Metabolic kill switch
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. The mechanism by which ROS kill cells is not fully understood , partly because they react quickly with contaminating metals to form more reactive species that obscure their own role in oxidation damage(Farr and Kogama, 1991), however prolonged exposure and or high levels of ROS triggers apoptosis like mechanisms(Held, 2015).
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 to determine the robustness of the kill switches over time.
Method
The following samples were tested for phototoxicity by exposing them to 12 W/m2 of white light for 6 hrs. Samples were then spread plated and CFUs 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.
Henceforth samples will be refered to as:
- KillerOrange induced
- KillerOrange not induced
- KillerRed induced
- KillerRed not induced
- RFP
- Control: BL21 (DE3)
See protocol for detailed method.
KillerRed is excited by green/yellow light (540-580 nm), KillerOrange by blue light (460-490 nm). We needed to provide light at these wavelengths at a reasonable intensity. We chose a light source consisting of a 4x8 LED array. With help from Ryan Edgington, we used an Ocean Optics spectrometer (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) to measure relative spectra and intensity. We constructed a box around the light to prevent ambient light entering. Access to inside the box was gained through an opening cut in the front.
Light box and thermocouple
The average temperature in the light box during a 6 hr experiment was 38.6 °C
A graph displaying the amount of light detected in the light box with the LEDs switched off compared to the amount of light detected inside the control (dark condition) sample wrapped in tin foil with the LEDs switched on. No significant difference is observed, this shows no light entered the samples covered in tin foil.
The light spectra emited from our light source. Peaks occur in the optimum ranges for KillerRed and KillerOrange excitation.
Metabolic Kill Switch: Results
Characterisation experiment
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. 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. 1. Percentage viable cells after 6 hrs in the light box. BL21 (DE3) transformed with pSB1C3 containing an RFP maker is compared to a control with no plasmid. 10 ml falcon tubes containing 4.5 ml of sample were covered in tin foil before being placed in the light box.
Fig. 2. Percentage viable cells after 6 hrs in the light box. BL21 (DE3) transformed with pSB1C3 containing an RFP maker is compared to a control with no plasmid. 10 ml falcon tubes containing 4.5 ml of sample were placed label down in the light box to allow maximum exposure to the light.
Fig. 3. Percentage viable cells after 6 hrs in the light box. BL21 (DE3) transformed with KillerRed (BBa_K1914003) is compared to a control with no plasmid. 10 ml falcon tubes containing 4.5 ml of sample were covered in tin foil before being placed in the light box.
Fig. 4. Percentage viable cells after 6 hrs in the light box. BL21 (DE3) transformed with KillerRed (BBa_K1914003) is compared to a control with no plasmid. 10 ml falcon tubes containing 4.5 ml of sample were placed label down in the light box to allow maximum exposure to the light.
Fig. 5. Percentage viable cells after 6 hrs in the light box. BL21 (DE3) transformed with KillerOrange (BBa_K1914001) is compared to a control with no plasmid. 10 ml falcon tubes containing 4.5 ml of sample were covered in tin foil before being placed in the light box.
Fig. 6.Percentage viable cells after 6 hrs in the light box. BL21 (DE3) transformed with KillerOrange (BBa_K1914001) is compared to a control with no plasmid. 10 ml falcon tubes containing 4.5 ml of sample were placed label down in the light box to allow maximum exposure to the light.
Ministat experiment
All samples from the ministat were tested using the KillerRed, KillerOrange protocol found here. Glycerol stocks were made of the samples taken at each time interval, testing was done using these glycerol stocks.
Fig.7,9 Average number of colonies after 0 h, 24 h, 120 h and 168 h of continuous culture. 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. 9,10 Data 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.
Fig. 7. Comparison of CFUs formed by KillerRed exposed to light and kept in the dark.
Fig. 8. Percentage viable cells of KillerRed exposed to light.
Fig. 9. Comparison of CFUS of KillerOrange exposed to light and in the dark.
Fig. 10. Percentage viable cells of KillerOrange exposed to light.
Enzymatic kill switch
Lysozymes are a group of enzymes that are an important part of the immune response against bacteria (Myrnes et al, 2013). They are defined as 1,4-fl-N-acetylmuramidases that cleave the glycosidic bond between the carbon 1 of N-acetylmuramic acid and the C-4 of N-acetylglucosamine in the peptidoglycan that makes up a bacterial cell wall (Jollès and Jollès, 1984)
Lysozymes are commonly used in mass spectrometry for protein mass calibration and are also effective lysing agents against gram-positive and gram- negative bacteria(Sigma aaldrich, 2016). Many previous iGEM teams have also used these enzymes and other lysis mechanisms as kill switches. For these reasons we thought Lysozyme C (Gallus gallus) would be a suitable candidate to test the effectiveness of lysis as a kill switch mechanism and investigate the potential for horizontal gene transfer if lysis is successful. We added an OmpA signal peptide to Lysozyme C which targets the periplasm, this was to ensure that the enzyme would be translocated to the cell wall where it would be most effective.
Method
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
Enzymatic Kill switch: Results
Characterisation
No difference in CFUs was observed between the control and the samples producing lysozyme. The results of the EnzCheck lysozyme assay were inconclusive.
Horizontal Gene Transfer
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.
DNase
DNAse I is a nonspecific deoxyribonuclease originally extracted from bovine pancreatic tissue. It degrades both double-stranded and single-stranded DNA resulting in the release of di-, tri- and oligonucleotide products with 5´-phosphorylated and 3´-hydroxylated ends (Vanecko, 1961). DNAse I has also been shown to work on chromatin and DNA:RNA hybrids (Kunitz, 1950).
DNAse I degrades these target polymer molecules through the hydrolytic cleavage of phosphodiester linkages in their backbone (Suck, 1986).
For a kill switch to be effective as a bio-containment device, the release of synthetic DNA must be mitigate. We aimed to do this is our project using the expression of DNase I. Dnase I is commonly used in a laboratory setting to degrade unwanted DNA. It was shown by Worrall and Connolly (1990) that expression of DNase I is possible in E. coli as long as it is under the control of a promoter with a strong off state. We constructed a part with DNase I under the control of the T7 promoter. Unfortunately no transformations were successful and all colonies produced contained empty plasmid backbone. Worral and Connolly reported that a promoter which is less tightly regulated (pKK223-3) would result in transformation failure. As was shown in our metabolic kill switch, the T7 promoter we used to control expression of the CDS is very leaky. This is likely the reason why transformations were unsuccesful. As immediately after transformation, production of DNase I would commence killing all the cells. If this is the case future work on a system that uses DNase I as a kill switch but under much tighter control may prove very effective.
Discussion
Metabolic Kill Switch: KillerRed and KillerOrange.
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.
Enzymatic Kill Switch: Lysozyme
The Enzcheck fluorescence assay used to determine lysozyme activity produced values that were not consistent with the CFU count of the plated sample. The HGT experiment we conducted showed that 50 µl of 500 U/ml lysozyme C normally used in the EnzCheck assay to produce a standard curve would effectively kill all the cells in a 50 µl sample of DH5a. The assay showed that 20x diluted sample produced near 500 U/ml activity readings yet this culture would still produce a lawn of bacteria when 200 µl was plated onto chloramphenicol. It is noted that the standard curve was of poor quality. The samples of lysozyme were assayed in the same way after continuous culture and did show a decrease in lysozyme activity, however the original readings that were used as a comparison have an error of sufficient size that this is not conclusive. The CFU count for lysozyme showed no difference from the control. Lysozyme added to a sample extra-cellularly was shown to lyse all the cells in our HGT experiment, even though gram-negative bacteria are partially protected from its action due to their outer membrane(Callewaert, 2008). Yet lysozyme produced intra-cellularly and targeted to the periplasm was not effective. There may have been issues with translocation of the protein to the target area, however this seems unlikely due to the effectiveness of its extracellular action. Another explanation may be that lysozyme as a kill switch mechanism is inherently ineffective, as any cells that are resistant will proliferate and a population that are not affected by its production very quickly develops. Regardless of the ability of lysozyme to kill cells effectively, we have shown that HGT is a concern with using this form of kill switch. Anti-biotic resistance markers commonly used in synthetic biology can be transferred to a different strain of E. coli and in principle any wild type organisms outside the lab.
