Bio-safety is an important issue in synthetic biology. Concerns surrounding synthetic organisms escaping into the environment have prompted the development of novel methods of bio-containment. Many iGEM projects that require an organism to be released from the lab use kill switches to address concerns about the effect of GMOs on the environment. Unfortunately, kill switches - inducible genetic devices that cause cell death - are poorly categorised in the standard registry of genetic parts. There is a distinct lack of quantitative data which prevents them being used with confidence.
Our initial concerns surrounding the use of kill switches as bio-safety devices were centred around their efficiency, robustness over time and ability to prevent transfer of synthetic DNA into the wild population. We contacted Dr Markus Gershater, chief scientific officer at Synthace Ltd, to ask him what the application of kill switches might be in an industrial setting and what evidence would be satisfactory for their use. Dr Gershater gave the view that kill switches would not be as effective or economical as the physical and chemical bio-containment methods that Synthace currently employ. One of his concerns was that any leakiness in a system would provide a strong selection pressure against cells with fully functional kill switches. In order for Dr Gershater to be satisfied, the kill switch would need to be tested across a wide range of environmental conditions and still remain effective. He would also need to see high efficiency levels, as in the context of large culture vessel, even a low proportion of cell survival would result in a large population lacking an effective kill switch. We also contacted Dr Tom Ellis of Imperial College London and asked his opinion on kill switch reliability. He gave the view that combining multiple mechanisms could greatly reduce organism escape rates after kill switch induction. Combining several kill switches was an approach we had discussed during the development phase of our project. Dr Gershater advised that the different systems would need to be truly orthogonal. For example, in an industrial setting two different kill switches that both rely on protein production could potentially be circumvented by the over expression of a useful enzyme that is being commercially produced. After talking to individuals from industry and academia about the strengths and limitations of kill switches, we decided to investigate the effectiveness of different types of kill switch, to quantify their robustness after several generations and investigate the possibility for horizontal gene transfer. We aimed to test if multiple kill switches in a system would reduce failure rate and if integration into the genome would increase stability. We have developed three types of kill switch to cover a broad range of strategies that may be employed in kill switch design: A metabolic kill switch that uses the production of reactive oxygen species to kill the cell, an enzymatic kill switch that uses the production of lysozyme C (Gallus gallus). We designed two systems that target DNA. A CRISPR based kill switch based on the work of Caliando and Voigt (2015) that targets specific sites for degradation, and a broad spectrum system that non-specifically degrades DNA mediated by expression of DNase 1.
Before starting the project we spoke to Prof. Robert Beardmore EPSRC Leadership Fellow in the Mathematical Biosciences at Exeter University. Much of his research has been into antibiotic resistance. We discussed how high selection pressure is applied by prolonged use of antibiotics and how kill switches may be analogous to this. It is clear that cells which develop a mutation that inactivates the kill switch would be strongly selected for. It was estimated that functional loss of the kill switch would occur in a short amount of time as a result, and if this was the case, could have strong implications for kill switch longevity. To test this we decided to use a ministat to perform a continuous culture. The ministat was developed in the Dunham lab at the University of Washington (Miller et al). Each ministat chamber is fed from its own media container via a peristaltic pump, with the culture volume set by the height of the effluent needle in the chamber. Air is bubbled through flasks of water to hydrate it and then used to agitate the culture. Chambers were inoculated with freshly transformed E. coli BL21 (DE3) and samples taken to test if the kill switches were still viable.
A growth curve was performed for E. coli BL21 (DE3) to determine the maximum specific growth rate. Dilution rate is equal to maximum specific growth rate. The dilution rate of the culture was calculated by measuring flow rate at a setting of 7.5 rpm on the peristaltic pump. For practical reasons the pump could not be run faster than this due to the amount of media needed. The dilution rate was set at 0.2 which produced cultures that grew at average OD of.... readings were taken with a Bug Lab OD scanner. When the same sample was measured in a tecan infinite 200 pro plate reader the Bug Lab showed approximately a reading three times higher, however the difference between cultures was consistent.
Our metabolic kill switches build on previous iGEM projects which have used the expression of highly phototoxic fluorescent proteins to kill the cells by exposing the culture to light. KillerRed and KillerOrange are homologues of GFP which, when irradiated with green and blue light respectively, generate reactive oxygen species (ROS). KillerRed has been shown to effectively kill cells when exposed to green light (540–580 nm) and is much less effective under blue light (460–490 nm) (Bulina, 2006). KillerOrange effectively kills cells when exposed to 450-495nm (Sarkisyan 2015), the range that KillerRed does not. Firstly we aimed to improve KillerRed, an existing registry part, by codon optimising it for E. coli and improving its characterisation by exposing it to previously untested light intensity. We characterised KillerOrange in the same way. Once we had established the efficiency of KillerRed and KillerOrange, ministat chambers were inoculated with samples of each.
The following samples were tested for phototoxicity by exposing them to 12 W/m2 for 6 hrs. Samples were then spread plated and CFU were counted. All parts were carried on the pSB1C3 plasmid and transformed into E. coli BL21 (DE3). Samples that were induced were done so with IPTG to a final concentration of 0.2 nM
See protocol for detailed method.
Fig.1-6 The average percentage viable cells for induced and uninduced samples after 6 hrs of exposure to 12 W/m2 of white light. Intensity was measured using an Ocean Optics USB2000+VIS-NIR-ES spectrometer, connected to a CC3 cosine corrector with a 3.9 mm collection diameter attached to a 0.55 mm diameter optical fibre. CFU count for the control condition was treated as 100 % and viable cells calculated as a proportion of that value. CFUs were not counted above 300, any lawns were assigned the value of 300. Error bars represent the standard error of the mean.The average temperature in the light box was 38.63 °C
Fig.7,9 Average number of colonies after 0 h, 24 h, 120 h and 168 h. Values were averaged across three biological repeats. A max value of 300 colonies is set as any plate with more than 300 colonies was not counted and assigned the max value. All samples were induced to a final concentration of 0.2 nM IPTG. All samples were diluted 1000 times in a final volume of 4.5 ml LB. Error bars represent the standard error of the mean
Fig. 9,10Data from Fig.7,8 represented as percentage viable cells over time. 100% viable is given when the CFU count for the kill switch condition equaled the control. Error bars represent the standard error of the mean.
Lysozyme is a common enzyme used in laboratories the Gallus gallus form is the basis of our enzymatic kill switch. It is bacteriolytic when transported into the periplasm of gram negative bacteria, hydrolysing the glycosidic bonds connecting N-acetylmuramic acid and N-acetylglucosamine. Under the control of a T7 promoter we induce lysis of the cell by adding IPTG.
To show the activity of lysozyme, a molecular probes EnzCheck lysozyme assay kit from Thermo fisher scientific was used. The CDS contains an OmpA signal peptide targeting it to the perisplasm therfore lysozyme will only be detectable if the cells have lysed. The kit uses a substrate containing Micrococcus lysodeikticus cell walls labelled with fluorescein to such as degree that fluorescence is quenched. The presence of lysozyme causes a sharp increase in fluorescence (AU) by easing the quenching. The increase in fluorescence is proportional to lysozyme activity in the sample. The fluorescence assay was used to measure the activity of the freshly transformed kill switch and that of the cultures grown in the ministat. CFUs were also used as a measure of efficiency by comparing number of colonies to a control. 5 ml ovenights of E. coli BL21 (DE3) transformed with pSB1C3 lysozyme were used to inoculate 250 ml Erlenmeyer flasks containing 50 ml of LB laced with 35 µg/ml chloramphenicol. Once an OD of 0.23 was reached IPTG was added to a final concentration of 0.2 nM. Protein production was allowed to proceed for 2 hrs. The sample was serially diluted (10-2,10-3,10-4). 200 µl of each dilution factor was spread plated and incubated at 37 °C overnight. CFUs were then compared to a control treated in the same way.
The potential for horizontal gene transfer was tested using the lysozyme C (Gallus gallus) provided in the EnsCheck lysosyme assay kit from molecular probes. Cells were lysed, the enzyme inactivated and then transformation of the resulting lysate performed. For a detailed protocol see HGT protocol
No difference in CFUs was observed between the control and the samples producing lysozyme. The results of the EnzCheck lysozyme assay were inconclusive
The HGT experiment showed that DNA present in lysate can be successfully transformed into a different E. coli strain with an average of 4 colonies per transformation (stdev=3.38). The BL21 (DE3) competent cells all gained the antibiotic resistance and RFP marker from the plasmid present in the lysed DH5a. 2 colonies from each plate were cultured over night and showed a fluorescence value concordant with that of the original culture. The starting cultures of DH5a had an average starting OD of 1.11 and fluorescence value of 258 before lysis. The BL21 (DE3) cultures transformed with the lysate had an average OD of 0.75 and average fluorescence of 306. None of the spread plated lysate produced any colonies, showing that all cells were killed in the lysis reaction.
We have shown that KillerRed and KillerOrange can effectively kill cells under much lower light intensity than is used in the literature (reference). On investigation into the kind of light source that was needed to produce the 1 W/cm2 of previous experiments (Bulina et al, 2005), it became clear that 1 W/cm2 was impractically bright. We decided to use an LED array that produces 0.0012 W/cm2 normally used for growing plants and expose our samples to light for a greater length of time. We showed that this was still effective with an average survival rate in the + IPTG condition of 2.2% for KillerRed and 12.7 % for KillerOrange. A wider range of exposure times and light intensities would greatly improve the characterisation of these parts, unfortunately time limitations prevented us from testing this. There was no (statistical) difference between the + IPTG condition and – IPTG condition. CFU counts for + IPTG conditions were within the standard error of – IPTG. For KillerRed the induced kill switch appears to be more effective whereas the uninduced switch is more effective in killer orange. The leakiness of the T7 promoter has likely lead to near equal expression both conditions, possibly exacerbated by the length of time that the cultures were left to grow in order for the protein to fully mature. The literature showed that cells had been kept in a cold room at 4 °C for 24 hrs before exposing the samples to light (reference), the reason given for this was to allow the protein to fully mature. We tested the validity of this as cultures were incubated at 37 °C 220 rpm overnight not 4 °C and the phototoxicity of KillerRed and KillerOrange was still evident. The light box itself had a negative effect on E. coli growth. Each sample was first diluted to 10-3,10-4 and 10-5 before exposure to light. The control showed fewer colonies at each dilution factor as would be expected, with the CFU count at a 10-3 dilution still being a lawn of bacteria. However in the dark condition, the control sample grew to a lawn of E. coli regardless of the starting dilution factor.
The continuous culture of KillerRed showed a 15 fold increase in the percentage of viable cells after 168 hrs. The average fluorescence reading for 0 hr KillerRed samples was 506.3 (recorded at an average OD of of 0.745). After 168 hrs the average fluorescence reading was 436 (at an average OD of 0.96). It seems unlikely due to the readings being similar that a mutation has occurred in the kill switch itself. As fluorescence is proportional to the amount of ROS being produced, up regulation of native E. coli enzymes that mitigate the effects of ROS may be the cause of the increase in cell survival. Future transcriptome analysis could provide interesting data on the mechanism of this change, this was unfortunately beyond the scope of this project.
One area that we were unable to explore was the incorporating multiple kill switches into the same system. Initially we aimed to construct an operon that contained KillerRed and KillerOrange. This was unfeasible with the cloning strategy that we were using as the overhangs that join the RBS to the CDS would not differentiate between KillerRed and KillerOrange. Constructing KillerRed and KillerOrange on plasmid backbones with different antibiotic resistance markers would allow both to be transformed together. This is a simpler way to test the hypothesis and would be interesting for the future.
While plasmids are widely used to carry genetic parts, integration into the host genome could prove a more robust approach to introducing genes into organisms. Genome integration removes the need for a selectable antibiotic resistance marker as parts will be faithfully replicated and the variability of copy number is removed. We aimed to investigate whether integration into the E. coli genome will affect the efficiency of our kill switches and whether they will remain functional for longer in a continuous culture. We aimed to use the lambda red recombination method to integrate our parts into the arsB locus using the pKD4 plasmid as a vector. Integrating at arsB has been shown not to affect E. coli growth (reference kiko paper). However the pKD4 plasmid contained illegal EcoRI and XBal restriction sites. To resolve this we decided to carry out site directed mutagenesis to change one nucleotide base pair in each sequence of the restriction sites. Primers were designed for use with the Q5 site directed mutagenesis kit. The first attempt using this kit involved a 2 step PCR reaction, this was shown by gel electrophoresis of the product to have been unsuccessful. The protocol was changed to a 3 step PCR reaction and a successful product was produced. The PCR product underwent a KLD reaction and was transformed into E. coli DH5a. The transformation was unsuccessful and so mutagenesis was carried out again and re-transformed. Each time the transformation was unsuccessful. Another mutagenesis kit, QC multi, which used all forward and reverse primers in separate reactions and could produce multiple mutations at once. Unfortunately, this kit was also unsuccessful. Therefore we decided to focus our efforts on other tasks within the lab.
We had hoped to develop a CRISPR based kill switch building on the work of Caliando and Voigt (2015). We designed the spacer array to target three essential genes polA, rpoC and topA using the deskgen platform. We selected three protospacers within the CDS of each essential gene. The cleavage sites were designed to be in the first third, the centre third and the final third of the CDS. The spacer array was designed to be carried on the pSB1C3 plasmid under the control of a constituitive promoter (BBa_J23100). Our aim was investigate how many essential genes would be needed for the kill switch to be effective, whether some genes were more effective targets than others and whether targeting multiple protospacers simultaneously was more effective than a single cleavage site. We designed primers to obtain the Cas9 and tracr RNA sequence from BBa_K1218011 provided in the distribution kit. After several attempts transformations remained unsuccessful. The spacer array could also not be produced as a G-block due to the high number of repeating sequences. A CRISPR based kill switch was shown by Caliando and Voigt (2015) to be stable for many months when integrated into the genome at multiple loci. Making this system available to iGEM teams could greatly improve on the short comings we have shown in the stability of toxic protein based switched carried on plasmids.
The modularity of the ministat allows several environmental conditions to be tested simultaneously. Future studies that would build on the work started in this project should include the testing of different media types, growth over different temperature ranges and cultures grown at a range of dilution rates.
References Bulina, M. E. et al., 2006. A genetically encoded photosensitizer. Nature Biotechnology.24(1). Sarkisyan, K. S. et al, 2015. KillerOrange, a Genetically Encoded Photosensitizer Activated by Blue and Green Light. PLoS ONE.10(12) Miller, A. W. et al, 2013. Design and Use of Multiplexed Chemostat Arrays. Journal of Visualised Experiments. (72).