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ON switch

Figure 1: A simplified overview of biosynthetic pathways of meso-diaminopimelate and lysine. Figure adapted from Born, T. L., & Blanchard, J. S. (1999) [2].


To ensure that our bacteria can survive only in the place of application and cannot spread through the soil in an uncontrollable manner, we designed a biocontainment strategy that makes the survival of our bacteria dependent on the presence of an externally delivered substrate.

In order not to overburden the metabolism of our bacteria with the production of additional proteins, as it is in the case of toxin-antitoxin systems, we decided to follow a different approach. We set out to knock out the dapB gene and thus construct a strain auxotrophic for diaminopimelate (DAP) and lysine which are compounds essential for the survival of bacteria [1]. dapB encodes for dihydrodipicolinate reductase, an enzyme playing a key role in the biosynthesis of DAP which serves as a direct precursor of lysine (Figure 1) [2].

The dependency on an externally delivered "ON switch" compound can be achieved by expressing dapB on a plasmid under the control of an inducible promoter. After assessing several compounds, cuminic acid was chosen as the best candidate, as it is soluble in water, inexpensive, considered not to be metabolised by microorganisms, non-toxic to plants in low concentrations needed for the induction of the system, and absent in the soil [3-5]. Moreover, it enables for tightly controlled gene expression depending on the concentration of cuminic acid [4, 5]. A simplified mechanism of action of this system is presented in Figure 2.

Figure 2: A schematic representation of the cuminic acid-inducible system. CymR is constitutively expressed under the control of a strong PJ23100 promoter. In its native state, the protein acts as a repressor of Pcym inhibiting dapB expression. Upon binding to a cuminic acid molecule, CymR cannot interact with Pcym, thus enabling the expression of dapB.

Experimental design and results

Growth and toxicity assays

In order to determine the applicability of a cuminic acid-inducible system for our project, we checked whether this compound can be used by P. fluorescens SBW25 for growth or has a toxic effect on the cells. To assess that, we performed a series of optical density (OD600) measurements using a plate reader. Both assays were performed on M9 medium. In the growth experiments, both glucose (positive control) and cuminic acid were used in the concentration of 20 mM (Figure 3). For the toxicity assay, concentrations of 0 (negative control), 0.1, 0.5, 20, and 100 mM we chosen (Figure 4).

Figure 3: Plate reader growth experiment with M9 medium and 20 mM cuminic acid. As a positive control, we used P. fluorescens SBW25 growing on M9 with 20 mM glucose.

Figure 4: Plate reader toxicity assay performed on M9 medium with 20 mM glucose and cuminic acid in the concentrations 0.1, 0.5, 20, and 100 mM. As a negative control, we used P. fluorescens SBW25 growing on M9 without the addition of cuminic acid.

The outcomes suggest that cuminic acid cannot be effectively metabolised by P. fluorescens SBW25 to sustain growth. Moreover, it has no toxic effect in concentrations up to 0.5 mM which is significantly higher than the concentrations needed to for the maximal induction of the expression system. These results show that cuminic acid is suitable for the purpose of this project.


The protocol used for creating the knockout was adopted from Wirth et al. (2019) [6]. In brief, this method makes use of DNA recombination forced by double-stranded cuts introduced by I-SceI meganuclease and it consists of two main steps. First, a suicide plasmid pGNW harbouring two homology arms (H1 and H2) flanked by I-SceI sites is introduced into the genome as a result of a single recombination event (Figure 5A). Successful integration can be easily visualised, as the plasmid also contains a sfgfp transcriptional unit. In the next step, the cells are transformed with pQURE6-H carrying I-SceI meganuclease. Upon induction by toluic acid, this enzyme creates lethal double-strand breaks and thus forces a second recombination event. This can result either in the restoration of the wild-type genotype or deletion of the gene of interest (Figure 5B).

Figure 5: Schematic overview of the knockout protocol. A: Integration of the suicide plasmid pGNW into the genome via a single recombination event occurring between either one of the homology arms. B: Genome modification by I-SceI meganuclease delivered to the cell on the pQURE6-H plasmid. H1 and H2 represent the homology arms, red blocks represent I-SceI sites recognized by the enzyme.

Figure 6: E. coli DH5α λpir containing pGNW-dapB-KO. Fluorescence of the cells suggests correct plasmid assembly.

To construct the suicide plasmid, the sequences corresponding to 500 bp upstream and downstream of dapB were amplified from the genome of P. fluorescens SBW25 and cloned into the pGNW backbone between two I-SceI sites. The plasmid was assembled via Golden Gate and transformed into E. coli DH5α λpir via chemical transformation (Figure 6). Fluorescent colonies growing on LB with kanamycin were screened via colony PCR and the correct assembly was confirmed by Sanger sequencing.

The suicide plasmid was then successfully introduced into the genome via tri-parental mating, a form of conjugation in which a helper strain facilitates the transfer of a plasmid between two other bacteria. In this experiment, these were P. fluorescens SBW25, E. coli DH5α λpir containing pGNW-dapB-KO plasmid, and a helper strain E. coli HB101 (Figure 7A). The confirmation was achieved by performing colony PCR (Figure 7B) with the primers annealing to the 3' end of H1 (forward) and 5' end of H2 (reverse). The double bands correspond to the bacterial chromosome after a single recombination event as shown in Figure 5A.

Figure 7: A: Fluorescent colonies of P. fluorescens with pGNW-dapB-KO incorporated into the genome. B: Agarose gel visualising the results of a colony PCR of P. fluorescens after transformation with pGNW-dapB-KO. Two bands of the expected size proved that the plasmid was successfully incorporated into the genome.

To construct the suicide plasmid, the sequences corresponding to 500 bp upstream and downstream of dapB were amplified from the genome of P. fluorescens SBW25 and cloned into the pGNW backbone between two I-SceI sites. The plasmid was assembled via Golden Gate and transformed into E. coli DH5α λpir via chemical transformation (Figure 6). Fluorescent colonies growing on LB with kanamycin were screened via colony PCR and the correct assembly was confirmed by Sanger sequencing.

Figure 8: Red fluorescent colonies of P. fluorescens after transformation with pQURE6-H.

Red colonies were chosen for genotyping via colony PCR (Figure 9). The primers used were annealing right outside the homology arms. The expected size of the bands was 1850 for a wild-type, and 1043 in the case of a mutant.

Figure 9: Agarose gel visualising the results of a colony PCR after transformation of P. fluorescens containing pGNW-dapB-KO integrated into the genome with pQURE6-H. The size of the bands indicates that all the colonies have a wild-type genotype.

Given the unsuccessful outcome, we modified our approach. Instead of supplementing the cells with DAP and lysine from the medium, we transformed them with pSEVAb33 overexpressing dapB under the control of a strong promoter J23100 (BBa_J23100) and then followed through the knockout procedure. The plasmid was assembled via Golden Gate and transformed into E. coli DH10β. After confirmation of the correct assembly, the plasmid was delivered to P. fluorescens containing pGNW-dapB-KO. Afterwards, the cells were transformed with pQURE6-H via tri-parental mating using P. fluorescens with pGNW-dapB-KO incorporated into the genome as an acceptor, E. coli DH10β containing pQURE6-H as a donor, and E. coli HB101 as a helper strain. The transformation was successful, however, colony PCR once again shown that all the colonies had a wild-type genotype. Despite going through several design-build-test-learn cycles, we were not able to knock out the dapB gene with the methods used.


The fact that we could not obtain a knockout even when introducing dapB on a plasmid suggests that the genomic copy remained essential for the survival of the cell. One possible explanation is that the high levels of enzyme resulting from overexpression were too burdensome for the cells. This could force the plasmid to mutate making it unable to sustain the growth of cells with a knockout. To check if this hypothesis is true, we could sequence the overexpression plasmid and check for mutations affecting the expression dapB. If such mutations were discovered, a possible solution would be to exchange the promoter and/or RBS used with another combination that results in lower expression levels.Alternatively, the same procedure could be applied to knocking out another essential gene than dapB, which could result in creating a strain auxotrophic for other compounds than DAP and lysine.

Off switch


For our product to be both safe and convenient to use, we set out to implement a system allowing the farmers to interrupt its activity at any given moment. For this reason, our bacteria will be equipped with a kill switch activated by the presence of an externally delivered “OFF switch” substrate. Considering the immense number of bacteria present in the soil, we aimed to create a kill switch that would limit the risk of native species taking up the modified genetic material during horizontal gene transfer (HGT). Therefore, we decided to employ a system based on a nuclease that degrades the DNA of the modified bacteria, thus reducing its availability to the surrounding microorganisms.

Experimental design

Killing mechanism

CRISPR (clustered regularly interspaced short palindromic repeats)- Cas (CRISPR-associated) system serves as a procaryotic adaptive immune system that protects the cell by cleaving foreign nucleic acids originating from plasmids or viral infections [1]. SuCas12a2 is a type V Cas nuclease originating from Sulfuricurvum sp. PC08-66. The study published at the beginning of this year by Dmytrenko et al [2] revealed that upon recognition of RNA complementary to crRNA, Cas12a2 triggers an abortive infection by cleaving ds- and ss-DNA, as well as ss-RNA in a non-specific manner (Figure 1). This feature makes SuCas12a2 a perfect candidate to be used in our project as an OFF switch. We received the plasmid with the nuclease from the Helmholtz Institute for RNA-based Infection Research.

Figure 1: The effect of a purified SuCas12a2–crRNA complex on different FAM-labelled nucleic acids. Figure adapted from Dmytrenko et al. (2023) [2].

In the case of our bacteria, the RNA target originates from the barcode sequence introduced to the genome of P. fluorescens SBW25 for detection purposes. Within this sequence, there are four PAMs recognizable by Cas12a2 (Figure 2), based on which we designed the spacers (Table 1).

Figure 2: The barcode sequence knocked into the genome of P. fluorescens SBW25.

Table 1: The spacer design for the CRISPR array.

Spacer Sequence (5’ - 3’)


Given the novelty and complexity of the chosen killing mechanism, and the resulting potential difficulties with its implementation, we decided to settle for a better-researched inducible system. Therefore, we identified three systems that could be suitable for use in this project: the lactose-/IPTG-inducible LacI-Plac, the arabinose-inducible araC-ParaBAD, and the rhamnose-inducible rhaSR-PrhaBAD. The main advantage of the first one is that both compounds that can activate it are naturally absent from the soil. However, due to its leakiness, we rejected the idea of using it in the killing mechanism. The choice between the other two systems was made based on the results published by Meisner J, Goldberg JB. (2016) [3] who compared their performance in Pseudomonas aeruginosa, a relative of P. fluorescens. The study shown significant differences in the efficiency of both systems proving that rhaSR-PrhaBAD enables much tighter control over gene expression and is eighty times more sensitive to the inducer.

Therefore, we chose this system for our experiments. One additional advantage of this system is that besides L-rhamnose, which may be found in soil (in low concentrations) and is known to be metabolised by some bacteria, it can also be induced by L-mannose. Kelly et al. (2016) [4] proved that this sugar acts as an analogue of L-rhamnose exerting the same inducing effect on the rhaBAD promoter, while not being metabolised by the cell (E. coli). A simplified mechanism of action of this system is shown in Figure 3.


We began with the characterization of the rhaSR-PrhaBAD expression system by performing fluorescence assays. To obtain the inducible gene expression in P. fluorescens, which does not have the rha operon, the entire rhaSR-PrhaBAD sequence has to be introduced to the cell. Additionally, as a result of expressing the entire system from a plasmid, its sensitivity significantly increases. To get a more complete picture of how rhaSR-PrhaBAD system works in P. fluorescens SBW25 we performed the assays using both inducers (Figure 4).

Figure 3: A simplified mechanism of the rhaSR-PrhaBAD expression system. L-Rhamnose binds to the transcription factors RhaR and RhaS. Next, the former binds upstream of its own promoter PrhaSR, and the latter activates the PrhaBAD promoter controlling the expression of the gene of interest (GOI).

Figure 4: Corrected fluorescence (GFP/OD600) of P. fluorescens with rhaSR-PrhaBAD inducible expression system after 24 hours of exposure to the inducer. A- System induced by L-rhamnose, B- System induced by L-mannose.

The results show that the system is indeed inducible by both L-rhamnose and L-mannose. The curves indicate a switch-like response, which is very promising for use in our project. However, in the end, we did not manage to use this system for the induction of the killing mechanism, as we struggled with assembling the complete plasmid. The sequencing results not only showed that the inserts were not cloned correctly, but also revealed many mutations in the backbone. One of the possible reasons may be the big size of both inserts, and/or the possible toxic effects of SuCas12a2. Due to time constraints, we were not able to experimentally determine the exact reasons.

Barcode detection LAMP assay


With the purpose of detecting and distinguishing our engineered P. fluorescens bacterium apart from the wild type, that ubiquitously lives in the soil, we sought to barcode our engineered bacteria. Once released into the environment, a designed Loop-mediated Isothermal Amplification (LAMP) assay allows for the detection of the bacterium through specific targeting of the barcode. Ultimately, this allows farmers to assess if colonisation and eradication actions of our GMOs in the soil are successful. Therefore, the creation of the LAMP test also prevents false frost protection expectations in the change of unsuccessful soil colonisation. Additionally, the barcode indirectly enhances the bacterium’s biosafety, as the barcode also represents the target sequence of the Cas12a2-based kill mechanism.

Stage 1 - Barcoding

A barcode should be unique, bio-orthogonal (devoid of any biological significance) and not present a burden to the cell. Barcodes allow for barcoded bacteria to be distinguished from its peers and have the ability to convey information on the GMO’s properties and origin [1].

We have inserted a barcoded construct with some additional features in P. fluorescens SBW25 (Figure 1) in the innocuous attT7N transposon site downstream of the glmS gene. The barcode construct includes a rhamnose promoter, necessary for the induced kill-switch mechanism, in which, through rhamnose induction, the transcribed Barcode’s mRNA represents the target sequence for a Cas12a2-based kill-switch. Moreover, the rhamnose promoter and the barcode were flanked by two strong terminators to ensure no unwanted transcription of the barcode. Lastly, two homology arms of 500 bps were placed on either side of the construct with the purpose of homology recombination in P. fluroescens.

The barcode fragment was obtained by annealing complementary synthetic oligonucleotide strands together. The homology arms were amplified from the P. fluorescens SBW25 strain and the remaining parts were amplified SEVA plasmids present in the laboratory. The used primers for the amplifications were designed for Gibson assembly of these in mind. Once ligated the linear construct was then inserted in the integrative pGNW backbone and transformed into E. coli DH5α λpir.

For the genomic insertion of the construct in the attT7N site in P. fluorescens, a triparental conjugation protocol was executed with the strains: E.coli HB101 (containing helper plasmid), E. coli DH5α λpir (containing barcode pGNW) and P. fluorescens SBW25. After successful conjugation, P. fluorescens was transformed with the pQURE6-H plasmid to promote the recombination event and cure the bacteria from pGNW. The integrated barcode was confirmed through Sanger sequencing.

Barcode construct

Figure 1: Knocked-in gene Barcode in P. fluorescens SBW25. As the barcode should be devoid of biological significance we defined the barcode as “iGEM-Wageningen-23” (as read through the genetic code).

Barcode sequence

Figure 2: Barcode construct assembled through Gibson assembly. Rhamnose promoter and barcode are flanked with strong terminators, and these are flanked by the recombination homology arms.

Stage 2 - LAMP assay

The LAMP assay is an alternative amplification technique to PCR [2]. Due to its isothermal character, it can be conveyed without the use of thermocyclers, making it easy to use outside of laboratory environment, e.g., for the farmers. The reaction is isothermal due to the use of a strand-displacing polymerase. Typically, six primers are used to anneal and initiate elongation resulting in a complex mix of amplicons forming an amplification loop.

This method is becoming more established in many fields due to its simplicity, speed, increased specificity, and sensitivity, without the requirement for expensive reagents or instruments [3]. The overall sensitivity and specificity of the LAMP method (96.6% and 95.6%) are very comparable with the ones found in PCR (97.6% and 98.7%) [4]. Although the produced amplicons are not suited for cloning it is very compatible with point-of-care (POC) diagnostics. Due to these factors, it has become very useful in fields such as agriculture, medicine, and food industries, [5].

The design of a functional LAMP test in this project would allow farmers to easily sample soil samples of their fruit tree fields and test in situ for the successful colonisation or eradication of our engineered P. fluorescens through the detection of the Barcode sequence.

For the design of a functional LAMP assay, the software tools NEB LAMP primer design tool and Primer explorer V5 were used to predict possible sets of primers that could isothermally amplify the knocked-in barcode [6, 7]. From these, three possible working LAMP sets of primers were predicted. Although the primer sets were tested to a number of gDNA extraction treatments (direct colony boiling, PBS washing of medium culture followed by boiling and LiOAc-SDS gDNA extraction), no clear positive samples produced the desired fluorescence. This unfortunately indicates that none of the tested primer sets had the necessary specificity to the Barcode.

pseupomona package image

Figure 3: LAMP reaction tubes visualisation under UV light. On the left negative LAMP samples and on the right positive LAMP reaction [8]. The addition of SYBR green dye allows to distinguish negative and positive samples with naked eye.


Despite not being able to find and showcase a working LAMP assay towards our Barcode, further studies should not have any limitation in the ability to demonstrate a functional assay to detect our modified P. fluorescens. Future research iterations could assay more primer sets in the described or other barcodes. Alternatively, a target sequence to already have a described functional LAMP test could also be incorporated into the barcode instead. LAMP assays’s are easier, have lower-cost, higher specificity and sensitivity, and have an elevated point-of-care capability [4,5]. These perfectly encompass the desired properties for an in situ detection method that would elevate the certainty of containment of PseuPomona [1], while being simple enough for farmers to test soil samples of their orchards. Check our applied design page to check how we propose the LAMP detection for farmers!

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