Thiol disulfide exchange catalysed by thioredoxin 2 and Disulfide Binding proteins in E. coli

  • Wyns, Lode (Co-Promotor)
  • Roos, Goedele, (Administrative Promotor)

Project Details

Description

Introduction Reactions within redox systems are based on thiol-disulfide exchange between enzymes and/or pairs of cysteines within one protein. In Escherichia coli (E. coli), thioredoxins and Dsb-(DiSulfide Bindings) proteins are involved1. In a first part of this project, we will study the Zn2+-binding redox enzyme thioredoxin 2 (Ec_Trx 2) of E. coli. In the more elaborated second part, we will focus on oxidative folding in the periplasm of E. coli catalysed by Dsb-proteins. Thioredoxin 2 E. coli Trx2 (Ec_Trx2)2 is a member of the thioredoxin-family and has a reaction mechanism analogous to the previously characterised E. coli Trx1 (Ec_Trx1)3. Its conserved WCGPC active site motif switches continuously between the oxidized and reduced state. Ec_Trx2 binds Zn2+ with a very high affinity (Ka > 1018 M-1)4. The zinc atom is coordinated by the four cysteines of the two N-terminal CXXC motifs2. Circular dichroism and electrophoresis experiments suggest a major conformational change induced by oxidation of the Zn2+-binding site4. Until today, no native substrates of Ec_Trx2 are known. The structural organization of the Zn2+-binding site in respect to the active site has to be revealed. The central themes in this part of the project are the identification of the native substrates, the exact function of Zn2+, the structural and biophysical characterization of Ec_Trx2. Native substrates of Ec_Trx2 The native substrates of Ec_Trx2 will be identified by using an E. coli trxC- (gene name of Ec_Trx2) strain, in which the CPGS active site mutant will be expressed. Natural substrates will be trapped in disulfides, analyzed by Western blot and identified by mass spectrometry or N-terminal sequence analysis. In vitro, the interaction of Trx2 with the newly identified substrates will be tested in a kinetic cascade assay. Biological function of Zn2+ Research on heat shock protein Hsp33 suggests that structural, zinc-coordinating cysteine centers are able to serve as regulatory units. They activate the enzyme by conformational changes in response to oxidative stress5. Does the Zn2+-binding site of Ec_Trx2 serves as regulatory unit? Therefore, the activity of wild type and zinc-free Ec_Trx2, in which the cysteines of the Zn2+-bindingsite are replaced by serine (Ec_Trx2_Zn-), will be measured with native substrates in the presence of oxidative stress (H2O2,...). Structure Since all crystallization experiments failed, the NMR spectra to build the structure will be collected in collaboration with Dr. Jane Dyson (The Scripps research institute, La Jolla, USA). We will calculate the NMR structure. Influence of Zn2+ on pKa, redox potential and stability In collaboration with Dr. J.-F. Collet with who the research group of Dr. Joris Messens has created the "Brussels Center for Redox Biology" (http://redox.vub.ac.be/), the function of the Zn2+-ion will be investigated. The redox potential of the active site disulfide and the pKa of the nucleophilic cysteine will be determined6,7 with wild type Ec_Trx2 and Ec_Trx2_Zn-. With chemical and thermal unfolding experiments, the stability of reduced and oxidized Ec_Trx2 with and without Zn2+-binding site will be measured using differential scanning-calorimetrie (DSC) and spectroscopic techniques (fluorescence, circular dichroism). My quantum chemical expertise obtained during my PhD will be helpful to theoretically investigate redox potentials of thiol/disulfide redox couples. Descriptors like hardness8, softness8, electrophilicity9 and nucleofugality10 used in a HSAB-principle11 context and defined by conceptual density functional theory (DFT)11 will be applied to study the influence of the properties (hardness) of the present metal ion on the redox potential of disulfide formation. The influence of the surrounding ligands on the hardness of the central metal ion will be investigated, inspired on how ligands influence the stability of Re and Te complexes12. Oxidative folding A functional protein is more than simply ribosomal translated RNA. The polypeptide chain has to be correctly folded. The work of Christian Anfinsen on pancreas' ribonulease A has enlightened the relationship between the amino acid sequence and the protein conformation13. Despite more than 50 years intensive research, it is not known how the information embedded in the amino acid sequence is translated into correctly folded protein. Therefore, protein folding is considered as the last great problem of the molecular biology. A better understanding of the mechanism behind folding can reveal this mystery. In this second part of the project the focus is on oxidative folding. Oxidative folding refers to the composite process by which a protein recovers both its native disulfide bonds and native structure. In the periplasm of E. coli, the oxidative folding is directed by the oxidase DsbA and the isomerase DsbC (Figure). The themes of this section are the development of a new in vitro protein folding technology with RNase I as model system and the folding kinetics of RNase I Figure: DsbA introduces disulfides in proteins transported to the periplasm14. DsbC corrects malformed disulfides15. DsbA is re-oxidized by DsbB. The transported electrons will finally reduce O2. After the isomerisation reaction, oxidized DsbC is reduced by DsbD16, a transmebrane protein that shuttles electrons from the cytoplasm to the periplasm. The shuttling mechanism of DsbD is not known. Structural characterization of RNase I As a new model protein to study oxidative protein folding, RNase I is chosen. It is an endogenous periplasmatic E. coli protein with three consecutive and one nonconsecutive disulfide bond17. The protein structure has recently been solved in our lab17 at a resolution of 1.4 Å. A question to be solved is how the active site of RNase I recognizes RNA as substrate. To reveal the crucial residues for substrate interaction, the crystal structure of RNase I in complex with an oligo-deoxynucleotide as substrate analogue will be solved (X-ray data set available). Mutations of these residues will be made and the KM will be measured with RNA as substrate18. In vivo folding of RNase I To determine the essential Dsb-proteins in vivo, RNase I will be expressed in wild type E. coli and in E. coli strains in which the Dsb-proteins are eliminated (dsbA-, dsbC-, dsbA-C-, and dsbA-D-). On the periplasmatic extracts of these cells, the RNase I activity will be spectrophotometrically measured with intercalated methylene blue RNA as substrate20. To answer this question correctly, it will be necessary to check whether all native disulfides have to be present to ensure catalytic activity. Therefore, the four disulfides present in native RNase I will be eliminated one by one by mutating the cysteines of the respective disulfides to serines. In vitro folding of RNase I The ideal folding conditions (redox potentials, pH, protein concentration, salt,...) of RNase I will be determined by incubating denaturated RNase I (dRNase I) or RNase I from Inclusion Bodies (IB_RNase) in the presence of Dsb-proteins (DsbA, DsbC). dRNase I will be prepared from wild type RNase I after reduction (with DTT) and unfolding (with GdnHCl). IB_RNase will be prepared by expressing RNase I without the signal peptide. The IB_RNase fraction will contain protein aggregates that are partially structured 20. In order to develop a general applicable folding method for cyteine rich proteins for which the current expression technology fails or results in low yield, the optimal conditions for RNase I folding will be used to rescue these proteins from IB. Folding intermediates The folding pathway of RNase I will be elucidated by capturing and characterizing the intermediates along the folding pathway. Are non-native disulfides involved? In which order are disulfides introduced? What is the stability of the folding intermediates? dRNase I will be refolded in the presence and the absence of Dsb-proteins to study the catalyzed and uncatalyzed folding. In function of the folding time, the folding intermediates will be trapped by blocking the disulfide formation21, separated with high-resolution separation techniques (HPLC or 2D-electrophoresis), and identified by mass spectrometry identification. As such, the nature of the disulfide intermediates during the folding will be determined. Further, we will measure catalytic activity and thermodynamic stability of the major intermediates. Folding kinetics The kinetics of both the catalyzed and uncatalyzed folding will be determined by measuring the RNase I activity in function of the folding time in a stopped/quenched-flow experiment22,23,24. The problem of possibly active intermediates will be bypassed by fluorescence and/or circular dichroism experiments22,23,24. Fluoresence experiments might require the presence of a fluorophore in the RNase I. This will offer perspectives for the development of a method to study protein folding in vivo with fluorescence microscopy. The stopped-flow circular dichroism experiments will be performed in collaboration with Dr. André Matagne (ULG, Centre for protein engineering).
AcronymFWOTM451
StatusFinished
Effective start/end date1/10/0731/10/14

Keywords

  • chemistry

Flemish discipline codes

  • Biological sciences