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Revision as of 13:04, 27 September 2016
Introduction
Biosafety is an important issue facing synthetic biology and concerns surrounding synthetic organisms escaping into the environment has 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 project seeks to investigate the effectiveness of different types of kill switch, to quantify their robustness after several generations and compare the stability of these devices when integrated into the genome or carried on a plasmid. We hope to improve the standard of biosafety in the synthetic biology community and provide valuable data for the field to move forward. We have developed three types of kill switch; an improved metabolic kill switch which builds on a previous iGEM project , an enzymatic kill switch that uses the production of lysozyme to kill the cell, and DNA degradation mediated by DNAse production.
Design and Discussion
Metabolic kill switch
KillerRed and KillerOrange are fluorescent proteins 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 is a mutant of KillerRed that is excited from 420-530nm that has been shown to work alongside KillerRed (Sarkisyan,2015). Our project has improved the characterisation of a KillerRed codon optimised for E. coli. We are also characterising KillerOrange in the same way. We aim to include both KillerRed and KillerOrange in the same system so as to be a more phototoxic. High levels of ROS lyse the cell but can also damage the DNA, this is an attractive prospect when developing a kill switch to reduce the risk of horizontal gene transfer (HGT).
Method
Preliminary experiments were performed to calibrate the tecan reader. An OD of 0.4 on the cuvette reader was used as the optimum level of growth to induce protein production, this corresponded to 0.26 on the tecan reader.
5ml overnights of the transformed E. coli with the KillerOrange and KillerRed kill switches were used to inoculate five 250ml erlenmeyer flasks covered in tin foil containing 50ml of LB 35µg/ml Chloramphenicol. The five flasks were inoculated with the following conditions.
- KillerOrange induced
- KillerOrange not induced
- KillerRed induced
- KillerRed not induced
- pSB1C3 RFP
The optical density of each culture was measured every 1.5 hours until it had reached 0.26 on the tecan plate reader, then 100µl of 0.1M IPTG was added to induce the protein production in the desired cultures. The cultures are then incubated at 37℃ and 220rpm overnight. A serial dilution was then performed to make 10-3,10-4 and 10-5 dilutions of the cultures. 4.5ml of each dilution factor was placed in 10ml falcon tubes one set were placed on their side label down in the light box, the other set were covered in tin foil and also placed in the light box. The temperature was taken periodically inside the box using a thermocouple. After 6hrs of irradiation, spread plates were performed for all the samples. CFU’s were then counted and the dark and light conditions were compared.
Results
Enzymatic
Lysozyme is a common enzyme used in laboratories and the gallus gallus form is basis of our enzymatic kill switch. It is bacteriolytic when transported into the periplasm of gram negative bacteria, hydrolysing the glycosidic bond connecting N-acetylmuramic acid and N-acetylglucosamine. Under the control of a T7 promoter we can induce lysis of the cell by adding IPTG. Simply lysing the cell does not prevent HGT it may even exacerbate the problem. We have tested the liklihood of HGT by lysing a culture of cells producing RFP, incubating the lysate in a 90 degree water bath to inactivate the lysozyme. Then growing competent E. coli with the lysate will show if the competent cells take up the RFP gene. Lysozyme EnzChek assaying kit is used to measure lysozyme activity in solution where an increase in fluorescence is proportional to lysozyme activity.
Method
Results
DNA degradation
DNA degradation
DNase 1 is an endonuclease that non-specifically cleaves DNA. We are creating a kill switch with DNase 1 to address the foremost problem associated with Biosafety - lateral gene transfer. The DNase 1 kill switch, on induction, will degrade DNA and kill the cell. Another biosafety ‘kill switch’ we aimed to test was DNase I. We received a G-block of DNase I DNA and cloned it using the MoClo method. Each time the product from the PCR reaction was transformed we encountered issues. Many times no colonies would form, and in cases when colonies were produced DNase was not present. This could be because colonies containing DNase I has their DNA immediately destroyed. However production of Lysozyme was successful so we continue work with this as a kill switch instead.
Continuous culture
We performed a continuous culture in a ministat developed in the Dunham lab at the University of Washington (Miller et al). This was to test the longevity of our kill switches. Each ministat chamber is fed from its own media container via a peristaltic pump. The culture volume is set by the height of the effluent needle in the chamber. Our preliminary experiment used E. coli expressing RFP, we tested it under different media conditions using LB with and without salt. The flow rate was set to around 4ml/hr.
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).
Future
Genome integration
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 the parts will be faithfully replicated and the variability of copy number is removed. We are investigating 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 have used the lambda red recombination method to integrate our parts into the arsB locus. Integrating in this locus does not affect E. coli growth (reference kiko paper). We used the pKD4 plasmid as a vector to carry our parts, this had a pst1 site and two xba1 sites. Using site Q5 site directed mutagenesis kit we removed these illegal restriction sites. We planned to test how the robustness of a kill switch that was integrated into the genome was different to the same kill switch gene that was incorporated on a plasmid. To do this we aimed to use the Lambda red genome knock in method, using the pKD4 plasmid to integrate KillerRed and KillerOrange into the E.coli DH5α genome at the arsB location. However the pKD4 plasmid we had available contained EcoRI and XBal restriction sites corresponding to the enzymes used by iGEM teams for parts. To resolve this we decided to carry out site directed mutagenesis to change one nucleotide base pair in each sequence of the restriction sites so as to remove them. Primers were designed for use with the Q5 kit. The first attempt using this kit involved at 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 DH5α E.coli. The transformation was unsuccessful and so mutagenesis was carried out again and re-transformed. Each time the transformation was unsuccessful. We decided to use 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.
