Module 1: Laccase production

Reduction and transformation of soil organic matter is a problem of great significance when it comes to forest fires’ consequences. A severe and intense fire not only causes volatilization of organic compounds, resulting in loss of organic matter but also promotes the formation of aromatic structures such as polycyclic aromatic hydrocarbons (PAHs), which are toxic to plants and increase soil’s hydrophobicity. ^{1, 2, 3} Moreover, studies suggest that in many cases phenolic compounds, lignin and tannin-like molecules increase due to thermal degradation caused by wildfires and can be phytotoxic or cause problems to soil’s nitrification. ^{4, 5}

Degradation of such compounds is a time-consuming and difficult process to be done naturally, which makes forest regeneration very slow. ^{1, 6} Our goal is to speed up this procedure. After a literature study, as well as conducting with Mrs. Pappa, Associate Professor of Genetics - Molecular Microbial Genetics in National and Kapodistrian University of Athens, we learned that White Rot Fungi are one of the few microorganisms that can degrade phenolic, lignin and hydrophobic compounds in general by secreting enzymes, such as laccases and peroxidases. ^{7, 8}

Choosing the host microorganism

While white rot fungi seemed to be a promising potential for being part of Euphoresis, due to their unique features, engineering such a fungus would be beyond challenging for us. As Mrs. Effimia Papaptheodorou, a professor at the School of Biology at Aristotle University of Thessaloniki, specialized in soil microbiology, explained to us, engineering fungal organisms in the lab can be a difficult and challenging situation. This would mean that fungal transformation attempts could probably be a failed attempt, and we would neither assure a desirable laccase production, nor improve the microorganisms’ biosafety aspects (inserting a kill-switch circuit, for instance). Our initial thought was to insert a fungal enzyme gene into a bacterial strain, but as Mrs. Pappa informed us, this would be a rather tricky process since it involves the insertion of an eukaryotic organism’s gene into a prokaryotic one. ^{9, 10} Moreover, upon searching on papers and articles, we came across the fact that fungal laccases are glycosylated molecules, but their glycosylation can only occur in fungal cells, thus it wouldn’t be possible in a bacterium, post-translationally.⁸

Fortunately, it didn’t take us long to realize that bacterial lignin-degrading enzymes do exist as well, while diving into the beautiful chaos of bibliographical knowledge.⁸ Mrs. Pappa also reassured us that those enzymes function in a similar way like the fungal ones and they can be effective in the same way in our project. Bacterial laccases, for example, can thrive in temperatures as high as 70° C. There is a wide variety of them, functional in different pH ranges.⁸ Taking those factors into account, we realized that engineering a bacterial strain to produce and secrete such enzymes in soil would be an ideal concept in our case.

At the end of the day, we chose Bacillus subtilis to serve as the host microorganism. B. subtilis, the Microbe of the Year 2023, is a gram-positive, aerobic bacterium found in soil. It has an optimal growth temperature of 30-35°C, making it capable of surviving in soil temperatures, and a doubling time of 20 minutes.^{11, 12, 13} It has also been used as a booster of plant growth (PGPR).

Initially, our strain of choice was the B. subtilis 3601 strain, which was available in the University’s Department of Biology. However, during the experimental process we did not detect the desired enzyme production. One of our concerns was that the initial strain choice was not suitable for the experimental design, since it relied solely on the strain’s availability in the laboratory. Before making new decisions, we conducted Mr. Alexiou, a PhD Candidate in Biotechnology and Synthetic Biology and a B. subtilis expert, who suggested that we use a protease-free strain, such as the WB800. Our final decision was indeed to utilize the WB800 strain, which does not produce extracellular proteases that may cause degradation of the ligninolytic enzymes, since their respective genes have been genetically knocked-out. ¹⁴

All those properties made B. subtilis the most suitable candidate to contribute to Euphoresis’ effort to reduce soil hydrophobicity and create a more suitable environment for the growth of new plant life.

Laccases or peroxidases? Choosing the most suitable enzyme for the job…

With the B. subtilis WB800 strain in our hands, we then needed to determine which of the enzymes we encountered in literature would fit our aim and purpose. To make a proper decision, we set the following criteria:

1. 1. The enzyme needs to show resistance in high temperatures, because soil after wildfire seems to be vulnerable to direct sunlight which causes soil temperature rise. A 2010 study in a Colorado burned area showed that factors, such as forest canopy or duff layer, that cooled down soil temperatures by 5°C did not affect soil temperature post-fire. [15]
2. 2. The enzyme must be functional in alkaline-pH environments, since forest soil after a wildfire has alkaline pH in numerous cases, due to soil deposition and the resulting release of alkaline compounds (see Integrated Human Practices, Mr. Dimitrakopoulos Alexandros).1
3. 3. Due to B. subtilis’ encapsulation in microspheres inside the hydrogel, with small pores, we needed our enzyme to be of a proper size and capable of exiting the microspheres upon secretion to be released in soil.

Two groups of enzymes were the final candidates: Dye-decolorizing Peroxidases (DyPs) and Laccases.

DyPs are multifunctional enzymes, with the ability to oxidize multiple phenolic and non-phenolic aromatics, such as syringyl and guaiacyl-type phenolics, lignin, lignin-phenols, veratryl alcohol etc. B. subtilis also produces such enzymes. However, DyPs function mainly in acidic pH, meaning that they would not be an appropriate choice for the alkaline soil environment. ^{7, 16, 17}

Laccases, on the other hand, are multicopper enzymes that catalyze, as well, oxidation reactions on various phenolic and non-phenolic aromatic substrates, and they are characterized by broad selectivity.^{8, 18} Their function is based on a combination of the oxidation reactions with the reduction of molecular oxygen to water. Four copper ions give laccases their catalytic role, by their distribution into the enzyme’s three binding sites.⁸ Bacterial laccases, as mentioned before, are more stable compared to fungal laccases and can work under higher temperature and various pH conditions.⁸

For all these reasons, we concluded that laccases would “get the role”, as they fulfilled all the proposed criteria. After further narrowing the frames, we decided to use the laccase 7 (Lac7), produced by the silA gene of Streptomyces ipomoeae CEST 3341.^{[19, 20]} Lac7 falls under the category of small laccases, meaning it can easily surpass the microsphere pores and reach the soil aromatic compounds, to take action.^[19] Moreover, it is thermostable, making it suitable for possibly extreme soil conditions, while it functions in alkaline pH conditions as well. Therefore, it matches with soil alkaline pH.^[19]

How will Laccase exit the host?

Bacterial laccases are intracellular enzymes⁸, so we needed to find a way to make them exit the bacterial cell, since B. subtilis is encapsulated inside the hydrogel, making it impossible to get in touch with soil surface. We fixed this problem by adding a LipA sequence before the laccase coding region. LipA is an esterase produced by B. subtilis and secreted after recognition from the Tat pathway.²¹ The Tat or twin-arginine translocation pathway regulates the export of folded proteins off the cytoplasmic membrane. It consists of numerous redox enzymes and cofactors.²² By adding the LipA sequence we expect the Tat pathway to promote protein excretion, thus enabling the laccase proteins to exit the bacteria.

The promoter

The laccase gene promoter used in Euphoresis is the PgroES promoter. PgroES is a strong, constitutive promoter of B. subtilis that is originally found before heat-shock proteins.²³

Module 2: Nitrogen fixation

The dramatically reduced soil organic and inorganic matter after severe wildfires affects the bioavailable nitrogenous compounds, as well.²⁴ Nitrogenous compounds are important for forest regeneration, since they are required for plant and microbial growth. Although it is implied by various studies that nitrogen (N) availability and concentration of NH4+ ­­and NO3- increase after wildfire, the bioavailable nitrogen forms become less and less over a short period of time, due to N immobilization in heterocyclic structures. Furthermore, fire’s intensity tends to promote the formation of inorganic N compounds, reducing, in this way, the soil organic N compounds.^[4] As for the soil microbiota capable of fixing or consuming nitrogenous compounds, metabolizing them into more bioavailable molecules, it seems to be seriously decreased over a year span after the wildfire and its recovery is very slow (years).^{24, 25} This justifies our choice to include in our biopolymer a bacterial strain able to fixate N and provide the soil with various bioavailable compounds, such as ammonia molecules.

Employ filamentous cyanobacteria to the game

Nitrogen fixation is a complex biochemical procedure that can not be easily engineered from scratch using synthetic biology. Therefore, we decided to enhance the function of an already existing nitrogen-fixing bacterium using synthetic biology tools. Symbiotic bacteria were opted out due to the absence of an extensive vegetation and root system.²⁶ Cyanobacteria, which are gram negative (-), photosynthetic bacteria, gained our interest, due to their ability to thrive using only atmospheric carbon dioxide, nitrogen, water and light, but mainly due to their ability to fixate atmospheric nitrogen.^{27, 28} The enzyme nitrogenase catalyzes the nitrogen fixation reactions. Since the enzyme is vulnerable to oxygen, nitrogen fixation can be only performed in anaerobic environments.²⁹ Cyanobacteria have developed various mechanisms to protect nitrogenase and perform atmospheric nitrogen fixation.³⁰.

Taking those mechanisms under consideration, we decided to utilize a filamentous cyanobacterial strain, Nostoc oryzae TAU-MAC 2710.³¹ This strain forms filaments composed of vegetative cells that perform photosynthesis, but also form special cells, called heterocytes. Heterocytes are differentiated cells in the filament that have created a protective polysaccharide layer and other mechanisms to exclude O2 for nitrogenase protection, in order to fix atmospheric nitrogen in a hypoxic environment, even if the filament lives under aerobic conditions.³²

HetR overexpression

Soil’s needs in nitrogenous compounds after a wildfire occurs are big, therefore our desire was to find a way to boost the nitrogen fixation reaction pathway and maximize the possible amounts of bioavailable N produced by our system. Upon research, we found that overexpression of the HetR would lead to a higher percentage of heterocytes per filament, meaning that a higher nitrogenase activity would be detected. HetR is a serine-type protease that regulates heterocyte differentiation, as the master molecule.^{33, 34, 35}

To achieve differentiation of more heterocyte per filament, we inserted the HetR gene, along with the strong, light-inducible promoter P_psbA1 .³⁵

Module 3: “The Bakill Switch”

Biosafety is an important factor for the proper function of Euphoresis, meaning that the release of a genetically modified microorganism, such as our B. subtilis, should not be a concept. That is why we wanted to create a kill switch mechanism for the B. subtilis to die before the hydrogel’s degradation. Unfortunately, literature research on this field didn’t show any results that would satisfy the needs of a highly biosafe product.

To develop our kill switch mechanism, we got inspiration from the “Deadman” circuit ³⁶ and Wageningen UR 2014 iGEM project.^[37] Combining their data and information we came up with the idea of a circuit that will be activated in the absence of IPTG.

The IPTG

IPTG, or Isopropyl β-D-1-thiogalactopyranoside is a widely used molecular biology reagent. It is an analog of allolactose, which naturally induces the activation of the lac operon. In E. coli, the lac operon consists of three genes, regulated by an operator region which, in the absence of lactose or the presence of glucose in the medium, is occupied by a lac repressor molecule, lacI, that prohibits the expression of the three genes. When present in the medium, allolactose, therefore IPTG as well, binds to the repressor and releases it from the operator region, enabling the three genes to be transcribed.^{38, 39}

The TetR

Apart from IPTG induction, we also utilize the function of Tet repressor proteins. TetR proteins are part of bacterial resistance mechanisms to tetracycline (Tc). In the absence of the antibiotic, TetR binds to the operator region of the tetA gene, which encodes a membrane protein that pumps out Tc before it reaches ribosomes. In a similar way to the IPTG induction, Tc binds to the TetR proteins and releases them from the tetA operator region, allowing the respective gene to be expressed.⁴⁰

How it functions?

Our BaKill Switch is a simple genetic circuit that consists of the following genes and promoters: A lacI gene with a constitutive promoter (BBa_J23106), a TetR gene (BBa_C0040) with the P_grac promoter (with lacO operator sites) and a bsrG type I toxin gene with a promoter that can be repressed by the TetR molecule - PlTetO1 (BBa_K3332034). ^{41, 42}

Alive state

This state is characterized by the presence of IPTG in the medium that surrounds the encapsulated Euphoresis microorganisms. IPTG binds to the LacI and prohibits it from suppressing TetR expression, whose gene expression is regulated by the P_grac promoter, which contains LacO operator sites. This way, TetR is freely expressed and binds to the operator sites of the PlTetO promoter, which regulates the production of the toxin gene, BsrG, suppressing gene expression. In this manner, with the presence of IPTG in the medium, which means that B. subtilis is still encapsulated in the microspheres inside the hydrogel, B. subtilis stays alive and continues with the laccase production.

Deceased state

The IPTG reserves in the medium will be depleted before the hydrogel’s and the microspheres’ degradation. In the absence of IPTG, lacI will be able to bind to the lacO operator of the P_grac promoter that regulates tetR production and suppresses the expression of the tetR protein. This will lead to the production of the BsrG toxin, that will lead the B. subtilis cells into cell lysis, as there will be no tetR protein to bind to the operator sites of the PlTetO promoter, located before the toxin gene.

Pgrac or PtrC-2?

Since our desire was to regulate the expression of the tetR by a promoter induced by IPTG, our initial choice was to use a PtrC-2-derived, IPTG-inducible promoter. The promoter sequence contains three operator sites that cause lactose molecules to bind tightly to them. The operator sites are 20-base long, inverted repeat sequences of the natural lactose operator’s first half. ^{36, 43} Despite the promoter’s efficiency, its functionality was questionable, because the promoter was derived from the E. coli. For this reason, we replaced the PtrC-2-derived promoter with the B. subtilis P_grac promoter in a second experimental cycle. The P_grac consists of a PgroE promoter sequence (native to B. subtilis) along with a lacI operator cite, meaning that is an IPTG-inducible promoter as well. After conducting our experiments, we realized that the P_grac led to a gene expression increase, so we decided to include it in the final Euphoresis design.⁴²

How does BaKill Switch promote biosafety?

Our kill switch system reassures the death of B. subtilis population before the biopolymer degrades, because life and death states are regulated by the amount of IPTG we add in the medium. This means that B. subtilis cells will never get in direct contact with the environment and will fulfill their mission behind the hydrogel’s and microspheres’ “bars”. Moreover, as it is already mentioned, IPTG, is an analog of allolactose, which is a lactose derivative, resulting as a reaction byproduct of the break of lactose to glucose and galactose in E. coli.⁴⁴ Lactose is a sugar found naturally only in mammal milk.⁴⁵ This means that B. subtilis would have little to no chance of surviving in forest soils, since neither lactose nor its analogs are expected to be detected in such environments.

Module 4: To kill a cyanobacterium

In the same way we didn’t want our genetically modified B. subtilis to be released into the ecosystem, our desire was to develop a mechanism to ensure the same thing for our cyanobacterial strain. Doing a literature search, we came across cyanobacterial toxin-antitoxin systems, but we also wanted to form a relationship between such system’s activation and the death of B. subtilis, to make sure that our Nostoc strain will die at the same time – before the biopolymer’s degradation – as well. In the same period, we were conducting research on quorum sensing systems that develop between bacterial strains to achieve a kind of communication, so our idea was to combine those two systems to create a reliable and effective mechanism for the on-time death of our cyanobacterial filaments.

The VapBC15 Toxin-Antitoxin system

The VapBC15 toxin-antitoxin system, which we utilize in our project, is found in the model cyanobacterium Synechocystis sp. PCC 6803. It is produced by the operon vapBC15 of its chromosomal DNA. When produced, the antitoxin VapB15 upregulates the operon’s function by directly binding to the operator, and at the same time destabilizes the toxin, VapC15, by directly binding to it, leading to the formation of a toxin-antitoxin complex (TA). When the cell is under stress conditions, the antitoxin levels drop due to the action of proteases and the ability of VapC15 to destabilize the bond between the VapB15 and the promoter region. This means that less TA complexes are formed and the VapC15 which belongs to the VapC-family toxins, acts as a ribonuclease and either prohibits cell growth, or even leads to cell death.⁴⁶

The Quorum Sensing system

A quorum sensing system (QS) is a signaling process between bacterial cells. QS systems function thanks to the production and secretion of small, diffusible signal molecules, which are referred as autoinducers. One molecule with such properties is the Acyl-Homoserine Lactone (AHL). AHL is produced by an enzyme, named AHL-synthase, that is in the bacterial strain that “transmits” the molecular signal, and is encoded by the LuxI gene. When AHL molecules are produced, they surpass the membranes and access the receiver cells. When inside the receivers, AHL molecules can bind to several transcriptional activators, such as the products from the LuxR, LasR and TraR genes and form complexes that regulate the production of some gene of interest. For instance, 3-oxo-hexanoyl-HSL, which is the AHL molecule produced by the LuxI AHL synthase, binds to a hydrophobic pocket in the N-terminus of the transcriptional factor encoded by the LuxR gene. In this way, the LuxR protein gets stabilized and becomes capable of binding to promoters and activating gene expression. It is understood that the production of the signaling AHL molecule heavily affects the production of the respective gene of interest. ⁴⁷

Mixing them up

Our mechanism for the death of Nostoc oryzae TAU-MAC 2710 is based upon both the VapBC15 Toxin-Antitoxin system and the AHL-regulated QS system. We transformed our B. subtilis WB800 strain with the LuxI gene along with the strong, IPTG inducible P_grac promoter. The P_grac was chosen instead from the initial P_groe promoter for biosafety reasons - an IPTG induced promoter would regulate the stop of luxI transcription, when IPTG is no longer available in the medium, even if the BaKill Switch would not function. The Nostoc strain has also been transformed with the insertion of the LuxR gene, which is regulated by the PtrC, IPTG inducible promoter.⁴⁷ We chose the LuxR gene because, according to Kokarakis et al. (2023), it was more effective and non-toxic to the cyanobacterial strains, which means that we would be able to observe gene expression. We also chose the IPTG inducible promoter in order to regulate LuxR expression with different concentration of IPTG and achieve a desirable LuxR production for the function of the toxin-antitoxin system, which will be described below.

In the cyanobacterial strain we also included the toxin VapC15 gene, with the P_psbA1, strong light inducible promoter, as well as the antitoxin VapB15 gene, regulated by the P_QS promoter. The P_QS promoter induces gene expression upon binding to the 3OC6-HSL (LuxR-AHL) transcriptional factor complex.⁴⁷ Upon the insertion of those genes, two states can develop for the cyanobacterium:

Alive state

When B. subtilis is still alive, it produces and secretes AHL molecules, which enter the Nostoc cells and bind to the steadily produced LuxR gene products. The formed complex acts as a transcriptional factor and binds to the VapB15’s P_QS promoter, inducing the production of the antitoxin molecule, VapB15. The VapB15 forms TA complexes with the VapC15 toxin, which is steadily produced under the P_psbA1 cyanobacterial promoter. In this way, the toxin molecule cannot function and the Nostoc cells stay alive, continuing their nitrogen fixation activities.

Deceased state

Eventually, IPTG will no longer be available in the medium, a situation that will lead to the death of B. subtilis cells. That means that AHL molecules will no longer be produced, resulting in the LuxR product being unable to induce the transcription of the antitoxin molecule. The VapC15 toxin, on the other side, is being steadily produced due to the function of the P_psbA1 promoter, which will cause a rise of toxin levels, and the eventual death of the Nostoc cells.

The backbone

Our main criteria for choosing the appropriate plasmid backbone for the Euphoresis microorganisms’ transformations were the following:

1. The plasmid backbone should contain an antibiotic resistance gene for an antibiotic that affects both bacterial strains. In this way, we ensure that the bacterial colonies that did not receive a plasmid would not survive in the solid bacterial cultures plates, since they would not express the respective antibiotic resistance. Kanamycin is an antibiotic both strains are vulnerable to.^{[48, 49]}]
2. The plasmid backbone should be compatible with the BioBrick assembly, meaning that it should contain the proper prefix and suffix regions.
3. The plasmid backbone should include a fluorescent protein sequence that will be excised from the plasmid during the assembly between the prefix and suffix regions. In this way, the bacterial colonies that have received recombinant plasmids, thus they do not include the FP sequence, will not be fluorescent.
4. A satisfactory plasmid copy number was desirable (medium copy number).
5. The plasmid needed to be of a proper size, so that it is amenable to the genes’ assembly. The plasmid size is also important for the bacterial transformation processes, since a large-size plasmid might not easily enter the B. subtilis and the N. oryzae cells.

Taking those parameters into account, as well as conducting NanoDrops to determine our shortlisted plasmids’ purity (see Results), we finally chose the iGEM Kit’s pJUMP29-1D'(sfGFP), as it met with all of the above criteria, both for the project’s final design, and the experimental design, too.^[50]

Experimental design

Breaking Euphoresis into pieces…

In the final Euphoresis product, two plasmids were constructed, in order to be inserted in the B. subtilis and in the Nostoc oryzae, respectively. However, in order to further investigate and comprehend the function of the individual parts and devices. In the following image the 9 experimental plasmids are shown.