Team:Wisconsin/Project

The above images and video give the general features of lignin peroxidase (lip), which includes 4 disulfide bonds and 2 Ca2+ that are conserved through all lip. The heme access channels are divided into a larger hydrophobic channel and a smaller hydrophilic channel. Hydrogen peroxide must travel through the hydrophilic channel since the heme is embedded inside of the protein. At the end of the channel, hydrogen peroxide turns into water by oxidizing 2 electrons from the heme at the hydrogen peroxide active site. This allows Lip to oxidize substrates (veratryl alcohol for example.) Since bulky molecules are not able to reduce the embedded heme directly, they must be oxidized at the surface. A Tryptophan with a hydroxyl group on its beta carbon is the center of the surface active site and allows the substrate's electrons to transfer through the protein into the center and reduce the heme, allowing the whole system to start over again. However, much is still unknown about Lip, such as the exact mechanism for lignin degradation or veratryl alcohol's precise role in increasing efficiency.

In researching for an appropriate assay in analyzing the amount of lignin peroxidase produced by our E. coli strain, we found that the veratryl alcohol assay, developed in 1984, is currently the most widely used. It works by viewing the absorbance spectrum of the reaction solution. As veratryl alcohol is oxidized by lignin peroxidase over time, the optical density (OD) and wavelength of the absorbance peak change. By observing the disappearance of the substrate (the rate of change), we can determine the amount of active enzyme in the solution.

In doing additional research we came across a newer and seemingly more effective assay—the Azure B assay. Research showed that Azure B (a blue textile dye) has a higher specificity to lignin peroxidase compared to veratryl alcohol, as veratryl alcohol can be oxidized by fungal enyzmes other than lignin peroxidase. The Azure B assay works the same way as the veratryl alcohol assay in that the disappearance of the dye is measured to determine the amount of lignin peroxidase. The reaction solution comprises of a tartrate buffer (50 mM at pH 4.5), the dye Azure B (32 μM), hydrogen peroxide (100 μM), and the enzyme to be measured, lignin peroxidase.

Figure 6: [http://www.sigmaaldrich.com/catalog/search/ProductDetail/SIGMA/A4043 Azure B. Sigma Aldrich.] (2008). Retrieved Oct 18.

Lignin peroxidase has a few disulfide bonds, which are unable to form within E. coli. To combat this problem we used a strain (any of the Origami strains) available from Novagen that is a knockout for both thioredoxin reductase and glutathione reductase. This prevents the cell from reducing disulfide bonds as they form, allowing for eukaryotic proteins containing disulfide bonds to be expressed in the active form.

Since the extremely large plant cell wall complex cannot be imported into the cell we had to figure out a way to get lignin peroxidase out of the cell. This could either be done by lysing the cell and recovering the enzyme or by exporting the enzyme from the cell using a native export system. We looked into using two native export pathways in E. coli, the Sec pathway and the Twin Arginine Translocation (Tat) pathway. Proteins are recognized for transportation across the membrane by signal peptide sequences. The Sec pathway exports unfolded proteins, which then fold once in the periplasm. The Tat pathway exports already folded proteins. Since the cytoplasm within the Origami strains was an appropriate place for the protein to fold correctly we focused on the Tat pathway. Export of lignin peroxidase could be achieved by fusing sequences encoding for Tat signal peptides in frame to the lignin peroxidase gene.

Figure 7:TAT pathways can export entire folded proteins, while Sec involves an unfolding step, export of the unfolded protein and a refolding of the unfolded protein. Both pathways result in a folded protein in the periplasm.

Another difficulty is the crucial addition of the heme group. Producing lignin peroxidase will only give us nonfunctional polypeptides if we do not have enough heme. We decided to focus up regulation on aminolevulinic acid (ALA) synthesis since it is the rate limiting step for heme production. However, instead of the time consuming up regulation of enzymes responsible for ALA, we decided to simply add ALA to the media, which would then be taken up by e coli. This chemical approach cuts out the rate limiting step entirely and is a cheap method that is realistic for the short time scale of iGEM research.

Sorbitol Biosynthesis

Sorbitol anabolism is part of the glycolysis pathway in E.coli. In the pathway, interconversion between fructose and sorbitol is catalyzed by sorbitol dehydrogenase(SDH), a monomeric enzyme that uses NADH in the process. Natural expression of SDH is generally up-regulated only in the presence of sorbitol as a carbon source. The initial idea to stimulate sorbitol over-production was to simply insert a plasmid containing the gene which codes for SDH and grow the cells in an environment lacking sorbitol thus pushing the metabolic reaction governing fructose and sorbitol interconversion towards sorbitol.

Sorbitol Dehydrogenase NADH binding site (purple wire frame), sorbitol active site (red wire frame), sorbitol hydrogen bonding amino acids (orange wire frame, doesn't include hydrogen bonding from the sorbitol active site) This image was made with PDB file 1K2W on pymol.

The gene encoding for SDH, srlD, is part of the srl operon in E.coli (operon pictured below). Information on the SDH assembly pathway and enzymatic activity is limited. Consequently we decided that insertion of both the srlD gene and srl operon into vectors would take more time initially but would potentially be beneficial due to the chance that the gene alone would not result in effective SDH production.

Figure 6: SRL Operon

Increased cellular production of SDH would most likely increase the amount of sorbitol produced by the cell, however it could not alone account for the type of mass production that would make sorbitol biosynthesis commercially useful. In order to determine other methods of increasing sorbitol production, our group turned to computer modeling.

Phosphofructokinase Knockout

Figure 7: PFK Knockout in the metabolic pathway.

The idea behind a phosphofructokinase (PFK) knockout is to literally stop the glycolysis pathway after fructose-6-phosphate. The backup of metabolic products caused by the knockout should pool and form a favorable pathway for increased sorbitol production. The obvious problem with this procedure is that knocking out a part of the glycolysis pathway under normal circumstances kills the cell. This is also why modeling didn't prove useful as the program showed the cell would be dead. We set out to determine how to bypass this normal cellular function in order to use it to increase sorbitol production.

Experiments have been successfully done to create a viable PFK chromosomal knockouts that contain inducible plasmids encoded with the pfk gene. After obtaining this strain we have done countless growth curves with varying media to create ideal conditions for cell growth. After the optimal media is obtained, the addition of a plasmid containing srlD or the srl operon will be introduced to create what we hope will be this metabolic pathway(right).

Triose Phosphate Isomerase Knockout

Figure 8: TPI Knockout in the metabolic pathway.

Computer modeling was done to determine gene knockouts that increased cellular flux towards sorbitol production. The model was given to us by Dr. Jennie Reed of University of Wisconsin-Madison. More extensive information about modeling can be found at our modeling page. Through the use of our model we determined that knocking out the gene encoding for triose phosphate isomerase would be the most effective at increasing sorbitol production when combined with srl gene up-regulation.

Triose phosphate isomerase(TPI) is another enzyme in E.coli's glycolysis pathway. Its function is to catalyze the interconversion of 3-Phosphoglyceraldehyde to Dihidroxy-acetone-phosphate and back. Knocking out the gene which encodes TPI stops this interconversion, a process which removes products of glycolysis from the cellular reaction mixture shifting the overall reaction downwards towards pyruvate. In essence, knocking out the TPI gene backs up glycolysis and produces a shift towards sorbitol (Pictured right).

TPI is a nonessential gene and as such we were able to obtain it from the Keio collection, a collection of non-essential single gene knockouts from the E.coli K-12 derivative strain BW25113. The increased cellular flux modeled in a TPI knockout combined with increased intracellular SDH production from a plasmid containing either the srlD gene or srl operon will hopefully result in a large scale production of cellular sorbitol.

Even with the promising modeling results, our group worked on alternative routes for increased cellular sorbitol. One of these routes, knocking out phosphofructokinase, was an additional project that our group undertook.

Approach

Figure 9: 1. Wild Type E.coli MG1655 is grown on an LB plate

2. A colony is selected and grown overnight
3. The chromosomal DNA is harvested from the over night culture
4. srlD gene is cloned and amplified via a PCR reaction adding restriction sites for Xba1 and Spe1 at either end
5. The biobrick vector pSB1AK3 as well as the srlD PCR product are digested with Xba1 and Spe1
6. The two digestion products are then ligated together
7. The ligation product is then transformed into competent E.coli
8. E. coli is then plated on LB plates containing Amp/Kan to select for transformants

References

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Guzman L., Belin D., CarsonM., and Beckwith L. (1995). Tight regulation, modulation and high-level expression by vectors containing the arabinose pBAD promoter. Journal of Bacteriology. 177(14): 4121-4130. Thomas Choinowski, Wolfgang Blodig, Kaspar H. Winterhalter and Klaus piontek (1999) The crystal structure of lignin peroxidase at 1.70A resolution reveals a hydrozy group on the Cb of tryptophan171: A novel radical site formed during the redox cycle. Journal of Molecular Biology 286, 809-827 Thomas L Poulos, Steven L Edwards, Hiroyuki Wariishii, and Michael H Gold (1993) Crystallographic reginement of lignin peroxidase at 2A. The journal of biological chemistry 268, 4429-4440 Wendy A Doyel, Wolfgang Blodig, Nigel C Veitch, Klaus Piontek and Andrew T smith (1998) Two substrate intereaction sites in lignin peroxidas e revealed by site directed mutagenesis. Biochemistry 37, 15097-15105

Wolfgang Blodig, Andrew T smith, Wendy A dDoyel, and Klaus Piontek (2001) Crystal Structures of Pristiena dn oxidatively processed lignin peroxidase expressed in escherichia coli and of the w171f variant that eliminates the redox activite tryptophan 171. implications for the reaction mechanisim. Journal Molecular Biology (2001) 305, 851-861