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<li><a id="links" style="margin:10px 0 30px 2px;padding:0;font-size:1.8vh;" href="https://2016.igem.org/Team:Exeter/Project">Lab Project</a></li>

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Introduction </div>

<p id="pp">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.</p>

<p id="pp">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 (<i>Gallus gallus</i>). 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.</p>

<p id="pp">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 <i>et al</i>, 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 <i>E. coli</i> 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 <a href="#MinistatProt">here</a>

<div class="col-xs-6"><span class="caption" style="padding: 5px 30% 5px 30%;">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. </span></div>

<p id="pp">Our own growth curve was performed to determine the maximum specific growth rate of <i>E. coli </i> 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 OD of 3.47 for KillerRed samples, 3.64 for KillerOrange samples and 3.17 for lysozyme samples. The ministat must be set to flow rate at which dilution rate is below the maximum specific growth rate. This prevents the culture being washed out of the chamber. 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 </div>

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<p id="pp">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 <i>E. coli</i> 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.</p>

<p id="pp">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 <i>E. coli</i> BL21 (DE3). Samples that were induced were done so with IPTG to a final concentration of 0.2 nM.</p>

<p id="pp">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 <a href="#">Ryan Edgington</a>, 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.</p>

             <span class="caption">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.</span>

             <span class="caption">The light spectra emited from our light source. Peaks occur in the optimum ranges for KillerRed and KillerOrange excitation.</span>

<p id="pp"><b>Fig.1-6</b> 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

             <span class="caption">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.</span>

             <span class="caption">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.</span>

             <span class="caption">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.</span>

             <span class="caption">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.</span>

             <span class="caption">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.</span>

             <span class="caption">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.</span>

<p id="pp">All samples from the ministat were tested using the KillerRed, KillerOrange protocol found <a href="#KRKOProt">here</a>. Glycerol stocks were made of the samples taken at each time interval, testing was done using these glycerol stocks.</p>

<p id="pp"><b>Fig.7,9</b> 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</p>

<p id="pp"> <b>Fig. 9,10</b> 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.</p>

<p id="pp">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.

<p id="pp">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 <i>Micrococcus lysodeikticus</i> 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 <i>E. coli</i> 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.</p>  

<p id="pp">The potential for horizontal gene transfer was tested using the lysozyme C (<i>Gallus gallus</i>) 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</p>

<p id="pp">No difference in CFUs was observed between the control and the samples producing lysozyme. The results of the EnzCheck lysozyme assay were inconclusive. </p>

<p id ="pp">The HGT experiment showed that DNA present in lysate can be successfully transformed into a different <i>E. coli</i> 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.</p>

<p id="pp">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 <i>E. coli</i> 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. <p>

<p id="pp">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 <i>et al</i>, 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 <i>E. coli</i> 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 <i>E. coli</i> regardless of the starting dilution factor.</p>

<p id="pp">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.</p>

<p id="pp">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 <i>E. coli</i> and in principle any wild type organisms outside the lab.</p>

<p id="pp"> 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.</p>

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 <i>ars</i>B locus using the pKD4 plasmid as a vector. Integrating at <i>ars</i>B has been shown not to affect <i>E. coli</i> growth (Sabri <i>et al</i>, 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 <i>E. coli</i> 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 <i>E. coli</i> that we had available would be able to replicate the pKD4 plasmid and this was the reason for our failed transformations.  </p>

<p id="pp">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 <i>polA</i>, <i>rpoC</i> and <i>topA</i> 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.</p>  

<p id="pp">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.</p>

Sarkisyan, K. S. et al, (2015).<i> KillerOrange, a Genetically Encoded Photosensitizer Activated by Blue and Green Light</i>. PLoS ONE.10(12)

integration of large DNA sequences, including whole metabolic pathways, onto the Escherichia coli chromosome at well-characterised loci</i>. Microbial Cell Factories, 12:60.

Miller, A. W. et al, (2013).<i> Design and Use of Multiplexed Chemostat Arrays</i>. Journal of Visualised Experiments. (72).</p>

             <li>Defrost one 100 µl (or 200 µl if available) aliquot of <i>E. coli</i> competent cells per transformation, plus one extra as a control (use whole tube and do not refreeze).

             <li>Add 1-5 µl of plasmid DNA (depending on concentration) to competent cells, mix by rolling or flicking the eppendorf.

             <li>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.</li>

             <li>Prepare 5 ml overnight cultures of KillerRed, KillerOrange and RFP transformed into <i>E.coli</i> BL21 (DE3). This can be done from a glycerol stock or fresh transformation. Place in 37 °C 220 rpm incubator overnight.</li>

             <li>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.

             <li>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. </li>

             <li>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.</li>

             <li>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 the 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.</li>

             <li>Add media to the containers. Add 100 µl of inoculum to the sterile culture chamber. Connect the media container to the culture chamber.</li>

             <li>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.</li>

             <li>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</li>

             <li>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. </li>

             <li>Each day take a sample in a sterile container for testing. Glycerol stocks should be taken. These can then be tested later.</li>

             <li>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. </li>

             <li>Add 50 µl of working solution to 50 µl of overnight culture in a PCR tube incubate at room temp for 30 mins</li>

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