Overview
We aim to independently construct epigenetically regulated kill switches in S.cerevisiae and E.coli. In a given environment, epigenetic modifications suppress the expression of toxic genes. Upon leaving the specified environment, these epigenetic modifications gradually dissipate, leading to the expression of toxic genes and subsequent cell death. We will employ epigenetic modifications and CRISPR technology to engineer these kill switches in both Saccharomyces cerevisiae and E.coli. Furthermore, we intend to develop a detection circuit within the wind-up cell and create test strips, enabling the wind-up cell to function during the delayed death phase. This page will provide an exposition of the design details and the iterative process of our project.
Kill Switch
S.cerevisiae Cycle 1
In S.cerevisiae, our initial approach involves the construction of a fusion protein consisting of dCas9 and a deacetylase enzyme as an inhibitory tool. This fusion protein is designed to specifically suppress the expression of toxic genes, with the expectation that when the organism departs from a specific environment, toxic gene expression will be enabled, ultimately leading to cell death.
Design
We aim to design and construct two types of plasmids. One type of plasmid consists of constitutive promoters and toxic genes specific to Saccharomyces cerevisiae. We chose two toxic genes, MazF and EcoRⅠ, and constructed two plasmids, pR_DK02 (containing MazF expression cassette BBa_K4703028) and pR_DK03 (containing EcoRⅠ expression cassetteBBa_K4703029). The other type is a tool plasmid based on epigenetic modifications, which involves the fusion of dCas9 and histone deacetylase (HDAC) to create the tool plasmid pR_DK01 (containing dCas9 and HDAC fusion protein expression cassette BBa_K4703031).
By employing gRNA specifically targeting the promoters of toxic genes, the dCas9 carrying histone deacetylase suppresses toxic gene expression through epigenetic modifications.
More details
Figure 1. Schematic diagram of the gene pathway of dCas9 and sir2 fusion protein.
Figure 2. Schematic diagram of the gene pathway of MazF/EcoRI.
Build
Tool Plasmid: We designed two primers using the computational tool and used PCR to obtain the sir2 deacetylase sequence. Subsequently, we employed Gibson assembly to connect this sequence with a linker and the dCas9 sequence on the pRS42H vector, resulting in the construction of the tool plasmid pR_DK01.
Toxic Gene Plasmids: We obtained the MazF sequence from the Escherichia coli genome using PCR. We also synthesized a codon-optimized EcoRⅠ enzyme from a commercial source. Subsequently, we inserted these two types of toxic genes into the pRS415 vector downstream of a constitutive promoter. This process led to the generation of two distinct toxic gene plasmids, named pR_DK02 and pR_DK03.
More detailsTest
We initially transformed the tool plasmid pR_DK01 into the Saccharomyces cerevisiae strain BY4742. Upon successful validation of the transformants, we designed primers using a computational tool and used PCR to split each toxic gene plasmid into two segments. Subsequently, both segments of the plasmids were simultaneously transformed into the previously validated Saccharomyces cerevisiae strain. This method was expected to delay the expression of the toxic genes, allowing the tool plasmid ample time for targeted suppression.
The transformants, confirmed to contain the toxic genes, were cultivated in selective liquid SC medium lacking leucine to induce loss of the tool plasmid through selection pressure.
Figure 3. Validation of toxic gene effects. a. Transferring toxic gene EcoRⅠ into Saccharomyces cerevisiae BY4742 resulted in sterile colony growth. b. Transferring empty vector pRS415 into Saccharomyces cerevisiae BY4742 resulted in abundant colony growth.
More detailsLearn
Based on the results, it is hypothesized that the lethality effect of the toxic genes may be excessively strong, or the suppression effect of the fusion proteins is relatively weak. Subsequent plans include optimizing the tool to achieve a higher level of suppression, aiming to construct a functional kill switch.
Saccharomyces cerevisiae Cycle 2
(This achievement is distinct from Contribution for Bronze)
In order to enhance the suppression of toxic genes more effectively, we conducted a literature review and discovered a novel approach. Certain specific sequences have been found to recruit silencing factors. We anticipate achieving successful suppression of the toxic genes through this method.
Design
We conducted research and selected HML-I and HML-E silencers, known for their ability to recruit other silencing factors, particularly the repressive Sir2/Sir3/Sir4 complex, in the establishment of heterochromatin-like structures at the HM loci. Our aim is to incorporate these sequences at both ends of the toxic genes to achieve a stronger suppression effect.
Figure 4. Schematic diagram of the gene pathway of MazF/EcoRI with HML-I and HML-E silencers inserted at both ends.
More detailsBuild
Initially, we designed two primers using a computational tool. Subsequently, we utilized PCR to extract the HML-I (BBa_K4703006) and HML-E (BBa_K4703007) sequences from the Saccharomyces cerevisiae genome, while adding 20bp homologous arms to the fragments. We employed the Gibson assembly method to insert the HML-I and HML-E silencers into both ends of the toxic genes in plasmids pR_DK02 and pR_DK03. This resulted in the construction of plasmids pR_DK04 (containing BBa_K4703025) and pR_DK05 (containing BBa_K4703026).
More detailsTest
Plasmids pR_DK04 and pR_DK05 were separately introduced into the Saccharomyces cerevisiae strain BY4742, resulting in the successful generation of transformants. Subsequently, we performed colony PCR and sequencing on multiple transformants, obtaining several strains with correct sequences. This provided evidence that the method can effectively suppress the expression of toxic genes to some extent.
Subsequently, we subjected several of these transformants to colony PCR and sequencing. We obtained multiple strains with correct sequences, thereby validating the successful suppression using this method.
Figure 5. The growth status of strains containing the kill switch. a. Transferred the pR_DK03 plasmid into Saccharomyces cerevisiae containing pR_DK01. b. Transferred the empty vector pRS415 into Saccharomyces cerevisiae containing pR_DK01 as a control.
Figure 6. Colony PCR verification to validate the above-mentioned transformants. a. Colony PCR verification to validate the pR_DK05. b. Colony PCR verification to validate the pR_DK04.
More detailsLearn
By incorporating HML-I and HML-E silencers on both sides of the toxic gene, recruiting deacetylation-associated proteins, we successfully achieved the suppression of toxic gene expression through epigenetic modifications.
Saccharomyces cerevisiae Cycle 3
Due to the immediate cell death upon leakage and expression of toxic genes, characterizing the suppression efficiency of regulatory elements becomes challenging. To assess the suppression effect of our regulatory elements on downstream genes, we replaced the toxic gene with sfGFP for characterization purposes. Firstly, we tested the suppression effect on downstream fluorescent proteins using the fusion proteins constructed as mentioned above. Subsequently, we plan to employ two promoters, CHO and CYC1, to evaluate the suppression effect on downstream genes induced by different promoters.
Figure 7. Schematic diagram of the gene pathway of sfGFP with CHO promoter (a) or CYC1 promoter (b).
Build
We employed Saccharomyces cerevisiae homologous recombination to integrate CHO-sfGFP and CYC1-sfGFP into the Saccharomyces cerevisiae genome.
Test
Subsequently, we performed colony PCR and sequencing on multiple transformants, obtaining several strains with correct sequences. After 24 hours of cultivation, we conducted Microplate Reader to assess gene expression. The results demonstrated a significant suppression effect of our regulatory elements on downstream fluorescent protein genes. Furthermore, it was evident that different promoters had a notable impact on the suppression of downstream genes.
Learn
The aforementioned experiments have substantiated the reliability of our epigenetic modification elements. The experimental results have inspired us to explore the impact of different promoters on the suppression of downstream genes. Consequently, we have relayed these findings to the dry lab team, who will further elucidate the underlying reasons for these differences through modeling. The discovery we have made holds significant implications for our subsequent experiments.
E.coli Cycle 1
(This achievement is distinct from Contribution for Bronze)
After conducting a thorough investigation, we have identified DNA methylation as a prevalent epigenetic modification mechanism in Escherichia coli. By employing methylation on promoters with specific sequences, regulation of downstream gene expression can be achieved.
The Dam methylase can recognize and bind to Dam methylation sites on the mioC promoter and dnaAP2 promoter.
We harnessed the Dam methylase from the Escherichia coli genome to investigate the inhibitory effects of methylation on two promoters by conducting suppression tests on fluorescent protein gene.
More details
Figure 8. Schematic diagram of the gene pathway of RFP with mioC promoter (a) or DnaAP2 (b) promoter.
Build
We obtained fragments of the mioC promoter (BBa_K4703004) and dnaAP2 promoter (BBa_K4703003) through primer annealing using custom-synthesized primers.
Subsequently, we employed Gibson assembly to connect the mioC promoter and dnaAP2 promoter upstream of the RFP fluorescent protein gene, and these constructs were integrated into the pETDuet-1 plasmid, resulting in the generation of pR_DK06 (containing the mioC-RFP expression cassette BBa_K4703020) and pR_DK07 (containing the dnaAP2-RFP expression cassette BBa_K4703019) plasmids.
We procured the JM110 strain with a commercial exogenous Dam gene from a supplier, while the MG1655 strain, containing the endogenous Dam methylase, was obtained from our laboratory.
More detailsTest
The pR_DK06 and pR_DK07 plasmids were transformed into the JM110 strain lacking the endogenous Dam methyltransferase and the MG1655 strain harboring the endogenous Dam methylase.
After 12 hours of cultivation, fluorescence intensity was measured using a Microplate Reader.
It is evident that the fluorescence intensity of JM110 transformants carrying pR_DK06 was lower than that of the MG1655 transformants. Conversely, the fluorescence intensity of JM110 transformants containing pR_DK07 was higher than that of the MG1655 transformants.
Figure 9. The growth status of MG1655 and JM110 strains containing the pR_DK06 plasmid or the pR_DK07 plasmid.
Figure 10. Fluorescence intensity of JM110 and MG1655 cultured for 12 hours.
More detailsLearn
Experimental results have indeed demonstrated that the Dam methylase can suppress downstream gene expression of mioC promoter. However, it was unable to achieve suppression of downstream gene expression when acting on the dnaAP2 promoter. Consequently, for subsequent experiments, we have chosen to focus on further investigations involving the mioC promoter.
The Dam methylation of the mioC promoter has exhibited a certain degree of inhibitory effect on downstream genes. We aim to enhance the suppression effect through further research and explore methods to assist epigenetic modifications in achieving better repression outcomes.
E.coli Cycle 2
After conducting research, we have identified CRISPRi as a commonly used tool for inhibiting gene expression. Therefore, we intend to utilize CRISPRi to complement epigenetic modifications for enhanced suppression of downstream genes.
Design
To avoid excessive plasmid introductions, we plan to combine the dCas9 plasmid and sgRNA scaffold plasmid from the commonly used CRISPRi system into a single plasmid. Transcriptional expression of dCas9 will be controlled using a lactose promoter to achieve control over the kill switch.
Considering the issue of plasmid enrichment, we have selected ccdB (BBa_K4244056) as the toxic gene because ccdB-tolerant strains can be purchased. To achieve better targeted inhibition, we have designed multiple sgRNAs, each targeting the ccdB promoter and open reading frame, respectively.
Figure 11. Schematic diagram of the gene pathway of dCas9 with trc promoter.
More detailsBuild
We employed Gibson assembly to connect the sgRNA scaffold to the plasmid regulated by the trc promoter (BBa_K3121004) for dCas9 (BBa_K4703000). Subsequently, we used the Golden Gate method to replace different sgRNAs, resulting in the generation of various plasmids. The mioC promoter and ccdB toxic gene were commercially synthesized and assembled onto a vector to obtain the plasmid mioC-ccdB (pR_DK11).
The mioC promoter and ccdB toxic gene were commercially synthesized and assembled onto a vector to obtain the plasmid mioC-ccdB (pR_DK11).
To enrich the pR_DK11 plasmid, we purchased the DB3.1 strain, which is tolerant to the ccdB protein, and performed the plasmid enrichment in DB3.1.
More detailsTest
We validated the toxicity of ccdB using MG1655 and found that MG1655 strains containing the ccdB gene exhibited minimal growth.
The constructed tool plasmids were transformed into MG1655. After selecting the correct transformants, susceptible cultures were prepared under 1000X induction. Then, the pR_DK06 plasmid was introduced, and the cultures were incubated for 18 hours. Fluorescence intensity was measured using a fluorescence quantitative plate reader.
Figure 12. Fluorescence values measured under different conditions after 18 hours of cultivation.
More detailsLearn
CRISPRi has indeed demonstrated a pronounced inhibitory effect. In the next step, we aim to combine CRISPRi with enhanced epigenetic modifications to achieve stronger suppression of downstream genes.
E.coli Cycle 3
We plan to enhance the suppressive effects of epigenetic modifications and combine them with CRISPRi to achieve a combined inhibition of downstream gene expression.
Design
We plan to introduce exogenous Dam methylase to co-express with endogenous Dam, reinforcing the suppressive effects of epigenetic modifications on gene expression.
We have designed two strategies combining exogenously introduced Dam with CRISPRi:
Approach 1: Dam and the toxic gene are constructed on the same plasmid, and their expression is controlled by the T7-lac promoter (BBa_R0187). The same inducer is utilized for simultaneous control of epigenetic modifications and CRISPRi.
Approach 2: A fusion protein of dCas9 and DAM (BBa_K4703002) is constructed. Guided by sgRNA, this fusion protein targets the specific sequence, achieving the inhibitory effect on the target site.
Figure 13. Schematic diagram of the gene pathway of our Option.
More detailsBuild
Approach 1: The dam gene, obtained from the Escherichia coli genome, was assembled onto the mioC-RFP plasmid using Gibson assembly, resulting in the plasmid mioC-RFP-Dam (pR_DK10).
Approach 2: Through Gibson assembly, Dam was connected downstream of dCas9 to construct the fusion protein plasmid pR_DK08.
More detailsTest
We transferred the plasmids constructed using Gibson assembly into Escherichia coli. We successfully constructed pR_DK08 and obtained transformants containing the pR_DK08 plasmid.
More detailsLearn
We transferred the plasmids constructed using Gibson assembly into Escherichia coli. We successfully constructed pR_DK08 and obtained transformants containing the pR_DK08 plasmid.
Whole-cell test paper
(This achievement is distinct from Contribution for Bronze)
We plan to create a whole-cell test paper based on the wind-up cell containing a kill switch as the core. When the test paper is activated, the kill switch is triggered immediately, initiating a countdown for cell death. The wind-up cell will function during the delayed cell death period, ensuring safer substance detection.
Design
We intend to utilize plasmids consisting of IPTG-inducible promoters coupled with the RFP fluorescent protein gene and TET-inducible promoters coupled with the GFP fluorescent protein gene as the detection circuit for our whole-cell assay.
To explore the detection threshold of the whole-cell assay, we plan to perform gradient dilutions of inducers to test the fluorescence intensity at different concentrations.
Furthermore, considering that the usage environment of the assay may not be strictly sterile, in order to better reflect the practical working conditions of the assay, we intend to conduct experiments both in sterile and non-sterile environments.
More detailsBuild
We obtained the lac-RFP (pR_DK13) plasmid and tet-GFP (pR_DK12) plasmid from the laboratory. The transformants of the former emit red fluorescence under IPTG induction, while the transformants of the latter emit green fluorescence under doxycycline induction.
Figure 14. Schematic diagram of the gene pathway of pR_DK12 (a) pR_DK13 (b) and pR_DK14 (c).
We constructed the tet-RFP (pR_DK14) plasmid through enzyme digestion and ligation. The transformants of this plasmid emit red fluorescence under doxycycline induction.
Cell immobilization: We fixed the engineered bacteria on the test paper through a crosslinking reaction with a solution of sodium alginate and calcium chloride to form the detection zone.
More detailsTest
We placed the test paper into a petri dish and followed the protocol to immobilize suitable bacterial cells on the test paper. After induction, we recorded the fluorescence under a blue light illuminator at specified time intervals.
Figure 15. Test paper result chart of lac-RFP.
Figure 16. Test paper result chart of tet-RFP.
To meet user’s requirements, we repeated the aforementioned detection procedures in a non-sterile environment. The results indicate that, in comparison to test paper produced under aseptic conditions, there was no significant reduction in the fluorescence intensity of the test paper.
More detailsLearn
We successfully fabricated whole-cell test paper. Among them, the whole-cell test paper containing pR_DK13 and pR_DK12 plasmids exhibited fluorescence upon induction, while those containing pR_DK14 plasmid did not. Therefore, we chose pR_DK13 and pR_DK12 plasmids for subsequent testing.
Our whole-cell test paper displayed fluorescence in a contaminated environment outside the clean bench that was not significantly different from the results obtained within the clean bench. Thus, the whole-cell test paper we produced are suitable for use in both contaminated and sterile environments.
Model
In the course of our project development, our team iteratively refined the dry experimentation models using the design-build-test-learn methodology. Serving as the core of our software and model development, we engaged in iterative processes of algorithms and models. We meticulously compared the computational data generated by our models with the results from wet lab experiments, conducting a thorough comparative analysis to assess the accuracy of our models. This iterative cycle persisted, culminating in the successful creation of two pivotal models: a toxic gene expression model and a Cell Death Simulation Model.
To explore our software toolkit, please visit our Gitlab repository.
Toxic gene expression model Cycle 1
(This achievement is distinct from Contribution for Bronze)
The toxic gene expression model emerged as a product of our meticulous alignment between dry and wet experimentation processes.
Design
We engaged in in-depth, face-to-face discussions with the wet lab team, thoroughly understanding their experimental procedures. Subsequently, we conducted extensive literature reviews, comprehensively considering various influencing factors relevant to the model. Through this process, we precisely defined the direction and objectives for establishing the toxic gene expression model.
Build
Building upon intensive research, we initially formulated multiple dynamic models, employing systems of multivariate ordinary differential equations to solve key parameters such as toxic gene transcription product concentration, epigenetic enzyme concentration, promoter modification ratio, cell death rate, and toxic gene expression levels.
Test
During the testing phase, we endeavored to integrate these models using computer modeling tools. Initially, we attempted to describe toxic gene expression levels using the Hill function, generating corresponding visual representations. However, significant disparities were observed between these outcomes and the observations from wet lab experiments.
Learn
We recognized the pivotal role of specific model selection in accurately depicting the experimental process. To enhance precision in our descriptions, we discerned the imperative need for the model involving promoter modification by methyltransferase to more closely align with our wet lab operational procedures and the established toxic gene expression model.
This meticulous approach and continuous refinement facilitated the development of our robust toxic gene expression model, bridging the gap between theoretical frameworks and real-world experimental nuances.
Toxic gene expression model Cycle 2
Design
In order to precisely delineate the processes of epigenetic modification, toxic gene expression, and cell growth, we reengineered the models associated with these aspects. This redesign aimed at more accurately predicting toxic gene expression levels and the dynamics of cell growth and death.
Build
Using computer modeling tools, we meticulously modeled the new set of equations, generating corresponding code. Subsequently, we developed a application (app) tailored for model manipulation and analysis, ensuring seamless integration and functionality.
Test
The newly formulated model code was coupled with specific parameter values to generate graphical representations. Comparative analysis was then conducted, aligning the model outputs with experimental results. Through iterative optimization, the model outputs gradually converged with the experimental data. Additionally, wet lab experiments revealed a correlation between the strength of epigenetic modifications and promoter activity.
Learn
To accurately capture the relationship between promoter activity and epigenetic modification effects, we recognized the need to introduce a standardized model linking promoter activity with epigenetic modifications. This step was essential for a comprehensive understanding of their intricate interplay, forming the foundation for further refinement.
Toxic gene expression model Cycle 3
(This achievement is distinct from Contribution for Bronze)
Design
The introduction of the parameter alpha, correlated with the inherent characteristics of the tools used, facilitated a more nuanced depiction of the relationship between promoter activity and epigenetic modification effects. This inclusion allowed for a finer-grained analysis of their mutual interactions.
Build
During application development, we seamlessly incorporated new code into the existing app, adjusting its controls. This effort resulted in the creation of an updated version of the application, enhancing its capability to meet our evolving requirements. The update primarily focused on the refinement and optimization of model parameters.
Test
Comprehensive testing was conducted under varied conditions: maintaining consistent promoter activity while altering epigenetic tools, and vice versa. These tests yielded invaluable experimental data, enabling a profound analysis of the model's performance and behavior in different scenarios
Learn
The testing process underscored the significance of optimizing model outputs to achieve a higher degree of accuracy and reliability. This realization serves as a pivotal guiding principle for our subsequent research and improvement endeavors, directing our focus towards continual enhancements and advancements in our modeling approach.
Cell Death Simulation Model
(This achievement is distinct from Contribution for Bronze)
Design
To simulate the macroscopic process of our test paper entering the environment to be tested - the death of toxic protein-expressing cells, we established a cell death simulation model.
Build
We used Python to write code for the cell death model, including the cell distribution cellular automaton model, inducer diffusion model, and association model.
Test
We performed 200 iterations of simulation using the cellular automaton model, obtaining simulated results for cell death on the test paper.
Learn
The predictive results of our model reflected some shortcomings in the wet lab experiment design. We proposed some improvement solutions and demonstrated the feasibility of these solutions through model simulations, providing new options for the iteration of wet lab experiments.
