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 genetically modified organisms (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 (HGT). 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)
And 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
A broad spectrum system that non-specifically degrades DNA mediated by expression of DNase 1
Continuous culture
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, 2013). 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.
By simulating in miniature how a kill switch might behave in an industrial setting, the ministat provides a proof
of concept for how a kill switch might be maintained in larger chemostats during a continuous culture. A protocol
for running experiments in the ministat can be found here
Media container used to feed a single ministat chamber.
Peristaltic pump
Ministat chambers in heatblock and 1 litre Duran bottle used to collect effluent
Ministat running a preliminary
experiment to calibrate parameters such as dilution rate and temperature of the heat block. 50 ml burettes used
here to accurately measure effluent levels. 1 litre Duran bottles were used for effluent collection in the main
experiment due to greater volumes of effluent.
Our own growth curve was performed to determine the maximum specific growth rate of E. coli
BL21 (DE3) in our lab, but could not be conducted for a sufficient length of time to be accurate. A maximum specific
growth rate value of 1.730 was used (Cox, 2004). The ministat must be set to a flow rate at which dilution rate is less
than maximum specific growth rate. This prevents the culture being washed out of the growth chambers. 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 an average optical density (OD) of 3.47 for KillerRed samples, 3.64 for KillerOrange samples and 3.17 for
lysozyme samples. OD was measured daily with a Bug Lab OD scanner. When the same
sample was measured in a tecan infinite 200 pro plate reader the Bug Lab showed reading approximately three times higher.
The difference between the samples was consistent regardless of the method used to measure OD.
Method and Results
Metabolic kill switch
KillerRed and KillerOrange are homologues of Green fluorescence protein (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 et al, 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). Prolonged exposure and or high levels of ROS triggers apoptosis like mechanisms (Held, 2015). 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. In 2013, the iGEM team from Carnegie Mellon developed a phage delivery system of the KillerRed gene, which was then expressed in the infected bacteria, killing it on exposure to light. Carnegie Mellon 2014 continued characterisation of KillerRed and its monomeric form Supernova by analysing their photobleaching characteristics.
Firstly we improved KillerRed, an existing registry part, by codon optimising it for E. coli and
improved its characterisation by exposing cultures expressing the protein to previously untested light intensity. We designed and submitted KillerOrange as a new part and tested it in the same way. We compared the phototoxicity of KillerRed and KillerOrange to the commonly used Red fluorescence protein (RFP). 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 white light from a 4x8 LED array for 6 hrs. Samples were then spread plated and colony forming units (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 Isopropyl β-D-1-thiogalactopyranoside (IPTG) to a final concentration of 0.2 nM.
KillerRed is excited by green/yellow light (540-580 nm) and KillerOrange by blue light (460-490 nm). We constructed a box around the LED array to prevent ambient light entering, and attached acetate colour filters to provide the desired excitation frequency. Access to the inside of the box was gained through an opening cut in the front. With help from Ryan Edginton, we used a portable spectrometer (Ocean Optics USB2000+VIS-NIR-ES, connected to a CC3 cosine corrector with a 3.9 mm collection diameter attached to a 0.55 mm diameter optical fibre) to measure light spectra and absolute intensity in the visible range.
Light box and thermocouple
The average temperature in the light box during a 6 hr experiment was 38.6 °C
Metabolic Kill Switch: Results
Characterisation experiment
Fig.2-7 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. 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 covered in tin foil before being placed in the light box.
Fig. 3. 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. 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 covered in tin foil before being placed in the light box.
Fig. 5. 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. 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 covered in tin foil before being placed in the light box.
Fig. 7.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.8,10 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 liquid broth (LB). Error bars represent the standard error of the mean
Fig. 9,11 Data from Fig.8,10 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. 8. Comparison of CFUs formed by KillerRed exposed to light and kept in the dark.
Fig. 9. Percentage viable cells of KillerRed 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 HGT if lysis is successful. We added an OmpA signal peptide to Lysozyme C which
targets it to the periplasm, this was to ensure that the enzyme would be translocated to the cell wall where it would be
most effective. Our model predicted the complete degradation of the cell wall to be within the first generation of E. coli (details of the model can be found here). This gave us confidence that lysozyme would be effective.
Method
To show the activity of lysozyme, a molecular probes EnzCheck lysozyme assay kit from Thermo fisher
scientific was used. The coding sequence (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 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 HGT 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 mitigated.
We aimed to do this in 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 unsuccessful 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 (Fig. 5,7) under much lower light intensity
than is used in the literature (Sarkisyan et al, 2015). 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 a much less powerful LED array that produces 0.0012 W/cm2 at the wavelengths most effective for KillerRed and KillerOrange (Fig. 1) 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 (Fig. 5) and 12.7 % for KillerOrange (Fig. 7). 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. The leakiness of the T7 promoter used in our kill switches was quantified in our system by comparing protein production in the +IPTG condition and -IPTG condition. A one tail t test assuming equal variance was performed for the mean fluorescence values of the cultures tested in the light box. Fluorescence was used as a measure of protein production. No statistically significant difference was found between the + IPTG condition and – IPTG condition (p value for KillerRed 0.18, p value for KillerOrange 0.16). CFU counts
for + IPTG conditions were within the standard error of – IPTG (Fig. 5,7). 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 (Sarkisyan et al, 2015), the reason given for this was to "increase the fraction of mature protein". 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 (Fig. 9). A similar pattern is shown for KillerOrange but with around a two fold increase (Fig. 11). Both KillerRed and KillerOrange show greater numbers of colonies forming over time (Fig. 8,10). This number approaches the amount produced in the dark condition by 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.
Future
Failsafe
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 ribosome binding site (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.
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 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 (Sabri et al, 2013). 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. We attempted to use an Agilent Quick change multi site-directed mutagenesis kit to
remove the illegal sites. Unfortunately, this kit was also unsuccessful. We then learned that no strain of E.
coli that we had available would be able to replicate the pKD4 plasmid and this was the reason for our failed
transformations.
DNA degradation
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.
Ministat
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.
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stably carried in the host genome. Nature communications, 6:6989.
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Protocols
Transformation of competent E. coli cells
Defrost one 100 µl (or 200 µl if available) aliquot of E. coli competent cells per transformation,
plus one extra as a control (use whole tube and do not refreeze).
Add 1-5 µl of plasmid DNA (depending on concentration) to competent cells, mix by rolling or flicking
the eppendorf.
Incubate on ice for 40 mins.
Heat shock in eppendorf at 42 °C for 2 mins.
Incubate on ice for 5 mins.
Add 0.7 ml of pre-warmed LB medium, incubate at 37 °C, 200-220 rpm for 45-60 mins.
Spin down cells at 8000 rpm for 5 mins.
Remove 0.5 ml of the supernatant.
Resuspend cells in remaining supernatant.
Plate out 200 µl on agar laced with 100 µg/ml ampicillin, 50 µg/ml kanamycin, 35 µg/ml chloramphenicol
or 15 µg/ml tetracycline as appropriate.
Incubate at 37 °C overnight.
KillerRed KillerOrange protocol
Prepare 5 ml overnight cultures of KillerRed, KillerOrange and RFP transformed into E.coli
BL21 (DE3). This can be done from a glycerol stock or fresh transformation. Place in 37 °C 220 rpm
incubator overnight.
Prepare 250 ml erlenmeyer flasks with 50 ml of LB broth and add 50 µl of 35mg/ml chloramphenicol
to a final concentration of 35 µg/ml. Cover the flasks in tin foil.
Measure the OD of the overnight cultures and inoculate the 250 ml flasks to a starting OD of 0.1
on a tecan infinite 200 pro plate reader.
Once an OD of 0.23 has been reached, induce + IPTG samples with 100 µl of 0.1 M IPTG to a final
concentration of 0.2 nM. Incubate at 37 °C 220 rpm overnight.
Measure the OD and fluorescence of each culture.
Perform a serial dilution of the samples by adding 5 µl of undiluted culture to 4995 µl of LB
broth in a 10 ml falcon tube and invert 5-10 times, this is the 10-3 culture. Take 500
µl of the 10-3 and add it to 4500 µl of LB broth, invert the tube 5-10 times, this is t
he 10-4. Finally take 500 µl of the 10-4 and add it to 4500 µl of LB broth,
invert the tube 5-10 times, this is the 10-5 culture. Remove 500 µl from the 10-5
culture, all falcon tubes will contain 4500 µl of sample. Repeat this twice for all the samples
(KillerRed induced, KillerRed not induced, KillerOrange induced, KillerOrange not induced, RFP and control:
BL21 (DE3). One set of samples should be covered in tin foil, the other set left uncovered.
Place the falcon tubes in the light box, label down for the uncovered samples.
Expose for 6 hrs, take the temperature inside the box periodically using a thermocouple.
Spread plate 200 µl of each sample and incubate at 37 °C overnight.
Count the number of colonies on each plate.
Ministat Protocol
Prepare the desired number of media containers and culture chambers ready to be autoclaved.
Add media to the containers. Add 100 µl of inoculum to the sterile culture chamber. Connect the
media container to the culture chamber.
Arrange the ministat array in the heat block and set the effluent volume to around 35 ml by moving
the effluent needle up to the desired height in the chamber.
Set the peristaltic pump to 90 rpm and allow the chambers to fill. Once at 35 ml turn off the
peristaltic pump. Turn on the air pump to start aeration and agitation. Let the culture grow for 24 hrs
Turn on the peristaltic pump to 7.5 rpm this will give a flow rate of approx 4 ml/hr. Ensure
that the effluent tubes are uncovered and inside a collecting vessel.
Each day take a sample in a sterile container for testing. Glycerol stocks should be taken.
These can then be tested later.
HGT Protocol
Prepare a 5 ml overnight of pSB1C3 RFP transformed into E. coli DH5a in a 10 ml falcon tube.
Prepare working solution of lysozyme in reaction buffer from a molecular probes EnzCheck lysozyme
assay kit by diluting 1000 U/ml lysozyme C (Galus Galus) two fold in reaction buffer. 50 µl of working
solution is needed per reaction.
Add 50 µl of working solution to 50 µl of overnight culture in a PCR tube incubate at room temp
for 30 mins
Incubate overnight at 55 °C (this step is critical to ensure all lysozyme is inactivated)
Perform full transformation protocol using 3 µl of the lysate with BL21 (DE3) competent cells.
Plate the remaining lysate onto a chloramphenicol plate as a negative control.
Count colonies, overnight these and measure fluorescence.
