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D 1282 OULU 2015 UNIVERSITY OF OUL U P.O. Box 8000 FI-90014 UNIVERSITY OF OULU FINLA ND U N I V E R S I TAT I S O U L U E N S I S ACTA A C TA D 1282 ACTA U N I V E R S I T AT I S O U L U E N S I S Van Dat Nguyen University Lecturer Santeri Palviainen Postdoctoral research fellow Sanna Taskila Van Dat Nguyen Professor Esa Hohtola MECHANISMS AND APPLICATIONS OF DISULFIDE BOND FORMATION Professor Olli Vuolteenaho University Lecturer Veli-Matti Ulvinen Director Sinikka Eskelinen Professor Jari Juga University Lecturer Anu Soikkeli Professor Olli Vuolteenaho Publications Editor Kirsti Nurkkala ISBN 978-952-62-0724-7 (Paperback) ISBN 978-952-62-0725-4 (PDF) ISSN 0355-3221 (Print) ISSN 1796-2234 (Online) UNIVERSITY OF OULU GRADUATE SCHOOL; UNIVERSITY OF OULU, FACULTY OF BIOCHEMISTRY AND MOLECULAR MEDICINE; BIOCENTER OULU D MEDICA ACTA UNIVERSITATIS OULUENSIS D Medica 1282 VAN DAT NGUYEN MECHANISMS AND APPLICATIONS OF DISULFIDE BOND FORMATION Academic dissertation to be presented with the assent of the Doctoral Training Committee of Health and Biosciences of the University of Oulu for public defence in Auditorium F101 of the Department of Physiology (Aapistie 7), on 6 February 2015, at 12 noon. U N I VE R S I T Y O F O U L U , O U L U 2 0 1 5 Copyright © 2015 Acta Univ. Oul. D 1282, 2015 Supervised by Professor Lloyd Ruddock Reviewed by Professor Neil Bulleid Doctor Renaud Vincentelli Opponent Professor Jan Riemer ISBN 978-952-62-0724-7 (Paperback) ISBN 978-952-62-0725-4 (PDF) ISSN 0355-3221 (Printed) ISSN 1796-2234 (Online) Cover Design Raimo Ahonen JUVENES PRINT TAMPERE 2015 Nguyen, Van Dat, Mechanisms and applications of disulfide bond formation. University of Oulu Graduate School; University of Oulu, Faculty of Biochemistry and Molecular Medicine; Biocenter Oulu Acta Univ. Oul. D 1282, 2015 University of Oulu, P.O. Box 8000, FI-90014 University of Oulu, Finland Abstract About one-third of mammalian proteins are secreted proteins and membrane proteins. Most of these proteins contain disulfide bonds in their native state, covalent links formed between the thiol groups of cysteine residues. In many proteins, disulfide bonds play an essential role in folding, stabilizing structure and the function of the protein. Therefore, understanding the pathways of disulfide bond formation is crucial for a wide range of medical processes and therapies. Disulfide bond formation is catalyzed by the Protein Disulfide Isomerase (PDI) family. To date the mechanisms of the PDIs in disulfide bond formation and pathways for disulfide bond formation have not been fully characterized. Here the structure of the substrate binding b’x domain of human PDI was determined. The structure shows that the b' domain has a typical thioredoxin fold and that the x region can interact with the substrate binding site of the b' domain. Specifically, the x region of PDI can adopt alternative conformations during the functional cycle of PDI action and that these are linked to the ability of PDI to interact with folding substrates. In addition, this study showed that two human proteins, GPx7 and GPx8 are involved in disulfide bond formation. The addition of GPx7 or GPx8 to a folding protein along with PDI and peroxide allows the efficient oxidative refolding of a reduced denatured substrate protein. Finally, this thesis includes the development of a system for the efficient production of disulfide bond containing proteins in the cytoplasm of E. coli. It showed that the introduction of Erv1p, a sulfhydryl oxidase and FAD-dependent catalyst of disulfide bond, allows the formation of native disulfide bonds in the cytoplasm of E. coli even without the disruption of genes involved in disulfide bond reduction. Introduction of Erv1p and a disulfide isomerase, e.g. PDI, allows the efficient formation of natively folded eukaryotic proteins with multiple disulfide bonds in the cytoplasm of E. coli. This system is able to express high levels of complex disulfide bonded eukaryotic proteins. Keywords: disulfide bond formation, Erv1p, GPx7, GPx8, hydrogen peroxide and protein disulfide isomerase Nguyen, Van Dat, Disulfidisidosten muodostumisen mekanismit ja sovellukset. Oulun yliopiston tutkijakoulu; Oulun yliopisto, Biokemian ja molekyylilääketieteen tiedekunta; Biocenter Oulu Acta Univ. Oul. D 1282, 2015 Oulun yliopisto, PL 8000, 90014 Oulun yliopisto Tiivistelmä Noin kolmasosa kaikista nisäkkäiden proteiineista on solun ulkopuolelle eritettäviä proteiineja ja kalvoproteiineja. Monet näistä proteiineista sisältävät natiivissa konformaatiossaan disulfidisidoksia, jotka ovat kovalenttisia sidoksia kysteiinitähteiden tioliryhmien välillä. Useissa proteiineissa näillä disulfidisidoksilla on keskeinen rooli proteiinin laskostumisessa, kolmiulotteisen rakenteen stabiloinnissa sekä proteiinin toiminnassa. Disulfidisidosten muodostumisen taustalla olevien mekanismien tunteminen onkin tärkeää monien lääketieteellisten prosessien ja hoitomenetelmien kannalta. Disulfidisidosten muodostumista katalysoivat proteiinidisulfidi-isomeraasi (PDI) -perheeseen kuuluvat entsyymit. PDI entsyymien toimintamekanismeja ja disulfidisidosten muodostumisen reaktioreittejä ei kuitenkaan vielä tunneta tarkasti. Tässä väitöskirjassa selvitettiin ihmisen PDI entsyymin substraattia sitovan b’x alayksikön rakenne. Rakenteesta voidaan todeta b’ alayksikön laskostuminen tyypilliseen tioredoksiini muotoon sekä x alueen interaktio b’ alayksikön substraattia sitovan kohdan kanssa. PDI entsyymin katalysoiman reaktioketjun aikana x alayksikkö voi muuttaa konformaatiotaan mahdollistaen PDI entsyymin interaktion laskostuvien substraattiproteiinien kanssa. Tässä tutkimuksessa osoitettiin myös kahden ihmisen proteiinin, GPx7 ja GPx8 osallistuminen disulfidisidosten muodostumista katalysoiviin reaktioihin. GPx7 ja GPx8 entsyymien lisäys laskostumisreaktioon yhdessä PDI:n ja vetyperoksidin kanssa mahdollistaa pelkistetyn, denaturoidun substraattiproteiinin tehokkaan, hapettaviin reaktioihin perustuvan uudelleenlaskostumisen natiiviin muotoonsa. Osana tätä väitöstutkimusta kehitettiin menetelmä, joka mahdollistaa disulfideja sisältävien proteiinien tehokkaan tuoton E.colin solulimassa. Menetelmässä sulfhydryylioksidaasina ja FAD:sta riippuvana disulfidisidosten muodostumisen katalysaattorina toimiva Erv1p mahdollistaa disulfidisidosten muodostumisen E.colin solulimassa myös ilman solun pelkistävien reaktioreittien geneettistä poistamista. Erv1p yhdessä disulfidi-isomeraasin, kuten PDI, kanssa mahdollistaa oikein laskostuneiden, useita disulfidisidoksia sisältävien eukaryoottisten proteiinien tehokkaan tuotannon E.colin solulimassa. Menetelmällä pystytään tuottamaan suuria määriä monimutkaisia disulfidisidoksellisia proteiineja. Asiasanat: disulfidisidos, vetyperoksidi Erv1p, GPx7, Gpx8, proteiinidisulfidi-isomeraasi, To my family 8 Acknowledgements The present thesis was carried in the research group of Professor Lloyd Ruddock, Biocenter Oulu, Faculty of Biochemistry and Molecular Medicine, University of Oulu and was supported by Academy of Finland, Department of Biochemistry, Biocenter Oulu, Oulu University Scholarship Foundation, University of Oulu. First of all, I would like to express my sincere gratitude to my supervisor Professor Lloyd Ruddock for giving me the great opportunity to learn and explore science in his research group. His outstanding supervision and his endless supports and encouragements were crucial to the progress of this thesis. I would like to thank Professor Kalervo Hiltunen, the head of the department for providing well facilitated research environment. All members of Department of Biochemistry are warmly acknowledged for creating such a good working atmosphere at the department. Special thanks to Professor Prof. Rikkert K. Wierenga and Dr. Antti Haapalainen for teaching me protein crystallography. I also would like to thank Professor Lars Ellgaard who was my external supervisor. I would like to thank the co-authors of the publications that form part of this thesis. Many thanks to my thesis committee group members: Professor Olli Vuolteenaho, Dr. Ilya Skovorodkin, Dr. Jingdong Shan, Dr. Jussi Vuoristo, Dr. Pekka Kilpeläinen. I would like thank to the current and former members of the LR-group: Kirsi Salo, Dr. Anna-Kaisa Lappi, Anna-Riikka Karala, Dr. Mirva Saaranen, Heli Alanen, Dr. Feras Hatahet, Dr. Annamari Ruddock, Dr. Irina Raykhel, Kati Korhonen, Marja Luukas, Herkko Sikkila for being very good friends and colleagues. Special thanks to Anna-Kaisa Lappi and Kirsi Salo for their encouragements and for pushing me forward with thesis writing. I would like thank to Dr. Emmy Verschuren and Dr. Liisa Kauppi for their encouragements during thesis writing. I am deeply grateful to my parents, my parents in law, my brother and my sister to their love and encouragements. Finally, My deepest gratitude I want to express to my wife Quynh and our beloved sons Truong, Nhat, Quoc. Helsinki, December 2014 Van Dat Nguyen 9 10 Abbreviations aa ACN Arg Asp BPTI CCD Cys D DHA DNA DNTB DTT EDTA ER FAD Glu Grx GSH GSSG IAA kDa LB Lys M MIMS MHC NNADPH PCR PDB PDI Q R ROS Amino acid Acetonitrile Arginine Aspartic acid Bovine pancreatic trypsin inhibitor Carboxy (terminal)Circular dichroism Cysteine Aspartic acid Dehydroascorbate Deoxyribonucleic acid 5,5-dithio-bis(2-nitrobenzoic acid) Dithiothreitol Ethylenediamidetetraacetic acid Endoplasmic reticulum Flavin adenine nucleotide Glutamic acid Glutaredoxin Reduced glutathione Oxidized glutathione Iodoacetamide Kilo Dalton Luria-Bertani Lysine Methionine Mitochondrial intermembrane space Major histocompatibility complex Amino (terminal)Nicotinamide adenine dinucleotide phosphate Polymerase chain reaction Protein data bank Protein disulfide isomerase Glutamine Arginine Reactive oxygen species 11 S SDS-PAGE Trx UV V W X 12 Serine Sodium dodecyl sulphate polyacrylamide gel electrophoresis Thioredoxin Ultraviolet Voltage Tryptophan Any amino acid List of original publications This thesis is based on the following publications, which are referred throughout the text by their Roman numerals: I Nguyen VD*, Wallis K*, Howard MJ, Haapalainen AM, Salo KE, Saaranen MJ, Sidhu A, Wierenga RK, Freedman RB, Ruddock LW, Williamson RA (2008) Alternative conformations of the x region of human protein disulphide-isomerase modulate exposure of the substrate binding b' domain. J Mol Biol 383(5):1144-55. II Nguyen VD, Saaranen MJ, Karala AR, Lappi AK, Wang L, Raykhel IB, Alanen HI, Salo KE, Wang CC, Ruddock LW (2011) Two endoplasmic reticulum PDI peroxidases increase the efficiency of the use of peroxide during disulfide bond formation. J Mol Biol 406(3):503-15. III Hatahet F, Nguyen VD, Salo KE, Ruddock LW (2010) Disruption of reducing pathways is not essential for efficient disulfide bond formation in the cytoplasm of E. coli. Microb Cell Fact 9(67):1-9. IV Nguyen VD, Hatahet F, Salo KE, Enlund E, Zhang C, Ruddock LW (2011) Preexpression of a sulfhydryl oxidase significantly increases the yields of eukaryotic disulfide bond containing proteins expressed in the cytoplasm of E. coli. Microb Cell Fact 10(1):1-13. * equal contribution 13 14 Contents Abstract Tiivistelmä Acknowledgements 9 Abbreviations 11 List of original publications 13 Contents 15 1 Introduction 17 2 Review of the literature 19 2.1 Protein folding......................................................................................... 19 2.2 Oxidative folding of proteins .................................................................. 21 2.3 Disulfide bond formation in the endoplasmic reticulum ......................... 22 2.3.1 Structure and function of human protein disulfide isomerase family members ........................................................... 23 2.3.2 Reactions of protein disulfide isomerase ...................................... 29 2.3.3 Disulfide bond generation in the endoplasmic reticulum ............. 31 2.4 Disulfide bond formation in the mitochondrial inter-membrane space ........................................................................................................ 37 2.5 Disulfide bond formation in the prokaryotic periplasm .......................... 39 2.6 Disulfide bond formation in the engineered cytoplasm of E. coli ........... 41 3 Aims of the present work 43 4 Materials and methods 45 5 Results 47 6 Discussion 49 6.1 Structure of the substrate binding domain of human protein disulfide isomerase and the functional significance of a novel structural interaction between the b’ domain and the x linker ................ 49 6.2 GPx7 and GPx8 are involved in disulfide bond formation in the endoplasmic reticulum ............................................................................ 51 6.3 Applications of disulfide bond formation ............................................... 54 7 Conclusions 57 References 59 Original articles 75 15 16 1 Introduction Disulfide bonds are covalent linkages formed between two cysteine residues in proteins as a result of oxidation reactions. Disulfide bonds are a crucial post translational modification for most secreted proteins and outer membrane proteins. Most of them require disulfide bonds for their stability, folding, and function. Catalysis of disulfide bond formation naturally occurs in the endoplasmic reticulum (ER) of eurokaryotes, the inter-membrane space of mitochrondria, and the periplasm of prokaryotes. These compartments have optimal environments, catalysts, and oxidation pathways for folding of native disulfide bond-containing proteins. Broadly speaking two steps are involved in native disulfide bond formations: i) oxidation reactions to form disulfide bonds and ii) subsequent rearrangement or isomerization of incorrect or non-native disulfide bonds to obtain native disulfide bonds. Native disulfide bond formation in human cells is catalyzed by the PDIfamily of proteins which contains 20 members. The archetypal member, PDI (PDIA1), is the most studied member in the family. It is a multifunctional enzyme whose most well-characterized function is as a catalyst of native disulfide bond formation. PDI contain four domains plus two short regions, whose architecture is defined: a-b-b’-x-a’-c. a and a’ are catalytic domains containing active sites, the b and b’ domains lack active sites and are non-catalytic, x is a linker region and c is a C-terminal acidic tail. The b’ domain is essential for substrate binding of PDI. It is needed for catalysis of protein disulfide isomerization and for non-covalent ligand binding. Ero1 proteins are also key factors in disulfide bond formation pathways in the ER. Ero1 works in conjunction with PDI and generates hydrogen peroxide as a by-product when producing a disulfide bond. The hydrogen peroxide produced could become a toxic product to the cell if it accumulates. However, potentially hydrogen peroxide can be used to generate disulfide bonds. Hence possibly, Ero1 can generate two disulfide bonds in each catalytic cycle. The question is what is the pathway in the ER that can convert hydrogen peroxide to disulfide bonds most efficiently? Native disulfide bond formation is often the rate-limiting step in protein folding in vitro and in vivo. Hence the production of recombinant disulfide bonded proteins is expensive on an industrial scale. In the present study a number of parallel approaches were adopted to understand the mechanisms of disulfide bond formation in the ER and to apply 17 knowledge of natural mechanisms for the efficient production of disulfide bond containing proteins in the cytoplasm of prokaryotes. Specifically, to examine in more detail the mechanism of action of PDI in catalyzing folding of disulfide bond-containing proteins a fluorescence screening of 75 mutants of the b’x domain was carried out and mutants that stabilize the b’x domain in different conformational forms were identified. The crystal structure of the I272A mutant of b’x domain was reported in this study, a structure that showed the x linker caps the substrate binding site of the b’ domain. This report also presents evidence that the two conformers are also significant in full-length PDI. In addition, this thesis showed that two ER resident proteins, GPx7 and GPx8 together with PDI form an efficient pathway to generate disulfide bonds from hydrogen peroxide in the ER. Finally, the present study innovated new efficient methodologies for making disulfide bond containing proteins by introduction of Erv1p, along with a catalyst of disulfide bond isomerization, into the cytoplasm of E. coli. 18 2 Review of the literature 2.1 Protein folding Proteins are made by a concerted process, with DNA being transcribed to mRNAs and mRNAs translated into the amino acid sequence of proteins. The sequence of a protein determines its native structure and function (Anfinsen CB 1973). In nature, proteins are synthesized as linear polypeptides, the so called primary structure, and then fold into their final secondary, tertiary, and quaternary native structure depending on the primary structure of protein and on its biophysical environment. Protein folding is driven by a number of non-covalent interactions such as hydrogen bonding involving the polypeptide backbone or polar side groups, electrostatic or ionic interactions between charged side groups, Van der Waals forces between all atoms, and hydrophobic packing of nonpolar side groups to exclude water. After folding the number of hydrophobic residues exposed on the surface of a protein is normally minimized. Hence a correctly folded protein usually has a closely packed hydrophobic core and hydrophilic residues, such as charged and polar side chains, displayed on the surface of protein (Pace et al. 1996). In the cell, the process of folding often initiates cotranslationally, so that the N-terminus of the protein begins to fold while the Cterminus of the protein is still being synthesized by the ribosome (Alberts et al. 2002). Across different species proteins fold in multiple cellular compartments including the cytoplasm, the ER, the intermembrane space and matrix of mitochondria and chloroplasts, and the extracytoplasmic space (e.g. the periplasm). In eukaryotic cells, more than a half of the total proteins are folded in the cytoplasm. This compartment, in both eukaryotes and prokaryotes, does not have mechanisms for oxidative folding i.e. the formation of disulfide bonds between the side chains of cysteine residues. In contrast, lysosomal proteins, membrane proteins and secreted proteins are mainly folded in the secretory pathway, specifically in the ER, in which oxidative folding does occur, in addition to posttranslational modifications such as N-glycosylation. Both disulfide bond formation and N-glycosylation make proteins more stable (Wickner & Schekman 2005). Proteins form into oligomers either after monomers of proteins have been folded completely into native domains, or when the subunit is not properly folded (Hurtley & Helenius 1989, Hartl & Hayer-Hartl 2009). Some large complex proteins continue folding events after they have exported from the ER, for 19 example the Von Willebrand Factor (Sadler et al. 2009), mucins etc (Adler et al. 2013). A cell may contain hundreds of thousands of proteins. For example the human cell has more than twenty thousand protein-coding genes (Pennisi et al. 2012) and alternate splicing, alternate initiation of translation and post translational modifications further increase the diversity. The total number of proteins in a human cell is estimated to be between 250,000 to one million (http://proteomics.cancer.gov/whatisproteomics). These proteins need to fold in a complex environment, for example the protein concentration in the cytoplasm has been estimated at being around 100 mg/ml (Zeskind et al. 2007). Inside the cell, many proteins are synthesized simultaneously by ribosomes at very high speed. A single eukaryotic protein is produced at a speed of 2-8 residues per second (Schroder & Kaufman 2005, Kleizen et al. 2005), while in prokaryotes the rate of transcription (around 60 nucleotides per second) is linked to the rate of translation (around 20 amino acids per second) (Alberts et al. 2002). During folding, the exposure of hydrophobic side-chains of newly synthesized polypeptides has to be minimized otherwise inappropriate aggregative events will occur (Pace et al. 1996). Aggregation is an autocatalytic event, which will have severe deleterious effects on the cell – including cell death. Hence protein synthesis and folding in the cell needs to be well organized and well controlled, so that proteins are folded accurately and efficiently. In cells the folding of proteins is assisted by pathways which control the speed of protein folding, and chaperones that prevent unwanted interactions between nascent proteins and hence avoid aggregation of newly synthesized proteins. Cells also have protein folding quality control pathways to make sure that all proteins are folded correctly before they reach their destinations where they act. Some proteins require posttranslational modifications such as disulfide bond formation, phosphorylation, glycosylation, etc, before attaining their native, functional state. Proteins that do not reach their native structure have to be refolded or degraded. The cell needs to maintain the balance between protein synthesis, protein folding and protein degradation/ turnover to avoid accumulation of protein aggregates caused by excess unfolded protein that leads to cell stress and disease. For example, accumulation of aggregated proteins leads to amyloidrelated illnesses such as Alzheimer’s (Harper et al. 1997, Knowles et al. 2014) and familial amyloid cardiomyopathy or polyneuropathy (Hund et al. 2002), prion infections (Burny et al. 1997), Huntington's and Parkinson's disease (Jucker & Walker 2011). Similarly a high rate of degradation causes a number of 20 proteopathy diseases such as antitrypsin-associated emphysema (Silverman & Sandhaus 2009), cystic fibrosis (Ratjen & Döring 2003) and the lysosomal storage diseases (Hollak & Wijburg 2014). 2.2 Oxidative folding of proteins Oxidative folding is the formation of disulfide bonds between cysteine residues in proteins, i.e. the formation of a covalent linkage between two thiol groups. This is a reversible reaction (Fig. 1). The disulfide bond is strong, but still it is a relatively weak covalent bond. Disulfide bonds play an important role in the folding and stability of some proteins, usually secreted proteins, outer-membrane proteins, and lysosomal proteins (Gross et al. 2002, Winther & Thorpe 2014). It is predicted that the majority of proteins which enter the eukaryotic secretory pathway are disulfide bonded proteins (Chen et al. 2005). Oxidative folding occurs in the periplasmic space of bacteria and in the ER, chloroplasts, and mitochondria of eukaryotic cells. It is also reported to occur in the cytoplasm of some extremophiles (Madonna et al. 2006, Limauno et al. 2014). Fig. 1. Schematic presentation of reduction, oxidation and isomerization reactions. Oxidative protein folding can be achieved by multiple pathways (Fig. 2) (Ruddock 2012, Oka & Bulleid 2013). For example, dithiol groups can be 21 oxidized directly by oxygen, or by low molecular weight biochemical compounds such as GSSG (Freedman 1995), by reactive oxygen species (ROS) (Cumming et al. 2004), by dehydroascorbate (DHA) (Saaraanen et al. 2010), by sulfhydryl oxidases such as the Ero1 family (via Protein Disulfide Isomerase (PDI); Frand et al. 1999) or QSox family (Alon et al. 2012), by Vitamin K linked oxidoreductases (Rutkevich & Williams 2012). Fig. 2. Disulfide bond formation pathways in the ER. Dotted arrows represent potential redox routes that had not been characterized experimentally in vivo before this study started. 2.3 Disulfide bond formation in the endoplasmic reticulum The ER is the second major organelle for protein folding in the eukaryote cell after the cytosol. One third of all human proteins are translocated to the ER, where they are folded before getting secreted through the Golgi apparatus (Chen et al. 2005). Proteins that fold in the ER are often modified by disulfide bond formation and glycosylation (Helenius & Aebi 2004). The ER has catalysts and chaperones that assist protein folding, including oxidative protein folding. It is estimated that there are about 1700 human proteins that are retained in the ER 22 (http://locate.imb.uq.edu.au/). The lumen of the ER has an optimal environment for oxidative folding of proteins and has efficient oxidative pathways to generate disulfide bonds including PDI-family members to assist native oxidative folding. 2.3.1 Structure and function of human protein disulfide isomerase family members PDI-family members are present in all eukaryotic species. To date, 20 members of the human PDI-family have been identified (Fig. 3). They are characterized by containing at least one thioredoxin fold containing domain, and having ER localization (Hatahet & Ruddock 2009). Most PDIs catalyze the formation, breakage and rearrangement of disulfide bonds, allowing proteins to quickly find the correct arrangement of disulfide bonds in their fully folded states; they might also have chaperone-like activities (Ellgaard & Ruddock 2005). Since there are only 20 PDI proteins, but about 7000 proteins are folded in the human ER, each member in the family probably catalyzes disulfide formation in multiple substrates and different family members may either play different roles in the process, or be involved in folding of different set of substrates i.e. have different substrate specificities. PDI-family members probably contribute to different stages of oxidative folding and work cooperatively to ensure the efficient production of multi-disulfide proteins in the ER. So far the functions of individual PDI-family members have not been fully characterized. The most well characterized protein of the PDI-family is PDI or P4HB. It comprises two homologous catalytic domains, a at the N-terminal and a’ at the Cterminal, which are connected by non-catalytic domains b and b’, The b’ domain is connected with the a’ domain by a 19 amino acid long linker named x. The protein also has a highly acidic C-terminal extension named c, and an ER retention motif at the C-terminal –KDEL (Ellgaard & Ruddock 2005). It is a highly abundant multifunctional enzyme. It was first shown to have oxidative protein folding activity 50 years ago (Goldberger et al. 1964). It is also the beta subunit of prolyl 4-hydroxylase (P4H), an enzyme that is involved in hydroxylation of prolyl residues in preprocollagen (Pihlajaniemi et al. 1987) and a subunit of microsomal triglyceride transfer protein complex (MTTP) (Pons et al. 2011). Other known functions include i) being a chaperone; ii) ER associated degradation (Molinari et al. 2002); iii) regulation of NAD(P)H oxidase (Janiszewski et al. 2005, Liberman et al. 2008). ; iv) PDI is involved in loading antigenic peptides into MHC class I molecules (Lee et al. 2009); v) PDI has been 23 shown to assist HIV entry by reducing a disulfide bond on the HIV gp120 protein during HIV infection of CD4 positive cells (Ryser et al. 2005); vi) PDI was identified to play an important role in glioma cell invasion, specifically deficiency of PDI inhibited tumor cell migration and invasion (Goplen et al. 2006). Fig. 3. The human PDI family (Hatahet & Ruddock 2009). Catalytic domains and noncatalytic domains with a thioredoxin fold are shown as elliptical circles, The active site motif is indicated in each catalytic domain. 24 Structures of the a and b, domains have been solved by NMR (Kemmink et al. 1996, Kemmink et al. 1999), an NMR structure of a’ domain has been deposited to the Protein Data Bank (PDB entry code: 1X5C) and, as part of this project, the structure of the b’x domain has been studied (Nguyen et al. 2008). All four domains have a thioredoxin fold. The catalytic a and a’ domains include the active site motif -CGHC-, that catalyzes disulfide reduction, oxidation, and isomerization reactions (section 2.3.2). The b’ domain plays a role in substrate recognition and binding (Klappa et al. 1998). The b domain plays a structural role, connecting and orientating the a and b’ domains. The a and a’ domains share nearly 37% identity with each other and are highly similar to thioredoxin. The structure of full length yeast PDI (Pdi1p) (Tian et al. 2006) and the structure of ERp57 in complex with tapasin (Dong et al. 2009) have been released. The structure of Pdi1p shows that the four domains of PDI (a, b, b’, a’) form an Ushape in which active site motifs of the a and a’ domains face each other. The structure of ERp57 shows high similarity to that of Pdi1p. Structures of other full length mature PDI family members have been solved including ERp44 (Wang et al. 2008) and ERp29 (Barak et al. 2009). The structure of AGR-3 has been deposited by the Ruddock group to Protein Data Bank (PDB entry code: 3PH9). AGR-3 displays a thioredoxin fold with an active site –CQYS- exposed on the surface of protein. The structure of the bb’ domains of ERp72 was published and shows significant similarity with the structure of the bb’ domains of ERp57 (Kozlov et al. 2009). Recently, structures of human PDI in an oxidized and reduced form have been characterized (Wang et al. 2013; Fig. 4). These show high similarity to the structure of Pdi1p and ERp57. Wang’s study showed that in the reduced PDI, domains a, b, and b' stay in the same plane, but the domain a' rotates ∼45° out, and the distance between the active sites is 27.6 Å. Whereas the four domains in the oxidized PDI stay in the same plane, and the distance between the active sites is 40.3 Å. Reduced PDI shows a closed conformation, whereas oxidized PDI exists in an opened conformation with more exposed hydrophobic areas and a larger cleft with potential for substrate binding. 25 Fig. 4. The overall structures of human PDI in oxidized form (4EL1), and in reduced form (4EKZ) (Wang et al. 2013). The structures show the a-b-b’-x-a’ domain architectures. The active site cysteines are shown in red, x linker is shown in blue. In both the reduced and oxidized forms, the a-b-b’-x-a’ domains are arranged as a horseshoe shape with two CGHC active sites in the a and a' facing each other. In the human PDI-family, Erp57, PDIp, and PDI share the same domain architecture of a-b-b’-a’-c, but appear to have significant other differences (Ellgaard & Ruddock 2005). For example, while PDI recognizes non-native proteins directly, ERp57 assists disulfide bond formation in glycoproteins by interacting with the ER-resident lectins calnexin or calreticulin (Leach et al. 2002, Urade et al. 2004). ERp57 also associates with Tapasin and is part of the major histocompatibility complex (MHC) class I peptide-loading complex, which is essential for formation and export of the antigen from the endoplasmic reticulum to the cell surface (Garbi et al. 2006, Peaper et al. 2005). ERp57 also interacts with another PDI-family member, ERp27 (Alanen et al. 2006), but the cellular function of this interaction and that of ERp27 is still unknown. Human PDIp has the same active site motif (CXXC) and domain architecture as human PDI and Erp57 (a-b-b’-a’) (Ellgaard & Ruddock 2005). It shares 46% amino acid sequence identity to human PDI (Desilva et al. 1997). PDIp is a glycoprotein specifically expressed in pancreatic acinar cells (Desilva et al. 1997), suggesting that PDIp may function as a protein-folding catalyst for secretory digestive enzymes (Fu et al. 2009). PDIp can bind 17β-oestradiol, and is reported to be the predominant intracellular oestrogen storage protein in the pancreas (Fu et al. 2012). 26 Erp72 is the second largest protein in the PDI family after ERdj5, and the only protein in this family that contains five thioredoxin-like domains with three catalytic domains, a structural domain and a substrate binding domain (Hatahet & Ruddock 2009). In addition, ERp72 contains an N-terminal stretch of 17 aspartate and glutamate residues, which is believed to bind Ca2+ (Van et al. 1993). ERp72 is localized in the ER, but also has been detected on cell membranes (Chen et al. 2008). ERp72 can catalyze thiol-disulfide exchange reactions (Alanen et al. 2003a). ERp72 is associated with several PDI family members such as PDI, P5, ERdj3 (Hatahet & Ruddock 2009). P5 or ERp5 is one of the less well studied member of the PDI family (Hatahet & Ruddock 2009). It contains two catalytic domains at N-terminal and a noncatalytic domain at C-terminal. It has been shown to be associated with Binding immunoglobulin protein (BiP) (Jessop et al. 2009), PDI and ERp72 (Stockon et al. 2003). ERp5 presents mainly on platelet intracellular membranes, but it is rapidly recruited to the cell surface in response to a range of platelet agonists (Jordan et al. 2005). ERp44 contains three thioredoxin like domain (a-b-b’), two non-catalytic domains and a catalytic domain with an unusual active site motif (CXXS) that lacks the C-terminal cysteine usually found in other family members (Anelli et al. 2002). The three domains (a-b-b′) of ERp44 form a V-shaped molecule (Wang et al. 2008), with a flexible C-terminal tail that could have a regulatory role in binding and releasing its substrate during the catalytic cycle (Wang et al. 2008). It is highly expressed in secretory tissues, and is localized mainly to the ER-Golgi intermediate compartment (Wang et al. 2007). This protein is associated with Ero1 (Anelli et al. 2003), and inositol 1,4,5-triphosphate receptor type I (Higo et al. 2005). ERdj5 is the largest protein in the PDI-family. It contains six thioredoxin like domains, five catalytic domains and a non-catalytic domain (Hatahet & Ruddock 2009). ERdj5 has been shown to have a reductase activity that cleaves the disulfide bonds of misfolded proteins. Hence it is involved in the degradation of misfolded ER proteins (Ushioda et al. 2008). ERdj5 catalyzes the removal of nonnative disulfides and correct folding of the LDL receptor (Oka et al. 2013). ERp29 or ERp28 has two domains, but lacks a catalytic domain (Ferrari et al. 1998, Barak et al. 2009). It is widely expressed (Demmer et al. 1997). The Nterminal domain of ERp29 is a b-domain like with thioredoxin fold, whereas the C-terminal domain shows an all α-helical fold (Barak et al. 2009). ERp29 has been shown to play a role in thyroglobulin processing and a role in the entry of 27 polyomavirus into cells (Rainey-Barger et al. 2007). This protein has been shown to promote ATF6 transport from ER to Golgi apparatus under ER stress conditions (Hirsch et al. 2014). ERp27 has only two non-catalytic domains, a b-like domain at the N-terminal of the protein and a b’-like domain at the C-terminal. This protein is up-regulated during ER stress. The b’-like domain of ERp27 can bind to peptides unfolded substrates (Alanen et al. 2006), but does not interact with folded substrates (Kober et al. 2013). The substrate-binding cleft of ERp27 on the b’-like domain is homologous to PDI, but is able to adapt in size and hydrophobicity (Kober et al. 2013). ERp27 interacts with ERp57 (Alanen et al. 2006) and presumably presents unfolded substrates to ERp57. ERp46 has three catalytic domains separated by long linkers, and forms an opened V-shape (Kojima et al. 2014). It plays a role in protecting cells against hypoxia (Sullivan et al. 2003). ERp46 can efficiently catalyze the formation of disulfide bonds but lacks the ability to introduce native disulfide bonds selectively (Sato et al. 2013). This protein probably acts during early oxidative protein folding events in the ER (Kojima et al. 2014). PDIr contains four domains, three C-terminal catalytic domains, and an Nterminal non-catalytic domain (Hayano et al. 1995). It has been shown to be involved in the folding of α1-antitrypsin and N-linked glycoproteins (Horibe et al. 2004). The non-catalytic domain of PDIr plays a key role in the recognition of protein partners and misfolded proteins (Vinaik et al. 2013). PDILT contains four thioredoxin like domains two of which have unusual active-site motifs SSKQS in the N-terminal domain a domain and WSKKC in the C-terminal a’ domain. This protein is expressed highly in postmeiotic male germ cells. PDILT associates with Ero1α (Van Lith et al. 2005), calmegin (Van Lith et al. 2007), and the testis-specific homologue of calnexin that is required for sperm fertility (Ikawa et al. 1997). Recently, it has been showed to interact with CALR3 in testicular germ cells and plays a crucial role in the disulfide formation in ADAM3 (Tokuhiro et al. 2012). ERp18 is one of the smallest of the human PDI-family members, containing a single catalytic domain with a CGAC active site motif (Alanen et al. 2003b). It shows low activity in catalyzing disulfide exchange reactions (Alanen et al. 2003b, Jeong et al. 2008). Erp18 has been shown to associate with hVKORC1 indicating its involvement in oxidative folding via vitamin K metabolic pathway (Schulman et al. 2010). 28 AGR-2 (anterior gradient homolog 2), and AGR-3 (anterior gradient homolog 3), both are small single-domain proteins that show sequence homology with ERp18, but they have very different active-site motifs, WCGAC (ERp18), ECPHS (AGR-2), and DCQYS (AGR-3). The crystal structure of AGR-3 has been solved (PDB entry code: 3PH9) and is highly similar to solution structure of ERp18 (PDB entry code: 2K8V) (Rowe et al. 2009). The functions of both proteins are not clear. However, AGR-2 has been shown to be localized within the ER of intestinal secretory epithelial cells and is essential for in vivo production of the intestinal mucin MUC2, a large cysteine rich glycoprotein that forms the protective mucus gel lining the intestine (Park et al. 2009). Transmembrane proteins (TMX), including TMX, TMX2, TMX3, TMX4, TMX5. TMX, TMX2, TMX4, TMX5 contain only one thioredoxin-like domain with unusual active site motifs while TMX3 contains three domains with a Nterminal catalytic domain (Hatahet & Ruddock 2009). TMX has been shown to associate with MHC class I heavy chain and protect it from degradation (Matsuo et al. 2009). Most of PDI family members except TMX and TMX4 were shown to form a mixed disulfide bond with Ero1α and Ero1β, indicating their involvements in oxidative folding (Benham et al. 2000, Appenzeller-Herzog et al. 2010). Association of TMX, TMX4 with hVKORC1 indicates their involvement in oxidative folding via vitamin K metabolism pathway (Schulman et al. 2010). 2.3.2 Reactions of protein disulfide isomerase The active sites (CXXC) of PDI naturally exist in multiple different states; a reduced dithiol state, an oxidized intramolecular disulfide state and an intermolecular mixed disulfide state. The reduced state can catalyze the reduction/isomerization of disulfide bonds in substrate proteins, while the oxidized state is able to catalyze oxidation of dithiols in substrates (Gane et al. 1995, Chivers et al. 1997). PDI may catalyze isomerization reactions through cycles of reduction and oxidation reaction, or directly though intramolecular disulfide rearrangement (Schwaller et al. 2003). In oxidation reactions (Fig. 5), two free thiols in substrate are oxidized by an oxidized active site of PDI to form a disulfide bond in the substrate and a reduced active site in PDI. In turn, PDI is re-oxidized to complete the catalytic cycle of PDI. In the ER the PDI is oxidized usually by Ero1 family, or oxidized glutathione, or DHA, or reactive oxygen species (see sections 2.2 and 2.3.3). 29 Fig. 5. Schematic presentation of the reactions of PDI. For clarity only one active site of PDI is shown. An oxidized PDI catalyses disulfide bond formation in a reduced substrate (oxidation); a reduced PDI reduces disulfide bonds in an oxidized substrate (reduction) or can catalyse native disulfide bond formation in an incorrectly folded substrate (isomerization). In reduction reactions (Fig. 5), a disulfide bond in substrate is reduced by the reduced active site of PDI which results in oxidation of the active site of PDI and the formation of two free cysteines in the substrate. In turn, PDI is reduced by reduced glutathione (GSH). Another function of PDI is to catalyze disulfide bond isomerization reactions (Fig. 5) in which disulfide bonds and free thiols in substrates are rearranged to gain native disulfide bonds. This reaction happens either via cycles of reduction 30 and oxidation reactions, or via a direct intramolecular disulfide rearrangement (Fig. 5). In the later, an incorrect disulfide bond in a misfolded protein is attacked by the N-terminal active site cysteine of PDI to form a mixed disulfide between a cysteine of the substrate and a cysteine of PDI’s active site. Next, the attack of a second cysteine residue from the substrate protein results in the formation of a more stable disulfide in the substrate protein. This is repeated until PDI is released. If at any time the C-terminal active site cysteine of PDI is the nucleophile, then the substrate undergoes a net reduction and PDI becomes oxidized and has to be reduced in order to play a catalytic role in reduction or direct isomerization. Disulfide bond formation depends heavily on the redox state and pH of the environment since the protonation state of thiol groups is heavily dependent on the pH of surroundings and only the deprotonated thiolate state is a good nucleophile. The pKa values of the active-site cysteine residues are very important for PDI activities. The redox potential of PDI depends on the pKa values of the two active site cysteine residues (Lappi et al. 2004). The pKa of the N-terminal cysteine in the active site is often very low and may not vary significantly during the catalytic cycle (Karala et al. 2010). In contrast, the pKa of the C-terminal cysteine is often high and may vary considerably during PDI’s reaction cycles. It has been shown that the pKa value of the C-terminal cysteine of a domain is modulated by the movement of the side chain of R120 (Lappi et al. 2004). This allow PDI to efficiently catalyze both oxidation and isomerization reactions (Karala et al. 2010). 2.3.3 Disulfide bond generation in the endoplasmic reticulum To date, several pathways for disulfide bond formation in the ER have been characterized (Fig. 2). The sulfhydryl oxidase Ero1 and PDI are thought to form the major pathway for protein disulfide bond formation in the ER (Fig. 6). Ero1 is a FAD-dependent sulfhydryl oxidase tightly associated with the luminal face of the ER membrane that uses molecular oxygen as an electron acceptor to oxidize itself. Oxidized Ero1 then transfers disulfide to folding secretory proteins via reduced PDIs (Frand et al. 1999, Benham et al. 2000). Ero1 family proteins are present in all eukaryotes. Many simple eukaryotes like invertebrates, fungi, and yeast encode a single Ero1, except S. pombe has two Ero1 isoforms (Frand et al. 1998, Kettner et al. 2004), whereas mammalian cells have two Ero1 family members, Ero1α, and 31 Ero1β (Cabibbo et al. 2000, Tu et al. 2000, Gross et al. 2006). The role of Ero1 in disulfide bond formation was first identified in yeast (Frand & Kaiser 1998, Pollard et al.1998). It has been shown that yeast Ero1p directly oxidizes Pdi1p, since it forms a disulfide bond with Pdi1p in vivo (Frand & Kaiser 1999). Yeasts that are mutated in Ero1p are extremely sensitive to the reducing agent DTT, whereas overexpression of Ero1p confers DTT resistance (Frand & Kaiser 1998, Pollard et al.1998). Later on, the roles of mammalian Ero1 family members, both Ero1α and Ero1β, in PDI oxidation in vivo were shown (Cabibbo et al. 2000, Pagani et al. 2000). Two mouse Ero1 isoforms have similar in vitro biochemical activities (Zito et al. 2010a). To date, no major differences in redox activities or substrate interactions have been identified between the Ero1 isoforms (Sevier 2010). However, there is different in tissue distribution between Ero1α and Ero1β. Ero1β presents selectively in the pancreas. Whereas, Ero1α appears in all tissues (Zito et al. 2010a). Mice lacking both Ero1α and Ero1β exhibit slightly decreased rates of disulfide bond formation (Zito et al. 2010a). A wide range of PDI family members were shown to associate with Ero1α and Ero1β (Benham et al. 2000, Appenzeller-Herzog et al. 2010). Fig. 6. Schematic representations for Ero1 activity. Activity of Ero1 controls redox homeostasis of the ER. Ero1 activity is regulated by the prescence of GSSG, oxidized PDIs and oxidized substrates in the ER. 32 Ero1 contains two pairs of catalytic cysteines, an active-site pair located at the C-terminus of Ero1 and a shuttle cysteine pair (Fig. 7) (Frand & Kaiser 2000). Ero1 uses molecular oxygen to oxidize its active site, which in turn transfers disulfide to the shuttle cysteine pair via internal dithiol-disulfide exchange, then the oxidized shuttle cysteine pair oxidizes reduced PDI which in turn oxidizes folding protein substrates (Frand & Kaiser 2000, Sevier & Kaiser 2006). Ero1 does not contain an ER retention signal, it is retained in the ER by associating with the PDI-family member ERp44 (Anelli et al. 2003). Ero1 activity is tightly regulated by non-catalytic regulatory cysteine pairs to control redox homeostasis of the endoplasmic reticulum (Sevier et al 2007, Appenzeller-Herzog et al. 2008, Tavender & Bulleid 2010a, Hansen et al. 2014). The regulatory disulfides are formed under oxidizing ER conditions and their formation is thought to limit the movement of the shuttle cysteines and hence limits the ability of Ero1 to shuttle disulfides to PDI. In contrast, the regulatory disulfides are reduced under reducing ER conditions leading to an increase of Ero1 activity (Sevier et al 2007, Appenzeller-Herzog et al. 2008, Hansen et al. 2014). Fig. 7. Schematic representation of the disulfide pattern in the active and inactive forms of human Ero1α with catalytic disulfides (solid lines) and regulatory disulfides (dashed lines) indicated (Appenzeller-Herzog et al. 2008, Inaba et al. 2010). Altogether the data suggests that Ero1 is an essential route for disulfide bond formation the ER of yeast, but in mammals Ero1 is non-essential (Zito et al. 2010a), but may still provide the major route for disulfide bond formation in the ER. What are the alternative routes for oxidative folding in the ER of mammalian cells? Another possible source for disulfide bond formation in the ER is via Quiescin Sulfhydryl Oxidases (QSOX) (Heckler et al. 2008). Like the Ero1 family they are FAD-dependent oxidases which can produce disulfides with the 33 corresponding reduction of molecular oxygen to hydrogen peroxide (Kodali & Thorpe 2010). QSOX family members are present in most eukaryotes, but absent in yeast and fungi (Thorpe et al. 2002, Kodali & Thorpe 2010). Human has two QSOX orthologs, QSOX1 and QSOX2. QSOX1 and QSOX2 share 41% identity. These proteins have been detected in the ER, the Golgi, and extracellularly, but are primarily in the Golgi apparatus and secreted fluids (Kodali & Thorpe 2010). QSOXs contain an FAD binding Erv/ALR sulfhydryl domain at the C-terminal, and a thioredoxin domain with a CXXC active site (Alon et al. 2012). The domain combination in QSOX family members makes them very efficient at catalyzing disulfide bond formation in vitro (Kodali & Thorpe 2010). QSOX can introduce disulfide bonds directly to reduced unfolded proteins much more efficiently than it does to reduced active site of PDI (Hoober et al. 1999, Thorpe et al. 2002). In vitro, QSOX cooperatively assist PDI in oxidative folding of disulfide bond-containing protein (Rancy & Thorpe 2008). Sulfhydryl oxidases including Ero1 and QSOX produce hydrogen peroxide as a by-product during their catalytic cycles, with one molecule of hydrogen peroxide being produced for each disulfide bond made. Hydrogen peroxide is considered to be a toxic product to the cells when it accumulates. A secretory plasma cell can generate 100000 disulfide bonds per second (Cenci et al. 2011), which would result in the rapid generation of high concentrations of peroxide. The ER does not have a catalase, nor a similar enzyme that could remove the peroxide, raising the question which mechanism is used to remove hydrogen peroxide produced in the ER during disulfide bond formation? In principal hydrogen peroxide can react directly with the free thiol group of cysteine in folding proteins or PDI to form cysteine sulfenic acid. In turn the cysteine sulfenic acid can react with another free thiol of cysteine to form a disulfide or react with another cysteine sulfenic acid to generate cysteine sulfinic acid (Fig. 8). Fig. 8. Formation of disulfide bonds by using peroxide. 34 In vitro it has been shown that hydrogen peroxide can be used to efficiently generate disulfide bonds in reduced folding substrates either directly or via PDI (Karala et al. 2009). Hence, in vivo, the ER may use hydrogen peroxide generated by Ero1 or QSOX to create disulfide bonds efficiently. If it is the case, for each catalytic cycle Ero1 will produce two disulfide bonds from a single oxygen molecule. That not only removes a potentially toxic byproduct, but is also more economical for the cell. Recently studies have shown that ER-localized peroxiredoxin IV can cooperate with PDI to convert hydrogen peroxide generated by Ero1 to disulfide bonds (Tavender et al. 2010, Zito et al. 2010b). Before the role of Ero1 in oxidative folding was discovered, oxidized glutathione (GSSG) had been considered the primary route for disulfide bond formation in the ER (Hwang et al 1992). Glutathione is the most abundant peptide in animal cells and exists primarily in a reduced state (GSH) or an oxidized form (GSSG). GSH is synthesized in cytoplasm, and imported into the ER by diffusion or active transport. In the ER, GSSG is generated from GSH by the oxidative folding machinery (Banhegyi et al. 1999, Appenzeller-Herzog 2011). GSH can act as an electron donor in the reduction of peroxides, which is catalyzed by glutathione peroxidases, or that of disulfides, a reaction catalyzed by glutaredoxins (Winterbourn & Metodiewa 1999). A deletion of glutathione leads to faults in native disulfide bond formation in the ER (Chakravarthi & Bullied 2004). GSH is required for the correct reduction or isomerization of non-native disulfide pairings during oxidative protein folding (Chakravarthi et al. 2006). The ratio and concentrations of GSH and GSSG determine the redox potential of intracellular compartments. GSH/GSSG forms the primary redox buffer in the ER (Gutscher et al. 2008). The ER is more oxidizing than the cytoplasm because of having higher levels of GSSG (Hwang et al. 1992, Fewell et al. 2001). GSSG is able to directly oxidize reduced PDI, or reduced folding proteins efficiently in vitro (Karala et al. 2009). All together the data suggests that glutathione plays an important role in oxidative folding in the ER, including in maintaining redox state of the ER. The metabolism of vitamin C and vitamin K are also considered as sources of oxidizing equivalents for disulfide bond formation in the ER. Dehydroascorbic acid (DHA) is an oxidized form of ascorbic acid (vitamin C). Cells contain ascorbic acid at millimolar concentration, including in the ER. It is a primary antioxidant in plasma and within cells. However, while DHA is actively imported into the endoplasmic reticulum of cells via glucose transporters, mainly by Glucose transporter 1 (Glut1), ascorbic acid cannot be imported (Welch et al. 35 1995), requiring the reduction of DHA to ascorbic acid in the ER lumen. This reaction is regarded to be catalyzed by several enzymes, including PDI (Ruddock 2012). However, recently studies in vitro has shown that DHA can react more rapidly with thiols in unfolded or partially folded proteins than with the thiols of reduced active site in PDI (Saaranen et al. 2010). DHA is able to react with thiols to form disulfide bonds and ascorbate. DHA can also be generated from the noncatalyzed antioxidant action of ascorbate in situ. Additionally, ascorbate in the ER reaches millimolar concentrations in most tissues (May 2012). Therefore, the ascorbate/DHA couple may be a significant route for disulfide bond formation in the ER (Zito et al. 2012). Another potential route for disulfide formation in the ER is via the vitamin K epoxide reductase (VKOR) family of proteins which are involved in vitamin K metabolism. Vitamin K is a class of closely related compounds. Vitamin K2 (menaquinone) is a cofactor of DsbB, a key disulfide bond producer in the bacterial periplasm (Kadokura & Beckwith 2010; section 2.5). VKOR family members are present in vertebrates, insects, plants, bacteria and archaea (Goodstadt & Ponting 2004). Bacterial VKOR has been shown to catalyze disulfide bond formation in the periplasm (Dutton et al. 2008). Mammals have two VKOR family members, VKORC1 and VKORCL1. Both VKORs are ERresident transmembrane proteins with cysteine active site residues pointing toward the ER lumen (Schulman et al. 2010). VKORC1 has been shown to be involved in γ-carboxylation of glutamate residues of some secreted proteins including blood clotting factors (Jin et al. 2007, Wallin et al. 2008). A CXXC motif of VKOR is oxidized during this process (Wajih et al. 2005). Recently VKOR has been shown to contribute to oxidation and disulfide bond formation within the ER (Rutkevich & Williams 2012). In contrast to VKORC1, VKORC1L1 has been poorly characterized to date. Altogether the data clearly show that the mammalian VKOR could function as per the bacterial homologue in disulfide bond formation. The question is, which protein in the ER can be a redox partner of VKOR? PDI was initially thought to be a partner since it has been shown to associate with VKOR (Wajih et al. 2007). However, a later study has shown that other PDI-family members, TMX, TMX4, and ERp18, but not PDI, crosslink to VKOR (Schulman et al. 2010). 36 2.4 Disulfide bond formation in the mitochondrial inter-membrane space While disulfide bond formation was originally thought to be limited to the ER in eukaryotes, more recently it has been identified that disulfide bond formation occurs in other eukaryotic cellular compartments as well, the best well characterized of which is the inter-membrane space of mitochondria (Riemer et al. 2009). Mitochondria primary function is in the production of cellular energy, and hence they are essential for cell growth and viability. More than 1400 proteins are present in human mitochondria (Taylor et al. 2003, http://locate.imb.uq.edu.au/). Of these only 13 proteins are encoded by mitochondrial DNA (Taylor & Turnbull 2005). Therefore most of proteins in mitochondria are encoded by nuclear DNA, are synthesized in the cytoplasm and then are imported into mitochondria. Proteins that are intended to be inside mitochondria normally have mitochondrial signal sequences at their N-terminal (Herrmann et al. 2000). The mitochondrial targeted proteins are recognized by TOM (Translocase of the Outer Membrane). TOM cooperates with TIM (Translocase of the Inner Membrane) to translocate proteins into the mitochondria (Koehler et al. 2004). Some proteins in mitochondria were identified to contain disulfide bonds (Mesecke et al. 2005, Webb et al. 2006, Herrmann & Kohl 2007). Hence mitochondria require machineries to assist oxidative folding (Fig. 9). 37 Fig. 9. Oxidative protein folding in the IMS of mammalian mitochondria. Reduced unfolded proteins from the cytosol are imported to the IMS via the TOM complex in an reduced unfolded state. Mia40 is oxidized by ALR and traps reduced unfolded imported proteins by disulfide links. The imported proteins then are released from Mia40 in an oxidized folded state. Electrons are delivered from the flavin cofactor of ALR to Cytochrome C and further into the respiratory chain. The intermembrane space (IMS) of mitochondria has a disulfide relay system that includes Mia40 (mitochondrial intermembrane space import and assembly protein 40) and Erv1p in yeast, or Mia40 and ALR (FAD-linked sulfhydryl oxidase) in mammals. This system promotes the import and folding of small cysteine-containing proteins (e.g. small TIM, Cox-17, Cox-19) into the IMS (Tokatlidis 2005, Daithankar et al. 2010). In the IMS, oxidized Mia40 forms mixed disulfide bridges with reduced newly imported proteins. Subsequent isomerization of these disulfide bridges permits the imported protein to be folded in the IMS (Allen et al. 2005, Mesecke et al. 2005). There are a number of similarities between oxidative protein folding in the ER and IMS. For example, Mia40 acts like PDI in that it catalyzes the formation and rearrangement of 38 disulfides, however it does not have thioredoxin folded domain, and it has CXC active site motif instead of CXXC (Banci et al. 2009). Erv1 and ALR are FADdependent sulfhydryl oxidases like Ero1. Also like Ero1 both contain two pairs of catalytic cysteines, an active site cysteine pair, and a shuttle cysteine pair. Erv1 transfers a disulfide to reduced Mia40 which in turn oxidizes reduced substrates, analogous to the Ero1-PDI-substrate protein relay in the ER (section 2.3.3). Like other sulfhydryl oxidases, Erv1 can use molecular oxygen to oxidize its active site and produce a hydrogen peroxide molecule. However, in vivo Erv1 activity is strongly associated with the respiratory chain of mitochondria, therefore preventing production of H2O2. In this pathway, electrons are passed from Erv1 to oxygen molecule via cytochrome c and cytochrome c oxidase (Bihlmaieret et al. 2007). 2.5 Disulfide bond formation in the prokaryotic periplasm In prokaryotes, the mechanism of oxidative folding is best characterized in the periplasm of gram-negative bacteria, especially E. coli. Most proteins in E. coli are folded in the cytoplasm, however some periplasmic and membrane proteins of E. coli are folded in the periplasm space. It is estimated that more than 250 proteins enter the periplasm of E. coli and gain disulfide bonds there (Dutton et al. 2008) e.g. PhoA, AppA. These proteins are synthesized in the cytoplasm, then are imported to the periplasm as reduced unfolded proteins – in an analogous manner to protein import and oxidative folding in the ER or IMS. Similarly, the periplasm has oxidative folding pathways to support oxidative folding of proteins. In the periplasmic space of E. coli native disulfide bonds are generated by two pathways; i) an oxidation pathway comprised of DsbB and DsbA and ii) an isomerization pathway linked to reduction pathways formed by DsbD, DsbC and DsbG (Ito et al. 2008) (Fig. 10). 39 Fig. 10. Pathways for disulfide bond formation in the periplasm of E. coli. DsbA is oxidized by DsbB. Oxidized DsbA oxidizes secretory proteins. DsbC and DsbG catalyst disulfide bond isomerization to form native disulfide bonds. DsbD shuttles electrons via thioredoxin from the cytoplasm to reduce DsbC and DsbG. DsbB is a transmembrane protein with four transmembrane helices. It has two critical cysteine pairs located in two periplasmic loops, the N-terminal active site cysteine pair C41/C44 and the C-terminal shuffle cysteine pair C104/C130 (Kobayashi et al. 2001). During the catalytic cycle of DsbB, the C-terminal shuttle cysteine pair is oxidized by the N-terminal active site cysteine pair, resulting in a reduced active site cysteine pair. The oxidized shuttle cysteine pair then transfers a disulfide to the soluble protein DsbA which in turn oxidizes reduced folding substrates (Kadokura & Beckwith 2002, Zhou et al, 2008). This is analogous to the mechanisms of disulfide bond formation in the ER and IMS (sections 2.3.3 and 2.4) i.e. the primary catalyst of disulfide bond formation does not act directly on folding substrates. Under aerobic conditions DsbB is oxidized by oxidized ubiquinone, which then is oxidized by respiratory chain cytochrome oxidases, finally cytochrome oxidases reduce oxygen. In anaerobic conditions DsbB is oxidized by menaquinone, which in turn is oxidized by fumarate reductase or nitrate reductase (Takahashi et al. 2004). The DsbB/DsbA system introduces disulfide bonds into folding proteins, however in many substrate proteins, especially those which contain nonsequential disulfides, the disulfide bonds introduced by DsbA may not be native 40 disulfides. Therefore, in order to reach the native state those incorrect disulfide bonds need to be rearranged by a disulfide isomerization pathway which consists of DsbC, DsbD, and DsbG. DsbC and DsbG act as disulfide isomerases. Both form a V-shaped homodimer with two catalytic domains facing each other (McCarthy et al. 2000, Heras et al. 2004). Each monomer consists of an N-terminal dimerization domain and a C-terminal catalytic domain with a thioredoxin fold and CXXC active site (McCarthy et al. 2000, Heras et al. 2004). Both DsbC and DsbG have a substrate binding cleft located between two arms of the V-shaped homodimer (Heras et al. 2004). However, the substrate binding cleft of DsbG is bigger than one of DsbC, suggesting that DsbG works on different substrates than DsbC (McCarthy et al. 2000, Heras et al. 2004). In the isomerization pathway, the reduced active site of DsbC or DsbG attacks an incorrect disulfide in a misfolded protein and forms a mixed disulfide between DsbC and folding protein. This is analogous to the isomerization reaction catalyzed by PDI, (see Fig. 5 and Fig. 10). In order to act as isomerases, DsbC and DsbG have to be kept in their reduced state. Oxidized DsbC and oxidized DsbG are reduced by the transmembrane protein DsbD (Katzen et al. 2000), which in turn is reduced by cytoplasmic thioredoxins. Thioredoxins are in turn reduced by cytoplasmic thioredoxin reductase (Goldstone et al. 2001, Cho & Beckwith 2009). Some bacterial species such as Mycobacterium tuberculosis and Streptomyces coelicolor, do not have DsbB in their periplasmic oxidative pathway although disulfide bond formation is detected in their periplasmic space. Instead their periplasmic spaces have bacterial homolog of the eukaryotic enzyme vitamin K epoxide reductase (VKOR) that can generate disulfide bonds (Dutton et al. 2008, Dutton et al. 2010). 2.6 Disulfide bond formation in the engineered cytoplasm of E. coli Naturally the cytoplasm of most bacteria has to be maintained as a reducing environment to support specific reactions, e.g. reaction of ribonucleotide reductase which catalyzes the reduction of ribonucleotides to deoxyribonucleotides (Elledge et al. 1992). The cytoplasm of E. coli contains two main pathways (Fig. 11) that reduce disulfides, the thioredoxin (Trx) / thioredoxin reductase (TrxR) pathway and the glutathione / glutaredoxin (Grx) / glutathione reductase (GSR) pathway (Prinz et al. 1997). 41 Fig. 11. Disulfide reducing pathways of the cytoplasm in E. coli. NADPH reduces TrxR which in turn reduces the thioredoxins TrxA and TrxC. Reduced thioredoxins reduce disulfide bonds in substrate proteins e.g. RNR. NADPH is also used to reduce glutathione reductase which in turn reduces oxidized glutathione (GSSG) to reduced glutathione (GSH). GSH can either directly reduce disulfide bonds or can reduce thioredoxins or glutaredoxins which in turn reduce disulfide bonds in substrates (Ritz & Beckwith 2001). E .coli has been widely used to produce recombinant proteins both on a laboratory and an industrial scale. However, many proteins of interest to both academics and for biopharmaceutical companies contain disulfide bonds. Production of disulfide containing proteins is not possible in the normal cytoplasm of E. coli. However, it has been shown that the disruption of both main reducing pathways in the cytoplasm through a knockout of glutathione reductase (gor) and thioredoxin reductase (trxB) combined with a lethality suppressor mutation in peroxiredoxin alkyl hydroperoxidase (ahpC) leads to a significant increase in the production of active disulfide bonded proteins such as vtPA, a protein that contains 9 predominatly non-sequential disulfide bonds (Bessette et al. 1999). Although those genetic modified strains can produce significant yields of active disulfide bonded proteins in their cytoplasm, the level and the quality of disulfide bonded protein production is far from that required by industry, probably due to the lack of an active catalyst of disulfide bond formation in this system. Therefore, for the cytoplasm of E. coli to be used as an efficient cell factory for the production of disulfide bond containing proteins a better system is needed. 42 3 Aims of the present work The aims of the thesis were to study mechanism of action of human PDI in disulfide bond formation, disulfide bond formation pathways in the ER and to generate an efficient system to produce disulfide bond containing proteins. My specific goals were: 1. 2. 3. to identify protein structure of the substrate binding domain (b’x) of PDI. to characterize the function of two ER resident glutathione peroxidases (GPx7 and GPx8), and their involvement in disulfide bond formation pathways. to develop systems for efficient folding of disulfide bonded proteins in the cytoplasm of E. coli using current knowledge of disulfide bond formation pathways. 43 44 4 Materials and methods Detailed description of the materials and methods used in this study are found in the original articles I, II, III, IV. Method Original article Vector construction II, III, IV Protein expression I, II, III, IV Protein purification I, II, III, IV Mutagenesis I Spectroscopy I NMR spectroscopy I Crystallization, Data collection, structure determination and refinement I, II Subcellular localization II GPx activity measurements II BPTI refolding II Kinetics of GPx8 oxidation followed by stopped-flow fluorescence II Bimolecular fluorescence complementation II Oxygen consumption assay II Glutathione peroxidase activity measurements II Protein analysis and activity measurements from lysates III, IV Analysis of purified proteins IV 45 46 5 Results Detailed description of the results in this study are found in the original articles I, II, III, IV. Result Original article Preparations of recombinant human PDI b’x contain at least two conformers I Screening mutations that trap a single conformation of PDI b’x I NMR studies of PDI b’x constructs I Crystallography of mutant PDI b’x I Evidence for alternative conformations of the x region in full-length PDI I Human GPx7 and GPx8 are ER-resident proteins II Stopped-flow kinetics for the peroxide oxidation of GPx8 II GPx7 and GPx8 are PDI-peroxidases and not glutathione peroxidases II GPx7 and GPx8 combined with PDI are efficient for peroxide-mediated oxidative protein II folding Crystal structure of human GPx8 II GPx7 and GPx8 interact with Ero1α in vivo and modulate Ero1α activity in vitro II Alkaline phosphatase production III Phytase production III Production of a tissue plasminogen activator fragment IV Production of a tissue plasminogen activator fragment fusion protein IV Development of a pre-expression vector for Erv1p and a disulfide isomerase IV Media effects on the yield of active vtPA IV Production of other eukaryotic proteins with multiple disulfide bonds IV 47 48 6 Discussion 6.1 Structure of the substrate binding domain of human protein disulfide isomerase and the functional significance of a novel structural interaction between the b’ domain and the x linker Human PDI is a key enzyme in the ER in the catalysis of disulfide bond formation in secretory proteins. PDI contains four domains a, b, b’, and a’, an x linker between domains b’ and a’. When this study was carried out, structures of full length human PDI and of the b’ domain had not been determined, only the structure of the individual thioredoxin domains a, b, and a’ had been solved by NMR (Kemmink et al. 1996, Kemmink et al. 1999, unpublished PDB entry code: 1X5C). The structure of full length yeast Pdi1p was published while this work was ongoing (Tian et al. 2006). The structure of full-length human PDI and that of the b’ domain by crystallization and NMR had not been successfully determined probably due to the high conformational flexibility of these proteins. The goal of my research was to study the structure of the b’ domain of PDI. This study should lead to increased understanding of the mechanisms of action of PDI as b’ is the major substrate binding domain of PDI, and the only domain whose structure had not previously been solved. In this study we showed that the b’x exists in at least two conformational states, which clearly differ in the structural relationship between the x region and the b’ domain (paper I). We hypothesized it might be possible to generate mutants that favorably stabilize one state and abolish the conformational heterogeneity and by doing so to overcome the complexities introduced by the existence of these states. Mutations that stabilize one state of conformation may also be suitable for structural studies. To screen for mutants that have single conformational state we used Trp fluorescence of purified recombinant mutants. Our study demonstrated that recombinant b’x exists in two distinct conformations that differ in tertiary structure, specifically in the environment of the unique W347 residue in the x region. Mutants that well adopted capped and uncapped conformation were identified. The I272A mutant of b’x has been crystallized and the structure determined. In the crystal structure of b’x I272A, the x linker caps the b’ domain and the side chain of W347 is buried in a hydrophobic pocket in the b’ domain. The structure of b’ is highly similar to the structure of b’ domains of 49 other PDI-family members, including the bb’ fragment of human ERp57 (PDB entry code: 2H81), the b’ domain of yeast Pdi1p (PDB entry code: 2BE5), the bb’ fragment from human PDI (PDB entry code: 2K18) and the b’ domain from the thermophilic fungus Humicola insolens (PDB entry code: 2DJK). While the crystal structure of b’x was being refined, a structure of full-length yeast PDI (Pdi1p) was released, in which the four thioredoxin domains a, b, b’, a’ form an U-shape structure (Tian et al. 2006). The structure of Pdi1p displays a highly hydrophobic pocket on the surface of the b’ domain and there are also hydrophobic patches on the surface of a, b, and a’ domains. These hydrophobic patches on a, b, and a’ domains and the hydrophobic pocket on the b’ domain form a continuous hydrophobic surface on PDI, which is thought to be important for substrate recognition of PDI. In the crystal structure one molecule of Pdi1p is occupying the central cavity of the U-shape structure of another Pdi1p molecule i.e. it is acting like a substrate. Hence the yeast structure represents a conformation in which a protein ligand is bound, displacing the x linker. The b’ domains of human PDI and yeast PDI show a similar structure fold and homologous hydrophobic pocket. However, one significant difference between the human and yeast PDI structures is in the orientation of the x region, that in the x region caps the b’ domain of the human PDI, but the x region of Pdi1p does not cap the hydrophobic pocket of the b’ domain (I Fig. 6). The crystal structure of human PDI-family member (ERp44) also was published while this work was ongoing (PDB entry code: 2R2J) (Wang et al. 2008). The ERp44 structure displayed three thioredoxin domains with the architecture a-b-b’ and with a Cterminal tail which caps a hydrophobic pocket in the b’ domain. This is highly similar to the x region capping the b’ domain of human PDI (I Fig. 7), in particular, L361 in ERp44 interacts with the hydrophobic pocket of b’, as W347 does in PDI. Previous studies have identified mutations that inhibit substrate binding of b'x or full-length PDI using a combination of modelling and mutagenesis studies (Klappa et al. 1998, Pirneskoski et al. 2004). However, the binding site identified by the previous studies does not match that identified in our b’x crystal structure or by b’ of Pdi1p (Tian et al. 2006). All of the mutations that were previously reported to inhibit substrate binding instead shift the equilibrium towards the capped conformer. They are not located in the hydrophobic pocket of the b’ crystal structure. Therefore, these mutations rather act indirectly to inhibit substrate binding by decreasing the accessibility of the substrate binding site. Analogous to this, studies on ERp44 showed the role of C-terminal tail in 50 regulating the activity of ERp44, in particular deletion of the C-terminal tail in ERp44 increased the activity of ERp44 and its ability to act as a molecular chaperone (Wang et al. 2008). The conformational exchange, which caps the substrate binding site, does not only happen in the isolated b’ domain. Studies of mutants in full length PDI demonstrated that a similar conformational exchange happens in the full-length protein with the x region capable of interacting reversibly with the principal substrate binding site on b’ domain (paper I). The x region can adopt different conformations and can potentially exchange between the capped conformation and the uncapped conformation. In other words, the substrate binding site is unavailable to bind substrates when it is capped by x, so that inhibits the activities of PDI towards protein substrates. The conformational exchange in the b' domain can be modulated by the addition of peptide ligands that can compete with x binding at the substrate binding site (Byrne et al. 2009). Recently, the crystal structures of hPDI containing all four domains (abb'xa') in both the reduced and oxidized states of the active sites (Wang et al. 2013) have been published. Those studies reveal a closed conformation of reduced hPDI, and an open conformation of oxidized hPDI with more exposed substrate binding sites. The redox-regulated conformational changes in hPDI are mainly located in the b’xa’ region. The flexibility of this region is mainly dependent on the x linker. The x linker in the crystal structure of reduced hPDI is close to the b’ domain; it is highly similar to our b’x I272A structure. The C-terminal half of the x linker and the a’ domain in oxidized hPDI twists about 45o in comparison to that of the reduced hPDI. Hence the x linker is required for conformational changes of PDI, which in turn depends on the redox-state of PDI. Altogether the published data suggests that the x region of PDI can adopt alternative conformations during the functional cycle of PDI and that these are critical for access to the substrate binding site. 6.2 GPx7 and GPx8 are involved in disulfide bond formation in the endoplasmic reticulum Currently the major route for disulfide bond formation in the ER is thought to be via Ero1 family members. This route, in which the flow of oxidizing equivalents is from molecular oxygen to Ero1 family members and from these to PDI and from PDI to substrates, is highly regulated (Sevier et al. 2007, AppenzellerHerzog et al. 2008, Baker et al. 2008). Regulation of Ero1 activity is important to 51 keep the optimal redox environment for oxidative folding of proteins. Specifically, oxidative folding requires not only oxidation, but also isomerization and reduction of incorrect disulfide bonds to reach the native state. If the redox environment is too oxidizing isomerization and reduction are inhibited as both need PDI in the reduced state. Each catalytic cycle, Ero1 family members generate one disulfide bond and one molecule of hydrogen peroxide (Gross et al. 2006). Hydrogen peroxide production as a by-product would result in cellular oxidative stress unless it is utilized efficiently or degraded. The ER does not have a recognizable catalase or a similar enzyme for peroxide degradation. In addition, It has been shown that disulfide bond formation does not induce oxidative stress, but newly synthesized misfolded proteins in the ER lumen, activates the unfolded protein response (UPR), causes oxidative stress, and induces apoptosis in vitro and in vivo (Malhotra et al. 2008). These combined suggest peroxide may be used functionally. A previous study from our group using an in vitro BPTI refolding assay showed that peroxide can efficiently oxidize dithiols in folding proteins to the native state under physiologically relevant conditions. These studies demonstrated that no significant oxidative side reactions occur so long as the peroxide concentration is kept at submillimolar concentrations (Karala et al. 2009). This productive oxidative folding can occur either via the direct reaction of peroxide with a free thiol group on the folding protein to form a cysteine sulfenic acid with subsequent formation of a disulfide bond or via a reaction with a thiol group in the active site of PDI and subsequent transfer of the disulfide made from PDI to the substrate protein. Therefore, potentially the ER should have pathways that can use hydrogen peroxide to generate disulfide bonds. If it is the case, two disulfide bonds can be generated per molecule of molecular oxygen with no net production of harmful species. While the previous in vitro studies showed that peroxide could be used to make native disulfide bonds, the kinetics of the process were relatively slow and required relatively high concentrations of peroxide (up to 0.5mM) to be efficient. To be utilized in vivo a catalyst of this process might be expected. The results from the study reported here (paper II) demonstrate a novel pathway for utilizing the peroxide generated by Ero1 for native disulfide bond formation. Two human glutathione peroxidase (GPx) family members, GPx7 and GPx8 were identified as containing sequences that directed them to the ER and ER retention signals. Both 52 GPx7 and GPx8 were shown to be localized in the ER, with GPx7 being a soluble protein and GPx8 being an ER transmembrane protein. Mature purified recombinant GPx7 or the luminal domain of GPx8 both gave only a marginally faster reaction than the non-catalyzed reaction in the coupled GPx assay, indicating that neither GPx7 nor GPx8 are efficient glutathione peroxidases. An amino acid sequence alignment of the human GPx family and comparison of the GPx7 protein structure (unpublished PDB entry code: 2P31) and our GPx8 crystal structure (PDB entry code: 3KIJ) with the GPx1 structure (unpublished PDB entry code: 2F8A) shows that GPx7 and GPx8 lack the loop that determines glutathione specificity in the glutathione peroxidase family (II Fig. 1). This suggests that both may be members of the thioredoxin peroxidase or TGPx family (Toppo et al. 2008). To date, the PDI-family contains the only known thioredoxin superfamily proteins that are present in the ER. Therefore, GPx7 and GPx8 may be PDI peroxidases that catalyze oxidation of reduced PDI by peroxide. Indeed, this study (paper II) demonstrates that both proteins have PDI peroxidase activity in vitro and hence presumably in vivo. The addition of either GPx7 or GPx8 to a mixture of reduced denatured BPTI, PDI, and peroxide increases the rate of oxidative folding of BPTI. Additionally, by monitoring oxygen consumption in vitro this study indicates that the presence of GPx7 and GPx8 increases Ero1α activity, suggesting that these PDI peroxidases interact with Ero1 and could directly utilize peroxide generated by Ero1. Our data (paper II) also shows interaction between GPx7 and GPx8 with Ero1α in the ER. Furthermore, a recent study shows that GPx7 promotes oxidative protein folding in the ER by directly utilizing Ero1α-generated H2O2. H2O2 oxidizes Cys57 of GPx7 to sulfenic acid, which can be resolved by Cys86 of GPx7 to form an intramolecular disulfide bond. In turn both intramolecular disulfide form and sulfenic acid form of GPx7 can oxidize PDI (Wang et al. 2014). Additionally, GPx8 has recently been shown to rapidly clear hydrogen peroxide produced by Ero1α, hence it prevents leakage of hydrogen peroxide from the ER (Ramming et al. 2014). GPx8 was identified as a cellular substrate of the hepatitis C virus (HCV) NS3-4A protease, and it is cleaved by the HCV NS3-4A protease at Cysteine 11 (Morikawa et al. 2014). This study also shows that GPx8 is involved in HCV particle production, but not in HCV entry or RNA replication. Recently, human peroxiredoxin IV (Prx IV), an ER localized protein was shown to protect cells by removing peroxide produced during disulfide bond formation (Tavender & Bulleid 2010b, Zito et al. 2010b, Tavender et al. 2010) with data suggesting that Prx IV may provide a parallel pathway to GPx7 and 53 GPx8 for peroxide utilization. However, the oxidation of a substrate protein to the native state by Prx IV + PDI + peroxide in vitro (Zito et al. 2010b) is significantly slower than that reported in this study (Paper II) for GPx7 or GPx8 + PDI (II Fig. 5d and f). Even though the assay systems were not same, this suggests that the PDI peroxidases are at least as efficient as Prx IV at converting peroxide into disulfide bond formation and their association with Ero1 suggests that they may provide the major route for peroxide utilization. All together strongly suggest that PDI peroxidases and Prx IV effectively utilize any peroxide produced during disulfide bond formation for disulfide bond formation in the ER. 6.3 Applications of disulfide bond formation Industrial recombinant protein production is a multi-billion dollar a year market. Many of the FDA approved biologics are disulfide bonded proteins, including anticancer drugs, insulin, interleukins, interferons, growth factors, antibody and antibody fragments etc and the number of such products is growing. The most common route for the industrial scale production of proteins that contain disulfide bonds was to express these proteins in the cytoplasm of E. coli as inclusion bodies, since it is low cost and high yield. The insoluble proteins generated are then refolded in vitro to get correct folded products. However this step is often inefficient, time consuming and costly. E. coli can naturally make disulfide bonded proteins in the periplasm (Collet et al. 2002). However, the volume of periplasm is rather small compared to the volume of cytoplasm, typically comprising approximately 10% of the total cell volume (Ghuysen & Hakenbeck 1994). Furthermore, the secretion and folding machinery can be easily saturated. This means that yields of proteins produced in the periplasm are often 10-fold or more lower than equivalent yields in the cytoplasm. The problems with the high level production of disulfide bond containing proteins in E. coli has led to a shift in the past decade towards the use of eukaryotic production systems such as CHO cells (Palomares et al. 2004). While the ease of use has increased and associated costs of using eukaryotic cells has decreased significantly over recent years, they are still more costly, more difficult to scale up and less regulatory authority friendly than the use of E. coli. To get around the limitations of the use of E. coli, researchers have been trying to create E. coli strains that can produce native disulfide bonded proteins in the cytoplasm. The cytoplasm of E. coli has been commonly used to produce 54 recombinant proteins at industrial and laboratory scales, but it is not suitable for the direct production of disulfide bonded proteins as it lacks enzyme systems to generate disulfide bonds. While strains have been produced which lack the reducing pathways normally found in the cytoplasm and which can produce much higher levels of disulfide bonded proteins than wild-type strains (Bessette et al. 1999), the yields and/or quality are often not optimal even when a disulfide isomerase is co-expressed. In contrast, eukaryotic cells have organelles in which disulfide bond formation occurs, such as the ER (Hatahet & Ruddock 2009) and mitochondrial inter-membrane space (Herrmann et al. 2007). These both have catalytic systems that use sulfhydryl oxidases to make disulfide bonds. The sulfhydryl oxidases use molecular oxygen to catalyze the oxidation of a dithiol to a disulfide. Hence we thought, it might be possible to create an efficient system to produce disulfide bonded recombinant proteins in the cytoplasm of E. coli by introducing sulfhydryl oxidases into this compartment. For proteins that contain a few disulfide bonds such as PhoA and AppA, co-expression of the sulfhydryl oxidase Erv1p gave yields of active disulfide-bonded protein produced up to 1000-fold over the wild-type E. coli control. Moreover, the production of active PhoA and AppA is more effective than the production in ∆gor ∆trxB background strain (paper III). For proteins that are more difficult to be folded correctly because they contain many disulfide bonds or which contain non-sequential disulfide bonds, such as vtPA and AppA, may require a catalyst of the other step of native disulfide bond formation, disulfide isomerization e.g. the use of PDI or DsbC. In addition to the requirement for catalysts of disulfide bond formation and isomerization, proteins like vtPA which contain a large number of disulfide bonds may require longer for the system to catalyze the correct folding of those proteins. This in turn may lead to the expressed proteins forming inclusion bodies rather than soluble natively folded states due to the aggregation process being faster than the folding process. To avoid the issues associated with folding intermediate aggregation, highly soluble fusion proteins, e.g. MBP, have been widely use to increase the solubility of the folding intermediates. The use of a MBP fusion protein, in conjunction with catalysts of disulfide bond formation and isomerization, significantly increases the yields of disulfide bonded proteins in our system (paper IV). Furthermore, pre-expression of the folding factors so they are available before the protein of interest is expressed, and the correct choice of media also result in higher yields of active complex disulfide bonded proteins. In combination, yields of active vtPA about 800 fold higher than previously reported 55 using DsbC co-expression in a Δgor ΔtrxB strain were obtained in our system (paper IV). However, homogeneous natively folded vtPA is not observed. Other disulfide bonded proteins such as resistin, BPTI, CSF3 and BMP4 have also been produced by this system (paper IV). Those proteins were expressed in high yields of soluble, and/or active proteins. However, only proteins with a few disulfide bonds were produced homogenously in high yields in this system. The most complex protein we were able to produce to near homogeneity during this work was Resistin that contains 5 intramolecular disulfide bonds per monomer plus an inter-molecular disulfide bond between subunits in the dimer. Coexpression of Erv1 in E. coli cytoplasm also significantly increases the yield and quality of expressed Camelidae single domain antibodies (Veggiani et al. 2011). Even though our system is significantly better than Δgor ΔtrxB strain in the production of disulfide bonded proteins, it still needs to be optimized for proteins which contain a high number of disulfide bonds, especially a high number of non -sequential disulfide bonds. Subsequent optimization with the group has significantly improved the processes and now the system is able to generate g/L amounts of homogeneously folded disulfide bond containing eukaryotic proteins in the cytoplasm of E. coli in fermentation (HGH, scFv, interleukins; unpublished results) and more than 100mg/L of correctly folded scFv and Fab antibody fragments from shake flasks (unpublished results). These optimized systems are all based on the work undertaken as part of this study (papers III and IV). 56 7 Conclusions Human PDI is a key enzyme involved in disulfide bond formation in the ER. Structural information will be very useful to understand the mechanisms of action of the PDI. Prior to this work the structure of three domain (a, b, a’) were solved, but the structure of the substrate binding b’x domain was not. It was shown that the b’x is not suitable for structural studies due to the flexibility of the x linker. Using a screen of 75 mutants, this study has identified mutants of b’x that exclusively adopt one of these alternative conformations. The crystal structure of one of these mutants, I272A, was determined. This structure reveals a hydrophobic pocket in the b’ domain which is capped by the x region. This study also showed that the two conformers are also significant in full-length PDI. These alternative conformations are linked to the ability of PDI to interact with folding substrates and hence to the mechanisms of action of this folding catalyst. Pathways for disulfide bond formation in the ER are not yet fully understood. Until recently the Ero1 family has been considered to be the main source of oxidation in the ER. These proteins produce disulfide bonds and hydrogen peroxide as a byproduct. This raises the question: what pathways the ER uses to remove hydrogen peroxide efficiently, as it will cause cell toxicity if it accumulates. This thesis has characterized two ER resident proteins, GPx7 and GPx8, that are involved in eliminating hydrogen peroxide. Human GPx7 and GPx8 were noted as secreted GPxs. This study showed that they are actually located in the ER and associate with the peroxide-producing Ero1α. Furthermore, neither has significant GPx activity, but instead have PDI peroxidase activity. It is shown in this thesis that the addition of PDI and GPx7 or GPx8 and peroxide to a reduced unfolded protein results in the very rapid achievement of the folded disulfide-bonded state. Furthermore, the addition of GPx7 to a mixture of purified Ero1α, PDI, and GSH resulted in a significant increase in oxygen consumption by Ero1α. Hence, these proteins, together with PDI, form an efficient way to prevent hydrogen peroxide accumulation in the ER by utilizing hydrogen peroxide to form a disulfide bond in substrate proteins. In parallel to these studies on understanding the mechanisms of disulfide bond formation in vivo, knowledge of these pathways was applied to the production of disulfide bond containing proteins. The cytoplasm of E. coli contains disulfide reducing pathways. Hence production of disulfide bond containing proteins in the cytoplasm of E. coli results in non-disulfide bonded proteins that normally form insoluble aggregates. It has been noted previously 57 that the disruption of at least one of the reducing pathways in cytoplasm of E. coli is essential for the efficient generation of disulfide bonds in proteins (Bessette et al. 1999). The study reported here has developed a system that showed efficient disulfide bond formation can be gained by exogenous expression of Erv1p in E. coli cytoplasm. This study refutes the current paradigm in the field that disruption of at least one of the reducing pathways is essential for the efficient production of disulfide bond containing proteins in the cytoplasm of E. coli and open up new possibilities for the use of E. coli as a microbial cell factory. The addition of PDI helps to increase the efficiency of production of multi-disulfide bonds containing proteins, in particular those containing non-sequential disulfide bonds. The yield of disulfide bond containing protein production is further increased significantly by fusion of the targeted protein to a highly soluble fusion protein, e.g. MBP, and by manipulating growth conditions. Various disulfide bond containing proteins that contain from one to multi-disulfide bonds have been successfully produced using this technology. 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III Hatahet F, Nguyen VD, Salo KE, Ruddock LW (2010) Disruption of reducing pathways is not essential for efficient disulfide bond formation in the cytoplasm of E. coli. Microb Cell Fact 9(67):1-9. IV Nguyen VD, Hatahet F, Salo KE, Enlund E, Zhang C, Ruddock LW (2011) Preexpression of a sulfhydryl oxidase significantly increases the yields of eukaryotic disulfide bond containing proteins expressed in the cytoplasm of E. coli. Microb Cell Fact 10(1):1-13. Reprinted with permission from Elsevier (I, II) and BioMed Central (III, IV). Original publications are not included in the electronic version of the dissertation. 75 76 ACTA UNIVERSITATIS OULUENSIS SERIES D MEDICA 1267. Kiviniemi, Marjo (2014) Mortality, disability, psychiatric treatment and medication in first-onset schizophrenia in Finland : the register linkage study 1268. Kallio, Merja (2014) Neurally adjusted ventilatory assist in pediatric intensive care 1269. Väyrynen, Juha (2014) Immune cell infiltration and inflammatory biomarkers in colorectal cancer 1270. Silvola, Anna-Sofia (2014) Effect of treatment of severe malocclusion and related factors on oral health-related quality of life 1271. Mäkelä, Tuomas (2014) Systemic transplantation of bone marrow stromal cells : an experimental animal study of biodistribution and tissue targeting 1272. Junttila, Sanna (2014) Studies of kidney induction in vitro using gene expression profiling and novel tissue manipulation technique 1273. Häkli, Sanna (2014) Childhood hearing impairment in northern Finland : prevalence, aetiology and additional disabilities 1274. Lehto, Liisa (2014) Interactive two-step training and management strategy for improvement of the quality of point-of-care testing by nurses : implementation of the strategy in blood glucose measurement 1275. Harjunen, Vanessa (2014) Skin stem cells and tumor growth : functions of collagen XVIII in hair follicle cycling and skin cancer, and Bmx tyrosine kinase in tumor angiogenesis 1276. Savela, Salla (2014) Physical activity in midlife and health-related quality of life, frailty, telomere length and mortality in old age 1277. Laitala, Anu (2014) Hypoxia-inducible factor prolyl 4-hydroxylases regulating erythropoiesis, and hypoxia-inducible lysyl oxidase regulating skeletal muscle development during embryogenesis 1278. Persson, Maria (2014) 3D woven scaffolds for bone tissue engineering 1279. Kallankari, Hanna (2014) Perinatal factors as predictors of brain damage and neurodevelopmental outcome : study of children born very preterm 1280. Pietilä, Ilkka (2015) The role of Dkk1 and Wnt5a in mammalian kidney development and disease 1281. Komulainen, Tuomas (2015) Disturbances in mitochondrial DNA maintenance in neuromuscular disorders and valproate-induced liver toxicity Book orders: Granum: Virtual book store http://granum.uta.fi/granum/ D 1282 OULU 2015 UNIVERSITY OF OUL U P.O. Box 8000 FI-90014 UNIVERSITY OF OULU FINLA ND U N I V E R S I TAT I S O U L U E N S I S ACTA A C TA D 1282 ACTA U N I V E R S I T AT I S O U L U E N S I S Van Dat Nguyen University Lecturer Santeri Palviainen Postdoctoral research fellow Sanna Taskila Van Dat Nguyen Professor Esa Hohtola MECHANISMS AND APPLICATIONS OF DISULFIDE BOND FORMATION Professor Olli Vuolteenaho University Lecturer Veli-Matti Ulvinen Director Sinikka Eskelinen Professor Jari Juga University Lecturer Anu Soikkeli Professor Olli Vuolteenaho Publications Editor Kirsti Nurkkala ISBN 978-952-62-0724-7 (Paperback) ISBN 978-952-62-0725-4 (PDF) ISSN 0355-3221 (Print) ISSN 1796-2234 (Online) UNIVERSITY OF OULU GRADUATE SCHOOL; UNIVERSITY OF OULU, FACULTY OF BIOCHEMISTRY AND MOLECULAR MEDICINE; BIOCENTER OULU D MEDICA
