Team:BrockU/Description
Project Abstract
Flip recombinase is a versatile and important recombinase enzyme with broad applications in molecular genetics. Flip recombinase has been used to induce genetic mutations in vivo in numerous model organisms including bacteria, Drosophila, Zebrafish, and mouse and human cells. However, Flip recombinase activity is binary and thus cannot be precisely activated in time and space. Utilizing light-sensitive protein interaction domains termed “magnets”, we have developed a light-sensitive optogenetic variant of Flip recombinase that can be controlled in Escherichia coli with exquisite spatiotemporal precision. We believe this Opto-Flip recombinase has the potential to be utilized in multiple model organisms and will provide a novel tool allowing for precise molecular-genetic control for numerous future research and industrial applications.
In silico Design of Opto-FLP
In order to avoid potential disruptions to flippase function, we have split flippase at a highly disordered region, which we hypothesize will increase the probability of optogenetic reconstitution of a functional flippase protein (Figure 1). These two portions termed FLPA and FLPB were conjugated to photosensitive proteins known as pMag and nMag, respectively. When the system is exposed to blue light (488 nm), pMag and nMag, which have been shown previously to specifically heterodimerize, will physically bring FLPA and FLPB together, resulting in reconstitution of an intact flippase protein, thereby stimulating flippase activity (Figure 2). Our hypothesis is that shining blue light on the system will render a functional flippase, allowing us to control its activity precisely in time and space. Each fragment (FLPA-pMag and nMag-FLPB) will be inserted into a separate multiple cloning site in the pCDF-Duet-1 vector, with each portion being transcribed by a unique T7 promoter sequence. This will allow for ratiometrically equivalent expression of both halves of Opto-FLP.
We have chosen to use the pMag and nMag photosensitive tags as they heterodimerize and show complete specificity for each other. Other optogenetic systems have demonstrated issues with clustering or homodimerization. However, the magnet system contains a charged pair of optogenetic tags, one being positive, the other negative. Tagging each half of Opto-FLP with different optogenetic magnets ensures that one FLPA protein will associate with one FLPB protein.
We have chosen to use the pMag and nMag photosensitive tags as they heterodimerize and show complete specificity for each other. Other optogenetic systems have demonstrated issues with clustering or homodimerization. However, the magnet system contains a charged pair of optogenetic tags, one being positive, the other negative. Tagging each half of Opto-FLP with different optogenetic magnets ensures that one FLPA protein will associate with one FLPB protein.
Figure 1: Identification of the "Split Site" of Flippase. A structural analysis of flippase (PDB ID: 5C73) revealed a highly disordered region (pink) between two distinct portions of flippase (purple). It is our belief that splitting flippase at this disordered region (red arrow) will produce two portions of flippase, FLPA and FLPB, that can be recombined to produce a functional flippase protein.
Figure 2: Predicted Mechanism of Opto-FLP. By separating flippase into two halves and tagging each half with an optogentic magnet, we will be able to control flippase activity with blue light with spatial and temporal precision.
Molecular Cloning of pCDF-Duet-1-FLPA-pMag-nMag-FLPB
A typical cloned construct has one gene that is inserted into a digested vector. The Opto-FLP construct requires the insertion of multiple fragments into a vector, which can be completed using a process known as Gibson Assembly (Figure 3).
Figure 3: Gibson Assembly of FLPA-pMag and nMag-FLPB. The genetic sequence of Opto-FLP will be cloned into the pCDF-Duet-1 vector using Gibson Assembly in two separate reactions.
Prior to insertion, primers must be designed that roughly have 20 base pairs of homology to the gene of interest as well as homology to the vector itself. This will allow the complementary strands to be annealed, allowing the DNA fragments to be ligated together. There are three steps that allow for proper fragment insertion during Gibson Assembly. First, an exonuclease chews back DNA from the 5' end, revealing additional homology. Next, the resulting single-stranded regions on adjacent DNA fragments anneal to one another. This step is made possible by the exposed single stranded homologous ends created by the exonuclease. The final step entails a DNA polymerase incorporating nucleotides to fill in any gaps that were caused by the exonuclease or other forms of degradation. DNA ligase then covalently joins the DNA of adjacent segments, thereby removing any "nicks" in the DNA. The final product will be a circular vector now containing the genes of interest; in this case it will be the Opto-FLP construct in pCDF-Duet-1 (Figure 4).
Figure 4: Gibson Assembly Product of pCDF-Duet-1-Opto-FLP. By cutting pCDF-Duet-1 (A) in each multiple cloning site following each T7 polymerase promoter, the genetic sequence of each half of Opto-FLP could be inserted into pCDF-Duet-1 (B). This would allow for a ratiometric equal expression of both halves of Opto-FLP.
Proof of Concept of Opto-FLP Functionality
In order to demonstrate that our Opto-FLP construct is functional and is responsive to blue light, we transfected pCDF-Duet-Opto-FLP into BL21 DE3 E. coli cells, which are a strain of competent E. coli cells that have a promoter for T7 polymerase, from which transcription can be induced in response to application of a common molecular biology reagent known as Isopropyl β-D-1-thiogalactopyranoside (IPTG). pCDF-Duet-Opto-FLP will be dual-transfected into the BL21 DE3 E. coli cells with a colour-switch plasmid we modified: pRSET-FRT-mScarlet-STOP-FRT-eGFP (Figure 5).
Figure 5: pRSET-FRT-mScarlet-STOP-FRT-eGFP Colour-Switch Plasmid. The colour-switch plasmid was designed such that the T7 promoter was upstream of the two FRT sites flanking the mScarlet sequence and stop codons. There were three stop codons, with each stop codon in a different reading frame to prevent any possible downstream translation. The eGFP was located downstream of the second FRT site, such that it would only be translated if the mScarlet-STOP sequence was excised.
Through dual-transfection of BL21 DE3 E. coli cells with pCDF-Duet-1-Opto-FLP and the pRSET-FRT-mScarlet-STOP-FRT-eGFP plasmids, we will be able to produce a system that can spatio-temporally control fluorescence expression. In a dark environment, Opto-FLP will not be active, thus mScarlet will be translated, but the stop codons will prevent eGFP expression. If we subject the bacteria to blue light, we will be able to spatially control where we express Opto-FLP. Opto-FLP will excise the mScarlet sequence and stop codons from the plasmid, allowing for eGFP expression (Figure 6). Over time, we should observe a change in the fluorescence produced by the bacteria, as mScarlet proteins are degraded and no longer transcribed, while eGFP becomes transcribed and translated by these cells.
Figure 6: Opto-FLP Colour-Switch Assay. The BL21 DE3 E. coli bacteria that contain both the Opto-FLP construct and the colour-switch plasmid will allow for the assessment of the functionality of the Opto-FLP protein. In the dark (A), Opto-FLP will not be active and the BL21 DE3 cells will only produce the mScarlet protein and will fluoresce red. When exposed to blue light (B), Opto-FLP will become a fully functional protein and excise the mScarlet and STOP codon sequence in between the two FRT sites, allowing for eGFP transcription and translation. Over time, the BL21 DE3 cells will increase in green fluoresce and decrease in red fluorescence. The change in fluorescence can be measured and used as a proxy for Opto-FLP functionality.