Since the beginnings of synthetic biology, there have always been questions to be raised when genetically modifying organisms: Do they pose a danger to humans, animals or nature?

Therefore, in almost all nations in the world the introduction of genetically modified organisms (GMOs) into the environment is strictly regulated (for more information have a look at our talk with Dr. Ulrich Ehlers) ^[1]. There are good reasons for this: Genetically modified, non-native organisms can have completely unforeseen effects when entering a novel ecosystem. As an example, an inserted antibiotic resistance might be passed on to other bacteria, leading to multiple resistances. This raises the important question: How do we prevent these organisms or their recombinant genetic material from entering the environment?

Most importantly, when working with GMOs, strict attention must be paid to the so-called biocontainment. In general, GMOs are not supposed to leave the designated facility. In the laboratory, disinfection or autoclaving of the equipment are the most common methods to efficiently neutralize all remaining microorganisms. Working inside of special rooms like cleanrooms or at a sterile bench can not only improve the quality and purity of a product, but can further reduce the risk of a GMO escaping the facility.

In contrast to the physical and chemical methods previously mentioned, there is also the option for a biological containment method, called kill switch. A kill switch is a genetic circuit used for selective elimination of a GMO ^[2]. The GMO is supposed to stay in its defined area. When it leaves the defined area, a genetic switch is triggered, leading to the death of the microorganism.

There is a need for biocontainment systems that couple environmental sensing with genetic switch-based control of cell viability. These systems are essential to prevent the escape of GMOs into the environment and thus the responsible application of biotechnology in direct contact to our environment. The genetic switches, generally referred to as kill switches, need to be reprogrammable, in order to change for example their environmental inputs, regulatory architecture and killing mechanism. In the scientific literature, only a few publications deal with kill switches. The best known publications are called "Deadman" and "Passcode" published by the group of James Collins^[2]. Both kill switches were designed for and constructed in Escherichia coli. The Deadman kill switch uses a transcription-based, monostable toggle switch system to ensure fast and robust killing of target cells. In the absence of an inducer not found in nature, e.g. anhydrotetracycline. In contrast, the Passcode kill switch utilizes hybrid LacI-GalR family transcription factors to implement complex environmental requirements for cell survival. Nevertheless, there are further different possibilities for the implementation of genetic kill switch systems. A popular and frequently used approach is the utilization of toxin-antitoxin systems:

This system, as the name suggests, consists of two components: The toxin and the corresponding antitoxin. The two genes that encode these components may be present on a plasmid. If the cells are not under selection pressure, the plasmid will be lost in the next generation and they will die, because the gene that encodes the antitoxin is not present. Thus, the antitoxin is not further produced, remaining antitoxins are degraded and the toxic proteins remains active and kills the new cell^[3].

A well-researched toxin-antitoxin system is the mazEF system in E. coli. But mazEF -like modules are also found in the genome of many other bacteria including pathogens ^[3]. The natural function of the mazEF system is to prevent the spread of a bacterial phage infections. Thereby MazF is a sequence-specific RNA endoribonuclease and thus a toxin that initiates a programmed cell death pathway. Its counterpart, the antitoxin MazE binds to MazF and renders it inactive, but also also prone for enzymatic degradation (see figure 1)^[4]. Various iGEM teams ^{[5, 6, 7]} have built on that the mazEF and implemented their own ideas. iGEM Wageningen 2019 , for example, has used the mazEF system together with the KaiABC system for circadian oscillations in gene expression ^[5]. One common problem with toxin-antitoxin systems is the basal expression of the toxin gene due to promoter leakiness and consequently unwanted cell death.A second example for the use of the mazEF system is provided by iGEM Munich 2012. They used the mazEF system in combination with sigma factor G of Bacillus subtilis to kill sporulating cells. However, the project struggled with too strong expression of the toxin and a consequent imbalance of toxin and antitoxin ^[6].

Another popular approach is engineering auxotrophy: One way to overcome the difficulties of the toxin-antitoxin system is to knock out an essential gene. Essential genes are genes that encode cellular components that are vital for the life of the cell. If essential genes are knocked out, the cell can only survive if it can compensate for the function of the missing component, e.g. synthesis of an essential metabolite, from other sources, e.g. uptake of supplemented metabolites from outside of the cell. Consequently, if bacteria escape into the environment, they cannot compensate for their auxotrophy due to the absence of supplemented essential molecules ^[8].

iGEM NCKU Tainan 2015 used synthetic auxotrophy to switch off the dapA gene encoding 4-hydroxy-tetrahydrodipicolinate synthase. This enzyme is a part of the diaminopimelate pathway and therefore plays an important role in cell wall synthesis ^[9]. If dapA is knocked out, diaminopimelate (DAP) cannot be synthesized and the cells can only grow by taking up external supply of DAP. If the cell leaves the controlled conditions into the environment, it will not survive ^[7].

However, engineering synthetic auxotrophy was not feasible for our project, as these components cannot be easily added in the wastewater treatment plant as the components would be flushed away, before they could enter the cells. Therefore, large amounts of essential molecules would have to be used, which is not cost-effective and not practically feasible. Furthermore, the auxotrophic molecules would then float in the water, and microorganisms could take them up after leaving the biofilm. Consequently, there would be no guarantee that the microorganisms would actually die after leaving the biofilm. We thought of something else: Our approach is based on the idea to use intercellular communication with an of an inducible gene switch to control an essential gene. The expression of the essential gene is genetically modified in a way that it can be controlled by a promoter induced by high cellular density. Within the biofilm, the molecules necessary for the transcription of the essential gene are present, the protein is synthesized and the cell is viable. If the cell leaves the biofilm, the necessary molecules inducing expression of the essential gene are missing and the cell consequently cannot synthesize the essential components and thus cannot survive.

We developed a concept for a kill switch model to hinder the wash-out of living organisms from the biofilm. Thereby our concept went through five phases. In this text, we present our reasoning, show which decisions were based on which essential parameters and how the design of our kill switch has evolved over the course of the project.

The final design or our "phase 5" in the evolution of our kill switch design can be seen in the text Final Kill Switch: Design and Function .

Phase 1

Our initial idea (figure 2) for the basic design of a kill switch consisted of joining two strains of B. subtilis in a communal biofilm. In each strain, an essential gene is knocked out in the genome and the lack of the gene is bridged by the expression of the respective gene from a plasmid. Essential gene expression is controlled by orthogonal promoter systems which are induced upon binding of small molecules produced by the respective communal partner strain. Consequently, the essential genes are only expressed if cells of the two strains are in close proximity to each other, which allows the transfer of inducing small molecules in the biofilm. Only if both inducing molecules are bound, the expression of the essential gene is induced. Cells leaving the biofilm leave an environment with inducer concentrations enabling essential gene expression, which will eventually lead to cell death.

The essential gene we utilized first was dnaA since it encodes a protein that plays an important role in initiating DNA replication in B. subtilis. During the phase 1 of our design creation process we switched to rpsB, because of several diverse factors. Read about our thought process in Which Essential Gene do we Knock Out? .

Our concept is challenged by bringing together cell growth from low to high cell densities and a kill switch design which is activated by high cell density. Although the design appears contrary on first view, we aim to enable biofilm growth under controllable conditions during which the lethal function of the kill switch cannot be activated.

The bacteria are then genetically altered by a quorum sensing signal upon crossing a threshold in cell density, what leads ultimately to the setup of the kill switch. This is accomplished by a novel cassette design:

It consists of the chosen essential gene, in our case encoding the ribosomal protein RpsB (Which Essential Gene do we Knock Out?) which is under control of an engineered promoter region. This promoter region encodes two promoters. One promoter is encoded on the sense strand, the second one is encoded on the antisense strand. Upon passing a cell density threshold, the promoters are ready to be flipped.
An inducer from the outside activates a recombinase which flips the promoter region and thus exchanges the promoter on the sense strand by the one previously encoded on the antisense strand.

By this, the essential gene can be controlled by to different promoters that can be interchanged through recombinase activity.

As mentioned above, the twist with this design approach is that the promoter region controlling the expression of rpsB can by inverted by the recombinase. We named the system “recombinase induced promoter exchange (RIPEX)”.

The RIPEX system and the engineered promoter region consist of two promoters: For the growth phase the constitutive promoter Pveg controls the essential gene rpsB. As already mentioned above (SEE: Evolution of our kill switch design), the growth of the bacteria can be controlled by this promoter and by adding inducer

The second promoter PdegQ naturally controls expression of the gene degQ, with the help of quorum sensing. In quorum sensing, ComA is phorsphorylated ... [hier Verweis auf den Com-system Text, aber nochmal kurz hier 2 sätze vllt dazu] by the phosphorylated regulatory protein ComA+P. PdegQ is located on the antisense strand in contrast to the constitutive promoter which is located on the sense strand inside of the engineered promoter region (NAME, see above). It is intended to be used by the bacteria as soon as the cell density has exceeded a certain threshold value and the inversion performed by a recombinase is induced by Xylose.

This is accomplished by using the Cre recombinase (Cre/lox-system) and two mutated recombination sites designed by the 2007 Paris iGEM Team lox66 (upstream of the desired region, BBa_I718016) and lox71 (downstream, BBa_I718017). After inversion, the sequences are altered and the recombinase does not recognize them as sites, which indicate a possible inversion.

In conclusion, this leads to an irreversible and unidirectional inversion of the promoter region. After the inversion, the essential gene is exclusively and irreversibly under the control of the quorum sensing promoter PdegQ. Consequently, the constitutive promoter Pveg, who is located on the antisense strand after inversion of the combined promoter region, should not lead to expression of the essential gene rpsB.

As stated before, there is another inducible promoter PXylA inside the cassette, controlling the expression of the recombinase. Through induction with Xylose, it can be used to specifically control, when the rotation of the combined promoter region and the switch from the constitutive to the quorum sensing promoter should take place. Or in other words: when the RIPEX system should be activated.

This is accomplished by using the Cre recombinase (Cre/lox-system) and two mutated recombination sites designed by the 2007 Paris iGEM Team lox66 (upstream of the desired region, BBa_I718016) and lox71 (downstream, BBa_I718017). After inversion, the sequences are altered and the recombinase does not recognize them as sites, which indicate a possible inversion. In conclusion, this leads to an irreversible and unidirectional inversion of the promoter region.

Through this novel technique, the problem of homology mentioned before (SEE: Evolution of our kill switch design) is overcome and the early growth phase can be survived by the bacteria in a controllable way. (before the cell density and therefore the concentration of the autoinducer ComX is high enough for the quorum sensing promoter to function)

After the inversion through the Cre/lox-system, the cells are exclusively depended on the high cell density and the community of the biofilm, because the cells receive only within the biofilm necessary for the expression of rpsB. When an individual cell leaves the biofilm, the concentration of ComX in its surrounding environment is lower than inside of the biofilm. The essential gene is not expressed as well as inside the living community and in the expected case the cell dies upon leaving the biofilm.

Since our kill switch is based on linking the availability of an essential gene (product) to the presence of a biofilm, this essential gene first had to be knocked out. In our first approach we chose the gene dnaA . Since it encodes a protein that plays an important role in initiating DNA replication in prokaryotes ^[1Jillian] , its absence leads to cell death. However, at the end of the first phase of our kill switch design we noticed that the choice of this gene caused a problem: dnaA is located in an operon containing another gene ^{[1Jillian], [2Jillian]} . In the case of a knockout by destroying the promoter, the entire operon would no longer be transcribed. This could have led to unpredictable complications. In addition, DnaA is only essential for cell division, so it is possible that the microorganisms do not die but go into dormancy instead.
For this reason, we searched for an easier accessible gene - and found rpsB. Its gene product is the ribosomal protein S2 in the 30S subunit of prokaryotic ribosomes. As one of the biggest ribosomal proteins in the small subunit ^[3Jillian] , RpsB is essential for the functionality of the ribosomes and thus for translation. When a bacterium leaves the biofilm, RpsB will no longer be produced due to our killswitch. This leads to the bacterium being unable to keep up metabolic function, resulting in cell death. Another advantage is that rpsB is the only gene under control of the corresponding operon ^[4Jillian] , therefore being much easier to control.

The implementation of additional, often heterologous, genes in organisms has always been a key step in Synthetic Biology. Several ways to insert new genes in organisms exist. Usually for prokaryotes, a plasmid with the genes of interest is introduced into the bacterium. With a corresponding origin of replication (ori), a fitting promoter region controlling the expression of the gene of interest, and an antibiotic resistance for selection, such a plasmid can often be very effective. Yet, several bacteria cannot robustly carry plasmids. In these cases, the genomic integration of the gene of interest is recommended.

Although B. subtilis can carry plasmids, its highly efficient homologous recombination machinery^[1Robert] allows the integration of our expression cassette in the genome. The cell normally uses this mechanism to repair DNA double-strand breaks, by using a homologous DNA fragment as a blueprint. This machinery also works without the generation of double-strand breaks. Therefore, genes of interest can be integrated directly into the genome of B. subtilis. Only a plasmid with double-stranded DNA, containing our genetic cassette, an antibiotic resistance for selection and homologous sequences to the genome, are required for insertion (see Figure XY)^[2Robert]. Theoretically a homology of ~70 base pairs is sufficient, however, we decided to use around 500 base pairs of homologies for both flanks after talking to our advisors from the Kabisch Lab.

We decided to use this technique to integrate our kill switch cassette into the genome of B. subtilis. This additionally prevents our cassette from being lost during replication, which is a major problem when using plasmids, as a constant selection pressure has to be maintained.

Our concept of a kill switch is solely theoretical, because we were not able to work on the kill switch in the wetlab due to the COVID-19 pandemic. For the practical continuation of this project, it is essential that we can test the functionality of the kill switch. A critical parameter of the kill switch to characterize is for example the threshold of cell density at which the kill switch system is activated.

Most importantly, we have to ensure that our kill switch does what it is supposed to do: Killing our genetically modified Bacillus subtilis cells upon leaving the biofilm environment. We expect that the cells will die, if the essential ribosomal gene rpsB is not expressed, because many bacterial antibiotics inhibit the protein biosynthesis or ribosomal proteins. Thereby, some antibiotics are able to kill cells by blocking the activity of these important factors. ^{[8Simone], [9Simone], [10Simone]} Nevertheless, we cannot be fully sure whether knocking out the essential gene kills the cells or only inhibits their growth. Therefore, we have to test in the laboratory, if the cells are viable without the expression of the essential gene. This can be done via live/dead staining. In our planned assay, cells with damaged membrane, which are considered dead or dying, show a red staining, while cells with intact membrane are stained green upon incubation with SYTO 9/propidium iodide stain.

Further, we have to show experimentally, at which cell density the cells produce enough of the autoinducer ComX  to be viable with the quorum sensing promoter P_deqQ solely controlling the expression of the essential rpsB gene. For this purpose, the cells are grown in separate test flasks up to a defined cell density. In the next step the expression of the recombinase is induced to initiate the irreversible inversion of the promoter region. Thereby the constitutive promoter in front of the gene is replaced by the ComX-dependent promoter. We hypothesize that if the cell density is not high enough, the induced switch of the promoter region would lead to the death of the cells. If it is the case, the cell density is high enough after the inducer has been set down for the growth phase, the cells should be able to live without a without an additional inducer. Cell viability and consequently the expression of the essential gene can be determined by live/dead staining, as already described above.