Direct manipulation: engineering of a cyanobacterium
Nutrient-rich freshwater lakes and ecosystems around the world are swarming with a vast variety of cyanobacteria, also known as blue-green algae. These microorganisms are characterized by the capacity to capture CO2 from their surroundings and convert it to useful molecules, a solar-powered process known as photosynthesis. This year, the KU Leuven iGEM team decided to harness this useful property of cyanobacteria for the development of sustainable biomanufacturing platforms, suitable for implementations in bioreactors.
In order to do this, a suitable cyanobacterial strain had to be found. The unicellular cyanobacteria Synechococcus elongatus UTEX 2973 has been reported to grow at a remarkably high rate, up to rates comparable to the yeast strain S. cerevisiae, a commonly used biomanufacturing chassis in industry. This strain caught our attention as a possible candidate for sustainable manufacturing purposes, as previous reports mentioned the high potential for targeted genetic modifications and conjugal DNA transfer. In this project, sfGFP as YFP were chosen as proof-of-principle proteins.
1. Engineering the desired plasmid via overhang PCR and Gibson assembly
For decades, plasmids have been a useful tool for the transfer of DNA into various types of microorganisms. A plasmid called pAM2991 has been described in literature as a cyanobacterial cloning vector for Synechococcus elongatus PCC 7942, which is closely related to our strain UTEX 2973. Therefore, it was chosen to host our proteins of interest - sfGFP and YFP - in subqesuent experiments. pAM2991 was first enzymatically digested with restriction enzymes EcoRI and BamHI. In an attempt to drive the secretion of these proteins by the bacteria, signal peptides were fused to the sfGFP and YFP genes. For the generation of the complete construct, YFP, sfGFP and three different types of signal peptides were separately amplified using tail-PCR with overhangs compatible for Gibson assembly and then joined with the plasmid by Gibson assembly. Successful amplification and ligation were verified at each step using gel electrophoresis.
2. Transformation via conjugation with E. coli
While some cyanobacterial strains actively take up DNA from their surroundings, the Synechococcus elongatus UTEX 2973 strain is not naturally competent and requires a conjugative DNA transfer. Triparental mating was performed by a Top10 strain containing mobilizer and helper vectors pRK24 and pRL528 respectively.
Figure 1: Triparental conjugation. Firstly, the constructed plasmid pAM2991, here denoted as the conjugative plasmid, has to be inserted into TOP10 E. coli using a heat shock protocol (not shown). Afterwards, the tra and mob genes lying on the helper plasmids allow for the formation of a conjugation pilus, forming a bridge between the two E. coli cells. This results in the conjugation of the constructed plasmid pAM2991 to the conjugative E. coli, now containing three plasmids. A second step in triparental conjugation then consists of transporting this plasmid to S. elongatus UTEX 2973 by again forming a conjugation pilus and transforming pAM2991 to the cyanobacterial strain. Only the cyanobacteria receiving this plasmid will then grow on selective medium.
Indirect manipulation: engineering cyanophage S-TIP37
In order to create a cyanophage that could produce the protein of interest upon infection, the strategy of choice was in vitro phage genome engineering. Through literature research, cyanophage S-TIP37 was identified as a suitable phage for our project. The selected phage had a genomic organization similar to that of coliphage T7, which is well studied and - due to its straight forward transcriptional scheme - attractive for genetic manipulation. Phage S-TIP37's natural host is Synechococcus sp. WH8109, a strain of marine cyanobacteria.
The experimental goal for this system was the expression of fluorescent protein YFP directly from the phage’s genome. In order to achieve this, a genomic insert was designed. A strong native phage promoter was put in front of the YFP gene, maximally driving transcription. Furthermore, in order to also maximize transcriptional efficiency, the YPF gene was codon optimized using the codon distribution of the phage’s capsid gene, one of the strongest expressed proteins of the phage’s genome. The complete insert cassette was synthesized as a whole with overhangs compatible for Gibson assembly.
Four insert cassettes were designed, all encoding the same regulatory elements and gene yet containing overhangs for integration into different locations of the phage's genome. This opens some possibilities for the optimization of protein expression, as the location on the phage genome might influence transcriptional efficiencies. The four engineering strategies are illustrated in Figure 2. Two approaches involve double Cas9 cutsites, enabling the deletion of either the lysogeny gene exclusively (A), or the entire putative lysogeny cassette (B). Deleting this gene or gene cassette may result in higher infection rates, as it presumably removes the lysogenic capacities of this phage. Subsequently more lytic cycles will occur. Alternatively, single cutsites were put after the last gene of the structural cassette (C) and the last gene on the phage genome (D).
Figure 2. Overview of different modification strategies on the S-TIP37 phage genome. A) Removal of the integrase and B) the entire putative lysogeny cassette by double cut sites to remove the option of lysogeny during phage infection. C) Single cut site for integration at the end of the structural cassette. D) Single cut site for integration at the end of the phage's genome.
1. In vitro phage genome engineering with Cas9 and Gibson assembly
Cas9 is an endonuclease enzyme associated with CRISPR and guided by RNA. It is commonly known in genome engineering as “molecular scissors” as it creates double stranded breaks in DNA. Single guide RNA can be specifically designed to bind to the sequence of interest in the DNA and facilitate the digestion by Cas9. In our project, a detailed bioinformatic analysis of the phage genome was performed and several digestion sites for the Cas9 enzyme were evaluated.
Figure 3: In vitro phage genome engineering with Cas9 and Gibson assembly. To modify the phage genome, the first step performed is purifying it using a phenol/chloroform extraction protocol. This results in the deletion of all bacterial DNA, as well as the capsid proteins from the phage particle. The naked DNA allows the Cas9 protein to reach it. It selects it target site for cutting using its single guide RNA, which is complementary to the DNA sequence. The cut will then be restored with designed inserts that have the overlaps of the region of insertion.
Following Cas9 specific cutting, Gibson assembly was used to join the fragments of the phage genome together with the insert cassette. Successful incorporation of the sequence into the phages genome was tested by PCR with specifically designed primers and gel electrophoresis.
2. Transformation via electroporation into cyanobacteria
Upon successful phage genome engineering, the linear DNA was to be transferred into cyanobacteria at their mid-exponential phase (OD750 = 0.5) electroporation. This procedure would then reboot the phages' lytic cycle, resulting in the synthesis of a few fist bacteriophages that can then infect the following bacteria. This way, an exponential amplification of the amount of phages will be formed after subsequent infections.
Figure 4: Jumpstarting a phage. To form new phage particles, the DNA has to be electroporated into the cyanobacterial host strain. By giving short bursts of high voltage, small holes will be formed in the cell wall of the bacterium, allowing for the passing of the phage DNA. Given the general structure of a phage genome, this will result in the new formation of phage particles.