Difference between revisions of "Team:Wageningen UR/Description/Biocontainment"
| Line 40: | Line 40: | ||
<br> | <br> | ||
<br> | <br> | ||
| − | The optogenetic kill switch is designed to exploit this mechanism. Rather than having both proteins expressed until the population encounters stress, the kill switch places MazF under control of pDawn. This means that in the darkness of the beehive, no toxin is produced, allowing the cell to remain stable. (Any leaky expression from the promoter can be countered by weakly expressed MazE). However, when the cell is exposed to (sun)light over a longer period of time, large amounts of un-countered MazF are produced, resulting in cell death. Like microbial vampires, any BeeT bacteria that make it outside will perish in the light of the sun. | + | The optogenetic kill switch is designed to exploit this mechanism. Rather than having both proteins expressed until the population encounters stress, the kill switch places MazF under control of pDawn. This means that in the darkness of the beehive, no toxin is produced, allowing the cell to remain stable. (Any leaky expression from the promoter can be countered by weakly expressed MazE). However, when the cell is exposed to (sun)light over a longer period of time, large amounts of un-countered MazF are produced, resulting in cell death. Like microbial vampires, any BeeT bacteria that make it outside will perish in the light of the sun. With a response time on the order of several hours for both toxin-antitoxin system and light sensor, the kill switch's design is well-suited to its function. Incidental exposure to light, such as opening the hive for beekeeping activities (as described by our beekeeper contacts), will not suffice to fully trigger the switch. |
<figure> | <figure> | ||
| Line 46: | Line 46: | ||
<figcaption>Basic design of optogenetic kill switch. Toxin MazF is kept repressed by pDawn, which is inactive in the dark. Leaky expression is countered by weakly expressed antitoxin MazE, forming equilibrium. In light, toxin expression greatly increases, causing cell death.</figcaption> | <figcaption>Basic design of optogenetic kill switch. Toxin MazF is kept repressed by pDawn, which is inactive in the dark. Leaky expression is countered by weakly expressed antitoxin MazE, forming equilibrium. In light, toxin expression greatly increases, causing cell death.</figcaption> | ||
</figure><br/> | </figure><br/> | ||
| − | |||
| − | |||
Revision as of 08:35, 14 October 2016
Appropriate biocontainment measures form a significant part of the BeeT project's Safety aspect. BeeT is intended to be used outside the lab, in beehives, where it will be in close contact with nature. As we cannot be sure about the effect on existing ecosystems if BeeT would be released in the environment, it must be engineered to die if it leaves the beehive. Our first measure to achieve this is a light-induced kill switch that relies on the balance of a bacterial toxin and an antitoxin that are expressed simultaneously. In the dark beehive, the system is unaffected. In the presence of blue light, a component of sunlight, the balance is disturbed in favour of the bacterial toxin. This will kill the bacterium.
As an additional safety measure, BeeT relies on the presence of a synthetic amino acid that is to be applied to the beehive. In the presence of the synthetic amino acid, catalytically dead Cas9 (dCas9) is produced that has the synthetic amino acid incorporated, at the expense of catalytically active Cas9. If BeeT escapes from the hive, the synthetic amino acid is no longer available, and dCas9 can no longer be formed. Instead, catalytically active Cas9 is produced which cuts the BeeT genome as well as any heterologous DNA that is present, thereby killing the bacterium and preventing horizontal gene transfer.
Optogenetic Kill Switch
Hijacking cellular control mechanisms
The optogenetic kill switch is the unification of two different genetic systems: pDawn, an artificially-created promoter system activated by light (the functioning of which is described here); and MazEF1, a toxin-antitoxin (TA) system native to E. coli. As the name implies, the latter system consists of two components: a toxin, MazF; and its complimentary antitoxin, MazE. The MazF protein functions as an endoribonuclease, cleaving mRNA and thereby inhibiting transcription, which is ultimately lethal for the cell. MazE forms a complex with MazF, preventing it from functioning. However, MazE is a more labile protein than MazF, degrading faster. For this reason, TA systems like MazEF are also known as addiction modules; the organism expressing the system becomes "addicted" to the antitoxin and is negatively affected should expression stop. In its native organisms, MazEF has a regulatory function. Under ordinary conditions, the proteins are co-expressed and the organism stays stable. However, under stressful conditions, such as nutrient starvation, this expression ceases. MazF is then free to cleave essential mRNAs, causing what is inferred to be a state of bacteriostasis.2 This process is reversible up until approximately 6 hours after taking effect, the so-called Point of No Return. Much like programmed cell death (PCD) in multicellular organisms, this prevents excess growth the bacterial population cannot afford, improving viability.
The optogenetic kill switch is designed to exploit this mechanism. Rather than having both proteins expressed until the population encounters stress, the kill switch places MazF under control of pDawn. This means that in the darkness of the beehive, no toxin is produced, allowing the cell to remain stable. (Any leaky expression from the promoter can be countered by weakly expressed MazE). However, when the cell is exposed to (sun)light over a longer period of time, large amounts of un-countered MazF are produced, resulting in cell death. Like microbial vampires, any BeeT bacteria that make it outside will perish in the light of the sun. With a response time on the order of several hours for both toxin-antitoxin system and light sensor, the kill switch's design is well-suited to its function. Incidental exposure to light, such as opening the hive for beekeeping activities (as described by our beekeeper contacts), will not suffice to fully trigger the switch.
Construction of kill switch genetic circuit
Unfortunately, creating a working version of the optogenetic kill switch proved too challenging to accomplish within the contest of the BeeT project. Particularly difficult was the creation of a genetic circuit actively expressing MazF. This possibly suggests that MazF's toxicity is simply too high for a chassis to handle without proper countermeasures. However, both modeling and wet lab experimentation proved the suitability of the pDawn system. For more information, see the Optogenetic Kill Switch notebook entry.
Cas9-based kill switch
To further minimize the risk of BeeT escaping the beehive, we aimed to include another safety mechanism in parallel with the light kill switch. An idea that came up was using a bacterial strain developed by Mandell and colleagues (2014)1 as a chassis for BeeT. This “biocontainment strain” can be confined to a certain area because it is auxotrophic for a synthetic amino acid, para-L-biphenylalanine (BipA). Several essential proteins of this bacterial strain were engineered to function only when BipA is incorporated in the active site, leading to death of the bacterium when the synthetic amino acid is not available. In our case BipA should be applied to the beehive, which will be the only place BeeT can survive given that the beekeeper continues to supply it.
A drawback of this system is that, even though the biocontained organism dies as soon as it is deprived of BipA, its heterologous DNA may remain in the environment. Since DNA is rather stable under certain circumstances2, there is a risk it is taken up by other bacteria through horizontal gene transfer3,4. In our project, we sought to complement the biocontainment strain by creating a switch to destroy heterologous DNA, depending on the presence of BipA. Additionally, the switch can be used to further strengthen the auxotrophy for BipA by targeting genomic DNA as well.
Using Cas9 to reinforce auxotrophy for a synthetic amino acid
To create the switch, we aimed to engineer Streptococcus pyogenes Cas9 to go from a catalytically dead form to a partially active form when BipA is not available. Cas9 is a well-studied nuclease that is involved in bacterial adaptive immunity (ref). Wild-type Cas9 can make double-strand DNA breaks because of two cleaving domains: a RuvC-like domain and a HNH-domain. While doing so, it is guided by a small RNA (guide RNA) that has a sequence complementary to the target DNA. When either the Asp10 residue in the RuvC domain or the His840 residue in the HNH domain are changed to alanine, Cas9 loses part of its activity. This partially active form, or nickase-Cas9 (nCas9) can make single-strand DNA breaks, or nicks. However, it was shown that nCas9 is able to cause double-strand DNA breaks when two spacers are available to nick DNA on opposite strands in close proximity6.
If both Ala10 and His840 are replaced by Alanine, all cleaving activity is lost resulting in a catalytically dead version of Cas9 (dCas9)5.
In this project, the codon for Asp10 (GAT) of the His840Ala-version of nCas9 is changed to the TAG stop codon. A new meaning is giving to this codon by providing the correct translation machinery for it: a tRNA that recognizes the TAG codon (tRNACUA) and an amino-acyl-RNA-synthetase (aaRS) that charges the tRNA either with BipA7 (resulting in dCas9) or Aspartate11 (resulting in nCas9). Normally the TAG stop codon is recognized by Release factor 1, thereby terminating translation (ref). For that reason, a genetically recoded strain that lacks any TAG stop codons as well as Release factor 18 was used, to make translation of the TAG codon more efficient.
In principle, BipA will be incorporated and a new version of dCas9, dCas9-Ala(GCT)10→ BipA(TAG), will be formed. To switch from expression of dCas9-Ala(GCT)10→ BipA(TAG) to our version of nCas9 (dCas9-Ala(GCT)10→ Asp(TAG)), dCas9-Ala(GCT)10→ BipA(TAG) is used to repress transcription9,10 of the aaRS/tRNACUA pair for Aspartate. This will happen as long as BeeT stays in the hive, where BipA is supplied. When BeeT leaves the hive no BipA-charged tRNACUA is available, the Cas9 transcript is not translated and dCas9-Ala(GCT)10→ BipA(TAG) is not formed. The result is that repression of the aaRS/tRNACUA pair for aspartate is released, the tRNACUA(Asp) is formed, and the Cas9 transcript can be translated again, this time resulting in formation of dCas9-Ala(GCT)10→ Asp(TAG). The dCas9-Ala(GCT)10→ Asp(TAG) is able to cut any DNA for which two suitable guide RNAs are offered, including all heterologous DNA.
Cloning and expression of Cas9 variants
The Ala(GCT)10→ BipA/Asp(TAG) mutation was introduced in dCas910. Wild-type Cas9, dCas9 and Cas9-Ala(GCT)10→ BipA/Asp(TAG) were cloned into the Expresso system for rhamnose induced expression from Lucigen12. This intoduced a C-terminal his-tag, which allowed for purification by Ni-NTA chromatography. A vector (pEVOL-BipA) containing the aaRS/tRNACUA pair for translating the TAG stop codon to incorporate BipA was already available from the biocontainment strain1. The alternative vector (pEVOL-Asp) containing the aaRS/tRNACUA pair for translating the TAG stop codon to Aspartate instead was constructed.
To test translation of the TAG stop codon and incorporation of BipA and Aspartate in Cas9, bacterial cultures transformed with Cas9-expresso constructs and pEVOL-vectors were used for production of the Cas9 variants. SDS-PAGE of the purified samples showed clear expression of both Cas9 and dCas9, and lower expression of dCas9-Ala(GCT)10→ BipA(TAG). A faint band could be seen for nCas9-Ala(GCT)10→ Asp(TAG), but since a control sample without any synthetic amino acid also displayed a faint band, this result is not conclusive (figure 2).
In vitro Cas9 cleavage assays
To assess the functionality of our Cas9 variants, in vitro Cas9 cleavage assays were performed. An PCR product containing the gene encoding RFP of 4140 bp was chosen as the substrate for cleaving to generate two fragments: one ~3100 bp fragment and one ~1040 bp fragment. All targets were chosen in close proximity. As a result, all cleavage products would be roughly the same length, and targets on opposite DNA strands should result in a double strand break when incubated with nCas9-Ala(GCT)10→ Asp(TAG).
From the assays, it can be concluded that our purified Cas9 is functional using several gRNAs targeting RFP, as the linear substrate was cleaved in all cases at least to some extend. As expected, dCas9 did not cleave DNA (one more assay, and I can have an indication that ala10BipA is also inactive). When two gRNAs that target the RFP gene on opposite DNA strands are offered to nCas9-Ala(GCT)10→ Asp(TAG), it is expected to see some cleaving caused by the two nicks in close proximity. Indeed, some cleaving was observed. However, also some cleaving was observed when offering only one gRNA. It has been shown that the His840Ala version of nCas913 (which is basically the same as nCas9-Ala(GCT)10→ Asp(TAG)) still has some residual double strand cleaving activity. Further testing is needed whether this caused the cleaving by nCas9-Ala(GCT)10→ Asp(TAG) in our case, or that it is an artefact of some sort (figure 3).
In conclusion, in this part of our project we showed that an artificial amino acid can be incorporated in Cas9 in response to the TAG stop codon (and that this version remains catalytically inactive → remains to be seen). We also tried to incorporate Aspartate in response to the TAG stop codon, and while we have some indications that this restored nickase activity, further testing is needed to verify whether this is the case.
References
References for optogenetic kill switch
1. Engelberg-Kulka, H., Hazan, R., Amitai, S. (2005). mazEF: a chromosomal toxin-antitoxin module that triggers programmed cell death in bacteria. Journal of Cell Science 118, 4327-4332. ↩2. Amitai, S., Yassin Y., Engelberg-Kulka, H. (2004) MazF-Mediated Cell Death in Escherichia coli: a Point of No Return. Journal of Bacteriology vol. 186 no. 24 8295-8300. ↩
3 Sat, B., Reches, M., Engelberg-Kulka, H. (2003) The Escherichia coli mazEF Suicide Module Mediates Thymineless Death. Journal of Bacteriology vol. 185 no. 6, 1803-1807. ↩
x Kitagawa, M., Ara, T., Arifuzzaman, M,, Ioka-Nakamichi, T., Inamoto, E., Toyonaga, H., Mori, H. (2005) Complete set of ORF clones of Escherichia coli ASKA library (a complete set of E. coli K-12 ORF archive): unique resources for biological research. DNA Research 12(5):291-9. ↩
References for Cas9-based kill switch
1. Mandell, D. J., Lajoie, M. J., Mee, M. T., Takeuchi, R., Kuznetsov, G., Norville, J. E., ... & Church, G. M. (2015). Biocontainment of genetically modified organisms by synthetic protein design. Nature, 518(7537), 55-60. ↩2. Romanowski, G., Lorenz, M. G., Sayler, G., & Wackernagel, W. (1992). Persistence of free plasmid DNA in soil monitored by various methods, including a transformation assay. Applied and Environmental Microbiology,58(9), 3012-3019. ↩
3. Thomas, C. M., & Nielsen, K. M. (2005). Mechanisms of, and barriers to, horizontal gene transfer between bacteria. Nature reviews microbiology, 3(9), 711-721. ↩
4. Smillie, C. S., Smith, M. B., Friedman, J., Cordero, O. X., David, L. A., & Alm, E. J. (2011). Ecology drives a global network of gene exchange connecting the human microbiome. Nature, 480(7376), 241-244. ↩
5. Mali, P., Aach, J., Stranges, P. B., Esvelt, K. M., Moosburner, M., Kosuri, S., ... & Church, G. M. (2013). CAS9 transcriptional activators for target specificity screening and paired nickases for cooperative genome engineering. Nature biotechnology, 31(9), 833-838. ↩
6. Shen, B., Zhang, W., Zhang, J., Zhou, J., Wang, J., Chen, L., ... & Skarnes, W. C. (2014). Efficient genome modification by CRISPR-Cas9 nickase with minimal off-target effects. Nature methods, 11(4), 399-402. ↩
7. Xie, J., Liu, W., & Schultz, P. G. (2007). A genetically encoded bidentate, Metal‐Binding amino acid. Angewandte Chemie, 119(48), 9399-9402. ↩
8. Lajoie, M. J., Rovner, A. J., Goodman, D. B., Aerni, H. R., Haimovich, A. D., Kuznetsov, G., ... & Rohland, N. (2013). Genomically recoded organisms expand biological functions. Science, 342(6156), 357-360. ↩
9. Qi, L. S., Larson, M. H., Gilbert, L. A., Doudna, J. A., Weissman, J. S., Arkin, A. P., & Lim, W. A. (2013). Repurposing CRISPR as an RNA-guided platform for sequence-specific control of gene expression. Cell, 152(5), 1173-1183. ↩
10. Bikard, D., Jiang, W., Samai, P., Hochschild, A., Zhang, F., & Marraffini, L. A. (2013). Programmable repression and activation of bacterial gene expression using an engineered CRISPR-Cas system. Nucleic acids research, 41(15), 7429-7437. ↩
11. Pastrnak, M., Magliery, T. J., & Schultz, P. G. (2000). A new orthogonal suppressor tRNA/aminoacyl-tRNA synthetase pair for evolving an organism with an expanded genetic code. Helvetica Chimica Acta, 83(9), 2277-2286. ↩
12. Expresso® Rhamnose Cloning & Protein Expression System ↩
13. Shen, B., Zhang, W., Zhang, J., Zhou, J., Wang, J., Chen, L., ... & Skarnes, W. C. (2014). Efficient genome modification by CRISPR-Cas9 nickase with minimal off-target effects. Nature methods, 11(4), 399-402.↩
