During our pursuit of creating a non-transmissible, modular DNA delivery system we designed a method to modify and amplify RP4 while simultaneously removing its origin of transfer. Our method consisted of a nearly entirely in-vitro pipeline, from beginning in a systematic process of diagnostic PCRs, gel imaging of large plasmids(see below), and ending in high fidelity homology-based end joining to assemble over 60 kb of large DNA fragments. All of our methods, primers, and reagents used are published in our experiments section as use for future teams, allowing significant ease of amplification and assembly of RP4.
Our primers that have been designed and optimized in multiple iterations are listed in our experiments section. Every primer (some with the use of additives and proper annealing temps, which are also listed), has been shown to produce amplicons with zero off target fragments. Furthermore, primers were created to be a modular set, allowing certain fragments to be ignored or incorporated, depending on the application, by selecting different sets. For example, using our “empty” set will allow a quicker assembly without including the modular golden gate site and kill switch and by removing the “oriT_remove” set the oriT can be left intact.
We have designed two additional cassettes for addition via our assembly, one with a golden gate compatible enzyme BaeI and another with a kill switch. Like the rest of our plasmid, certain primers can be used to include one, both, or neither.
Usage of RP4 as a method of transformation has shown to be extremely effective at DNA transfer between strains of E. coli. We’ve created a protocol for both fast and effective transformation between multiple plasmid carrying strains (up to three). We’ve also used it for movement of multiple plasmids multiple times into a selectable destination vector, allowing for far simpler cotransformation than conventional methods. An important note we discovered is that conjugation with RP4 is incredibly favorable in solid media vs liquid media. Our protocol is listed here
After a golden gate procedure combining the strong promoter PM4 with the GadE gene, the newly formed plasmid was transformed into E. coli Nissle. GadE, along with this promoter, strongly activates the expression of several acid resistance pathways in the Gad family, critical for creating acid fitness islands. After testing this new plasmid in E. coli Nissle, we found it to be beneficial to acid survival. This data can be used by future iGEM teams in efforts to increase the pH resistance in E. coli strains along with procedures for the assembly, transformation, and testing of GadE.
As part of the efforts in engineering an E. coli Nissle probiotic capable of colonizing the lower stomach line, we added the pBbB8k-csg-amylase plasmid into Nissle in order to upregulate curli amylase fibers thus promoting biofilm growth, stomach lining adhesion, and acid resistance. As outlined in the procedures, we successfully transformed pBbB8k-csg-amylase into E. coli Nissle creating an upregulation in curli fiber depression. Through both colorimetric plating and fluorescent liquid culture assays using the dye Congo Red, we provided procedures and data indicating these results. Future iGEM teams will be able to follow our procedures when using Curli fibers as a means of engineering biofilm constructs in E. coli.
Through our experiments in measuring biofilm growth on Congo Red plates, we have demonstrated for the first time to our knowledge, the expression of curli fibers and biofilm growth at 37°C rather than standard literature procedures growing at 30°C. In addition to curli expression on plates, we demonstrated that the addition of pBbB8k-csg-amylase increases curli fiber expression not only in stationary form, but in a liquid culture motile form.
Finally, we have demonstrated that curli fiber upregulation increases survivability in acidic liquid cultures, allowing for the survivability of E. coli Nissle in acidic environments with as low as pH 3.5. This finding demonstrates the potential for curli fibers in engineering the E. coli colonization.
We’ve successfully shown that Nissle 1917 can still express our specific targeting system despite its unique K5 capsid. Furthermore, we’ve shown that the tip of the neae is exposed and able to perform in binding using our anti-myc conjugated fluorophore assay. Alongside this, we optimized a yeast protocol for surface expression staining for E. coli. The protocol is listed here.
To increase functionalization and targeting breadth, we added BspQI binding sites flanking the neae (primers available in experiments), offering future teams the option to assemble on binders of their choice for surface expression.
We created a program to design protein binders without requiring advanced knowledge in protein engineering. From a single configuration file, EZBinder takes you from PDB code to working, scored binders in just one click. Our in silico results show better computational metrics than experimentally proven binders for the fluorescent protein mCherry with default settings and zero extra configuration. Check out our software page to learn more!
To image both RP4 and our fragments mid assembly, both our PI and online sources said we needed a pulsed field gel electrophoresis system, which is incredibly expensive. To fix this, we discovered a method of imaging the length of large plasmids (60 kb and under). Our protocol will allow teams who don’t have access to pulsed field gel electrophoresis to image large plasmids for a cheap cost. The protocol can be found here.
During the course of the summer, we had significant difficulties isolating RP4 from cells using a typical kit-based method. We have developed a method using a genomic extraction kit that does not extract any genomic DNA but does extract large plasmid DNA with high purity and low contamination. Furthermore, as a method available to teams with no access to more expensive genome extraction kits, we tested and successfully used an alkaline lysis protocol with fully in lab synthesized and mixed reagents.
We followed Chris Voigt’s protocol for our standard electroporation of E. coli Nissle 1917. We also discovered several ways to increase its efficiency(improving time constants). We have published a protocol for future iGEM teams to use, with notes on our suggestions for improvements. The protocol can be found here.
RP4 was highly successful at transforming into E. coli Nissle 1917. It was capable of moving both large plasmids and multiple small plasmids back and forth between Neb-10β, DH5-α, NEB turbo, and Nissle 1917. Since transformation efficiency of Nissle is already significantly lower than that of common cloning strains and based on our attempts, we believe cotransformation is incredibly difficult through standard methods. Using our conjugation protocol worked extremely well and generate colonies faster than standard transformation methods (~6 hours). The protocol can be found here.
Using colony stamping and antibiotic selection markers, we developed a method for easy selection of conjugation efficiency and colony counting. Our method creates replicates of each plate which allows for simple visualization and calculation of efficiency of conjugation using the formula η C = Y R G Δ t, where Y=transconjugants, R=recipients, G=donors, and Δ t=duration. Our protocol is listed here
We developed a method for generating a large scale library of variable expression and degradation genetic circuits for high throughput testing of optimal protein expression. By using a combinatorial golden gate assembly with interchangeable parts, we generated a library of 384 possible combinations of RBS of differing strengths, degradation tags of various rates with linkers, and different copy number backbones. This strategy is incredibly modular and can be used for expression of nearly any protein, serving as a method for high throughput screening of optimal expression and degradation rate of proteins designed for specific tasks. Throughout all of our work both in other areas of the project and in times outside of iGEM, we have discovered that a protein expressed at an unfavorable rate or a protein that lingers in the cytoplasm for too long can result in ineffectiveness or even cytotoxicity. We have more details and important notes located here which we hope will allow future iGEM teams to have higher throughput during expression optimization.
Positive selection markers tend to be viewed as unfavorable in comparison to negative selection markers due to their increased complexity, and difficulty in selection in many different scenarios. During the design of our conjugative assay we developed a method of using lack of antibiotic resistance as a selection marker via colony stamping. This is an extremely useful technique when combined with a multi-plasmid conjugation experiment. It allows easy identification of every single plasmid within each colony via replicate plating. You can see our protocol here.