Team:Tianjin/Experiment/Protein Engineering
Experiment of Protein Engineering
Motivation
The degradation of PET is completed in two steps by PETase and MHETase. PETase hydrolyze PET to mono(2-hydroxyethyl) terephthalic acid (MHET), which will be further decomposed by MHETase into two monomers, terephthalic acid (TPA) and ethylene glycol (EG . In this two-step reaction system, MHET, the product of PETase-mediated hydrolysis of PET, was found to be a very minor component, which reveals rapid MHET, indicating the rate determining step in this reaction is the first step, hydrolysis of PET. So in order to accelerate the whole PET degradation speed, increasing the activity of PETase is rather crucial. To enhance PETase hydrolysis activity, we first tried to understand the mechanism of the hydrolysis reaction by generally confirming active sites of PETase and 3 dementional structure simulation.
Fig.1 Simulated 3D structure for PETase
Serine-based Catalytic Triad Mechanism & 3D Model Simulation
Since there is no x-ray structure for PETase , the mechanism of PETase hydrolysis activity can not be exactly identified. But the binding site and catalytic site can be generally inferred according to α/β hydrolase mechanism. Based on the efforts have been made to identify and characterize bacterial cutinases[2], α/β hydrolase fold family contain a highly conserved characteristic GXSXG motif. With sequence analysis, PETase was also found to contain an accordant GWSMG motif. Meanwhile, we simulated a best fit model for PETase by SWISSMODEL, an automated comparative protein modeling server[3]. The template was Thc_Cut2, which shares 52％ sequence identity with PETase [4]. As expected, the homology model of PETase displays a canonical α/β hydrolase fold with a Ser161-His237-Asp237 catalytic triad and a preformed oxyanion hole (Fig.1), suggesting a classic serine hydrolase mechanism. Ribbon diagram of a predicted PETase model. The catalytic triad residues are shown as ball-and-sticks in green, formed by Ser161, His237 and Asp237, and the oxyanion hole binding site residues are in blue, formed by the main chain amides of Met161 and Tyr87.
Theoretical Background
1. The Cpx Regulation System[1]
In order to adapt to their changing environment，Escherichia coli bacterium need plenty of regulatory systems. The Cpx system is a three-component regulatory system which is kind of similar to the lactose operon.
The Cpx system consists of the histidine kinase CpxA, the response regulator CpxR and the periplasmic CpxP protein. CpxA is composed of a large periplasmic domain and a highly conserved cytosolic catalytic domain. Both domains are connected via two trans-membrane helices. CpxA has autophosphorylation, phosphor-transfer and phosphatase activities .Sensing envelope perturbation by an unknown feature, CpxA transmits a signal via a phosphorelay to CpxR, which in response acts as a transcription regulator of genes, whose products are mainly involved in envelope protein folding, detoxification and biofilm formation. The Cpx stress response is controlled by feedback inhibition CpxP acts at the initiation point of signal transduction by reducing CpxA auto-phosphorylation activity in the reconstituted CpxRA system.
The Cpx pathway is activated by a large number of different signals including elevated pH, increasing osmolarity, metals, altered membrane composition, overproduction of outer membrane lipoproteins and misfolded variants of maltose binding protein.
When the stress is at lower level, the CpxP protein combine with the CpxA to prevent CpxA from phosphorylating CpxR and when the stress changes at higher level, some signals lead to activation of the Cpx pathway. Particularly, misfolded protein such as MalE219 interacts directly with the periplasmic domain of CpxA, resulting in stimulation of CpxA phosphotransfer activity towards CpxR.
Fig.5. The Cpx inclusion body responding system in E.coli [1]
Fig.6. The function of ddpX in E.coli under starvation conditions[2]
2. DdpX cell lysis effect[2]
DdpX, namely D-alanyl-D-alanine dipeptidase, is a kind of peptidoglycan hydrolase which can hydrolyze the D-Ala-D-Ala part in peptidoglycan molecule. As we all know, the cell wall of bacterial mainly consists of peptidoglycan, so the ddpX can hydrolyze the cell wall of bacterial.
It cannot be more strange that many bacterial own this kind of seemly dangerous gene in their genome. In fact, this gene also has many benefits to bacterial. In gram-positive bacterial, Vancomycin, a kind of antibiotics, can cause cell lysis because it can combine the D-Ala-D-Ala residue of peptidoglycan in cell wall and block the cross-linking of peptidoglycan. Some gram-positive bacterial like Enterococcus faecalis and Streptomyces toyocaensis have developed the resistance to the vancomycin because they have VanX gene, the homologue of ddpX gene, which can hydrolyze the D-Ala-D-Ala and transfer the D-Ala-D-Ala residue of peptidoglycan to D-Ala-D-Lac residue so that the vancomycin cannot combine the peptidoglycan.
However, in gram-negative bacterial like E.coli, which own the robust outer membrane that can resist the vancomycin, the hydrolase ddpX with the same effects also exists. This is strange because the gram-negative bacterial have no necessity to own this kind of seemly dangerous hydrolase. Actually the ddpX in E.coli has another vital use when they are under starvation conditions. The ddpX can hydrolyze the D-Ala-D-Ala in their cell wall to produce the D-Ala as the carbon source to maintain their life. This mechanism is only carried out when they are under starvation conditions. If the ddpX gene is overexpressed, the cell wall will be damaged and cell lysis will occur.
3. TPA Positive Feedback Mechanism[3][4]
As we all know, PET is solid in normal condition. So it’s not easy for microorganisms to realize if there is any PET in the environment. For this reason, we designed the following regulating path.
We aim at finding a way to offer bacterial the ability to sense TPA so that it can produce more enzyme when TPA degraded by PETase exists in the environment. We find the similar mechanism in the Rhodococcus jostii RHA1, which can make use of TPA as carbon source. We speculate that there must be the pathway we want in the Rhodococcus jostii RHA1. By the way, RHA1 is also well used in microbial consortia part of our project. In Rhodococcus, the distinct expression patterns of the TPA gene clusters indicate that they are independently regulated. The cluster contains gene encoding putative regulatory protein, namely tpaR. This gene encodes the regulatory protein of the IclR family, based on the presence of a conserved signature region. The regulator has helix-turn-helix domain and encodes regulator for its respective operons, which is consistent with the case for IclR-type regulatory proteins for other aromatic catabolism pathways. IclR-type positive regulators bind a sequence before their promoter DNA in the existence of inductor and start the transcription of downstream gene, so we need to express the regulator too.[4] Then the gene followed the promoter will be regulated by TPA.
Fig.7. The TPA positive feedback effects found in Rhodococcus jostii RHA1[3]
We find a promoter from the upstream of a gene named tpaAa regulated by TPA. It will express 300 times more when TPA exist. So we plan to transform the three genes into Saccharomyces cerevisiae. They respectively encode TPA transporter, TPA regulation protein and RFP bonded with the TILS. Then we can detect the intension of the red signal to measure the expression of the protein in distinct concentrations of TPA.
Experiment Design
1. Construction of Reporting System
We use a common expression vector plasmid, pUC19, in E.coli to load our device, which consists of heterologous gene part (in this circumstance, PETase gene) and inclusion body reporting part. First of all, we transform the plasmid with part BBa_K339007 from the kit shipped to us using the protocol in the instruction from iGEM official website. Then we use PCR to amplify this part with restriction endonuclease cutting sites Xba1 and Pst1 respectively on sense and anti-sense primers. Then we use corresponding restriction endonuclease to cut the part and plasmid pUC19 and then use T4 DNA ligase to link them together. The next step is to transform the PETase gene into the same plasmid. The initial gene synthetized does not has promoter and terminator so it cannot express. We have to cut the PETase gene and plasmid pET21A with BamH1 and Sal1 enzyme and link them together to transform the PETase gene into pET21A and then use PCR to amplify the T7 promoter-PETase gene-T7 terminator fragment added the restriction endonuclease cutting sites EcoR1 and Sac1. In this way, after we cut the recombinant plasmid pUC19 and T7 promoter-PETase gene-T7 terminator fragment with corresponding restriction endonucleases and link them together, we can obtain the complete device we want.
Fig.8. The construction process of our reporting system
2. Verification of RFP in the part BBa_K339007
The verification of RFP is carried out by using PCR to amplify the RFP gene with restriction endonuclease cutting sites Xba1 and Sac1 added and then cut the RFP and plasmid pET21a with corresponding restriction endonuclease. Then the cut fragments are linked together and transformed into E.coli to express. Then we can detect the red fluorescence.
3. Method of Red Fluorescence Assay
The red fluorescence is detected by 96-well Microplate Reader. The excitation wavelength is set at 584nm and the emission wavelength is set at 607nm. Considering the RFP has an advantage that it can be directly observed by bare eyes, we also use centrifugation to precipitate the bacterial and observe the color of sediment. The red color can be observed if the RFP is expressed. All the experiment including the latter mentioned regulation system use this assay method.
4. Culture and Expression Condition of E.coli in this experiment
Tradition culture medium LB (5g/L yeast extracts, 10g/L peptone, 10g/L NaCl) is also used by us. Because of the ampicillin resistance gene in the plasmid pUC19 and pET21A, ampicillin (100μg/mL) is added to screen for the correctly transformed bacterial. 5mL bacterial are cultured in test tube at 37℃ with 200rpm shaking speed. IPTG is added to induce the expression of PETase gene after 6 hours.
Fig.9. The construction process of our cell lysis based regulation system
5. Construction of Cell Lysis Based Regulation System
This system has a great similarity to the reporting system above. Therefore it is easy to construct because we only need to change the RFP gene to the ddpX gene. However, there is no restriction endonuclease cutting site between the CpxR and RFP gene sequence according to the part map from the iGEM official website, so we have to use PCR to amplify the CpxR promoter solely and add restriction endonuclease cutting sites Xba1 and BamH1 respectively in both end. The ddpX gene is obtained from the E.coli genome using colony PCR and the BamH1 and EcoR1 restriction endonuclease cutting sites are added respectively to both end. Then the three fragments, CpxR promoter, ddpX gene, and cut plasmid pET21a are linked together. Then the whole part is amplified by PCR with Xba1 and Pst1 restriction endonuclease cutting sites added respectively to both end. This way, we can easily cut down the former CpxR-RFP fragment and add the new CpxR-ddpX fragment to the plasmid pUC19.
6. Verification of ddpX Gene Effect
Just like the verification of RFP mentioned before, the verification of ddpX is carried out in the similar way. The pET21a plasmid is cut by BamH1 and EcoR1 instead of Xba1 and EcoR1 , so that the ddpX can be linked to the cut plasmid pET21a solely. Then we can detect if the cell lysis occurs.
7. Method of Cell Lysis Assay
Cell lysis can be reflected by the OD600 of culture medium. The lower the value of OD600 is than the wild type E.coli at the same condition, the stronger the cell lysis effect will be. The OD600 is detected by 96-well Microplate Reader. In order to know the OD600 value continuously, the detection process works through the time of bacterial growth and we will obtain the OD600-Growing time curve.
8. Chassis selection for TPA Positive Feedback Based Regulation System
As the explanation before, the TPA positive feedback system is derived from the TPA degradation metabolic pathway in Rhodococcus jostii RHA1. Considering the difficulty of conducting gene-scale operation in this unusual organism, we directly synthetize all the gene including tpaK, tpaR, and TILS. At first we want to use E.coli to test this device because of the easy and familiar operation. However, in this situation, we have to transform at least 3 plasmids and this cannot be more difficult for E.coli. Therefore, we use another familiar organism, Saccharomyces cerevisiae, as the chassis. In the preliminary experiment, we successfully transform 3 plasmids into Saccharomyces cerevisiae.
9. Construction of TPA Positive Feedback Based Regulation System
We use common plasmids of Saccharomyces cerevisiae, pRS413, pRS415 and pYES2, to respectively load the TPA transporting protein gene, TPA regulation protein gene and TPA induced RFP gene. First of all, we use PCR to amplify all of these fragments and add different restriction endonuclease cutting sites. Then we cut the plasmids with corresponding restriction endonucleases. Then these cut fragments are linked according to the designed order and transformed into Saccharomyces cerevisiae. We screen for the correctly transformed cell by using the Sc-Ura-Leu-His plate.
Fig.10. The construction process of our TPA Positive Feedback Based regulation system
10. Culture and Expression Condition of Saccharomyces cerevisiae in this experiment
Traditional YPD culture medium (22g/L glucose, 20g/L peptone, 10g/L yeast extracts) is used by us. Sc-Ura-Leu-His culture medium (22g/L glucose, 6.7g/L yeast nitrogen base, 1.224g/L nutrient deficiency mixture without Ura, His, Leu and Trp, 5mg/L Trp) is used to screen for correctly transformed cell. All the cells are cultured in 5mL medium at 30℃ with shaking speed of 200rpm. To induce the expression of RFP, we add TPA with different concentration. We first make up TPA standard solution with TPA concentration of 5g/L. Then we respectively add 0, 1μL, 10μL, 100μL, 1mL standard solution to the culture medium.
Expected Results
PETase and MHETase are two key enzymes in our project. However, as heterologous proteins, the expression of these two enzymes face many problems just like expressing other heterologous proteins before including the formation of inclusion body, the lack of regulation pathway, etc. We design this R-R system in order to express the two enzymes visibly and regularly.
First, we hope to directly observe the expression condition of our enzyme by color, when the inclusion body form, which means the overexpressing, the red color can be observed.
Second, when inclusion body form, the normal way to solve this problem is to use lysozyme and ultrasonic to break the cell and purify the protein, which is complex and time-consuming. We expect the cell lysis will automatically occur when the inclusion body form by using ddpX gene.
Third, we expect the chassis organism can sense the existence of TPA, the hydrolyze product of PET and using TPA as the induction of PETase gene. Thus if the degradation process start, this process can be even enhanced until the PET is used up.
References
[1]Physiologie der Mikroorganismen, Humboldt Universitat zu Berlin, Chausseestr. Misfolded maltose binding protein MalE219 induces the CpxRA envelope stress response by stimulating phosphoryl transfer from CpxA to CpxR. Research in Microbiology 160 (2009) 396-400.
[2]Ivan A. D. Lwssard and Christopher T. Walsh. VanX, a bacterial D-alanyl-D-alanine dipeptidase: Resistance, immunity, or survival function? Proc. Natl. Acad. Sci. USA. Vol. 96, pp. 11028–11032, September, 1999.
[3]Hirofumi Hara, Lindsay D. Eltis, Julian E. Davies. Transcriptomic Analysis Reveals a Bifurcated Terephthalate
Degradation Pathway in Rhodococcus sp. Strain RHA1. Journal of Bacteriology, Mar. 2007, 189(5), 1641–1647.
[4]Molina-Henares, A. J., T. Krell, M. E. Guazzaroni, A. Segura, and J. L. Ramos. 2006. Members of the IclR family of bacterial transcriptional regulators function as activators and/or repressors. FEMS Microbiol. Rev. 30: 157–186.
