Lysis Team

Engineering Cycle 1: Creating Engineered Bacteria with aTc-Activated Lytic Genes

Goal

Use the lysis mechanism native to E. coli bacteriophages to burst open our probiotic and release the therapeutic. The benefits of lysing the probiotic are releasing larger proteins into the environment that are not easily excreted through the bacterial cell wall and controlling the population of our probiotic inside the gut microbiome.

Design

From literature review, we identified five proteins from bacteriophages that infect E. coli that induce bacterial lysis through different mechanisms. The lysis genes we identified were lambda phage Rz/Rz1, lambda phage holin, T4 phage lysozyme, T7 phage lysozyme, and ΦX174 phage gene E. We designed a control using mEGFP instead of a lysis gene. We inserted the lysis gene and positive control GFP under the control of the inducible Tet promoter for testing using the pDSG346 {avGFP} plasmid for the backbone.

Build

We used In-Fusion assembly to insert the lysis and control meGFP genes into a plasmid under a Tet promoter. We then transformed the plasmid into E. coli via heat shock. The T4 lysozyme gene was found on the T4 Lysozyme WT plasmid on AddGene. The ΦX174 gene E was found on the pCSaE500 plasmid on AddGene. The genetic sequences of lambda Rz/Rz1, lambda holin, and T7 lysozyme were synthesized from NCBI.

Test

To induce lysis, we added aTc to activate the promoter to transcribe the lysis genes and produce subsequent proteins. We added varying concentrations of aTc (50, 100, 200 ng/mL) to see what amount of aTc was optimal for the most complete lysis of the population. We initially measured cell viability by measuring the OD600 of the sample using a spectrophotometer every hour for 6-7 hours. However, we switched to measuring OD600 in a plate every 10 minutes over 18 hours so we could get a more continuous reading over a longer period of time.

Learn

We learned that ΦX174 gene E and lambda Rz/Rz1 were the most effective at lysing cells, resulting in a decrease in the population. We also learned approximately how long after the addition of the aTc inducer the cells started to lyse and when the population reached a stable minimum. We saw the T7 and T4 lysozymes were not effective at lysing, and through further literature review we learned that they are usually paired with holin genes to pass through the inner membrane and access the cell wall. We also observed that a concentration of 100 ng/mL aTc was the most effective for lysis

Engineering Cycle 2: Quantify How Much Protein Lysis Mechanism Releases

Goal

Compare how much GFP protein can be released through different lysis genes

Design

Next, we wanted to observe how much protein would be released when the E. coli was burst via the lysis proteins. To do so, we added green fluorescent protein (GFP) under a constitutive promoter (BBa_J23199) to our backbone. This would be an easily measurable marker of how much GFP was in the environment. We then added the two most effective lysis genes, lambda Rz/Rz1 and ΦX174 gene E, under the tet promoter in the same plasmid. We additionally designed a control by inserting mCherry instead of a lysis gene. When the bacteria was lysed, the GFP would be released into the supernatant.

Build

We ordered the J23199 promoter and GFP as a gene block and used In-Fusion assembly to insert the block into the plasmid. We also used In-Fusion assembly to insert the lysis and control mCherry genes into a plasmid under a Tet promoter. We then transformed the plasmid into E. coli via heat shock.

Test

We prepared an overnight culture so the bacteria could accumulate the constitutively produced GFP. The next day we first measured the starting OD600 in the spectrophotometer. Then we added aTc at varying concentrations (50, 100, 200 ng/mL) and let them incubate for 6 hours to fully induce lysis. After, we measured the OD600, spun down a 200ul to obtain the supernatant, then measured the GFP fluorescence in the plate reader. We measured the GFP fluorescence normalized to the OD600 versus time.

Learn

When adding 100 ng/mL aTc to the samples, the bacteria expressing Rz/Rz1 had a higher GFP fluorescence / OD600 than the control by ~6000 a.u. When adding 100 ng/mL aTc to the samples, the bacteria expressing protein E had a higher GFP fluorescence / OD600 than the control by ~112,000 a.u. We learned both of the lysis mechanisms allowed GFP to be released into the environment, but protein E was more successful in releasing higher amounts. This demonstrated that we could achieve a “high dosage” of our produced therapeutic protein by using ΦX174 gene E.

Engineering Cycle 3: Test How Our Lysing Probiotic Mechanism Interacts with Nascent Microbiome

Goal

See whether our lysing bacteria causes negative effects on other bacterial cells or mammalian cells to mimic how BoomColi may interact with other cells in the gut

Design

We wanted to determine whether our lysing mechanism would harm surrounding bacteria in the intestine. We tested how our lysate would affect other competent stellar cells as a model for how it would affect other gram-negative bacteria living in the gut.

Build

We took the cells transformed with the plasmid containing the lysis gene and grew them in an overnight culture. We then took two approaches to testing if the lysis mechanism affected other bacteria. Our first approach to test this idea was to co-cultured our engineered bacteria with the lysis gene under the Tet promoter with bacteria constitutively expressing GFP. Our second approach was to first lyse the engineered bacteria independently, extract the lysate, then add it to bacteria and measure their OD600 over time.

Test

For the first approach, we grew our engineered lysing bacteria and constitutively expressed GFP bacteria in separate overnight cultures. After, we diluted the samples to have the same starting optical density and added equal amounts in well plates. We added varying concentrations of 50, 100, and 200 ng/ul aTc to the wells and measured optical density and GFP fluorescence over 18 hours. For the second approach, we also grew our engineered lysing bacteria and neighboring bacteria in separate overnight cultures. After, we added varying concentrations of 50, 100, and 200 ng/ul aTc to the engineered lysing bacteria and let it incubate for 6 hours to lyse. To verify, we observed a decrease from initial OD indicating the bacteria lysed over the 6-hour time period. We spun down the bacteria and took the supernatant containing the lysis proteins and other cell contents. We then added equal amounts of the lysate and neighboring bacteria and measured the optical density over 18 hours.

Learn

From our first approach, we learned that it was difficult to normalize the GFP fluorescence data expressed by the neighboring bacteria over the optical density because the optical density contributed by the engineered vs. neighboring bacteria was indistinguishable. It was difficult to interpret the data collected from this experiment to correctly represent the interaction, so we decided not to include it. However, from our second approach, by growing the neighboring bacteria alone and just adding the lysate, we were able to gain clearer data. We found that there was no difference in the growth of the neighboring bacteria when the various lysates containing phiX174 protein E, lambda phage Rz/Rz1 proteins, and the control were added, supporting that the lysing mechanism would have little effect on nascent bacteria.

Engineering Cycle 4: Test How Our Lysing Probiotic Mechanism Interacts with Mammalian Cells

Goal

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Design

We wanted to test whether the lysate will cause any issues with neighboring mammalian cells because in vivo, our E. coli will be around gut cells.

Build

To do so, we designed an experiment in which we allowed our engineered cells to lyse, took their lysate, and put it onto HEK cells, then measured the cell count to see whether there is a significant loss of mammalian cells. We started by growing overnight cultures of mammalian cells using plasmids Lp 001, 002, 003, 004, 007. We then took 500uL of media and 500uL of lysate (supernatant). Meanwhile, we also cultured wildtype HEK293T cells.

Test

Once the bacterial lysate was ready, we added about 1.5 mL onto each of the 6 wells of the HEK293T mammalian cell plates. We waited for 24 hours and measured how many of the mammalian cells survived using a cell counter. We also used just LB media without any bacteria or lysate as a control.

Learn

We noticed that the amount of mammalian cells decreased a similar amount when adding lysate and normal LB media. We believe that even just LB, without the bacterial lysate made them unhappy, however, there was not a significant decrease in cells due to just the bacterial lysate. However, we wanted to better the results of this experiment by imaging the cells after rather than just counting the number of cells to see if there were any noticeable changes. We redid the experiment, testing the lysate of the plasmids LP001-LP004, LP007 on HEK cells and using LB as a control. However, unlike last time, we also imaged the mammalian cells after and observed if there were any changes to their appearance. We did not notice any observable differences and there was not a drastic change in the cell count.

Theraptuics Team

Engineering Cycle 1: Constructing plasmids containing signal peptide and IL-10 (TP002-TP004)

Goal

Construct plasmids TP002-TP004 to produce functional IL-10 E.coli

Design

In order to produce functional IL-10 in E.coli, a periplasmic signal peptide was fused to IL-10 and these genes were placed under the control of a constitutive promoter- ProD. Three different periplasmic signal peptides were tested for this purpose. The fused signal peptide + IL-10 gene fragments and the backbone TP001 plasmid containing mCherry were ordered from IDT.

Build

PCR was performed on TP001 (backbone plasmid) to amplify it and remove the mCherry sequence. PCR was also performed on the three ordered fused gene inserts to amplify them. Gel electrophoresis and PCR purification were performed to verify the desired fragments. This was followed Infusion assembly and chemical transformation to construct the final plasmids- TP002, TP003 and TP004

Test

We carried out two rounds of western blot experiments on cells containing the above plasmids and observed no IL-10 bands. With the exception of plasmid TP003 showing production of IL-10 in our first western blot. However, when a second round of western blot was performed, we did not see a band around 22 kDa as expected. We sent our plasmids for sequencing and compared them to our designed plasmids. Analysis of the gene sequences showed that there were stop codons in all three of our signal peptide sequences and no start codon directly before the signal peptide sequence.

Learn

Stop codons in our signal peptide sequences- DsbA, TorA and PhoA, and no start codon before these periplasmic signal sequence may have resulted in no IL-10 production. Removing these stop codons and adding a start codon before the periplasmic signal sequence should fix the issue.

Engineering Cycle 2: Removing Stop Codons and Editing the Periplasmic Signal Sequence

Goal

To remove the stop codons within the periplasmic signal sequences and add a start codon before the periplasmic signal sequence.

Design

New primers were designed and ordered to remove the stop codons within the three signal peptide sequences and add a start codon.

Build

PCR was performed on plasmids TP002, TP003 and TP004 using the newly ordered primers. Gel electrophoresis and PCR purification was performed to verify the presence of desired products. We were unsuccessful in cloning TP003 and hence decided not to continue with it. In-Fusion assembly and chemical transformation was carried out for plsmids TP002 and TP004 to construct our final desired plasmids.

Test

A final round of western blot was performed on cells containing plasmids TP002 and TP004. We observed desired bands of around 22 kDa for cells containing plasmids - TP002 and TP004, indicating these cells produced functional IL-10.

Learn

These results helped us conclude that we were successful in cloning E.coli to produce IL-10.

Engineering Cycle 3: Constructing plasmids containing signal peptide + IL-10 + linker and mCherry (TP005-TP007)

Goal

Construct plasmids TP005-TP007 to produce functional IL-10 in the periplasm of E.coli and to test the functionality of all three signal peptides - DsbA, TorA, and PhoA.

Design

A periplasmic signal peptide was fused to IL-10, a linker sequence, and mCherry which were placed under the control of a constitutive promoter, ProD. Three different plasmids were designed using three different signal peptide sequences—DsbA, TorA, and PhoA. The fused signal peptide + IL-10 + linker gene fragments and the backbone TP001 plasmid containing mCherry were ordered from IDT.

Build

PCR was performed on TP001 (backbone plasmid) and the three ordered fused gene inserts to amplify them. Gel electrophoresis and PCR purification were performed to verify the desired fragments. This was followed by Infusion assembly and chemical transformation to construct the final plasmids—TP005, TP006, and TP007.

Test

The plasmids were sent for sequencing, and we compared them to our designed plasmid sequence. The plasmid sequence was identical to our designed plasmids, but we noticed a stop codon after the IL-10 gene sequence that we forgot to remove.

Learn

It is important to double-check sequences obtained from online sources and edit them according to our experiment. We then worked to remove the stop codon in the next engineering cycle.

Engineering Cycle 4: IL-10 Sequence Editing

Goal

To remove the stop codon after the IL-10 gene sequence.

Design

New primers were designed and ordered to remove the stop codon after the IL-10 gene sequence.

Build

PCR was performed on plasmids TP005, TP006, and TP007 using the newly ordered primers. Gel electrophoresis and PCR purification was performed to verify the presence of desired products. In-Fusion assembly and chemical transformation was carried out for plasmids TP005, TP006, and TP007 to construct our final desired plasmids FTP005, FTP006, and FTP007.

Test

The newly constructed strains were observed under a confocal fluorescence microscope. We did not observe a fluorescent ring around each cell as expected. All three strains appeared fully fluorescent under the microscope, indicating that our protein is not restricted to just localizing in the periplasm. Plasmids FTP005, FTP006, and FTP007 were sent for sequencing. The results showed that we were successful in removing the stop codon after the IL-10 sequence in plasmids TP005 and TP007 only. TP006 still had the stop codon after its IL-10 sequence.

From this engineering cycle, we learned that the presence/absence of a stop codon after IL-10 did not make a significant difference, as each individual cell appeared bright under the microscope irrespective of a stop codon after IL-10. After analyzing the gene sequences once more, we found stop codons within the periplasmic signal sequences and also realized that these signal peptide sequences lacked a start codon near its start site. Additionally mCherry had its own start codon. In the next cycle we tried to remove these stop codons within the periplasmic signal sequences and add a start codon.

Engineering Cycle 5: Edit the Signal Peptide Sequence

Goal

To remove the stop codons within the periplasmic signal sequences (DsbA, TorA, and PhoA) and add a start codon before the signal peptide sequence.

Design

New primers were designed and ordered to remove the stop codons within the periplasmic signal sequences and add a start codon.

Build

PCR was performed on plasmids FTP005, FTP006, and FTP007 using the newly ordered primers. Gel electrophoresis and PCR purification was performed to verify the presence of desired products. In-Fusion assembly and chemical transformation was carried out for plasmids FTP005, FTP006, and FTP007 to construct our final desired plasmids.

Test

The newly constructed strains were observed under a confocal fluorescent microscope, and we did not see a fluorescent ring around each cell as expected. Our results were similar to those of the previous round of imaging.

Learn

As each individual cell appeared fluorescent under the microscope, we hypothesized that this could be due to the mCherry reporter gene having its own start codon. So our periplasmic signal sequence+IL-10+linker+mCherry was being produced as well, but we were unable to see it clearly due to the production of excess mCherry in the cytoplasm. We believe that removing this start codon before mCherry will allow us to observe a thin fluorescent ring around each individual cell as desired. Due to time constraints we were unable to do another round of molecular cloning.