Breakdown of Trimethylamine via Trimethylamine Dehydrogenase to Minimize Heart Disease Caused by Red Meat Consumption

Phase 1: Intro (Building the System)

Phase 1 mostly involves designing the recombinants and testing protein expression. In this phase the sequential characteristics of our parts were designed and optimized for our expression conditions. Then the insert/gene of interest was cloned into our vector using T4 ligation. After designing and cloning our parts we had to test the optimal conditions for protein expression. The goal of this phase was to design and build our parts for future testing.

Phase 1: Restriction Digest

Restriction Digestion is the process of cutting DNA into shorter fragments with Restriction Endonucleases.
What do we use it for? We utilize this essential process in order to cut up the plasmids. This year we used the enzymes EcoRI-HF and MluI-HF for restriction enzymes, which are both HF. This is the first year we are using MLU1, so we had to add primers to the gel.
General process: Restriction digest is accomplished by incubation of the target DNA molecule with restriction enzymes - enzymes that recognize and bind specific DNA sequences and cleave at specific nucleotides either within the recognition sequence or outside of the recognition sequence.
Applications: Restriction digest has numerous applications, particularly useful for testing for mutations and creating a new assembly of plasmids.

Phase 1: Designing the Recombinants

In order to quantify the efficiency of TMA and formaldehyde degradation, we designed 2 plasmids containing trimethylamine dehydrogenase (TMADH) and formaldehyde dehydrogenase (FDH).
The first construct we designed used the pBAD vector backbone which incorporated the AraC operon and an ampicillin-resistant gene to select for transformed bacteria. The AraC operon allows for the controlled induction of the protein that we want to test, TMADH, upon the addition of arabinose. The Strep-tag II serves as a method for purification of TMADH. Finally, a PhoA signal peptide allows for the secretion of the protein into the bacteria’s intermembrane space, eventually diffusing into the environment and degrading TMADH as intended. The HA-tag is an antigen that is encoded into our plasmid that can be targeted by anti-HA, an antibody that reacts to produce light. This gene allows for identification of the protein later on western blot.
The second construct shares the same pBAD vector design and therefore the same operons for gene expression, but secretes FDH instead of TMADH. The FLAG-tag is a unique antigen that will bind to the anti-FLAG antibody. This antibody has an additional antigen, Horseradish Peroxidase, that will produce light. This allows us to identify our protein of interest similar to how HA-tag is used for TMADH.

In the future, we plan to create a plasmid which expresses both the TMADH and FDH protein in one construct, once we have an understanding of the degradation levels of each individual protein. The combined system will be capable of eliminating the presence of TMA without the toxic by-products which are typically created when TMA is broken down. All of the genes found in the original plasmids will be conserved aside from the Strep-tag II because protein purification is not necessary. In order to confirm that our plasmid expresses the intended proteins, we will be utilizing two membranes with each of the plasmids on two separate western blotting membranes with an anti-HA tag and an anti-FLAG tag on the other to serve as a control.
We ordered the “inserts” from IDT, the two gBlock fragments that we ordered were the StrepII-TMADH-PhoA-HA and StrepII-FDH-PhoA-FLAG. Then we cut both the gBlock fragments and the pBAD vector that we had in the lab with MluI-HF and EcoRI-HF so that the two fragments would have compatible sticky ends and then we could ligate them to form a complete plasmid. Then, in order to confirm our clones we transformed them into bacteria on LB-Ampicillin plates, isolated the DNA, and sent the samples for Sanger sequencing.

Phase 1: Arabinose Induction

The purpose of this arabinose induction test is to test the optimal concentration of arabinose to induce the expression of our protein. In order to do this, first pBAD-FDH and pBAD-TMADH constructs were transformed into bacteria and cultured at four different concentrations arabinose. These concentrations were achieved via a serial dilution as follows: 0.1%, 0.01%, 0.001%, and 0% arabinose. After an induction period of 4 hours the bacterial pellet was lysed and the samples were run on an SDS-PAGE gel to separate the proteins by size. Then the proteins were transferred to a membrane then western blot was performed in order to identify the expression levels of the specific protein. The pBAD-FDH protein also has FLAG-tag which can be recognized by the anti-FLAG antibody; the pBAD-TMADH protein also has an HA-tag which is recognized by the anti-HA antibody. Both antibodies can be reacted, (in the case of anti-HA or indirectly through secondary antibodies in the case of anti-FLAG) with luminol to produce light that can be captured on an x-ray film. In this way it is possible to not only identify the expression of our protein but also roughly estimate the level of expression by qualitative observation of the brightness of the protein bands (as a higher concentration of protein should result in brighter bands.)

Phase 1: Western Blot

Western blot is a biological technique used to detect and analyze specific proteins in a mixture of various proteins. In our experiment, we analyzed FDH and TMADH.
The main purpose of western blot for our project was to confirm the induction efficiency of our proteins, we need to first prepare the bacteria via arabinose induction. A serial dilution of arabinose was added to the bacteria and incubated for four hours. Then 1 mL of each sample was centrifuged and the bacterial pellet was resuspended in SDS-PAGE sample buffer. The solution was heated to 95C in order to disintegrate the bacterial membrane and free the protein.

Gel Electrophoresis
The samples prepared in the previous step were then loaded into a SDS-PAGE gel. SDS-PAGE separates proteins according to their mass in kDa. This is a denaturing process as the SDS detergent added destroys the secondary and tertiary structures of proteins. SDS leads to equally negative-charged proteins, resulting in separation that depends only on the mass of the proteins (and not their charges). We loaded two gels each with identical samples, both had TMADH and FDH samples. This is so that when we blot the two membranes TMADH will serve as a negative control for FDH when anti-FLAG is added and vice versa.
Blotting the Membrane
During this stage, negatively charged proteins move away from the gel to the membrane and are transferred onto a solid nitrocellulose membrane. The advantage of nitrocellulose is that it limits background noise that are non-specific.
Two main methods of blotting are wet and semi-dry. Wet method is more efficient and prevents the gel from drying. Semi-dry method, although less efficient, is recommended for small, less complex proteins. For this experiment, the wet method was used.
Antibody Probing
After blotting, blocking buffer is added. Blocking buffer is a solution of proteins that bind to all remaining surfaces of the membrane that is not already covered by protein. This method is used in order to reduce any non-specific binding and decrease background noise.
Next, a solution with blocking buffer and the appropriate antibodies are added on top of the membrane so that the antibodies can bind to the proteins on the membrane. Two major types of antibodies added are primary antibodies and secondary antibodies. Primary antibodies specifically binds to the antigen of interest. Secondary antibodies bind to primary antibodies. For this experiment, anti-FLAG was the primary antibody to the FDH samples and anti-goat mouse FLAG was the secondary antibody to anti-FLAG; for TMADH samples anti-HA was the primary antibody and there was no secondary. When combined with certain substrates, the reporter enzymes in secondary antibodies produce color or light, making it easier for scientists to detect and image them. In our case, the substance luminol was used to produce light upon reaction with the antibodies on our membrane.
Detection
Chemiluminescent substrate produces light, which can be captured using film and imaging techniques such as the CCD (charge-coupled device). In order to detect the signal coming off the membrane we used x-ray film over a fifteen second exposure time and developed the film using a film processor.

Phase 2: Intro (Testing Our System)

Phase 2 constitutes the biological assays where we tested the expression level of TMADH and FDH on agar culture plates. The main goal of this phase was to observe the expression of each protein in E.coli and also see if the proteins could be manipulated and expressed in the culture environments that we had designed.

Phase 2: Endo Agar Plate Assay

In order to measure the degradation of trimethylamine (TMA) as a result of the secretion of trimethylamine dehydrogenase (TMADH) in bacteria. This assay is trying to detect and potentially quantify the secretion of TMADH, if proven successful this assay will also be used to differentiate between more or less successful colonies after each successive generation of mutagenesis during the enzyme evolution part of our project.
This assay, builds off the basis of the endo agar plates, which was originally used to test for gram positive versus gram negative bacteria. The plates rely on the present of the LacC operon in bacteria which produces aldehydes once absorbed into the cell, only gram negative bacteria would be able to absorb lactose that is in the plate and therefore secrete aldehydes. The aldehydes would react with the compounds in the plate, namely sodium sulfite and basic Fuchsin, and turn a darker pink color. We are using this mechanism of reacting in the presence of aldehydes to detect the formation of formaldehyde after the breakdown of TMA by TMADH, as formaldehyde is a byproduct of this reaction. However, in order to prevent non-specific uptake of lactose and secretion by other gram negative bacteria we used the strain DH5a, which although is a gram negative bacteria has a mutation in the LacC operon and therefore does not secrete aldehyde.

Phase 2: Testing TMADH on Endo Agar Plate: Expected Results

When testing TMADH on a separate plate, for the negative control the expected results is that there is bacteria growth on the plate while the surrounding gel color hasn’t changed. This is because the plate detects the formation of formaldehyde. We will test the plasmid which expresses TMADH using a plate which turns pink in the presence of formaldehyde. Since the breakdown of TMA via TMADH should leave formaldehyde as the by-product, we expect to see pink rings around each colony that grows and has transformed bacteria for the positive control. If the bacteria do not successfully express TMADH, we do not expect to see any change in the color of the plate, but there will be colonies growing. Our negative control will contain bacteria without the plasmid, in which we expect no bacterial growth.
When testing the positive control for TMADH on a separate plate, the expected results is that there is bacteria formation on the plate. Surrounding the plate, a pink color should form because TMA will produce formaldehyde if the TMADH is active.

Phase 2: HCHO Plate Assay: Expected Results

An HCHO assay is used to test the expression of formaldehyde resistance. This is used in our experiment during normal testing of the mutated E. coli bacteria and enzyme evolution.
There are four components to the HCHO plate assay in our experiment. The first component is LB, a nutrient for the E. coli bacterial growth. The agar allows for the solidification of the gel. The third component in the assay plate is Ampicillin, which will prevent the bacteria that do not contain the plasmid from proliferating. Finally, formaldehyde is added to test if the E. coli bacteria expressing FDH will exhaust its presence. In order to determine the plate conditions that would be optimal for our tests we plated non transformed DH5a cells onto the plates which has a serial dilution of formaldehyde on them. Then we were able to determine the concentration range that killed a significant majority of the cells, but not all of them. We determined the ideal range for testing was 0.05%-0.1% formaldehyde. We observed that it was necessary to make the formaldehyde plates after each trial because time decreases the effectiveness of the plate; formaldehyde has a low boiling point so it evaporates easily.

When plating our confirmed clones we expect that the bacteria expressing our FDH gene will survive much better on these formaldehyde positive plates than the negative control.

Phase 3: Intro (Evolution and Mutagenesis)

The goal of this phase was to induce thousands of mutations in our construct in order to select for the mutation over a series of generations. In order to accomplish this we first needed to calibrate the PCR conditions and quantify the mutation rate. The main goal of this phase was to perform the first round of mutagenesis.

Phase 3: Enzyme Evolution (Error Prone PCR)

Our goal is to generate several mutations of our constructs so that they can later be screened and selected for the best mutation. When repeated over several generations, a more efficient enzyme should be produced as an end result.
A typical reaction mix for PCR would include the DNA polymerase, dNTPs (deoxyribonucleotide triphosphates), the primers, template, and water. For error prone PCR, dPTP which is a mutagenic analog of dNTP and can cause mutations at a “ratio of 5:4:1:1 (A→G:T→C:G→A:C→T), with a total rate of mutagenesis of up to 19%” (Jena Bioscience). During testing, the test group was the PCR reaction with dPTP and Taq polymerase (as well as all the other components of a normal PCR reaction) and one without dPTP but with Taq polymerase, dNTPS, etc. this was in order to compare the mutated sample to a negative control while confirming the PCR products size on an agarose gel. The general strategy for this random mutagenesis, with the dPTP, requires that the PCR product from the first PCR was used as the template and that PCR be repeated without dNTP in order to remove the mutagenic dPTP from the sample.

Before we could use this procedure in our experiment we needed to calibrate some of the more technical conditions. Firstly, while we were testing the PCR conditions for the experiment we noticed that a gel purification step was required between the first and the second PCR otherwise the resulting PCR product of the second PCR was non specific. We also needed to determine the appropriate PCR conditions for our experiment, which is outlined in our results.

Team:TPHS San Diego/Experiments