, measured our plasmid concentration with Nanodrop and obtained a concentration of 29.6 ng/uL. It allowed us to calculate a required adjusted volume of 3.4 uL to add to our cells for the transformation, but it also failed. We measured our DNA sample again with a Qubit assay for dsDNA to obtain a reliable concentration. The assay indicated an actual concentration of 11 ng/uL for our sample, which was significantly lower than our previous measurements. We then knew we needed more plasmid DNA for the transformation to be successful. (For more details see [Notebook](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27notebook%27%29%20%7D%7D))

## 2nd cycle - Design and build

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To obtain a higher concentration of plasmid DNA, we extracted plasmids from 9 ml of overnight liquid cultures instead of the previously 3 ml we had used. Using a speed-vac device, we also concentrated our sample to a final concentration of over 100 ng/ul by partially evaporating our sample under reduced pressure. (see [Notebook](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27notebook%27%29%20%7D%7D))

Using the obtained concentrated DNA sample for transformation, we successfully transformed *P. protegens* and got transformant colonies on selective media . We learned that a very high concentration of plasmids was needed to transform successfully  *P. protegens*. Having this information at hand, we performed all further transformations of these bacteria with highly concentrated plasmid preparations.

## 2nd cycle - Test

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To check whether our plasmids were indeed transferred correctly into *P. protegens*, we performed colony PCRs and loaded the PCR products on a 1% gel. The PCR products were also sent for sequencing. We obtained amplicons of the expected size and sequencing results confirmed that the genes were correctly cloned into the plasmids (see [Notebook](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27notebook%27%29%20%7D%7D)).

![E.coli growth curve](https://static.igem.wiki/teams/4209/wiki/t-engineering-fitd-bild3.png)

**Figure 3:** Electrophoresis result of colony-PCR products of transformed P.protegens. Fragments are labeled and their sizes expected as follows: A amplified from inducible *fitG* plasmid, 3356bp; B amplified from constitutive *fitG* plasmid, 1968bp; C amplified from inducible *fitD* plasmid, 3480bp; D amplified from constitutive *fitD* plasmid, 3480bp. Ladder used: 1kb Plus DNA Ladder.

At that point, we had successfully engineered all four different constructs of transformed bacteria and could proceed to lyse them and test their efficacy in killing mussels. Our subsequent experiments with mussels showed that our *fitG* construct kills significantly more mussels than the wild-type *P. protegens* CHA0 strain. Detailed information on bacteria lysis and mussel-testing can be found under [Experiments](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27experiments%27%29%20%7D%7D) and [Measurement](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27measurement%27%29%20%7D%7D). The results of mussel-tests can be found under [Proof of concept](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27proof-of-concept%27%29%20%7D%7D).

## 3rd cycle Design

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We had successfully constructed 4 different strains whose toxicity we wished to test on mussels. In the next step, we needed to lyse the transformed bacteria for two reasons:

As we overexpress the protein FitD but not its transporters to the cellular membrane, some FitD is possibly not secreted and remains inside the cell, which may have less effect on the mussels (see [Design](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27design%27%29%20%7D%7D)). Lysing the bacteria would fix this problem because the cell content would be released in the solution.

Another reason for lysing the bacteria is that genetically modified organisms cannot be released into the environment. As we wanted to develop a product that is applicable in real-life, we wished to inactivate the bacteria after they produce the FitD toxin. The final product would contain dried lysed bacteria and FitD toxin and could be applied in mussel-infested pipes.

As we needed the FitD protein to stay stable, heat could not be considered as elimination method. Chemical treatments using alcohol, aldehydes, or chlorine cannot be considered because of their unknown effect on mussels and the environment. We could not use antibiotics either because of the antibiotic-resistance risks. After some research we found that sonication, a method based on applying ultrasonic frequencies to bacterial samples, would be efficient to lyse our bacteria without adversly affecting our protein (see [Experiments](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27experiments%27%29%20%7D%7D)).

## 3rd cycle Test and learn

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We tested if the inactivation by sonication worked by serially diluting and plating our sonicated samples on selective media to count colony-forming units (CFUs). In the first sonication program, we used a pulse of 1s on/ 1s off during 30s. Unfortunately, there were still too many surviving bacteria. By increasing the sonication period up to 40s, we hoped to eliminate the bacteria effectively. We had to recognize that by doing so, the samples began to heat, and we ran the risk that the proteins got damaged.

## 4th cycle design

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At that point, we needed to redesign our inactivation method and started looking for other techniques. We finally had the idea of using bead-beating: a mechanical way of killing bacteria by letting them collide with ceramic beads in a machine that stirs the sample.

## 4th cycle test

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The plating of the bead-beater samples on restrictive media confirmed that this method was effective in lysing the bacteria: by microscopy we observed that 98% of our bacteria are lysed (see [Experiments](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27experiments%27%29%20%7D%7D)). Therefore, we used bead-beating for further experiments.

After the experiments with mussels, it is confirmed that one of our constructs, the *fitG* constitutive transformation, actually kills significantly more mussels than the wild-type strain of  *P. protegens* CHA0 (see [Proof of concept](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27proof-of-concept%27%29%20%7D%7D)).

Engineering success Zosteric Acid

## Overview

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After extensive research on substances effective for combating mussels, we found that certain algae (e.g., eelgrass algae) naturally produce an anti-fouling substance called zosteric acid, which prevents the attachment of bivalves on surfaces. We, therefore, set out to modify bacteria to produce zosteric acid, a sulfate ester of the intermediate p-coumaric acid. (Jendersen *et al*.,2019)

When genetically modifying our bacteria, we followed engineering principles and underwent the following four stages: design, build, test, and learn.

## 1st cycle - Design

We based our design on the following pathway:

![E.coli growth curve](https://static.igem.wiki/teams/4209/wiki/t-engineering-za-bild1.png)

**Figure 4:** The zosteric acid microbial cell factory (Jendersen *et al*., 2019)

Production of zosteric acid can be achieved via a two-step enzymatic pathway. First, the tyrosine-ammonia lyase enzyme (TAL) forms p-coumaric acid by non-oxidative deamination of tyrosine. The sulfotransferase enzyme (PST) then converts p-coumaric acid to zosteric acid. We, therefore, wished to engineer the bacterium *E. coli* BL21 DE3 to express these two enzymes.

Since the production of zosteric acid requires much sulfate, we also aimed to modify our bacteria to increase uptake and activation of sulfate by overexpression of several enzymes, such as the Cys proteins. Ultimately, we designed our expression system as two plasmids: one catalyzing the transformation of tyrosine into zosteric acid (catalytic vector) and another plasmid for sulfate uptake (transport vector). The resulting plasmids were to be transformed into *E. coli* BL21 DE3, a strain with which zosteric acid has already been successfully produced (Jendersen *et al*.,2019).

## 1st cycle - Build

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To achieve the expression of zosteric acid by *E.coli* BL21 DE3, we built four different plasmids. Two of them were catalytic plasmids, and two were transport plasmids.

![E.coli growth curve](https://static.igem.wiki/teams/4209/wiki/t-engineering-za-bild2.png)

**Figure 5:** plasmid maps of vectors designed for the production of zosteric acid

To catalyze tyrosine into zosteric acid, two genes were required: *tal–fjo* and *SULT1A1*, encoding the TAL and PST enzymes, respectively. Since the gene topology may affect the production efficiency, we built two variant plasmids with different gene orders. The genes *tal-fjo* and *SULT1A1* were synthesized and subsequently amplified by PCR (see [experiments](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27experiments%27%29%20%7D%7D)). We registered the *tal-fjo* biobrick under the number BBa\_K4209000 in the registry for basic [Parts](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27parts%27%29%20%7D%7D).

For the transport plasmid, one gene was required for sulfate activation: the *cysDNCQ* gene that encodes the proteins CysDN, CysC and CysQ. An additional gene responsible for sulfate uptake was also required. For this, we designed two plasmids: one plasmid bearing the *cysPUWA* gene and another the *cysP* gene (corresponding to *cysZ* in the pathway) as the uptake gene. The *cys* genes were all amplified directly from the genome of *Bacillus subtilis*. (see [Notebook](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27notebook%27%29%20%7D%7D)). We registered the *cysP* biobrick under the number BBa\_K4209004 in the registry for basic [Parts](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27parts%27%29%20%7D%7D).

By loading the PCR products on an agarose gel, we wanted to confirm whether the amplification of our genes was successful.

![E.coli growth curve](https://static.igem.wiki/teams/4209/wiki/t-engineering-za-bild3.png)

**Figure 6:** Electrophoresis result of gene amplification for both catalytic and
transport plasmids. The expected sizes of the fragments are: *cysP* 3480bp, *cysQ* 3480bp, *cysPUWA* 3812bp, *cysDNC* 2943bp, *SULTA1* 3356bp and *Tal-Fjo* 1968bp. Ladder used: 1kb Plus DNA Ladder.

We obtained bands of expected sizes for all genes (for more details see [Results](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27results%27%29%20%7D%7D)).

For the assembly of these genes we needed backbones that were amplified by PCR from two different template plasmids, pCOLA and pET Duet (changed to pET17b later), chosen for their optimization for *E. Coli* and their IPTG inducible promoters and providing different antibiotic resistances. After amplification, the pET Duet backbone fragments needed to be purified by gel-purification and QIAGEN kit because of nonspecific primer binding. Using Gibson assembly technique, we finally cloned the amplified genes in the corresponding backbones.

## 1st cycle - Test

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Subsequently, *E. coli* DH5α was transformed with our assembled plasmids using electroporation (see [Experiments](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27experiments%27%29%20%7D%7D)). Colony PCR and sequencing verification of obtained transformants confirmed the successful assembly of both transport plasmids: the genes *cysDNCQ* as well as *cysP* or *cysPUWA* were cloned in the plasmid without mutation (see [Notebook](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27notebook%27%29%20%7D%7D)). We registered both transport plasmids with the numbers BBa\_K4209009 and BBa\_K4209010 in the registry for composite [Parts](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27parts%27%29%20%7D%7D).

Unfortunately, no colony of the bacteria transformed with the catalytic plasmids grew on selective media.

## 1st cycle – Learn

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After some electrophoresis tests, we found that the step of purification of the pETDuet backbone was responsible for the failure of the catalytic plasmids’ assembly: After gel-extraction and purification using QIAGEN kit, we performed electrophoresis which showed that after purification, no DNA was present in our sample. Despite multiple attempts, we were unable to perform the purification of the backbone fragments successfully. That is why we chose a new template plasmid for the backbone of the catalytic plasmids.

## 2nd cycle – Design

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We changed the backbone for the pET17b plasmid which allowed specific primer binding for the amplification of the backbone. Therefore, subsequent gel-purification was not necessary anymore before assembling the plasmids. However, the pET17b template plasmid does not contain the domain for Lac-regulation of the catalytic genes, so we decided to obtain this domain from the template plasmid pAND-MCS, donated by Schaerli-lab, and clone it into the new backbone pET17b.

## 2nd cycle - Build

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We performed PCR on the template pET17b to obtain the new backbone and on pAND-MCS to amplify the lacI and Ptac promoter region. Electrophoresis of the amplified fragments gave bands of the expected sizes:

![E.coli growth curve](https://static.igem.wiki/teams/4209/wiki/t-engineering-za-bild4.png)

**Figure 7:** Electrophoresis results from amplifying the different parts of the new backbone. The expected sizes are 3080bp for *pET17b* and 1563bp for *pAND-MCS*.
Ladder used: 1kb Plus DNA Ladder.

With the new backbone pET17 and lac-regulation domain, we assembled the catalytic plasmids using Gibson-technique. Subsequently, we transformed  *E. coli* DH5α with the different catalytic and transport plasmids. The transformed bacteria were incubated overnight on a selective media. The next day, we had colonies that grew on the plates (see [Notebook](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27notebook%27%29%20%7D%7D)).

## 2nd cycle – Test

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After performing colony-PCR on our transformed bacteria, we ran an agarose gel to see whether we got bands of the expected size, which was the case. Sequencing results showed us that all genes were present in the plasmid (see [Notebook](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27notebook%27%29%20%7D%7D)). We registered both transport plasmids with the numbers BBa\_K4209007 and BBa\_K4209008 in the registry for composite [Parts](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27parts%27%29%20%7D%7D).

At this point, we successfully engineered our different transport and catalytic plasmids. In the next step, we had to incorporate the different combinations of plasmids into *E. coli* BL21 DE3. The results of these transformations can be found under [Results](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27results%27%29%20%7D%7D).

## References

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1. Daniel P. Molloy, Denise A. Mayer, Michael J. Gaylo, John T. Morse, Kathleen T. Presti a, Paul M. Sawyko, Alexander Y. Karatayev, Lyubov E. Burlakova, Franck Laruelle, Kimi C. Nishikawa, Barbara H. Griffin. (2013, January 5). Pseudomonas fluorescens strain CL145A – A biopesticide for the control of zebra and quagga mussels (Bivalvia: Dreissenidae). *Journal of Invertebrate Pathology.*
2. Christian Bille Jendresen , Alex Toftgaard Nielsen. (6. September 2019). Production of zosteric acid and other sulfated phenolic biochemicals in microbial cell factories. *Nat Commun 10.*

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Experiments

## Strain and culture condition

All strains of bacteria and plasmids used in this project are listed in Table 1 and Table 2.

**Table 1:** Bacteria used for FitD and Zosteric acid experiments

| Strains | Characteristics | Reference or source |
| --- | --- | --- |
| *E. coli* DH5α | Cloning strain | Strain available in the lab |
| *Pseudomonas protegens* CHA0 | Contains *fitD*, strain we used to overexpress the toxin | Gift from Christoph Keel laboratory |
| *E. coli* BL21(DE3) | Strain we used to produce ZA | Strain available in the lab |
| *E. coli* DH5α cysPUWA-DNCQ | Cloning strain with plasmid encoding for sulfate intake genes | This project |
| *E. coli* DH5α cysP-DNCQ | Cloning strain with plasmid encoding for sulfate intake genes | This project |
| *E. coli* DH5α Tal-SULT1A1 | Cloning strain with plasmid encoding for ZA production catalytic enzymes | This project |
| *E. coli* DH5α SULT1A1-Tal | Cloning strain with plasmid encoding for ZA production catalytic enzymes | This project |
| *E. coli* BL21 (DE3) cysPUWA-DNCQ-T-S | Expression strain with plasmids encoding for *cysPUWA* and *Tal-SULT1A1* genes | This project |
| *E. coli* BL21 (DE3) cysPUWA-DNCQ-S-T | Expression strain with *cysPUWA* and *SULT1A1-Tal* genes | This project |
| *E. coli* BL21 (DE3) cysP-DNCQ-S-T | Testing strain with *cysP* and *SULT1A1-Tal* genes | This project |

**Table 2:** Plasmids used for FitD and Zosteric acid experiments

| Plasmids | Characteristics | Reference or source |
| --- | --- | --- |
| pSEVA2313 | Plasmid with constitutive EM7 promoter, for insertion of *fitD* and *fitG* | 1 |
| pSEVA234 | Plasmid with inducible trc-lac promoter, for insertion of *fitD* and *fitG* | 1 |
| pET17b | Plasmid backbone for insertion of *SULT1A1* and *tal-fjo* gene | Novagen |
| pCOLADuet | Plasmid backbone for insertion of *cysP/PUWA* and *cysDNCQ* genes | Novagen |
| pETDuet | Plasmid backbone for insertion of *SULT1A1* and *tal* genes | Gift from Stephan Gruber laboratory |

**Table 3:** Genes used for FitD and Zosteric acid experiments

| Gene or operon | Purpose | Reference or source |
| --- | --- | --- |
| *fit* operon | Contains a type 1 secretion system, *fitD* gene, outer membrane protein coding gene *fitE* and regulation genes *fitF*, *fitG* and *fitH* | 2 |
| *fitD* | Toxin belonging to a virulence operon with insecticidal and molluscicidal agent | 2 |
| *fitG* | Activator of *fitD* expression belonging to virulence operon | 2 |
| *cysP* | Gene for sulfate intake | 3 |
| *cysPUWA* | Gene for sulfate intake | 3 |
| *cysDNC* | Gene for sulfate intake | 3 |
| *cysQ* | Gene for sulfate intake | 3 |
| *SULT1A1* | Gene encoding for catalytic enzymes for the production of zosteric acid | 3 |
| *tal-fjo* | Gene encoding for catalytic enzymes for the production of zosteric acid | 3 |

*Pseudomonas protegens* and *Escherichia coli* strains were grown in liquid lysogeny broth (LB) medium at 35°C with 220 rpm shaking. Only for ZA production, *E. coli* BL21(DE3) strains were grown in M9 minimal media supplemented with 4 mM coumaric acid and 10 mM of sulfate (chemodex).
For culture on solid medium, all strains were grown on LB agar plates (1.5% (w/v) agar). Ampicillin (100 µM) and kanamycin (50 µM) antibiotics were added to the medium, where appropriate.
Induction of gene expression was achieved by the addition of Isopropyl β-D-1-thiogalactopyranoside (IPTG) in liquid cultures to a final concentration of 0.1 mM; at mid-logarithmic growth phase for FitD expression and directly after culture inoculation for zosteric acid (ZA) production.
To preserve relevant strains, glycerol stocks were prepared by mixing overnight cultures with glycerol (to a final concentration of 20%) in a cryotube and stored at -80°C.

## Preparation of electrocompetent strains

To make electrocompetent  *E. coli*  DH5α and  *E. coli*  BL21(DE3), we prepared a pre-culture and used it for inoculation of the main culture on the next day. The cells were grown overnight, then centrifuged and washed three times with H2O. The washed cells were then used to make glycerol stocks (10% v/v diluted in H2O) of competent cells.
To make electrocompetent *P. protegens*, we made an overnight culture of our bacteria at 35°C. Our cells were centrifuged and washed two times with 1mM MOPS buffer + 15 % glycerol for 5 minutes at 6000 rpm. The washed cells were then directly electroporated with our plasmids of interest as we were warned that we can’t make glycerol stocks of competent *P. protegens* cells.

## Cloning

Polymerase Chain Reactions (PCRs) were performed to amplify genes of interest using Phanta polymerase master mix (Vazyme) and the primers listed in Table 4. PCR programs were set according to provider instructions. PCR products were analyzed by gel electrophoresis on 1% (w/v) agarose (solved in Tris-borate-EDTA (TBE)) containing SYBR Safe DNA dye (running condition: 110 V, 40 min). Once we were sure that the obtained DNA fragments had a correct length, PCR products were purified using ReliaPrep DNA Clean-up and Concentration system kit (Promega) according to the manufacturer's instructions. If necessary, DNA fragments were extracted from agarose gels using QIAquick Gel extraction kit (Qiagen) according to the manufacturer’s instructions.
To construct our plasmids from the different DNA fragments produced via PCR, we performed Gibson assembly using HiFi mastermix (NEBuilder) that contains 5’ exonuclease. The Gibson assembly reaction mixes were incubated at 50°C for one hour. Electrocompetent *E. coli* DH5α were transformed by electroporating 1 µL of Gibson assembly reaction mix into 50 µL of competent bacteria in a 2 mm electroporation cuvette at 2500 V (time constant was around 5 and 6 ms). After electroporation, 500µL of SOC medium were added and the liquid culture was then recovered at 35°C. After the incubation time, the liquid cultures were then plated on LB agar plates with the appropriate antibiotic and incubated at 35°C for 24 hours.
Colony PCR were performed using primers listed in Table 3 to screen for the correct plasmid constructs. Single bacterial colonies were picked from agar plates and resuspended in 100 µL of LB with the corresponding antibiotic. 1 µL of the resulting solution was mixed with 20 µL of water, 2 µL of forward and reverse primer (Table 3) and 25 µL of Taq polymerase master mix. Colony PCRs were performed using the standard PCR program adapted based on the annealing temperature of the primers and target fragment length (1 minute per 1000 base pairs).
Once the correct constructs were identified, the corresponding colonies were grown in liquid LB media supplemented with the appropriate antibiotic at 35°C for 16 hours and plasmids were extracted using PureYield Plasmid Miniprep System kit (Promega) according to the manufacturer’s instructions. All of the obtained plasmid constructs were verified by Sanger sequencing (Microsynth, Switzerland). For large constructs (e.g. plasmids containing fitD gene), Nanopore sequencing (Plasmidsaurus, USA) were used for verification instead. For overexpression of FitD protein, correct plasmids were electroporated into *P. protegens* CHA0 strain. For production of Zosteric acid, combinations of plasmids (Table 1) were electroporated into *E. coli* BL21(DE3) strain.

## Sample preparation for mussel survivability assay

Starting cultures were prepared by growing  *P. protegens* strains in liquid LB media (50 mL) supplemented with kanamycin and 0.1 mM IPTG (if induction is necessary) overnight. Optical density (OD) of these cultures were determined by measuring absorbance at 600 nm. Once the OD is assessed, we proceed to wash the cultures three times by centrifugation at 4’400 rpm for 10 minutes and resuspension in 25 mL of 1x PBS. Then, the samples were divided into two, for the lysed and non-lysed sample preparation.
For the lysed sample preparation, cell suspensions were centrifuged at 4’400 rpm for 10 minutes to obtain cell pellets, and then these pellets were resuspended in 1 mL PBS. These concentrated cell suspensions were transferred into a 2 mL reaction tube, and glass beads (diameter 100 µm) were added into each sample to reach a final volume of 2 mL. The reaction tubes were then placed in the bead beating machine (FastPrep-24; MP Biomedicals) and a program of 8 cycles consisting of 30 seconds beating and 30 seconds cooling was applied. This experiment was performed inside a 5 °C cold room to ensure that the bead beating machine and the samples did not overheat. Light microscopy and serial dilutions were performed to assess the effectiveness of the cell lysis treatment (Figure 1). Approximately 98% of the cells were lysed after this treatment.

![explanation of FitD](https://static.igem.wiki/teams/4209/wiki/t-experiments-1.png)

**Figure 1:** Cells A. before and B. after bead-beating treatment.

After cell lysis treatment, the concentrated cell suspensions (including the glass beads) were transferred into a 50 mL Falcon tube and resuspended to reach the initial volume, in our case 50 ml. This will restore the initial concentration of lysed cells. Lysed samples were diluted with lake water to reach the desired OD (OD 1, 0.5, 0.25, 0.125). Finally, 10 mL of these cell suspensions were added to Petri dishes containing each one mussel.

## Keeping quagga mussels alive

Mussels were taken fresh from Lake Geneva prior to every experiment. The amount of mussels taken was dependent on our experimental design, on which we applied the 3R (Replace, Reduce, Refine) principle.
If you are interested in the procedure that we used to keep mussels alive in our laboratory you can check our page “[Contributions](https://2022.igem.wiki/unilausanne/contribution)”

## High Performance Liquid Chromatography

High performance liquid chromatography (HPLC) was performed to quantify the production of zosteric acid. In preparation for HPLC analysis, the overnight liquid cultures of *E. coli* BL21 (DE3) containing the zosteric acid production constructs and grown in M9 media were spun down for 10 min at 4’400rpm. The supernatant was collected and filtered to eliminate bacterial residues and sent for HPLC analysis. HPLC was performed at the EPFL facility with a HPLC Waters 1500 model using a C18 column (75mm x 4.6mm, particles of 3.5µm diameter) and the mobile phase consisted of MeOH 20% as solvent A and a tampon phosphate (pH=3 ; 0.0125M) as solvent B. Injection volume was 20 µL and flow rate was set to 0.8 ml/min. HPLC standard curves for the detection of zosteric acid and coumaric acid were prepared using pure samples obtained from Toronto Research Chemicals Inc and Chemodex, respectively.

**Table 4:** List of primers used for our experiments

| Primers | Sequence | Purpose |
| --- | --- | --- |
| D-D1/cPCR3 | cacgcccaagtcctacatcag | Colony PCR of fitD |
| D-2fr1 | catggagcagcacctggtg | Extract *fitD* from *Pseudomonas protegens* |
| D-2fr2 | caccaggtgctgctccatg | Extract *fitD* from *Pseudomonas protegens* |
| A1 | accgagctcgaattcgcg | Extract backbone from plasmid and do gibson assembly |
| A2 | cgtcgtgactgggaaaaccc | Extract backbone from plasmid and do gibson assembly |
| D-B1 | cgcgaattcgagctcggtatggcttttatgtccaaggacttcac | Extract *fitD* from *Pseudomonas protegens* |
| D-B2 | gggttttcccagtcacgacgtcaggtcagtgaaggcaccag | Extract *fitD* from *Pseudomonas protegens* |
| cPCR2 | gtcagccagttcagacgcac | Colony PCR of *fitD* and *fitG* |
| G-B1 | cgcgaattcgagctcggtatgcctaacttcgcagatctgg | Extract *fitG* from *Pseudomonas protegens* |
| G-B2 | gggttttcccagtcacgacgctagcgggtgtcctgggtc | Extract *fitG* from *Pseudomonas protegens* |
| G-cPCR1 | cttggtgtccaaccggcaag | Colony PCR of *fitG* |
| MM\_P001 | CTTTAATAAGGAGATATACCatggaattagccgctattttattta | *cysP* gene amplification with overhang BB |
| MM\_P002 | tacgattcccccgccttg | *cysP* gene amplification with overhang on interregion |
| MM\_IR001 | acaaggcgggggaatctgaTCGAACAGAAAGTAATCGTATTG | Interregion amplification with overhang on *cysP* gene |
| MM\_IR002 | gtaagtcgtatttgatccatATGTATATCTCCTTCTTATACTTAACT | Interregion amplification with overhang on *cysD* gene |
| MM\_DNC001 | atggatcaaatacgacttactca | *cysD* gene amplification |
| MM\_DNC002 | tcaggatctgataatatcgttctg | *cysC* gene amplification |
| MM\_Q001 | aacgatattatcagatcctgataacaccgctcacagagac | *cysQ* gene amplification with overhang on BB |
| MM\_Q002 | TGCTCAGCGGTGGCAGCAGttagtaaatagacactctgaacccc | *cysQ* gene amplification with overhang on *cysC* gene |
| FR\_PUWAfwd\_01 | CTTTAATAAGGAGATATACCatggccgttaacttactgaaaaagaac | *cysPUWA* gene amplification with overhang on BB |
| FR\_BBrev\_01 | GGTATATCTCCTTATTAAAGTTAAACA | BB amplification |
| FR\_PUWARev\_01 | TACGATTACTTTCTGTTCGAtcaggcgctttgtgcgag | *cysPUWA* gene amplification with overhang on interregion |
| FR\_INTERfwd\_001 | TCGAACAGAAAGTAATCGTATTG | Interregion amplification |
| FR\_BBfwd\_01 | CTGCTGCCACCGCTG | BB amplification |
| SD\_001 | ATGAACACCATCAACGAATATCTGAGC | *tal* gene amplification |
| SD\_002 | tcaATTGTTAATCAGGTGGTCTTTTACTTTCTG | *tal* gene amplification |
| SD\_003 | ATGGAATTCAGTCGCCCACCC | *SULT1A1* gene amplification |
| SD\_004 | TCAAAGCTCGCAGCGGAACTTG | *SULT1A1* gene amplification |
| SD\_005 | AGTTCCGCTGCGAGCTTTGAtaattaacctaggctgctgc | Backbone amplification |
| SD\_006 | TATTCGTTGATGGTGTTCATggtatatctccttcttaaagttaaaca | Backbone amplification |
| SD\_007 | ACCACCTGATTAACAATtgagcggccgcataatgcttaag | Interregion amplification |
| SD\_008 | GGTGGGCGACTGAATTCCATatgtatatctccttcttatacttaact | Interregion amplification |
| SD\_009 | ACCACCTGATTAACAATtgataattaacctaggctgctgc | Backbone amplification |
| SD\_010 | GGTGGGCGACTGAATTCCATggtatatctccttcttaaagttaaaca | Backbone amplification |
| SD\_011 | AGTTCCGCTGCGAGCTTTGAgcggccgcataatgcttaag | Interregion amplification |
| SD\_012 | TATTCGTTGATGGTGTTCATatgtatatctccttcttatacttaact | Interregion amplification |
| MM\_SeqP01 | CCCTGTAGAAATAATTTTGTTTAAC | *cysP* gene sequencing 1st primer |
| MM\_SeqP02 | GCCGTGTACAATACGATTAC | *cysP* gene sequencing 2nd prime |
| MM\_SeqDNCQ01 | TCCCCATCTTAGTATATTAGTTAAG | *cysDNCQ* gene seqencing 1st primer |
| MM\_SeqDNCQ02 | CAAATGCCTGAGGTTTCA | *cysDNCQ* gene seqencing 2nd primer |
| MM\_SeqDNCQ03 | tgattgaccgcgaccaggcg | *cysDNCQ* gene sequencing 3rd primer |
| MM\_SeqDNCQ04 | atgagatcgacatcagccgt | *cysDNCQ* gene sequencing 4th primer |
| MM\_SeqDNCQ05 | gcagaaattcatctcaatgg | *cysDNCQ* gene sequencing 5th primer |
| FR\_seqPUWAfwd\_001 | ctggctctatagcccgcagg | *cysPUWA* gene sequencing 1st primer |
| FR\_seqPUWAfwd\_002 | ccggaaatatcgcgtggaagac | *cysPUWA* gene sequencing 2nd primer |
| FR\_seqPUWAfwd\_003 | aaccctgtcgctgccgttac | *cysPUWA* gene sequencing 3d primer |
| FR\_seqPUWAfwd\_004 | cgcgaaccggcgacccg | *cysPUWA* gene sequencing 4th primer |
| SD\_013 | ttgtacacggccgcataatc | *tal* gene sequencing 1st primer |
| SD\_014 | cccattcgccaatccggatatag | *tal* gene sequencing 2nd primer |
| SD\_015 | ggatcgagatcgatctcgatcccg | *tal* gene sequencing 3rd primer |
| SD\_016 | atttcgattatgcggccgtgtaca | *tal* gene sequencing 4th primer |
| SD\_017 | ttgtacacggccgcataatc | *SULT1A1* gene sequencing |

## References

1. Silva-Rocha, R., Martínez-García, E., Calles, B., Chavarría, M., Arce-Rodríguez, A., de las Heras, A., … de Lorenzo, V. (2012). The Standard European Vector Architecture (SEVA): a coherent platform for the analysis and deployment of complex prokaryotic phenotypes. Nucleic Acids Research, 41(D1), D666–D675. <https://doi.org/10.1093/nar/gks1119>
2. Péchy-Tarr, M., Borel, N., Kupferschmied, P., Turner, V., Binggeli, O., Radovanovic, D., … Keel, C. (2013). Control and host-dependent activation of insect toxin expression in a root-associated biocontrol pseudomonad. Environmental Microbiology, 15(3), 736–750. https://doi.org/10.1111/1462-2920.12050
3. Jendresen, C.B. and Nielsen, A.T. (2019). Production of zosteric acid and other sulfated phenolic biochemicals in microbial cell factories. Nature Communications, 10(1). doi:10.1038/s41467-019-12022-x.

{% endblock %}

{% extends "layout.html" %}
{% block title %}{% endblock %}
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* [Results FitD](#topic1)
* [Results Zosteric Acid](#topic2)
* [References](#topic3)

Results FitD

## Overview

---

FitD is a toxin naturally produced by  *Pseudomonas protegens*. The *fitD* gene is part of a virulence operon which is regulated by three regulatory proteins, including an activator encoded by the *fitG* gene and a type I secretion system. The FitD toxin has been shown to have insecticidal activity by inducing the midgut epithelium's apoptosis and paralyzing and destroying insect phagocytes (Péchy-Tarr et al., 2013). Interestingly for our project, FitD is also known for its potential as a molluscicidal agent (Molloy et al., 2013).
Hence, we tried to overexpress the FitD protein in *P. protegens* using plasmids bearing either an extra copy of the *fitG* or *fitD* genes. By overexpressing *fitD* or its activator *fitG* in *P. protegens*, we expected that it would impact the survivability of quagga mussels.

![explanation of FitD](https://static.igem.wiki/teams/4209/wiki/results-fitd/t-results-fit-figure.png)

**Figure 1:** Plasmid maps of the constructed plasmids for the overexpression of the FitD toxin in *P. protegens*.

## Construction of the overexpression *fitD* and *fitG* plasmids

---

To improve FitD production in *P. protegens*, we thought of overexpressing the toxin by either (a) introducing simply an extra copy of its gene (*fitD*) on a plasmid, or (b) introducing an extra copy of its activator *fitG*. Considering that expressing extra copies of these genes might be toxic or represent a significant metabolic burden on our cells, we designed vectors bearing both genes under the regulation of a constitutive or inducible promoter (Figure 1). We chose pSEVA plasmids as backbones for our cloning as they are designed to be compatible with *Pseudomonas* bacteria. Gibson assembly was used to clone either the *fitD* or *fitG* gene in our vector of choice. The inducible plasmids are based on the promoter lacIq-Ptrc, which IPTG can induce. The constitutive plasmids are based on the promoter EM7. Ultimately, the following vectors were designed:

* pSEVA234 + *fitD*
* pSEVA234 + *fitG*
* pSEVA2313 + *fitD*
* pSEVA2313 + *fitG*

pSEVA234 refers to an IPTG inducible plasmid and pSEVA2313 refers to a constitutive plasmid.

The *fitD* and *fitG* genes were PCR amplified using the genomic DNA of *P. protegens* as template. The obtained gene fragments were cloned into their respective backbone through Gibson assembly and were transformed into *E. coli* DH5α. After plasmid extraction, the constructs were transformed into *P. protegens*.By performing colony PCR and subsequent sequencing on the resulting transformants, we also were able to confirm the successful cloning of all of our plasmids (Figure 2).

![explanation of FitD](https://static.igem.wiki/teams/4209/wiki/results-fitd/t-results-fitd-figure2.png)

**Figure 2:** Electrophoresis gel confirming the correct assembly of our *fitD* and *fitG* constructs in *P. protegens* by colony PCR scale is given by 1kb plus ladder, A) *P. protegens* pSEVA2313 *fitG* shows a band at the expected size of 1968 bp, B) *P. protegens* pSEVA234 *fitG* shows a band at the expected size of 3356 bp, C) *P. protegens* pSEVA2313 *fitD* shows a band at the expected size of 3480 bp, D) *P. protegens* pSEVA234 *fitD* shows a band at the expected size of 3480 bp.

## Validation of the killing activity of our engineered *P. protegens* against quagga mussels

---

After a trial and error process (see [Engineering](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27engineering%27%29%7D%7D)) we successfully transformed *P. protegens* CHA0 with our newly-built constructs. We then set out to test the efficacy of the obtained four engineered *P. protegens* strains in killing quagga mussels. After testing various conditions and troubleshooting [measurements](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27measurement%27%29%7D%7D), we initially assess the killing activity of our strains on 10 mussels using a final lysed bacterial concentration of 8·108 cells/mL of water (Figure 3).

![explanation of FitD](https://static.igem.wiki/teams/4209/wiki/results-fitd/t-results-fitd-figure3.png)

**Figure 3:** Lysed and engineered *P. protegens* kill quagga mussels. Kaplan-Meier graph shows probability of mussel survival over time when subjected to different lysed bacterial treatment applied to a concentration of 8·108 cells per mL. Gehan-Breslow-Wilcoxon test, \* significant for p-value < 0.05, n = 10 mussels.

The data obtained showed that our engineered *P. protegens* strains were efficiently killing quagga mussel (Figure 3). While the rate of killing appeared extremely similar between the wild–type *P. protegens* and our strain *P. protegens* pSEVA2313 *fitD*, we observed a trend suggesting our strain *P. protegens* pSEVA2313 *fitG* has better activity than the wild-type. However, the bacterial concentration used was relatively high (8·108 cells per mL) and 10 mussels was too little to have enough statistical power to detect significant differences.
We therefore decided to increase the number of mussels tested per condition to 45 as well as to test the efficacy of a lower concentration of cells. Since the workload was exponentially increasing with the number of mussels we were planning to use, we also chose to focus our experiments on the most promising strains only for further testing. As one of the strains is the inducible *fitD*, we wanted to use a strain without an inducer as another control. We ended up testing the following strains:

* P. protegens pSEVA234 *fitD* induced with IPTG
* P. protegens pSEVA234 *fitD* without IPTG
* P. protegens pSEVA2313 *fitG* (constitutive)
* Wild type (i.e containing *fitD* in the genome)

We also tested a lower concentration of cells applied to the mussel in order to see if we could limit the amount of product needed and still significantly improve the killing of mussels (Figure 4).

![explanation of FitD](https://static.igem.wiki/teams/4209/wiki/results-fitd/t-results-fitd-figure4.png)

**Figure 4:** Lysed *P. protegens* cells engineered to overexpress the *fitG* gene show improved killing of quagga mussels compared to wild-type *P. protegens*. Kaplan-Meier graph shows probability of mussel survival over time when subjected to different lysed bacterial treatment applied at a concentration of 2·108 cells per mL. Gehan-Breslow-Wilcoxon test, \* p-value < 0.05, n = 45 mussels.

We were pleased to observe that our engineered and lysed *P. protegens* cells overexpressing the *fitG* gene were able to significantly reduce the probability of survival of the tested mussels (p-value 0.0117; Figure 4). Indeed, our data show that the probability of survival of the quagga mussels decreases remarkably to 86 % after 80 hours when exposed to our engineered strain, compared to the wild-type *P. protegens* did not impact the mussel’s fitness. Although we could not measure a statistically significant difference in the probability of survival of the mussels between the wild-type *P. protegens* treatment and our engineered strain overexpressing the *fitD* gene, we noted that the latter, however, seems to reduce the mussel survival as well modestly. We would therefore argue that both our modified *P. protegens* strains, expressing *fitD* or *fitG*, are able to kill the invasive quagga mussels.
Importantly, at the lower final bacterial concentration of 2·108 cells per mL tested here, the wild-type *P. protegens* cells do not seem to affect quagga mussels at all, whereas our engineered cells demonstrate the killing of mussels. These results show that our constructs significantly improve the quagga mussel-killing capacity of *P. protegens* and would allow its use at low concentrations which could prove to provide a cost-effective alternative to the currently marketed Zequanox product (see [proof of concept](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27proof-of-concept%27%29%7D%7D)).

Results Zosteric Acid

## Overview

---

Zosteric Acid (ZA) has been proven to prevent foot attachment of mussels to surfaces (Ram et Al., 2012). We sought to produce heterologously ZA in *Escherichia coli* by introducing a two-step enzymatic pathway such that tyrosine is first converted into coumaric acid (pHCA) and then pHCA is converted to ZA by the addition of a sulfate group (Figure 1). Therefore, another set of genes, *cysDNC*, *cysQ*, *cysPUWA* and *cysP*, must additionally be added to the pathway as they are responsible for the entry and the transport of sulphate into the cell (cfr [design](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27design%27%29%7D%7D))(Jendresen & Nielsen, 2019).

![explanation of FitD](https://static.igem.wiki/teams/4209/wiki/t-results-za-figure-1.png)

**Figure 5:** Schematic representation of the ZA production pathway. The TAL enzyme converts the tyrosine, naturally present in the bacteria, into pHCA. CysP or CysPUWA are responsible for the uptake of sulphate into the cell. The CysDNC enzymes replace two phosphate groups of ATP by the sulphate. The sulphate carrier is used by SULT1A1 to incorporate sulphate into coumaric acid creating zosteric acid. CysQ molecule is essential to recycle ATP and allow for a second cycle to start.

## Construction of the zosteric acid biosynthesis plasmids

---

### a. Gene amplification:

![explanation of FitD](https://static.igem.wiki/teams/4209/wiki/t-results-za-figure-2.png)

**Figure 6:** Plasmid maps showing the constructed plasmids used for ZA production. A) and B) represent the 2 plasmids containing the catalytic enzymes *tal-fjo* for the conversion of tyrosine into pHCA and *SULT1A1* for the conversion of pHCA into ZA. C) and D) represent the 2 plasmids containing sulphate transporter genes *cysDNCQ* for both, responsible for the transport of sulphate inside the cell, and two different genes (*cysP* and *cysPUWA*) for the entry of sulphate inside the cell.

Ultimately, we designed our pathway to be split onto two separate vectors with one ‘transport’ plasmid, bearing the genes responsible for the intake of sulphate, and one ‘catalytic’ plasmid, bearing the genes responsible for the synthesis of ZA (Figure 2). We obtained the genes for our transport vector, *cysDNC*, *cysQ*, *cysPUWA* and *cysP*, by PCR-amplifying them all from the genome of *E. coli* but for the latter, which we got from the genome of *Bacillus subtilis*. The *tal-fjo* and *SULT1A1* genes necessary for the catalysis of the two conversion reactions (tyrosine to pHCA, and pHCA to ZA, respectively) were ordered from Twist Bioscience. The ‘transport’ genes were to be cloned into the pCola\_Duet vector, while the ‘catalytic’ genes were to be inserted into the pET17b plasmid ([Experiments](%7B%7Burl_for%28%27pages%27%2C%20page%3D%27experiments%27%29%7D%7D)).

![explanation of FitD](https://static.igem.wiki/teams/4209/wiki/t-results-za-figure-3.png)

**Figure 7:** A) agarose gel electrophoresis of amplified genes. Expected sizes: 1=c*ysP* (1017bp), 2=*cysQ* (741bp), 3=*cysPUWA* (3812bp), 4=*cysDNC* (2943bp), 5=*SULT1A1* (876bp), 6=*tal-fjo* (1520bp). B) agarose gel electrophoresis of amplified inter-regions: 1= Inter region of pCOLA (135bp), 2= Inter region of pET17b-S-T (157bp), 3=Inter region of pET17b-T-S (150bp). C) agarose gel electrophoresis of the amplified sequences: 1=pCola\_Duet inter-region (135bp), 2= pCola\_Duet backbone (3.3Kb). D) agarose gel electrophoresis of the amplified sequences: 1= pET17ST BB (Backbone for the insertion of *SULT1A1* and *tal-fjo*), 2=pET17TS BB (Backbone for the insertion of *tal-fjo* and *SULT1A1*), 3 = *LacI* gene for *SULT1A1* and *tal-fjo* plasmid, 4= *LacI* gene for *tal-fjo* and *SULT1A1* plasmid.

Ultimately, after thorough primer design for all the parts we wished to amplify, we successfully PCR-amplified all the genes and linear backbone fragments required to perform our cloning (Figure 3).

### b. Gene assembly:

The obtained gene fragments were cloned into their respective backbone through Gibson assembly and were transformed into *E. coli* DH5α. By performing colony PCR and subsequent sequencing on the resulting transformants, we also were able to confirm the successful cloning of all of our plasmids (Figure 4).

![explanation of FitD](https://static.igem.wiki/teams/4209/wiki/t-results-za-figure-4.png)

**Figure 8:** Gel electrophoresis of colony PCR performed on *E.coli* DH5a transformed with our plasmids. A) agarose electrophoresis of colony PCR performed on the second MCS of both transporter plasmids containing the genes *cysDNCQ*. Expected band size= 3.7Kb. Observed band size=~3.7 Kb. B) agarose electrophoresis of colony PCR performed on the first MCS of one of the two transporter plasmids containing the gene *cysP*. Expected band size= 1Kb. Observed band size=~1Kb. C) agarose electrophoresis of colony PCR performed on the first MCS of one of the two transporter plasmids containing the gene *cysPUWA*. Expected band size= 3.8Kb. Observed band size=~3.8Kb .

The newly-built plasmids were then co-transformed into *E. coli* BL21 (DE3), a strain more adapted for protein production. Indeed, we wished to generate four distinct strains, each containing a different combination of two plasmids, one bearing sulfate uptake genes and one bearing the catalytic enzymes. Unfortunately, we were not able to co-transform *E. coli* BL21 (DE3) containing *pCola\_Duet\_P\_DNCQ* and *pET\_17b\_Tal\_SULT1A1*. We, however, successfully obtained the three following co-transformant that we analyzed further for ZA production (Fig. 5).

* *pCola\_Duet\_PUWA\_DNCQ + pET\_17b\_Tal\_SULT1A1* (PUWA-TS)
* *pCola\_Duet\_P\_DNCQ + pET\_17b\_SULT1A1\_ Tal* (P-ST)
* *pCola\_Duet\_PUWA\_DNCQ + pET\_17b\_SULT1A1\_ Tal* (PUWA-ST)

## Zosteric acid production from our engineered strains

---

To test the ability of our engineered strains to produce ZA, we used HPLC to analyze the supernatant of our cells grown in liquid cultures to detect ZA and pHCA.

### a. Expected results from HPLC:

We first purchased the pure compounds ZA and pHCA and ran them into the HPLC as standards (Fig 5). This allowed us to confirm that we could detect ZA and pHCA with our method, with a retention time (RT) of 5 and 9 min, respectively.

![explanation of FitD](https://static.igem.wiki/teams/4209/wiki/t-results-za-figure-5.png)

**Figure 9:** HPLC chromatograms of standards for Za and pHCA. A) Zosteric acid and B) pHCA.

Next, we analyzed the supernatants of our three mutant strains to assess ZA production. To do so, we cultured overnight the strains in M9 medium supplemented with 4 mM of pHCA, 10 mM of K2SO4-, and 0.1 mM IPTG induction. When supplied with all molecules, we expected our engineered cells to utilize pHCA and produce ZA, resulting in an absorbance peak at 5 min RT corresponding to ZA and the reduction of the peak at 9 min RT for pHCA, for a non-induced control (Fig.6).
Indeed, as a reference control, we additionally analyzed the supernatant of cells grown with the substrates pHCA and K2SO4 (Figure 6) but without IPTG induction. In these conditions, we expected to observe no production of ZA but a significant amount of pHCA corresponding to the added initial concentration that cannot be utilized by the cells.
Lastly, we prepared a second control for each mutant strain where we added IPTG but no supplementation of pHCA and K2SO4. The rationale of this control was to assess the efficacy of the *tal-fjo* gene-encoded protein to convert tyrosine into pHCA effectively. Without added sulfate and pHCA, we reasoned that any detected pHCA would be the product of the *tal-fjo* protein activity. pHCA would most likely accumulate as no added sulfate would limit its further conversion to ZA. Here, we expected to observe the production of pHCA and virtually no ZA. In the case where the basal level of sulfate present in the M9 media used for our experiment would be sufficient for conversion of ZA, we would then expect minimal production of ZA and a reduced amount of pHCA (Figure 6).

![explanation of FitD](https://static.igem.wiki/teams/4209/wiki/t-results-za-figure-6.png)

**Figure 10:** Schematic representation of the testing conditions. The upper part of the image shows the *E. coli* strains that we constructed and tested. The table in the figure represents the specificity for each test and control condition. As we can see, in every test condition, the M9 media has been implemented with 4mM of pHCA, 10mM of K2SO4, and induced with 0.1mM. Control 1 media has been implemented with 4mM of pHCA, and 10 mM of K2SO4, but no IPTG was added; the plasmids are therefore not induced. Control 2 has no implementation of chemicals. Instead, the plasmids have been induced with 0.1mM of IPTG. The two last columns represent the expected results for each condition: ZA should be produced in Test and Control 2, and pHCA should be detected in Control 1.

### b.Observed HPLC results:

After a few preliminary analyses to determine the best conditions under which to perform the HPLC, the following results were obtained:

![explanation of FitD](https://static.igem.wiki/teams/4209/wiki/t-results-za-figure-7.png)

**Figure 11:** HPLC analyses of our modified *E. coli* strains’ supernatant after overnight culture in M9. A) Decrease of pHCA compared to the negative control. Analysis of each transformed BL21(DE3) strain’s (PUWA-TS, P-ST, PUWA-ST) supernatant in test condition : with addition of 4mM pHCA, 10mM SO4- and induction with 0.1mM IPTG. B) Negative control showing the amount of pHCA that was added at the beginning of the incubation. Analysis of each transformed BL21(DE3) strain’s (PUWA-TS, P-ST, PUWA-ST) supernatant with addition of 4mM pHCA, 10mM K2SO4- but no IPTG induction (negative CTRL). C) Production of coumaric acid. Analysis of each transformed BL21(DE3) strain’s (PUWA-TS, P-ST, PUWA-ST) supernatant without implementation of chemicals (pHCA, K2SO4-) but with 0.1mM IPTG induction.

Indeed, we analyzed the supernatant of our three different strains, each bearing a different combination of the transport and catalytic plasmids. We, however, observed the strongest signal with the P-ST and PUWA-ST co-transformants. For clarity, we decided to report only the results obtained for these strains (Fig. 7).
The absorption profile of the supernatant isolated from the PUWA-ST strain showed that in control 2 growth conditions, cells could convert tyrosine into pHCA as a great peak with the correct retention time can be observed while the molecule was not added here (Fig. 7C, PUWA-ST-CHEM). However, comparing the test and control 1 condition for this strain, no decrease of pHCA and detection of ZA can be seen, which suggests that no pHCA is being utilized. Altogether, these observations indicate that the *tal-fjo* is functional in producing pHCA but the PUWA does not appear to allow significant sulfate uptake to permit conversion of pHCA to ZA.
On the other hand, the P-ST strain showed no detectable pHCA in control 2 conditions, while we observed a clear decrease in pHCA between the test and control 1 condition when it was added in the culture. Although we do not detect a peak for ZA itself, we argue that this data suggests that ZA is being produced, albeit below the detection limit of the method used, as pHCA decreases specifically when the cells are induced. We thus considered a notable decrease in pHCA as a proxy for ZA production. Comparing our two strains, we can also conclude that the CysP protein seems more functional than the CysPUWA protein in sulfate uptake, which is effectively used to convert pHCA to ZA, as the decrease of pHCA could only be detected with the *cysP* gene.
In conclusion, we are pleased to report that our data so far suggest our P-ST strain can convert pHCA to ZA. Further work will, however, be carried out to improve our growing conditions and the detection of ZA to confirm this.

## References

1. Péchy-Tarr, M., Borel, N., Kupferschmied, P., Turner, V., Binggeli, O., Radovanovic, D., … Keel, C. (2013). Control and host-dependent activation of insect toxin expression in a root-associated biocontrol pseudomonad. *Environmental Microbiology*, 15(3), 736–750. <https://doi.org/10.1111/1462-2920.12050>
2. Molloy, D. P., Mayer, D. A., Gaylo, M. J., Morse, J. T., Presti, K. T., Sawyko, P. M., … Griffin, B. H. (2013). Pseudomonas fluorescens strain CL145A - a biopesticide for the control of zebra and quagga mussels (Bivalvia: Dreissenidae). *Journal of Invertebrate Pathology*, 113(1), 104–114. <https://doi.org/10.1016/j.jip.2012.12.012>
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