iGEM 2023 — Team uoregon

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# Engineering

Engineering in biology can be summarized in four key steps:

## Design Phase:

Applying engineering principles to biology, with a focus on developing plans and using parts to create constructs with a desired function. Modeling and previous literature is used to make predictions, design experiments, and make data driven decisions about experimental direction.

## Build phase:

Biological parts, such as proteins, DNA, or RNA are acquired, manipulated, and assembled to create parts. In the context of synthetic biology, this often includes cloning, transformation, assemblies, PCR, and various diagnostic procedures such as blotting, imaging, or colony counting. This phase also includes implementation of the construct into a chassis or cell-free system

## Testing phase:

The constructed parts are tested in various assays for proper function, additional effects, and efficacy. Data is gathered during this phase.

## Learn phase:

The data gathered during the testing phase is gathered and analyzed. The results can feed back into the models into the design phase to improve accuracy, point out flaws during the build phase, or highlight errors during experimentation. This cycle is repeated, often until success is achieved and there exists no desire for improvement.

Our team went through multiple different design-build-test-learn cycles throughout the course of the summer. Each aspect of the project achieved at least one full iteration of the engineering cycle.

## Acid resistance and colonization:

Our colonization engineering efforts began with several key modifications backed by reviewing the literature of stomach bacterial conjugation. Along with an abundance of acid resistance pathways, we determined cell aggregation and production of biofilms to be key abilities in the potential for stomach colonization. The expression of curli amyloid fibers is well known to be a prerequisite for biofilm growth, activated during the stationary phase of bacterial growth. We began with the pBbB8k-csg-amylase, encoding for the csg gene family creating the protein complex required for the production of extracellular curli fibers.

Our first steps in wetlab procedures transformed this plasmid into *E. coli* Nissle, opening the door for biofilm and colonization experiments. Our first experimental steps involved colorimetric biofilm assays using the Amyloid binding dye Congo Red, protocol [here](%E2%80%9Dhttps%3A//2023.igem.wiki/uoregon/experiments%E2%80%9D) From these experiments, several key results guided the rest of our colonization efforts.

First, we showed that uninduced *E. coli* Nissle carrying the curli plasmid grew stronger biofilms than DH5α carrying the same plasmid. From these results, we concluded that *E. coli* Nissle, without the upregulation of csg genes, already holds potential for effective colonization on solid environments. Secondly, we discovered a significant decrease in colony growth as concentrations of the inducer arabinose increased. Although initially surprising, further review of the literature indicated excessive curli fiber expression exhibiting toxicity. With this in mind, our next steps moved towards identifying the ideal concentration of the arabinose inducer to upregulate curli expression without decreasing fitness. We did this by running two separate experiments with a range on arabinose concentrations, one in a liquid culture set up for measuring fluorescence reads of Congo Red bound to curli fibers and a second experiment on solid agar plates using qualitative observations of the color of colony growth through accumulation of Congo Red dye. These experiments indicated an ideal concentration between 0.01% and 0.001% arabinose, overexpressing curli fibers while resulting in minimal to no decrease in survival or biofilm production. Not only this, but additional compelling results from comparing these two experiments arose. On the solid agar plates, colonies in stationary phase for both induced and control groups exhibited little to no observable difference in Congo Red accumulation. In contrast, the fluorescent reads from the liquid phase cultures showed extremely significant contrast between these two groups. This indicated a greater curli expression difference during liquid phase than during the stationary phase, an attribute that could increase a cultures ability to thrive in low pH liquid environments such as stomach acid.

From these results, and our induction metrics established, we decided to move forward with survivability testing in low pH environments. As outlined in the procedures, a series of plate read data yielded interesting results. Without inducing Curli upregulation, we found *E. coli* Nissle to retain survival, albeit with limited growth, as low as a pH of 4. However, after testing the survival of *E. coli* Nissle induced for the curli producing plasmid, we found significant growth in pHs as low as 3.5.

Fascinated by these new findings, our next objectives sought to combine curli fiber upregulation with constitutive expression of the gadE regulating gene known to activate key acid resistance pathways.

Our acid resistance engineering cycle began with design of constructs for expression of GadE. We set up a triple assembly pipeline that would allow us to perform all necessary data collection while ordering only one gene fragment from twist. The pipeline was designed with the first assembly inserting the GadE gene into a expression vector in front of a Lac promoter to enable IPTG-inducible expression to quantify ideal induction amounts, then into a second vector for reduction in copy number to mimic the genome, and then a promoter swap with a strong, OrthoRep-designed constitutive promoter for high expression. Each assembly was designed using benchling with golden gate assembly standards. The first golden gate used BsaI to assemble the construct(ordered from Twist Biosciences) into a freegenes vector; we selected a medium copy plasmid from the distribution. However, we were plagued with constant assembly failure. We attempted multiple times but could not get a proper assembly. We tested with multiple other fragments with compatible overhangs and could not achieve success either. After consultation with other iGEM teams who also experienced faults in the distribution, we decided to move to assembly two, which worked on the first try with a BbsI golden gate. Once confirmed successful with correct length on gel electrophoresis, we moved onto assembly three, which involved adding in a strong, constitutive promoter we discovered from recently published literature. Using a PaqCI golden gate assembly, we successfully performed a promoter swap, removing the IPTG-inducible Lac promoter. Once completed, we transformed into Nissle 1917 and ran a plate assay testing survival in different pH ranges, with supplemented glutamate (since one of the major acid resistant pathways activated by gadE requires glutamate) AND with curli to compare the resistance offered by both. We also performed a cotransformation with both of them to see if they exhibited potentially synergistic effects. After the plate read was completed, we analyzed the data and determined that GadE and curli coexpressed exhibited a synergistic effect and allowed survival at a pH of 2.0 for long term. From this data, we learned that supplementation with excess glutamate was required for this acid resistant phenotype. We also learned that GadE in the absence of curli does not exhibit strong acid resistance. Our next data driven step is to test for ability to perform tasks such as conjugation or chemotaxis in high levels of acid with coexpression of curli and GadE.

## Chemotaxis:

We spent significant time researching the best possible molecular gradient to elicit a chemotactic response. After extensive consideration of our target pathogens, the environment of the stomach and gut, and our potential methods of detection, we settled on using a small molecule induction due to permeability of the membrane, specifically urea, which forms a gradient in the stomach that leads to sites of *H. pylori* colonization. This allows us to completely bypass any form of engineering or modification of receptors or receptor pathways, which, while incredibly interesting, poses a significant challenge. Our design bypasses this obstacle by using intracellular detection via a transcription factor, UreR, that binds to Urea and activates a promoter. This engineered circuit allows us to express proteins at different rates that correspond to urea concentration within the cytoplasm. Simultaneous to this design process, our computational members began designing a protein binder to EfliG, a small protein that serves the role as a “switch” in chemotaxis flagellar motor rotation. Using mutants of the protein found in literature, a set of binders were designed to inhibit FliG and act as an inducer for tumbling in chemotaxis, which biases movement and inhibits straight movement. Our strategy was to combine this with our urea inducible expression circuit and induce tumbling chemotaxis in the presence of urea, which would bias motion of the bacteria to move down a urea gradient and towards sites of *H. pylori* colonies.

This was quite a lofty goal due to the precise regulation required for correct communication between urea detection and flagellar motion, so we decided to pursue a highly combinatorial, mass screening approach for expression and degradation optimization. We designed a library generating Golden Gate assembly that synthesized a massive number of combinations of RBS of differing strengths, different FliG binders designed in-silico by our computational team, different N-term and C-term degradation tags, and different copy number backbones. The total number of combinations successfully created was 384 via plate scraping after the assembly transformation. All variants were then cloned into Nissle and screened on our custom designed chemotaxis assay plate, which consisted of a absorbent paper soaked in urea solution embedded in soft agar to form a gradient as it diffused through the liquid. We then streaked our transformation in parallel lines and grew overnight to observe growth patterns. Unfortunately, we could not observe any patterns that suggested chemotactic mobility. After analysis of our construct design and testing method, we believe there were limitations to our assay method. We had issues with horizontal growth, and our plan is to add lanes to divide cells up. Furthermore, our current set up includes a rectangular plate, but we have ideas for using circular plates with the soaked urea paper plated in the middle or on the sides. Altering urea concentration or gel percentage is also of interest to us, to determine whether those variables could be responsible for our lack of success. Finally, using a reporter to determine the extent of variability or even functional expression of our FliG binder would be a useful diagnostic.

## Conjugation:

Our engineering cycle for this sector of the project was by far the most extensive in comparison to acid resistance and chemotaxis. The large nature of RP4 posed significant difficulties at almost every single phase and experimentation step as most common kits and protocols fail to account for large plasmids. Every single step needed to be planned out ahead of time, even from the smallest transformation to gel electrophoresis to modification. We sought out new protocols, bought new ladders, and researched nuances of procedures to produce the highest accuracy during each procedure. Due to transformation difficulties, we sought to take a unique approach involving complete in-vitro replication and modification. This included three rounds of primer design, with the first round not meeting qualifications for purchase, the second round getting purchased but resulting in significant failed amplification and extensive off target effects, and the third finally achieving success. During the first two rounds, fragments were selected based on gene similarity, i.e. keeping all genes involving the T4SS apparatus on the same segment and all of the partitioning proteins together, but flanking regions were not optimal for primer annealing. Primers were designed using genetic design tools (benchling and snapgene) automatic primer generation and analyzed and improved by hand. Important note is that these primers included extension tails (regions that didn’t anneal to the template). Primers were ordered and arrived, but did not work well, resulting in 1/14 perfect PCRs (no off targets, high yield of target amplicon). Primers were redesigned 100% manually, targeting ideal annealing sites throughout the plasmid. All amplification was done to create homologous ends between each fragment, and primer extensions were minimized to only four fragments and had optimal, long annealing regions with GC clamping. All primers were screened via alignments to check for off target annealing. All primers were further analyzed using IDT’s oligoanalyzer for hairpin/secondary structure formation, homodimer, and heterodimer, with a maximum ΔG of -4 Kcal/mol-1 for annealing regions. Once all primers met these qualifications they were ordered and tested. Optimization of these primers included multiple gradient and touchdown PCR methods that spanned a variety of temperatures, times, cycle numbers, and additives. Each reaction was screened with an agarose gel at 0.7% agarose. Additives tested were ETSSB, betaine, and 1,2-propanediol. Touchdown PCR was performed to one degree below the calculated Q5 annealing temp using NEB ™ calculator. Gradient PCR was performed from 61°C to 72°C. After analyzing results and getting successful yields and limited off target replication, ideal cycling conditions were recorded and repeated to generate higher yield of fragments. Fragments were then assembled using NEB HIFI assembly, which yielded successful lengths on gel electrophoresis. However, the yield from reactions have been far too small to allow any downstream applications such as sequencing or even transformation, which is difficult with such a large construct. We need to run more assemblies and combine them to reach a yield significant for transformation or sequencing.

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# RP4 amplification

For our complete in-vitro replication of GadE, we went through many rounds of primer optimization and testing. We ordered an initial round of primers that were found to have off target effects and poor yield. To improve, we redesigned completely by hand and performed alignments with each primer against RP4 to check for off target annealing. We scanned each primer for disadvantageous secondary structures or dimerization, and once every primer passed these tests it was ordered. Once primers arrived we ran both touchdown PCRs and large scale gradient PCRs for each amplification as well as a variety of different PCR additives such as ETSSB, betaine, and 1,2 propanediol. Through combinatorial testing, we were able to get fragments for all 11 parts of the RP4 assembly with extremely minimal off targets and high yields. We then performed NEB HIFI assembly on the fragments, confirming that size was not an interference with the homology-based assembly.

We have uploaded [the primers](https://static.igem.wiki/teams/4598/wiki/primers.csv) (BBa\_K4598013-BBa\_K4598036) that successfully amplify RP4 with minimal off-targets and quality yield. Our iGEM parts database list cycling conditions for each part, as well as additive amounts that worked well.

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# Conjugation Procedure

Before beginning acid survivability experiments, combining the curli and gadE upregulating plasmids was necessary. However, standard cotransformations utilizing electroporation yielded no success due to the constraints and inefficiencies associated with *E. coli* Nissle. Alternatively, we chose to proceed with a conjugative approach to combine these vectors. Starting with three separate overnight cultures of EcN + Curli, EcN + gadE, and EcN + RP4 (a conjugative plasmid). Each of these cultures was washed of antibiotics in PBS. After diluting each sample to an OD of 0.5, 1 mL of each was combined and set into a 37\*C shaker for 1 hour. As the gadE plasmid contained an oriT site allowing for conjugative transport via RP4, all combinations of the three plasmids should have occurred in the joint culture. We then plated on the antibiotic resistances associated with the gadE and curli plasmids (CAM and KAN respectively) to select for the target combinations.

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# Positive selection marker-stamping:

Immediately following the Conjugation Procedure listed above with desired plasmids ( One must contain an oriT site and one must be stationary) along with the RP$ conjugative plasmid, plate 1000x or greater dilutions on antibiotic free plates. Once single colonies form, use a felt colonies stamp to lightly stamp antibiotic free plates. This stamp is then used as a template to lightly stamp plates of every possible combination of antibiotic resistances found in desired plasmids. Recording original colony numbers and transferred colony numbers will provide data on conjugative efficiencies of each plasmid in the system.

![Diagram of positive selection marker-stamping](https://static.igem.wiki/teams/4598/wiki/pos-markerstamp.png)

Diagram of positive selection marker-stamping

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# Chemotaxis Assembly: 1. 0.12 pmol each fragment 2. 0.06 pmol backbone 3. 0.75 uL BsaI 4. 0.25 uL T4 DNA ligase 5. 3.5 uL T4 ligase buffer 6. 1 uL ATP 7. H20 to 25 uL 8. Cycling: 37C, 5 mins→16C, 5mins: 110 cycles --- Chemotaxis Assay: After completing combinatorial assembly of chemotaxic binders, all combinations in a low concentration sample were transformed via electroporation into neb10b. 50 mL samples of the newly transformed culture were then mira prepped to produce adequate amounts and concentrations of the chemotaxis binder library. This high concentration sample was then transformed into Nissle using the Nissle Electroporation Procedure previously outlined. Outgrowth was immediately plated onto 0.3 % agar gel with kanamycin resistance parallel to a previously established urea gradient starting at 20 mM concentration along with control Nissle. Data will be collected by analyzing growth patterns either selectively towards or away from the gradient. --- Large Size Gel electrophoresisReagents 1. 0.3% Agarose 2. 400mL TAE 3. 10 000x Sybr gold or other post stain compatible gel 4. 10ul Loading dye per sample 5. ~50ng Large DNA 6. 1:1 v:v H20 to DNA Pour extremely thick, low percent gel, 0.3% is suitable for most plasmids from 40-60 kb. We recommend aiming for an 80 mL gel if using a standard mini casting tray. Thicker gels can run higher voltages without melting. Since our gel is such a low percentage, it will melt incredibly easily. The gel should also be run in a cold room, which will further reduce melting risk. It is highly unfavorable to run a large gel, as they can develop heating inequality across the gel. The gel should be boiled thoroughly, as underboiling will severely harm visibility and cause smearing in the gel. Visibility is an incredibly difficult variable to maintain here, so every precaution must be taken. Post stain is highly advisable as well, as the presence of intercalators in a precast gel can cause DNA to run differently. Furthermore, post stains like SYBR gold are far more sensitive than ethidium bromide, GelRed, and SYBR safe, allowing less DNA to be run. Lastly, large plasmids can easily cause smearing if too much DNA is loaded, so be conservative. Gel voltage for mini-gel: 35 volts for ~5 hours. Ladder recommended is MidRange PFG marker from NEB or an alternative of similar size. --- Chemotaxis Assay: After completing combinatorial assembly of chemotaxic binders, all combinations in a low concentration sample were transformed via electroporation into neb10b. 50 mL samples of the newly transformed culture were then mira prepped to produce adequate amounts and concentrations of the chemotaxis binder library. This high concentration sample was then transformed into Nissle using the Nissle Electroporation Procedure previously outlined. Outgrowth was immediately plated onto 0.3 % agar gel with kanamycin resistance parallel to a previously established urea gradient starting at 20 mM concentration along with control Nissle. Data will be collected by analyzing growth patterns either selectively towards or away from the gradient. --- gadE and csgXCongo Red Plating Assays Arabinose Induction: Following the Congo Red and YESCA media Plating Protocol in Protocols, plates were made both with and without arabinose concentrations of 0.1%. These plates were used to run the following three experiments: The first comprised of two plates (one arabinose induced and one control) grown in 37\*C and two plates (one arabinose induced and one control) grown in 30\*C. On these plates, E. coli Nissle carrying the curli plasmid was plated in three dilutions of 1x, 10x, and 100x starting at an OD of 0.6. The goal of this experiment was to obtain qualitative data on colony size and color indicative of growth rates and accumulation of Congo Red Dye via binding to curli amyloid fibers. Images and results presented in the results section. Congo REd Plating Assay Nissle vs DH5α: Following the Congo Red and YESCA media Plating Protocol in Protocols, plates were made and used to run the following three experiments: The first comprised of two plates (one arabinose induced and one control) grown in 37\*C and two plates (one arabinose induced and one control) grown in 30\*C. On these plates, three dilutions of *E. coli* Nissle without the curli plasmid were added at dilutions of 1x, 10x, and 100x starting at an OD of 0.6. The goal of this experiment was to obtain qualitative data on colony size and color indicative of growth rates and accumulation of Congo Red Dye via binding to curli amyloid fibers. Images and results presented in the results section. --- Acid Survivability:Curli Three iterations of acid survivability experiments were conducted through plate read growth measurements. These experiments utilized m9 aliquots of pH spanning every half pH value from a pH of 7 to a pH of 1. The first two experiments focused on the curli plasmid expression using pH ranges of pH 7, pH 4, pH 3, pH2, pH1 and pH 4, pH 3.5, pH 3, pH 2.5, pH 2 respectively. These experiments began with cultures of *E. coli* Nissle carrying the curli plasmid and E. coli Nissle cultures without the plasmid. Both these cultures inoculated two new cultures induced with 0.0015% arabinose and 0% arabinose respectively. All four cultures grew for exactly 12 hours at which point they were each divided into 96-well plates in triplicates for each pH value. All controls were added alongside. Plates were then entered into the plate reader, a set for 24 hour procedures with OD:600 reads spaced every three minutes followed by shaking. --- Curli + gadE Conjugation Procedure Before beginning acid survivability experiments, combining the curli and gadE upregulating plasmids was necessary. However, standard cotransformations utilizing electroporation yielded no success due to the constraints and inefficiencies associated with E. coli Nissle. Alternatively, we chose to proceed with a conjugative approach to combine these vectors. Starting with three separate overnight cultures of EcN + Curli, EcN + gadE, and EcN + RP4 (a conjugative plasmid). Each of these cultures was washed of antibiotics in PBS. After diluting each sample to an OD of 0.5, 1 mL of each was combined and set into a 37\*C shaker for 1 hour. As the gadE plasmid contained an oriT site allowing for conjugative transport via RP4, all combinations of the three plasmids should have occurred in the joint culture. We then plated on the antibiotic resistances associated with the gadE and curli plasmids (CAM and KAN respectively) to select for the target combinations. The third iteration sought to examine the combined effects on acid survivability of upregulated gadE and Curli fiber expression. Following the same acidic media procedure as the previous plate read experiments using pHs of 4, 3.5, 3, 2.5, and 2, we plated the following combinations of plasmids and inducers in this configuration as shown below. Each plasmid-inducer combo along with controls were plated into a 384 well plate and set for a 24 hour procedure with OD:600 reads spaced every three minutes followed by shaking. --- Step-by-step Miraprep protocol. Protocol Title: Miracle-prep (Miraprep) PROTOCOL FOR: Plasmid DNA isolation Manuscript Title: The Miraprep: A protocol that uses a Miniprep kit and provides Maxiprep yields Reagents * GeneJet Plasmid Miniprep Kit (Thermo Scientific), or * QIAprep Spin Miniprep Kit (Qiagen) Procedure 1. Set up a 50 ml bacterial culture in appropriate selective media and incubate on a shaker (250 rpm) at 37°C overnight. 2. Transfer bacteria to a 50 ml tube and spin at 4000xg at 4°C for 10 min. 3. Discard supernatant and resuspend pellet in 2 ml resuspension buffer with 50 ug/ml RNase freshly added. 4. Add 2 ml of lysis buffer, invert 3-4 times and incubate for 3 min at RT. 5. Add 2 ml of neutralization buffer and invert 3-4 times. 6. Distribute lysate into 1.5 ml Eppendorf tubes (~4 tubes) by pouring, not pipetting. 7. Spin at 13,200xg at RT for 10 min. 8. Collect supernatants in a 15 ml tube and discard pellets. 9. Add 1x volume of 96% ethanol (~5 ml). 10. Mix thoroughly for 5 sec. 11. Load the sample-ethanol mix onto 5 spin-columns in three sequential ~700 µl aliquots--after the addition of each aliquot, spin the column 30 sec at 13,200xg. 12. Discard flow-through. 13. Repeat step 11 until the entire sample is run through the spin-columns. 14. Wash with 500µl washing buffer and spin at 13,200xg at RT for 30 sec. 15. Discard flow-through. 16. Repeat step 14-15. 17. Do a final spin at 13,200xg at RT for 1.5 min. 18. Discard the old tube and put the column onto a new tube. 19. Add 30-35μl of dH2O and incubate for 2 min at RT. 20. Spin at 13,200xg for 2 min to elute the DNA from column. 21. Combine the eluted DNA from all 5 columns in one tube (~175μl). 22. Measure DNA concentration. 23. Store samples at -20°C. --- Golden Gate Assembly: Updated: November 18, 2020 Materials * T4 DNA Ligase ([NEB #M0202](https://www.neb.com/products/m0202-t4-dna-ligase)) and Buffer * BsaI-HFv2 ([NEB #R3733](https://www.neb.com/products/r3733-bsai-hf-v2)) Notes: Super important. Do not freeze & thaw the large tube of T4 Ligase Buffer 10X. This has ATP which is sensitive and all ligations will start to fail. This large tube should be aliquoted into many small PCR tubes with ~8µL of buffer each. When doing a reaction only take one small PCR tube out. Pmol calculator(REMEMBER IT ASKS A VALUE IN MICROGRAMS, YOU MUST CONVERT FROM NANOGRAMS) <https://www.promega.com/resources/tools/biomath/> Calculate picomoles in one uL, then determine how many microliters to add to meet the below ratios | Reagent | Concentration | Amount (µL) | | --- | --- | --- | | Vector Backbone | 0.06 pmol | | insert | 0.12 pmol | | T4 ligase buffer | 10x | 2.5 | | T4 ligase | 2000 units/uL | 0.25 | | BsaI | 20 units/uL | 0.75 | | H20 | Add water to 25uL total | - | 1. Set up 25 µl assembly reactions as follows 2. Mix gently by pipetting up and down 4 times. 3. Briefly microcentrifuge (1 sec.) to bring material to the bottom of tube. 4. Transfer to thermocycler and program as follows: 1. 37°C for 5 min 2. 16°C for 5 min 3. go to step 1, 30X 4. 80°C for 20 min (inactivate both enzymes) 5. 12°C forever 6. Clean up reaction with a typical PCR cleanup kit(ELUTE WITH LESS elution buffer than normal, there will not be much DNA in the reaction) 7. Transform into cells and hope for the best 8. You may need to drop dialyze Low-salt LB medium 1. Deionized H2O, to 950 ml 2. Tryptone, 10 g 3. NaCl, 1g 4. Yeast extract, 5 g 5. To prepare low-salt LB, combine the above reagents and shake until the solutes have dissolved. Adjust the pH to 8.0 with 5 N NaOH. Adjust the volume of the solution to 1 l with deionized H2O. --- NISSLE Transformation 1. Grow overnight cultures of nissle-1917 in culture media (LB or TB) with respective antibiotic at correct temperature 2. Dilute cells 50-fold into 25 mL low salt LB media and grow until OD=0.55-0.6 3. Aliquot into 50 mL falcon tubes or tube of equivalent size 4. Centrifuge at 4,000 xg at 4C for 10 minutes 5. Discard supernatant 6. Resuspend in 5 mL ice cold glycerol --- Gel Electrophoresis: THIS IS FOR 1% GEL, YOU MAY NEED TO ADJUST DEPENDING ON SITUATION Materials 1. PCR samples 2. 240 mg agarose 3. 30 mL TAEr 4. 3 μL SYBR Safe 5. 5 μL dye for each ladder and sample Procedure 1. Combine 300 mg agarose and 30 mL TAE 2. Microwave solution in 20 seconds increments until agarose is fully dissolved, making sure to avoid boiling (a lower intensity setting can help with this) 3. Add 3 μL SYBR Safe into solution 4. Insert comb into tray 5. Cast gel into tray, gel should be less than 1 cm thick 6. Cover gel with tinfoil, wait 10–15 minutes to harden 7. Add 5 μL of dye to each 5 μL DNA sample 8. Prep ladders (1 kb and 100 bp) (4 μL H20, 1 μL ladder, 5 μL dye) 9. Transfer the full 10 μL volume of each dyed dna sample into a lane (if possible, avoid using the edge lanes and adjacent lanes) 10. Run gels at 100V 11. Weigh eppendorf tube and write empty mass on tube 12. Use razor blade to precisely cut out the band of the sample fragment 13. Store in marked eppendorf tube, & weigh again. Record full mass on tube as well. 14. Store in -20℃ overnight --- Mini-Prep: Very nice instructions for this on the instructions card, and everything is labeled. Essentially consists of spinning liquid with a centrifuge, adding something, then removing supernatant or sediment. Materials 1. 4 volumes of ≥ 95% ethanol per volume of Monarch Plasmid Wash Buffer 2 2. Culture to be miniprepped Procedure 1. Pellet 1–5 ml bacterial culture (not to exceed 15 OD units) by centrifugation(16000 rcf) for 30 seconds. Discard supernatant, or liquid, and keep the sediment. 2. Resuspend pellet in 200 μl Plasmid Resuspension Buffer (B1) (pink). Vortex or pipet to ensure cells are completely resuspended. There should be no visible clumps. 3. Lyse cells by adding 200 μl Plasmid Lysis Buffer (B2) (blue/green). Invert tube immediately and gently 5–6 times until color changes to dark pink and the solution is clear and viscous. Do not vortex! Incubate for one minute. 4. Neutralize the lysate by adding 400 μl of Plasmid Neutralization Buffer (B3) (yellow).(LOCATED IN FRIDGE, MAKE SURE TO KEEP REFRIGERATED) Gently invert tube until color is uniformly yellow and a precipitate forms. Do not vortex! Incubate for 2 minutes. 5. Clarify the lysate by spinning for 2–5 minutes at 16,000 RCF. 6. Carefully transfer supernatant to the spin column and centrifuge for 1 minute. Discard flow-through. 7. Re-insert column in the collection tube and add 200 μl of Plasmid Wash Buffer 1. Plasmid Wash Buffer 1 removes RNA, protein and endotoxin. (Add a 5 minute incubation step before centrifugation if the DNA will be used in transfection.) Centrifuge for 1 minute. Discarding the flow-through is optional. 8. Add 400 μl of Plasmid Wash Buffer 2 and centrifuge for 1 minute. 9. Transfer column to a clean 1.5 ml microfuge tube. Use care to ensure that the tip of the column has not come into contact with the flow-through. If there is any doubt, re-spin the column for 1 minute before inserting it into the clean microfuge tube. 10. Add ≥ 30 μl DNA Elution Buffer to the center of the matrix. Wait for 1 minute, then spin for 1 minute to elute DNA. --- Inoculation: 1. Circle colony and label it with a number/letter 2. Transfer 5 ml of media into a test tube 3. If media(LB) does not have an antibiotic, you must add 5 uL of your desired antibiotic(located in freezer in middle slider on the right) 4. Use a pipette tip to very gently tap on selected colony 5. Eject pipette tip into test tube with media and antibiotic 6. Re-parafilm plate and put back in fridge 7. Place tubes in 37°C shaking incubator for 16 hours, then take out and store in fridge --- Antibody Staining of pNeae for Plate Reader 1. Prepare 4 tubes PER induction amount. I recommend running at least three different induction levels. All steps should be performed in as little light as possible and in a cold room. Perform all centrifugation at 4C. Tubes per induction level: cells only, primary only, secondary only, primary and secondary together * Note: Ideal tubes are 1.5 mL microcentrifuge tubes * Note 2: primary is mouse α c-myc antibody (9E10), secondary is goat α mouse-AF488 fluorescent label. 2. Perform each step for each tube unless otherwise noted 3. Pellet 10^8 cells after induction(common for yeast is 10^6, but creates extremely weak signal if used for bacteria). Remove supernatant and resuspend in 0.5-1mL PBSA. * Note: Pellet by centrifuging at 12,000g for 30 s 4. Pellet cells again. Remove supernatant and resuspend in 50 μL PBSA * Keep on ice away from light 5. For primary only and primary and secondary together tubes: * Add 1.25 μL mouse α c-myc antibody. * Incubate (stir or agitate) at 4°C for 45 mins(need longer time or incredibly weak fluorescence (up to two hours). * Add 0.5-1mL PBSA, and pellet. Remove supernatant and resuspend in 50 μL PBSA * For secondary only and primary and secondary together tubes: * Add 0.625 μL 647 if using the plate reader. * Incubate at 4°C for 30 mins (will remain stable for up to two hours). * Add 0.5-1mL PBSA, and pellet. Remove supernatant * Repat. Resuspend in 50 μL PBSA * Fill each tube with PBSA to amount for plate reader, 96 well is recommended to use 200 uL --- Electroporation for Nissle: 1. Fill bucket with ice 2. Gather electrocompetent Nissle cells from –80 freezer(one with green handle) 3. Place Cuvette into the -80\*C freezer for ~10 minutes 4. Gather DNA prepared from 50 mL cultures through the Mira Prep Procedure. Concentrations should be >100 ng/uL 5. Combine 1-2 µL of your resuspended DNA with 100 µL of electrocompetent Nissle cells and place on ice 6. LABEL TUBE 7. Place plates with correct antibiotic in incubator so it's warm when you use it later 8. Mix gently and transfer to electroporation cuvette (between metal plates), place in cuvette holder and hit “pulse” 9. Immediately add transformation(WARMED) media(975 uL) (SOB) to the cuvette and transfer recovered cells to a test tube 10. Leave to shake in the shaking incubator for 30-60 minutes 11. Remove test tubes from incubator and rest on a tube rack 12. Label plates to correspond to the tubes 13. Transfer 100 µL of your transformation to its respective plate(100x and 1000x dilutions are best, so mix 1uL of transformant with 99 uL of sob for 100x, 0.1 uL of transformant with 99.9 uL of SOB for 1000x dilution), pour 10–15 glass beads onto the plate, close the lid, and shake the plate to cover all the afar with the cells (not the lid, just side to side shaking)(MAKE SURE PLATE IS NOT UPSIDE DOWN, SMALLER LID SHOULD BE ON THE BOTTOM) 14. Pour beads into a used beads bin 15. Replace lid on plate and place in the incubator upside-down (agar-side up) 16. Leave plates in the incubator for 12–16 hours 17. Time is specific, take out and parafilm the plates so they are sealed and place in fridge with the clear doors 18. Bleach tubes after plating cells --- Electroporation (Standard): 1. Fill bucket with ice 2. Gather cells from –80 freezer(one with green handle) 3. Gather DNA from distribution(see other protocol)(NOTE: DISTRIBUTION PLATES DO NOT NEED TO BE KEPT ONE ICE FOR THE COURSE OF THE PROCEDURE. DNA is pretty stable at room temp, but for long term storage it should go back in the -20 4. Combine 1 µL of your resuspended DNA with 25 µL of your cells and place on ice 5. LABEL TUBE 6. Place plates with correct antibiotic in incubator so it's warm when you use it later 7. Mix gently and transfer to electroporation cuvette (between metal plates), place in cuvette holder and hit “pulse” 8. Immediately add transformation(WARMED) media(975 uL) (SOB) to the cuvette and transfer recovered cells to a test tube 9. Leave to shake in the shaking incubator for 30-60 minutes 10. Remove test tubes from incubator and rest on a tube rack 11. Label plates to correspond to the tubes 12. Transfer 100 µL of your transformation to its respective plate(100x and 1000x dilutions are best, so mix 1uL of transformant with 99 uL of sob for 100x, 0.1 uL of transformant with 99.9 uL of SOB for 1000x dilution), pour 10–15 glass beads onto the plate, close the lid, and shake the plate to cover all the afar with the cells (not the lid, just side to side shaking)(MAKE SURE PLATE IS NOT UPSIDE DOWN, SMALLER LID SHOULD BE ON THE BOTTOM) 13. Pour beads into a used beads bin 14. Replace lid on plate and place in the incubator upside-down (agar-side up) 15. Leave plates in the incubator for 12–16 hours 16. Time is specific, take out and parafilm the plates so they are sealed and place in fridge with the clear doors 17. Bleach tubes after plating cells --- Transformation From Distribution: 1. Check [igem distribution airtable](%E2%80%9Dhttps%3A//airtable.com/shrUtW6RwzMabg8d1/tbls8zhnooNme6Eyg%E2%80%9D) and select target DNA you want more of 2. With a pipette tip, punch a hole through the foil cover into the well of the part that you want. Make sure you have properly oriented the plate. Do not remove the foil cover, as it could lead to cross-contamination between the wells. 3. Pipette 10 µL of dH2O (distilled water) into the well. Pipette up and down a few times and let sit for 5 minutes to make sure the dried DNA is fully resuspended. The resuspension will be red, as the dried DNA has cresol red dye. 4. Transform 1 µL of the resuspended DNA into your desired competent cells(use NEB 10b for training)(see other protocol for transformation), plate your transformation with the appropriate antibiotic(shown in igem distribution airtable), and grow overnight. 5. Pick a single colony and inoculate broth (again, with the correct antibiotic); grow for 16 hours. 6. Use the resulting culture to miniprep the DNA AND make your own glycerol stock. We recommend using the miniprepped DNA to run QC tests, such as restriction digests and sequencing. --- Plating:Materials needed: * Clean 2L plastic beaker with handle OR clean 1L or 2L glass bottle * BD Difco LB Broth, Miller (white plastic container with blue top) * BD Bacto Agar * Plastic petri dishes (100mm normal size, or 150mm large size, or 60mm mini size) * Magnetic stir bar * IR thermometer * Antibiotic 1000X solution Procedure: ([https://legacy.bd.com/europe/regulatory/Assets/IFU/Difco\_BBL/244620.pdf)](%E2%80%9Dhttps%3A//legacy.bd.com/europe/regulatory/Assets/IFU/Difco_BBL/244620.pdf%E2%80%9D) 1. Put a large magnetic stir bar into a: clean 2L plastic beaker with handle OR a 1L or 2L glass bottle. 2. Measure out 25g of LB Broth per Liter (50g for 2L) in a plastic tray on the scale. 3. Tare the scale again and measure out 15g of Bacto Agar per Liter. 4. Put the powder into the beaker or bottle and fill it with DI water to 1L or 2L. 5. If using beaker, place on cold hotplate and using magnetic stirring to dissolve powder. If using bottles, shake very well or use magnetic stirrer. 6. If using beaker, place aluminum foil over the entire top. If using bottles, place aluminum foil around the top cap such that you can take the top cap off and put in on the bench and the bottom of the cap will not touch the table, only the aluminum foil will. 7. Put autoclave tape on the aluminum foil. 8. Label contents and date on autoclave tape. 9. When putting top on bottle it should be loose, otherwise bottle may explode in the autoclave.(put tape over the lid so it doesn't fall off 10. Place the beaker or bottle(s) into an autoclave tray. 11. Autoclave the bottle(s). The duration depends on volume. Typically 1L needs 30min at 121C (not including ramp time). Larger volume needs longer time. You need to account for the fact that large volumes need longer time for entire volume to reach temperature because of the high specific capacity of water. Note that certain autoclave settings may cause the contents to boil over. 12. If using bottles, while waiting for autoclave prepare a 60C water bath. No need if using beaker(not really necessary) 13. Take tray out of autoclave (HOT! use gloves) 14. Make sure autoclave tape has dark stripes on it. If not something went wrong. 15. (you don't need to use a water bath, can just leave it out on benchtop)If using bottle place it into hot water bath, submerging about 75%. Don't let dirty bath water get near mouth of the bottle. Don't place the bottle into the water bath immediately after taking it out of the autoclave since it will still be very hot and can actually warm up the water bath. Wait for it to cool a bit ~65C. If using beaker place on cold hotplate, very low stirring ~40rpm (make sure stirring is not making bubbles). 16. Use the IR thermometer to check the temperature of the bottle or beaker. 17. Wait for the media to cool down to at least 55C to 60C before adding antibiotic. Using 1000X antibiotic add 1mL of antibiotic per Liter of media(if ur making 2L of media, add 2mL). When adding antibiotic make sure you use aseptic technique. So either do this near a flame or in a BSC(the hood in bacterial room). 18. After adding antibiotic magnetic stir for 1 min (<80rpm so no bubbles) after to make sure antibiotic is mixed in well. 19. Get your plates and start pouring the media into them, opening one at a time. Pour as little as you can. I usually pour not enough to cover entire petri dish then gently tilt dish so that media covers everything. Avoid bubbles! For regular 100mm plates I usually pour in stacks of 5 plates. If you stack plates it prevents water condensation on the lids. 20. Wait until plates have solidified and turn them upside down. 21. Let the plates dry at least 2 hr before using. Fresh plates can be left at RT for a few days or placed into plastic bags (these come with petri dishes) and put into the cold room or fridge. --- DPNI digestion of PCR products: 1. Add 1 uL of DPNI restriction enzyme to finished PCR reactions(50 uL) 2. Incubate at 37 for 30 minutes 3. Incubate at 80 for 20 minutes for inactivation --- NEB Hifi Assembly | Reagent | Length | Pmol/uL | Total pmol | Assembly amounts | Positive control amounts | | --- | --- | --- | --- | --- | --- | | Insert 1 | 2217 | 0.071 pmol | 0.0 | 0.7 uL | | | Insert 2 | 4351 | 0.021 pmol | 0.0525 | 2.5 uL | | | Insert 3 | 4811 | 0.015 pmol | 0.0495 | 3.3 uL | | | Fragment total | | | 0.03-0.3 for <3 fragments, 0.2-0.5 for >3 fragments | Less than ~20% of RXN if using straight PCR products | | | NEBuilder MM | | | | 10 uL | 10 uL | --- Curli Procedures:Transformation of pBbB8k-csg-amylase into E. coli Nissle 1917 through Electroporation:Preparation of Competent Cells: 1. A single colony of E. coli Nissle 1917 was picked from a fresh LB plate and inoculated into 5 ml of LB broth, which was incubated overnight at 37°C with shaking at 200 rpm. 2. The overnight culture was diluted 1:100 into fresh 50 ml LB medium and allowed to grow until it reached an OD600 of 0.5-0.6. 3. The cells were then chilled on ice for 20 minutes. 4. Cells were harvested by centrifugation at 4000 x g for 15 minutes at 4°C. 5. The pellet was washed three times with ice-cold, sterile 10% glycerol. 6. Finally, the cells were resuspended in 1 ml of 10% glycerol. Electroporation: 1. 50 μl of competent Nissle 1917 cells were mixed with 1-2 μl of pBbB8k-csg-amylase plasmid (previously purified). 2. The mixture was transferred to a pre-chilled electroporation cuvette. 3. Cells were pulsed using an electroporator set at 2.5 kV, 25 μF capacitance, and 200 Ω resistance.- Immediately after the pulse, 1 ml of LB broth was added to the cuvette. - Cells were recovered by incubating at 37°C with shaking for 1 hour. - Transformants were selected by spreading the recovered cells on LB agar plates containing the appropriate antibiotic (depending on the resistance gene on the pBbB8k-csg-amylase plasmid) and incubating overnight at 37°C. Induction with Arabinose: 1. Individual colonies were picked and grown in LB broth with the appropriate antibiotic till mid-log phase. 2. The culture was then induced with a final concentration of 0.001% arabinose and incubated for an additional 3-5 hours for curli fiber production. Curli Fiber Staining on Agar Plates using Congo Red Dye: 1. A 1% solution of Congo Red dye was prepared and filter-sterilized. 2. This dye solution was added to molten LB agar (cooled to around 50°C) to achieve a final concentration of 0.004%. 3. The agar was poured onto plates and allowed to solidify. 4. Transformed Nissle colonies were streaked onto the Congo Red-containing plates and incubated at 30°C for 48 hours. 5. Colonies producing curli fibers will take up the Congo Red dye, appearing red or pink on the plate. Curli Fiber Staining in Liquid Culture and Fluorescence Measurement: 1. Cultures induced with arabinose were grown to the desired time-point, and then 1 ml of the culture was centrifuged to pellet the bacteria. 2. The pellet was resuspended in 1 ml of a 0.004% Congo Red solution and incubated for 30 minutes at room temperature with gentle shaking. 3. The cells were washed three times with phosphate-buffered saline (PBS) to remove any unbound dye.- The stained cells were resuspended in 200 μl PBS. - The fluorescence of the stained cells was measured using a plate reader with excitation at 497 nm and emission at 614 nm. The relative fluorescence units (RFUs) were recorded for each sample. --- [Yesca Congo Red Agar Plates:](https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5324733/#:~:text=YESCA%20agar%20plates:%20YESCA%20agar,to%20make%20normal%20YESCA%20plates) **YESCA media**: YESCA agar plates: YESCA agar is used to induce curli production. Dissolve 1 g/L yeast extract, 10 g/L casamino acids, and 20 g/L agar in water. Pour 25 mL per Petri dish to make normal YESCA plates. For thin YESCA plates used in plug western blots, pour only 15 mL per Petri dish. **0.1% Arabinose:** 1mL of Stock (15% dilute Arabinose) in 150 mL of media **Congo Red**: 4uL Congo Red Stock solution per 1mL of media Quantification of CR fluorescence for bacteria prestained on YESCA CR plates. 1. Spot 4 μL of overnight bacterial culture on the YESCA CR plate. Incubate the plate at 26°C for 48 h. 2. Recover bacteria from the YESCA CR plate. Wash them in]==[ 50 mM KPi by centrifugation at 12,000 rpm for 1.5 min and resuspend them in 1 mL 50 mM KPi. Adjust the cell density to 1OD600/mL. 3. Load 100 μL bacteria suspensions onto a 96-well plate reader. Measure the CR fluorescence with a Spectra max M2 plate reader as described above. --- Conjugation Procedure Before beginning acid survivability experiments, combining the curli and gadE upregulating plasmids was necessary. However, standard cotransformations utilizing electroporation yielded no success due to the constraints and inefficiencies associated with *E. coli* Nissle. Alternatively, we chose to proceed with a conjugative approach to combine these vectors. Starting with three separate overnight cultures of EcN + Curli, EcN + gadE, and EcN + RP4 (a conjugative plasmid). Each of these cultures was washed of antibiotics in PBS. After diluting each sample to an OD of 0.5, 1 mL of each was combined and set into a 37\*C shaker for 1 hour. As the gadE plasmid contained an oriT site allowing for conjugative transport via RP4, all combinations of the three plasmids should have occurred in the joint culture. We then plated on the antibiotic resistances associated with the gadE and curli plasmids (CAM and KAN respectively) to select for the target combinations. --- Dry Lab ProtocolsBinder Creation Install and use EZBinder, adjusting parameters as specified in the documentation --- Recycling 1. Find the highest pLDDT structure (MONOMER, not complex) and upload it to the remote server 2. Create a SLURM script based off the standard header (from the EZBinder scripts) 3. Edit the following line for your particular binder - remember to have a newline between the header and this line, and a newline between this line and the end of the file. Replace `124-124/0` with the actual length of your protein in amino acids (this example is 124 residues). Replace `B662-875` with the actual chains you want to bind to (662-875 of chain B in this example) 1. `python /home/username/RFdiffusion/scripts/run_inference.py inference.input_pdb=/home/username/input_pdb/input.pdb 'contigmap.contigs=[124-124/0 B662-875]' diffuser.partial_T=15 inference.num_designs=300 inference.output_prefix=/home/username/output/RFD_out/1/prtldf_batch1_` 4. Switch to the RFDiffusion venv 5. Submit the RFDiffusion SLURM job 6. After the RFDiffusion job completes, create a ProteinMPNN script using the RFDiffusion output as the input directory 1. Edit an EZBinder generated ProteinMPNN script, replacing the `pbddir` argument with the RFDiffusion output directory specified above 7. Switch to the ProteinMPNN venv 8. Submit the ProteinMPNN SLURM job 9. After the ProteinMPNN job completes, create an ESMFold script using the ProteinMPNN output as the input directory 1. Edit an EZBinder generated ESMFold script, replacing the `pbddir` argument with the ProteinMPNN output directory if it is different than the EZBinder script 10. Upload `esm.py` 11. Switch to the ESMFold venv 12. Submit the ESMFold job 13. After the ESMFold job completes, download the ESMFold output csv --- Partial Diffusion Follow the instructions for recycling, but use a structure from RCSB as the input instead of the highest pLDDT structure from a previous run --- RosettaDock 1. Prepare your structures for docking by extracting the target into a single pdb and the monomer binder from ESMFold into a single pdb 2. Concatenate the two files using the prepare\_for\_dock() function from helpers.py 3. FastRelax the concatenated structure and upload it to the server 4. Edit the RosettaDock SLURM scripts 1. Use the header from a previous job (remember to switch the partition if your previous job used a GPU-intensive program) 2. Edit the prepack invocation line to use your complex pdb and edit docking:partners if you’re not docking chains A and B 1. `/path/to/username/rosetta_git/Rosetta/main/source/bin/docking_prepack_protocol.default.linuxgccrelease -s /home/username/complex.pdb -docking:partners A_B -docking:sc_min true` 1. Run the RosettaDock prepack script (this should be very fast, and create a file called `complex_0001.pdb`) 2. Edit the RosettaDock invocation line to use your packed complex pdb. 1. `/path/to/username/rosetta_git/Rosetta/main/source/bin/docking_protocol.default.linuxgccrelease -s /home/username/complex_0001.pdb -docking:partners A_B -use_input_sc true -nstruct 50000 -out:file:silent dock_rosetta.out` 3. Submit the RosettaDock SLURM job 4. When the RosettaDock SLURM job completes, download the dock\_rosetta.out silent file. 1. For later analysis, the lines that start with SCORE contain RosettaDock metrics for that structure (there should be 50000 for the above example) {% endblock %}

{% extends "layout.html" %}
{% block title %}Results{% endblock %}
{% block page\_content %}

# Curli

Our research team set out to explore the potential effects of introducing the pBbB8k-csg-amylase in conjunction with the arabinose inducer on curli fiber expression. Our initial hypothesis was that this plasmid would increase the expression of curli fibers and aid in biofilm growth. After experimentation and analysis, our results show promising data working towards our engineering goals.

After running the Congo Red Fluorescence Assay (found in protocols), we revealed several interesting results.

![Fluorescence Measurements from plate reader](https://static.igem.wiki/teams/4598/wiki/curlifig1.png)

Figure 1: Fluorescence measurements from plate reader

## pBbB8k-csg-amylase and Arabinose Induction:

Confirming our hypotheses, it was evident that the introduction of pBbB8k-csg-amylase alongside an arabinose inducer indeed elevates the expression of curli fibers. In figure one, we found more than twice the fluorescence of Congo Red in the arabinose induced cultures over controls. Additionally, the *E. coli* Nissle culture carrying the curli plasmid without the arabinose inducer shows wild type level curli fiber expression. In contrast to similar experiments measuring Congo Red amyloid binding to stationary phase cultures on solid agar, the difference is significantly stronger in liquid culture. This may indicate a lack of curli fiber expression during liquid growth phase which is compensated for by upregulation of the curli plasmid.

![Figure 2a: 0.1% Arabinose Induction”> <p class=](https://static.igem.wiki/teams/4598/wiki/curlifig2left.png)Figure 2a: 0.1% Arabinose Induction

![Figure 2b: 0% Arabinose Induction](https://static.igem.wiki/teams/4598/wiki/curlifig2right.png)

Figure 2b: 0% Arabinose Induction

## Arabinose Concentration Sensitivity:

Interestingly, while arabinose acts as an inducer, concentrations beyond 0.01% adversely impact curli production. Both colorimetric and fluorescence measurements, taken in varied environments of liquid and solid cultures, have demonstrated this phenomenon. It is postulated that higher concentrations of arabinose potentially compromise survivability, thereby negatively impacting curli fiber production. Looking at Figure 1., we found an inducer concentration of 0.01% produced the greatest binding of Congo Red to amyloid fibers, represented by fluorescence. Additionally, we can see in Figure 2. that cultures grown on solid agar induced with a 0.1% concentration of arabinose have significantly worse growth than induced controls Moving forward, this data was used as metrics to establish effect induction levels of curli expression.

![Figure 3a: 37°C](https://static.igem.wiki/teams/4598/wiki/curlifig3left.png)

Figure 3a: 37°C

![Figure 3a: 30°C](https://static.igem.wiki/teams/4598/wiki/curlifig3right.png)

Figure 3a: 30°C

## Nissle 202 Strain – A Suitable Candidate:

Our experiments unveiled that the Nissle 202 strain is proficient in curli expression and can support biofilm growth at both 30°C and 37°C. As seen in the images of Figure 3., we have documented for this first time to our knowledge, curli expression grown at a temperature of 37\*C. By observing minimal to no colorimetric difference between cultures grown at these two temperatures, we conclude that temperatures within this 7\*C range has no effect on Curli expression level and likely does not affect biofilm growth. However, cultures grown at 37\*C appears to have more cohesive and larger growth areas, possibly indicating a stronger metabolic burden of curli fibers at higher temperatures.

![Figure 4: Comparison with cloning strains](https://static.igem.wiki/teams/4598/wiki/curlifig4.png)

Figure 4: Comparison with cloning strains

## Comparison with Cloning Strains:

Figure 4. shows a comparative analysis with *E. coli* DH5α revealed that the *E. coli* Nissle strain has superior curli fiber production in a stationary growth phase on agar plates. This was determined by qualitative observations of the colony size and color indicating biofilm spread and accumulation of Congo Red dye respectively. When comparing color only, Nissle culture appears unmistakably scarlet red while DH5α appears as a white to pinkish colony completely distinct from Nissle. This trend held true regardless of whether pBbB8k-csg-amylase was present or whether arabinose induction wasapplied. Nissle's intrinsic ability to generate greater levels of curli fiber densities could offer considerable advantages in specific applications of stomach colonization.

![Figure 5: Plate read data with curli](https://static.igem.wiki/teams/4598/wiki/curlionly-plateread.png)

Figure 5: Plate read data with curli

## Curli Fibers Enhance Survival in Acidic Conditions:

The most promising results for further engineering goals is the enhanced survivability offered by curli fibers against low acidity environments. In experiments that compared the wild-type Nissle and curli-fiber-producing strains, it was found that the presence of curli fibers enabled survival at a pH level of 3.5. These results are reflected in the growth curves shown in Figure 5. In contrast, the wild-type Nissle strain could only endure up to a pH of 4. To our surprise, the over expression of curli fibers aided in acid resistance at a pH of 3.5.

![Figure 6: Plate read data with curli and GadE](https://static.igem.wiki/teams/4598/wiki/curligade-plateread.png)

Figure 6: Plate read data with curli and GadE

## csg and gadE Synergistic Effects:

Once completed, we transformed into Nissle 1917 and ran a plate assay testing survival in different pH ranges, with supplemented glutamate (since one of the major acid resistant pathways activated by gadE requires glutamate) AND with curli to compare the resistance offered by both. We also performed a cotransformation with both of them to see if they exhibited potentially synergistic effects. After the plate read was completed, we analyzed the data and determined that GadE and curli coexpressed exhibited a synergistic effect and allowed survival at a pH of 2.0 for long term. From this data, we learned that supplementation with excess glutamate was required for this acid resistant phenotype. We also learned that GadE in the absence of curli does not exhibit strong acid resistance. Our next data driven step is to test for ability to perform tasks such as conjugation or chemotaxis in high levels of acid with coexpression of curli and GadE. These results are reflected in Figure 6.

# Curli Discussion:

## *E. coli* Nissle 1917 and its Modification for Enhanced Stomach Colonization

The *Escherichia coli* Nissle 1917 (EcN) strain has proven to possess therapeutic attributes, such as the capacity to compete with and exclude pathogenic bacteria in the gut and the ability to produce antimicrobial substances. Typically, EcN flourishes in the small intestine. However, to enable its colonization and thriving in the stomach, a strategic genetic modification is proposed.

Central to this initiative is the expression of curli fibers. These proteinaceous surface appendages, produced mainly by enteric bacteria, serve pivotal roles in bacterial physiology. Curli fibers enable cell-cell interactions and support bacterial adhesion to various surfaces. This adherence is integral to biofilm formation—a structured, protective community of bacterial cells enclosed in a self-produced polymeric matrix. Beyond their role in biofilm architecture, curli fibers enhance bacterial resilience against host defenses and offer resistance against external challenges, like acidity, desiccation, and antimicrobials.

In the unique environment of the stomach, characterized by its mucosal lining and acidic pH, curli fibers can offer a strategic advantage. The lower stomach lining or the antrum is a region where mucus is less acidic and thick, potentially allowing for bacterial colonization. Curli fibers, with their adhesive properties, can facilitate the binding of the modified Nissle strain to the epithelial cells of the antrum. This binding may offer a stable niche, enabling the bacteria to withstand stomach peristalsis and remain adhered to the antrum, thereby enhancing its colonization potential.

Furthermore, *Helicobacter pylori* is a bacterium known to colonize regions of the stomach, particularly in the protective mucus layer, leading to chronic gastritis and increasing the risk of ulcer disease and gastric cancer. The curli-enhanced EcN can be employed as a strategic tool to target these regions. By promoting co-aggregation or even competitive exclusion, the modified Nissle can interact with *H. pylori*, potentially disrupting its colonization or even reducing its numbers. This interaction could offer a novel therapeutic angle in managing *H. pylori*-related diseases, by utilizing the probiotic potential of Nissle in conjunction with its enhanced adhesive properties via curli fibers.

## mCherry Binder Design

### Summary

We designed 154,019 binders to the fluorescent protein mCherry over the course of our project. Every binder design went through the same pipeline,beginning with RFdiffusion to generate plausible backbone structures, ProteinMPNN to generate sequences, and finally ESMFold to extrapolate structure from sequence. We automated this pipeline and released the automation as our team’s software, [EZBinder](https://2023.igem.wiki/uoregon/software).

Our mCherry binder design proceeded in 5 runs: Initial, Partial Diffusion, Rescue, Recycle Rescue, and Recycle Partial Diffusion.

* Initial: EZBinder outputs from 6IR2
* Partial Diffusion: RFDiffusion partial diffusion using 6IR2 as the initial motif
* Rescue: RFDiffusion partial diffusion using the highest pLDDT output from Initial as the initial motif
* Recycle Rescue: RFDiffusion partial diffusion using the highest pLDDT output from Rescue as the initial motif
* Recycle Partial Diffusion: RFDiffusion partial diffusion using the highest pLDDT output from Partial Diffusion as the initial motif

Every run except for the initial used 15 diffuser timesteps in RFDiffusion and no additional weights (in contrast to the EZBinder defaults: `potentials.guiding_potentials=[\"type:interface_ncontacts,weight:1,type:binder_ROG,weight:1\"] denoiser.noise_scale_ca=0.5 denoiser.noise_scale_frame=0.5`.

### ESMFold metrics

![Table of ESMFold Metrics ](https://static.igem.wiki/teams/4598/wiki/summarytable-1.png)

Summary statistics of our binders. The nanobodies “run” is evaluating the sequences for anti-mCherry nanobodies in RCSB with ESMFold, using the same evaluation method as our designed binders. Nanobodies used (PDB ID | Nanobody Name): 6IR2|Lam2, 6IR1|Lam4, 7SAJ|LaM2, 7SAK|LaM4, 7SAL|LaM6, 8ILX|LaM3, 8IM0|Lam8.

Each run of binders we generated produced a plausible protein structure with very high confidence in several important computational metrics outputted by ESMFold. Every single one of our runs outperformed the real nanobodies for mCherry in every metric except for percentage high confidence pTM, which is biased by the different sample sizes.

pLDDT is a measure of local accuracy, e.g. how far ESMFold believes that its predicted residue location will differ from the real location. It ranges from 0-100 with 100 as the best confidence. The cutoff for very high pLDDT varies between sources, we chose 90 for our analysis. Every run generated thousands of potential binder candidates that met this criterion, despite the fact that pLDDT was by far our most demanding filter for binder designs. Interestingly, recycling binder designs inspired by an actual structure through our protocol (Partial Diffusion, Recycle Partial Diffusion) reduced the percentage of structures with very high pLDDT, whereas recycling binder designs completely hallucinated by RFDiffusion (Rescue, Recycle Rescue), increase the share of very high pLDDTs relative to the initial run. Successive recycling through the protocol (Recycle Partial Diffusion, Recycle Rescue) reduced both the top pLDDT and the share of very high pLDDTs relative to the parent runs. From these results we can conclude that successive recycles don’t improve the quality of the generated structure.

PAE is predicted aligned error, e.g. How far ESMFold believes that a particular predicted residue location will differ when the entire predicted structure is aligned to the entire real structure. A PAE of less than 10 means that a design is a good candidate for wetlab experimentation and lower values are better. PAE universally increases as binders are recycled through the protocol.

pTM is the predicted TM score, a similar metric to PAE, but it takes a global view of the protein rather than a per-residue score like PAE and pLDDT. RFDiffusion recommends using cutoffs based on mean AlphaFold PAE, but ESMFold uses pTM as a global metric already so we’ve included both values for completeness. The share of high confidence (>0.7) pTM scores increases as binders are recycled through our protocol and the mean PAE decreases. PAE and pTM should have the same information, but our results show that they’re inversely correlated. Future work that uses ESMFold as a drop-in replacement for AlphaFold should take this behavior into account.

### Docking Funnels

We took the highest pLDDT binder from each run and docked it to mCherry using the default RosettaDock docking protocol. To evaluate docking effectiveness, we created a plot of energy in REU against rms from the input structure (commonly referred to as a docking funnel). We evaluated energy at the interface of the binder and target, as well as the total energy of the complexed structure. The relative y values should only be compared within plots, and the relative x values should be ignored. Rms is a measure of deviation from the initial input structure, which we created by concatenating pdb files. A tight column of points indicates a single docking position, and the presence of many low energy conformations indicates that the dock is good.

![Docking funnel of lowest pLDDT mCherry Binders, complex energy](https://static.igem.wiki/teams/4598/wiki/round3-total-zoomed.png)

Docking funnel of lowest pLDDT mCherry Binders, truncated to the region between 1200 to -1200. This plot uses the total complexed energy of the protein in REU (Rosetta Energy Units).

![Docking funnel of lowest pLDDT mCherry Binders, interface energy](https://static.igem.wiki/teams/4598/wiki/round3-interface-zoomed.png)

Docking funnel of lowest pLDDT mCherry Binders, truncated to the region between 0 and -30. This plot uses the interface energy of the protein in REU (Rosetta Energy Units).

We omitted recycle partial diffusion from this graphic because none of the docked structures had a negative score, meaning that the complexed energy was predicted to be higher than the energy of the binder and monomer separately, and the binder does not work *in silico*. Recycled partial diffusion created a protein that was computationally predicted to be stable, but it is not predicted to dock, which makes it useless as a binder.

The energetics at the interface and for the entire complex (both docking funnels above) show the characteristic “funnel” behavior of a good dock. As RMSD from the input structure increases, the energy increases, and there are many structures that are slightly different from one another that are all low-energy.

An experimentally validated mCherry binder, Lam2, is a real nanobody that has been shown to bind to mCherry experimentally. The top binder from each run shows comparable energetics to the real nanobody, and several have lower energy conformations than the nanobody. Our best docked structure is from Recycle Partial Diffusion, which has a lower energy conformation than the real nanobody, and a tighter spread of RMSD, suggesting that the docked structure has even less “wobble” than the experimentally verified binder.

### Data Availability

We have made all of our generated pdb ESMFold scores [available on our wiki](https://gitlab.igem.org/2023/uoregon/-/blob/4ae45dc7198a4c733027efcbc9fc46c6f10b4cb9/Binder%20Software/Data/mCherry_Binder_data.csv). We have archived all of our pdb structures as well, but the archive is too large to be hosted on the iGEM wiki. Please contact igem@uoregon.edu if you would like a copy of this archive. Each protein has a unique ID, the sha1sum of its amino acid sequence. This value is provided in the linked spreadsheet under the column `seq_hash`.

## FliG Binder Design

Another computational arm of the project focused on the creation of *E. coli* FliG protein for the purpose of chemotaxis towards our selected bacteria. The goal was to create an inhibitor that binds to *E. coli* FliG so the flagella will only rotate counterclockwise. The rotation will allow it move towards the selected bacteria. From Harshey et.al, they explore the requirement of YcgR for motility inhibition. YcgR is a c-di-GMP binding protein and binds to FliG and FliM. It shifts the motility of the cell and curates it towards rotating in a counterclockwise direction[1](#Ref1). The inhibitor was created using computational modeling. In figure 3D of the article, it depicts which residues correlate to a motility change in FliG when mutated. We used that information to determine which hotspot binding sites on FliG to use so the inhibitor would change the motility.

To begin, the sequence for E coli FliG was found using Ecocyc[2](#Ref2). The sequence was pasted into Alphafold2 on google colaboratory[3](#Ref3). The crystal structure of FliG from *H. pylori* in *Thermotoga maritima* with a pdb number of 1LKV was used as a template for AlphaFold2 to create *E. coli* FliG. Once the *E. coli* FliG protein was generated, it was cleaved to keep the c-terminal domain. Hotspots that Hatshey et. al. found from *E. coli* FliG that binds to YcgR were used as the hotspots that the created inhibitor would bind to. The pdb was relaxed using FastRelax. The protein was named Eflig and was run through RFDiffusion to generate the structure for the inhibitor to Eflig. It was initially run with no potentials and a denoiser scale of 0. The inhibitors were run through protein MPNN fast relax to generate the sequence. They were run through AlphaFold2 to predict if the proteins will take on the conformation and actually bind in the location desired.

AlphaFold2 had multiple results. The predicted local distance difference test (pLDDT) and predicted aligned error (PAE) of the designs were obtained using AlphaFold. A high pLDDT score means that there is a high likelihood that the protein designed is going to actually take on the structure it has[4](#Ref3). A low pLDDT score means that there might be disordered regions in the designed protein and it won’t fold well. A pLDDT score over 80 is high and below 50 is a low score. A PAE score predicts if the binder and the original protein will have aligned residues and bind well. A low PAE score means that the orientation and position of the binder are most likely going to bind well. A PAE below 10 is considered good. A high PAE score means that the binder won’t be aligned and the position of it is not great so the binder will most likely not bind at all.

Initially, in round 1, the designs were run with no potentials and a denoiser scale of 0. These were chosen to begin with initial values to test how the binder would turn out. There were 1362 binder designs made with those potentials and denoiser scale. The following figure shows the average pLDDT and the number of designs that have that pLDDT out of the initial design. The average pLDDT is between 60 and 74, meaning that the model has low confidence so the prediction can’t be trusted to fold. A majority of the pLDDTs were around 70. The designs do have potential because they were all above a pLDDT of 50 so they did not have tremendously low values. From this data, we can not make any claims that the binders will actually fold.

The PAE data for Round 1 is shown below. The average PAE was around 21, which is a high PAE. The lowest PAE was around 17, which is still a high PAE. The high PAE values signify that there is a low confidence that the binder and FliG protein would bind.

![pLDDT of Round 1 EFliG Binders](https://static.igem.wiki/teams/4598/wiki/round1-eflig-pld-2.png)

![PAE of Round 1 EFliG Binders](https://static.igem.wiki/teams/4598/wiki/round1-eflig-pae.png)

After looking at the data obtained from AlphaFold2, we had an idea of how to improve the EfliG designs. In RFDiffusion, adding potentials and a denoiser scale would improve the design of our specific proteins. The binder radius of gyration (ROG) is a potential that tells the protein how compact it should be. Adding a binder ROG would improve the folding and design of the protein. We added an ROG weight of 0.5 and an`interface_ncontacts` weight of 1. The RFdiffusion github notes that a denoiser scale at 0.5 improves binder design so we used a denoiser scale at 0.5. These new potentials and denoiser scales were incorporated into our second round of design.

From these designs, we obtained AlphaFold2 data, shown in the figure below. The pLDDTs from Round 2 AlphaFold2 were much higher. There was a larger spread of pLDDTs from this round. The values averaged around 75. There were also many above 80, meaning that many had the potential to actually fold into a working inhibitor. There were 153 binders above a pLDDT of 80 and 26 binders above a pLDDT of 90 out of 1793 designs. This round had actual designs that had the potential to fold compared to Round 1.

![pLDDT of Round 2 EFliG Binders](https://static.igem.wiki/teams/4598/wiki/round2-eflig-pld-3.png)

![PAE of Round 2 EFliG Binders](https://static.igem.wiki/teams/4598/wiki/round2-eflig-pae-3.png)

The PAE data from Round 2 was also gathered and is shown in the figure below. The average PAE from this set is around 16, which is lower than the first round of design. There are also 70 binders with a PAE lower than 10, meaning that there is a high likelihood for those binders to bind to the target protein.

In order to have a protein that actually folds well and binds to the target protein, both the pLDDT and PAE values have to be good. Of the 1793 proteins from Round 2, 38 were selected based on pLDDT and PAE scores and further analyzed. The highest pLDDT was 96 and the lowest was 80. The lowest PAE score was 5 and the highest 10.

![pLDDT of best round 2 binders](https://static.igem.wiki/teams/4598/wiki/roun2-best-pld.png)

![PAE of best round 2 binders](https://static.igem.wiki/teams/4598/wiki/round2-best-pae.png)

Of the 38 binders, six were selected based on pLDDT and PAE and ordered. The following table are the pLDDT and PAE values of the binders ordered. The highest pLDDT value ordered was 96.39, which is very high and the lowest was 89.98. The lowest PAE value was 5.12 and the highest was 6.3. For the binders with a lower pLDDT, they had a higher PAE.

![Table summarizing best round 2 binders](https://static.igem.wiki/teams/4598/wiki/hi-1.png)

## References:

1. Paul, K.; Nieto, V.; Carlquist, W. C.; Blair, D. F.; Harshey, R. M. The C-Di-GMP Binding Protein YcgR Controls Flagellar Motor Direction and Speed to Affect Chemotaxis by a “Backstop Brake” Mechanism. *Molecular Cell* **2010**, 38 (1), 128–139. <https://doi.org/10.1016/j.molcel.2010.03.001>.
2. Keseler, I. M.; Gama-Castro, S.; Mackie, A.; Billington, R.; Bonavides-Martínez, C.; Caspi, R.; Kothari, A.; Krummenacker, M.; Midford, P. E.; Muñiz-Rascado, L.; Ong, W. K.; Paley, S.; Santos-Zavaleta, A.; Subhraveti, P.; Tierrafría, V. H.; Wolfe, A. J.; Collado-Vides, J.; Paulsen, I. T.; Karp, P. D. The EcoCyc Database in 2021. *Front Microbiol* **2021**, *12*, 711077. <https://doi.org/10.3389/fmicb.2021.711077>. <https://biocyc.org/gene?orgid=ECOLI&id=EG11654>
3. Mirdita, M.; Schütze, K.; Moriwaki, Y.; Heo, L.; Ovchinnikov, S.; Steinegger, M. ColabFold: Making Protein Folding Accessible to All. *Nat Methods* **2022**, *19* (6), 679–682.<https://doi.org/10.1038/s41592-022-01488-1>.
4. Varadi, M.; Anyango, S.; Deshpande, M.; Nair, S.; Natassia, C.; Yordanova, G.; Yuan, D.; Stroe, O.; Wood, G.; Laydon, A.; Žídek, A.; Green, T.; Tunyasuvunakool, K.; Petersen, S.; Jumper, J.; Clancy, E.; Green, R.; Vora, A.; Lutfi, M.; Figurnov, M.; Cowie, A.; Hobbs, N.; Kohli, P.; Kleywegt, G.; Birney, E.; Hassabis, D.; Velankar, S. AlphaFold Protein Structure Database: Massively Expanding the Structural Coverage of Protein-Sequence Space with High-Accuracy Models. *Nucleic Acids Res* **2021**, *50* (D1), D439–D444. [https://doi.org/10.1093/nar/gkab1061.](https://doi.org/10.1093/nar/gkab1061)

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