Bacteriophages – TUDA iGEM 2021

Bacteriophages

Overview

Our goal was to enable a controlled phage induction through a quorum sensing signal and the release of a phage from a new host organism. The whole concept is summarized in Figure 1.

Figure 1. A schematic overview for the concept of the controlled phage induction. Our sleeper cell contains an integrated phage genome, a plasmid comprised of holin genes and the genetic circuit for controlled phage induction. Through the genetic circuit a repressor is produced, which keeps the phage in the lysogenic state and therefore our sleeper cell inactivated. After sensing the pathogen’s acyl homoserine lactone (AHL) molecules, a repressor-cleaving protease (or antirepressor) is produced. This protease cleaves the repressor leading to the production of phage and holin particles. The assembly of these particles leads to the release of functional phages out of our sleeper cell. Now, these phages can infect the sensed pathogen and therefore lyse it.

Background

Why we chose to work with phages

Bacteriophages are viruses that can only infect prokaryotes, which makes them harmless to humans.¹ They have co-evolved with their host bacteria for centuries and are therefore perfectly adapted to them. This is also the reason why they specifically attack only their target bacteria.² Due to fitness costs and/or trade-offs between phage resistance and bacterial growth, evolutionary theories suggest that the likelihood of a bacteria being resistant to both antibiotics and phages is significantly reduced.³This makes phages a promising tool in the fight against multi-drug-resistant germs.⁴ For these reasons, we think phages are a good way to protect a biofilm from infiltration by pathogenic bacteria.

Phages recognize their host by docking to its surface with their long tail fibers (Figure 2). This is followed by the secondary attachment through the short tail fibers, which then leads to the injection of the phage DNA (RNA) via contraction of the sheath.⁵ In the cell, control is then taken over by the phage, and the entire cellular machinery is switched to phage production. Phage production stops only when the cell is lysed by phage proteins and thus bursts open, releasing the newly produced phages into the environment and the lytic cycle starts all over again. In addition to the just described lytic cycle, phages can also induce a lysogenic cycle (Figure 3).⁶ In the lysogenic cycle, the phage DNA is incorporated into the bacterial genome and replicates with the host until it is triggered to switch into the lytic cycle. For our project, it is very important to implement a phage in our cell and only initiate its production when a pathogen is sensed. In consequence, we decided to use a lysogenic phage and exploit its lysogeny-lysis switch. In the interest to work with model organisms, we chose to use the well-studied Escherichia coli lambda phage, which is a lysogenic phage.⁶ In the next paragraphs we will first describe the native lambda switch and then the genetic circuit we designed to control it.

Figure 2. Schematic overview of a tailed bacteriophage and its morphological elements. Tailed bacteriophages are composed of a polyhedral head capsid and a baseplate with tail fibers. Both parts are connected by a contractile sheath. After the phage docks on the bacterial cell surface via its tail fibers, the viral DNA is transferred from its capsid head through the contractile sheath into the bacterial cell. Modified after Peltomaa et al. 2016.⁷

Figure 3. A schematic overview showing the lytic and lysogenic cycle of bacteriophages. Both cycles start with the phage attachment and the following infection of a host bacterium (white arrow). Upon the infection, the decision between the lysogenic (purple arrows) and lytic (pink arrows) cycle is made. For the lysogenic cycle, the phage genome integrates into the host genome and then replicates as a prophage with the host genome. Through a triggering signal, the lytic cycle is activated. In the lytic cycle, the host DNA is degraded and the genetic information of the phage is replicated. This leads to the production of phage particles, which assemble into a functional phage. Afterward, the cell is lysed by phage enzymes. The lysis results in the release of the phages and the cycle can start again. Modified after Markl et al. 2019.⁸

The Native Lambda Switch

Figure 4. The shift between lysogenic and lytic state is toggled by the operator region of the lambda phages genetic switch. a. The operator consists of three parts (O_R1, O_R2, O_R3) overlapping with the promoters P_RM and P_R. P_RM enables the expression of cI (lysogenic state) and P_Renables the expression of cro (lytic state). b. cI dimers are binding to O_R1 and O_R2, repressing the Cro production and enabling an optimal cI production by enhancing the binding affinity of the RNA-polymerase to P_RM. Thus, the lambda phage stays in the lysogenic cycle. c. Cro dimers bind to O_R3 and the production of cI is repressed, initiating the lytic cycle. Modified after Ptashne 2004.⁹

After infection of Escherichia coli, the lambda phage either enters the lytic or the lysogenic cycle. Entering the lysogenic cycle, only two genes are active. The one relevant to us, called cI or lambda repressor, is expressed by the RNA polymerase binding to the P_RM-promoter (Figure 4a.). To the right of cI the P_R-promoter and cro are located. Cro is expressed in the opposite direction.⁹ Both promoters do not overlap but are connected through the tripartite right operator O_R, consisting of O_R1, O_R2 and O_R3, where O_R1 and O_R3 are located within these promoters (Figure 4a.). cI forms dimers that bind to O_R. The first dimer binds to O_R1 repressing the Cro production and enabling the cI production. The second one binds to O_R2, enhancing the binding affinity of the RNA polymerase to P_RM (Figure 4b.).⁹ When the cI concentration reaches a threshold at which O_R3 is bound, its production is repressed in a self-regulating manner. Further, P_R is blocked and cro cannot be transcribed. This state is almost indefinitely stable.⁹

The lytic cycle can be activated by causing DNA damage, evoking the cell’s SOS response. This happens through ultraviolet light or carcinogens, for example modified benzopyrene.⁹ Thus RecA, which normally catalyzes recombination between DNA-molecules, gets activated as a specific protease that cleaves cI monomers. The cleaved cI cannot bind to the operator and cI is no longer produced.⁹ To switch into the lytic cycle, the production of Cro is necessary. Cro acts as an antagonist of cI and conducts only negative regulation. It binds the same operator sites as cI but with reverse affinity, repressing the synthesis of cI. Without cI binding to the operator, the RNA polymerase can bind to the stronger promoter P_R and Cro is produced (Figure 4c.). Thus, the lytic state is initiated.⁹

Phage Induction

Our genetic circuit for controlled phage induction

For our project, it is necessary that a functional phage is specifically expressed. Therefore, we engineered a switch that allows us to control the induction of the expression of our chosen lambda phage. Initially, we wanted to design a switch based on the production of the lambda repressor cI, leaving lambda in the lysogenic cycle.⁶ The next step would be the repression of cI, whereby it would no longer be produced and would therefore decay over time. The expert Prof. Dr. Grzegorz Węgrzyn drew our attention to the fact that such a switch would be too slow, as cl is a very stable protein, and its decay would take too long. Therefore, we revisited the natural switch of the lambda phage, which is triggered by the protein RecA. Normally, DNA damage is required for RecA to undergo a structural change enabling it to cleave cI, and thereby inactivating the repressor. Prof. Dr. Węgrzyn mentioned a constitutively active mutant of RecA, RecA730. This protein has a point mutation (E38K)¹⁰, which causes RecA730 to be in its active form as soon as it is expressed, thus cleaving cI independently of DNA damage. Targeted degradation of cI monomers by RecA730 should lead to a significant acceleration of our switch.

Our switch is designed to express cI located on a plasmid and herewith maintain the lambda phage in a lysogenic state (Figure 5b.). The second part of our genetic circuit is activated by recognition of quorum sensing molecules (QS molecules) of pathogens (Figure 5c.). This is accomplished by QscR (allosteric transcription factor) binding the QS molecules and the resulting complex activates the P_PA1897 promoter. Through activation of this promoter the expression of recA730 and the repressor tetR, which both act as countermeasures to cI, is induced. The produced TetR binds to the P_tetO promoter and herewith prevents production of cI. The produced RecA730 degrades the remaining cI, whereby the promoter of the lytic cycle in the genome of the lambda phage is no longer repressed and cro is expressed, thus leading to the activation of the lytic cycle. With the activation of the lytic cycle the expression of bacteriophages is induced. For our proof of concept, the QS-molecule induction was replaced by an IPTG induction and eGFP was used as a reporter protein for the induction of phage expression (Figure 5). For this, we placed operator sites O_R1-3 of the lambda phage also containing promoter sequences upstream of the gene of eGFP.

Figure 5. Schematic overview illustrating the design of our genetic circuit for phage induction. a. General order of the genetic elements in our construct and the native lambda switch. b. Our genetic circuit before induction through the QS-signal (inducer). The phage is kept in the lysogenic state by the expression of cI (lambda repressor), repressing the expression of cro. c. The genetic circuit is activated by the QscR-inducer-complex because the P_PA1897 promoter contains a QscR binding site. Activation of the P_PA1897 promoter on the one hand leads to the expression of recA730, cleaving cI. On the other hand, the expression of tetR is induced, which represses the expression of cI. This leads to the lytic cycle through the expression of cro.

Lab

To test our genetic switch, we first assembled the individual DNA parts together. We chose to use “Golden Gate Assembly”. This assembly method is based on type IIS restriction enzymes, that cut outside their recognition sequence. In this case, BsaI is used whereby a single-stranded overhang of 4 base pairs is created. We had to design these individual assembly sites in each of the four fragments and the backbone pDGB3_alpha_1 to perform directional ligation. The major advantage of Golden Gate Assembly is that the restriction and ligation of multiple fragments can be carried out in the same tube since the recognition sequences of the restriction enzyme are no longer present in case of a successful assembly (Figure 6).

Figure 6. Methods to demonstrate proof of concept of our genetic circuit for controlled phage induction. The assembly was carried out according to the Golden Gate protocol using multiple gene fragments and a vector. A restriction digest was performed to verify the assembly of our genetic circuit. E. coli BL21(DE3) were then transformed with the assembled plasmid. For verification of our concept a plate reader assay was performed to measure the presence of a fluorescent reporter gene (egfp) upon or without induction with IPTG.

The resulting antibiotic-resistant colonies (P1-P14) were used for inoculation of fresh medium and the plasmids were then isolated and analyzed by restriction digest to verify the successful transformation. The vector contains two EcoRI recognition sites up- and downstream of the insert. The incubated restriction digest was then analyzed using gel electrophoresis (Figure 7).

As a result, the assembled insert (ca. 3.6 kbp) should be separated from the vector (ca. 6.5 kbp) and two distinct fragments of the respective size should be visible. This is exactly what we observed with the samples from colonies P12 and P13, which were used for further testing.

These samples were sequenced to investigate if problems could occur further down the line with substitutions or frame-shift mutations in the sequence. The sequencing results showed two mutations in the 3742 bp sequence (99.95 % percentage identity). One is placed in the terminator sequence after tetR. This should not impose any problems since it only brings in an additional CG-base pair in a non-coding region without a recognition site. The second mutation on the other hand is located in the coding region of the lambda repressor cI. The amino acid Gly-54 is replaced by Asp-54. Since the new amino acid features a negatively charged side chain, this could result in a different folding behavior of one of the key enzymes in our genetic switch. However, we assume that the mutation has little or no effect on the binding affinity of cI to the P_R promoter, since the following assay showed the expected results.

Figure 7. Agarose-gel electrophoresis of the restriction digest to verify the successful insertion of the assembled phage induction module into the pDGB3_alpha_1 vector. The restriction digest was performed using EcoRI-HF (New England Biolabs). The separation was carried out at 120 V for 90 min on a 1 % agarose gel. As reference a 1 kb Plus DNA ladder (New England Biolabs) was used.

Figure 8. Ratio of the emitted fluorescence per optical density [FI/OD600] of our sleeper cell (E. coli) before and after the induction with IPTG. The measurements were carried out at 30 min intervals. Between these measuring points the samples were shaken at 225 rpm and 37°C. The induction with 0.5 mM IPTG took place after 195 min. Two biological replicates and five technical replicates per sample per measuring point have been measured.

The phage induction module was further characterized by investigating the fluorescence output of the respective culture before and after the induction with IPTG in place of the quorum sensing molecule AHL. For this we monitored the OD600 and the fluorescence of eGFP at 485 nm and the emission at 528 nm (Figure 8).

As expected, the ratio of fluorescence per optical density does increase rapidly after the induction (195 min) with IPTG. As a result, a nearly 12-fold increase in this regard was observed, strongly indicating that our composite part does work as intended. Furthermore, sample 12 without induction showed a stable baseline over a timeframe of more than 20 h. The behavior of sample 13 seems to be the result of a low optical density, since the raw fluorescent output is not much higher than the blank background. This suggest that our system is working with little to no significant basal expression. When implemented in the lysogen MG1655, this should result in a controlled phage induction. Nevertheless, it is necessary to mention that the informative value of our analysis is limited by the number of samples. While we did test biological duplicates, our number of technical replicates amounts to five per sample per measuring point. This has been conducted two times.

We tried to decelerate the increase in fluorescence to further strengthen our assumption, that these observed effects are achieved through our construct. Therefore, we inhibited the system through the addition of anhydrotetracycline (AHT), a molecule which shows a high affinity for TetR. The resulting AHT-TetR complex is not able to repress the expression of cI (Figure 9).

There is a visible difference between the P12+IPTG sample and the P12+IPTG+AHT t1 sample, which was inhibited at the same point as the induction took place (146 min). This is true for both biological replicates. In both cases the discrepancies are greater than their respective error margins. On the other hand, when inhibited later in the measurement (214 min), the effect is less striking. While there is a difference in the P12 samples, it is not outside the error margin. With P13, the measurements are nearly identical. The fitted curve indicates, that the AHT addition after 214 min leads to an increased expression of egfp. However, this is probably the consequence of the two measurement points after 367 min and 387 min, which can be explained by measurement errors. The samples (P12-P13) inhibited at t1 achieved only 53 % of the uninhibited fluorescence output after about 50 min upon induction. In contrast, the P12-P13 mean maximum fluorescence when inhibited at t1 is about 80 % of the regular induced sample. Overall, the comparison of the induced sample and the direct inhibition indicates that TetR was bound by AHT, which resulted in the renewed expression of cI. This shifts the cI-RecA730 equilibrium and results in a decreased expression of egfp. It is worth noticing, that even when inhibited at the same time as the induction occurs, the fluorescence output is still increasing at a greater speed than the optical density. This implies, that our switch and the respective assembly could be further simplified by erasing the TetR component, if one is willing to trade about 30 min of reaction time in the system. This could serve as a starting point for further engineering using an iterative approach to optimise our genetic switch.

Figure 9. Ratio of the emitted fluorescence per optical density [FI/OD₆₀₀] of our sleeper cell (E. coli) before and after the inhibition with AHT. The measurements were carried out at 20 min intervals. Between these measuring points the samples were shaken at 225 rpm and 37°C. The induction with 0.5 mM IPTG took place after 146 min. The inhibitions took place after 146 min (t1) and 214 min (t2). Two biological replicates and five technical replicates per sample per measuring point have been measured. a. Inhibition assay of P12. b. Inhibition assay of P13.

In conclusion, we were able to gather data which matched our expectations in nearly every sample including the controls. The only measurement which was unexpected was the little difference between the second point of inhibition compared to the induced sample without AHT. This makes us extremely optimistic about the further characterizations and applications of our composite part at a later point in time. A logical next step would be the induction of our circuit in a transformed lysogen.

Challenges

During our work on the project, we faced several challenges that had to be overcome in order to achieve the goal we set. The integration of the functioning lambda genome into Bacillus subtilis provides one of the biggest challenges. Its assembly and expression seem difficult since phages are dependent on specific host proteins and factors making the transfer of a phage into a different host a big challenge in general. B. subtilis is a Gram-positive organism and thus differs in the membrane composition from the Gram-negative E. coli¹¹, which may have major implications on our engineering approach, e.g., on the phage release.

First, we wanted to integrate the phage genome into B. subtilis, this specifically means integration of a huge DNA fragment (approximately 50 kbp¹²). For this purpose, a specific integration site was necessary so that the phage genome can get replicated within the host genome.¹³

In order to stay close to the naturally used integration sites of B. subtilis phages, we decided to create a new integration site for our phage in this area following the advice of Dr. Anna Dragoš. Another benefit of this approach is the independence of the amyE locus which would remain free for further modification.

Secondly, we had to think about possible solutions to the problems that may arise regarding the phage expression in the new host organism. For example, the strength of the native phage promoters may be insufficient in B. subtilis and thus limit the rate of phage production. In our opinion, the most significant challenge is the adaptation of the ribosomal binding sites (RBS) from lambda phage to B. subtilis. Without a well functional RBS, ribosomes cannot initiate translation.¹⁴ Our proposed solution for this issue is shown in our outlook.

Furthermore, the decision between lytic and lysogenic behavior was crucial to control the release of phages. We wanted to ensure the modularity of our system for it to be used in a broad spectrum of possible application. Therefore, we made a repressor-antirepressor like system, that could be applied to every phage with such a system and even further as shown through our switch, which is not a repressor-antirepressor system in the typical manner. Even though we tried to keep our system modular, we cannot evidently calculate the functionality of the phage expression in a new host organism. The expression of a phage (lambda) in a new host organism (Salmonella typhimurium) has been done before, but there was some modification necessary.¹⁵ Therefore, the limited research on this field suggests, that changing the host organism of a phage is possible though it might come with unexpected problems.

Another challenge, which Dr. Anna Dragoš pointed out to us, is the possibility that phages in the lysogenic state might produce substances that are toxic for B. subtilis. That may lead to premature death of the bacterial cell without the release of functional phages. In the specific case of the lambda phage Prof. Dr. Grzegorz Węgrzyn told us that this problem should not occur because there are only two genes active in the lysogenic state and none of them are toxic.

Outlook

To overcome some of the problems and shortcomings mentioned above, we thought about the next logical steps. Certainly, one of these shortcomings is the lack of tests in Bacillus subtilis since all experiments have been established using E. coli. This helped to achieve a realistic proof of concept for the limited time we had as a result of the prevailing pandemic but came with some major shortcomings regarding our proposed implementation. One way to overcome this hurdle is to establish our phage-induction system in B. subtilis using our proposed “lambda integration site” mechanism, that will be shown in the next paragraph. Proving the assembly of functional lambda phages in a phylogenetic distant organism would be a very important next step. To achieve this, it is probably necessary to implement B. subtilis compatible regulatory regions for control of gene expression e.g., ribosome binding sites into the lambda phage genome. One tool to enable the insertion of dozens of short sequences in one step is “Multiplex Automated Genome Engineering” (MAGE).¹⁶ Using this technology and degenerate RBS oligos, the authors were able to optimize 24 genes simultaneously and to increase the production of their pathway product up to four times. This will allow for a high throughput testing to finetune the necessary delicate balance of gene expression for phage assembly. This approach could provide a first solution for the implementation of a non-native bacteriophage into a synthetically redesigned host organism such as our B. subtilis sleeper cell.

Phage Integration Site

In order to integrate the large (ca. 50 kbp) lambda genome into B. subtilis without blocking one of the standard plasmid integration sites (e.g., the amyE locus) we decided to create a new integration site in B. subtilis. This idea came up in the interview with Dr. Dragoš and with whom we remained in contact to further develop this idea. This is just one of the examples how experts shaped the development of our project. For the purpose of creating a new integration site, we wanted to clone the attB site of lambda from E. coli and then introduce it instead of the spsM gene in B. subtilis. The attB site is the genetic sequence, that is recognized by the phage and used as integration site.¹⁷The spsM gene is naturally used as an integration site by the spß phage (B. subtilis phage), containing sporulation-promoting factors.¹⁸ By integrating the attB site into B. subtilis, the lambda’s own integrase should be able to perform the integration.¹⁷ Another factor that speaks for the success of this method is the fact that the lambda integration site works in vitro.¹⁹Although we could not test this approach in the lab it seems like a promising way to integrate the lambda phage into the genome of B. subtilis. Testing this theory in the laboratory should easily be possible by introducing the attB site through homologous recombination into B. subtilis and then transforming the modified B. subtilis with the lambda genome. This approach could be used as a modular platform for integration of large gene fragments by adding the attP site to them.

Phage Release

Releasing the phages out of the cell is not a trivial process, because there are various cell barriers that have to be overcome. As the release is highly crucial for the success of our work, we tried to find a solution by researching how this process naturally occurs. Therefore, we decided to focus on holins, well-known proteins that contribute to the destruction of the bacterial cell envelope (BCE).²⁰

The production of the holins takes place in the middle and last steps of the phage infection.²⁰ The detailed steps of holin assembly are shown in (Figure 10). The holins cause lesions to the cell membrane by getting aggregated and integrated in it. This gives other proteins like lysins the opportunity to attack the murein and then destroy it.²¹ This way holins trigger the host cell lysis by activating the endolysins and collapsing the proton motive force.²⁰

We focused our research on finding a proper holin that is suitable not only for E. coli but also for B. subtilis. We excluded native lambda phage holins, since holins of Gram-negativ bacteria are unable to penetrate the thick peptidoglycan layer of Gram-positive ones.²⁰ Therefore we decided to go for the holin of the B. subtilis phage Phi29 because it has a great similarity to the holins of lambda.

Figure 10. Schematic overview of the holin hole formation. (1) Inactive holin monomers are produced in the cytoplasm and consist of three transmembrane domains. Two of them have hydrophilic parts (orange striped) and hydrophobic parts (orange) and one connects them (pink). (2) After reaching the critical concentration, the holins are transported close to the cell membrane where they dimerize. (3) Those dimers polymerize further to the inactive tetramer. (4) Those tetramers polymerize further to the inactive oligomer. (5) Following accumulation and oligomerization of the molecules leads to the formation of pre holin ‘death rafts’ in the membrane. (6) The local depolarization of the cytoplasmic membrane occurs, which triggers conformational changes in the holins and results in further expansion of the lesion and finally the lysis of the cell. Modified after Harper et al. 2021.²²

The main factor, when choosing a suitable holin, was to find the protein encoding gene, which would have a high similarity with the lambda S-genes. That is where the evolutionary conserved initiation region for the holin expression comes into action. In order not to violate the system’s integrity of the lambda phage in B. subtilis we needed to find a holin with the same dual start translational motif. This implies, that the codon sequence Met-Lys-(X)-Met is located at the beginning of both necessary genes, that serve as translational starts for polypeptides with the opposite functions. The shorter polypeptide is the active holin, or lysis-effector, whereas the longer one acts as an inhibitor of holin function. All features of the dual start motif were identified in holin gene 14 of B. subtilis phage Phi29.^23,24 This would allow a simultaneous expression of lambda’s holins and the holins of Phi29, making the release of lambda phage from B. subtilis probable (Figure 11).

Figure 11. Sleeper cell controlled phage release using holins. The genetic information of the holin from phage Phi29, namely gene 14, and the lambda phage are implemented in B. subtilis. As a response to an induction signal, the sleeper cell starts the expression of phage (yellow) and holin (orange pink circle) genes. This results in the release of assembled lambda phages from B. subtilis, through the expressed B. subtilis holins.

We found out, that each gene encoding the holins of the lambda phage and phage Phi29 is preceded by two consecutive ribosome-binding sequences and has two potential start codons. Moreover, the gene arrangements of the corresponding holins and the murein hydrolase functions in both phages seem to be identical.²³

Furthermore, Dr. Wadim Weber discussed with us possible modifications of holins that could help us optimize a phage release. Thus, a possible approach could be implemented via promoters of different strength. In case of further development of this project, holins will provide an opportunity for a precisely controlled release of phages and subsequent lysis of the host cell.

The next logical step after the assembly and release of functional lambda phages in a non-natural host organism would be to achieve a highly modular system, expressing specific bacteriophages as a response to a variety of biofilm invading species. While phages themselves are very specific, their genetic elements evolved modular.²⁴ Interestingly, this also seems to be true for the system controlling the native transition of lambda phages into the lytic cycle. Since our synthetic switch is based on the lambda phage regulatory unit, our design should be compatible with a variety of bacteriophages with a minimal number of adaptations. This includes among others the exchange of the cI-RecA730 repressor-antirepressor like system with the native repressor-antirepressor system of the respective phage. This makes the modular approach a matter of genetic storage capacity for the bacteriophage genome.

References

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Team:TU Darmstadt/phages