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ISBN 978-952-62-2846-4 (Paperback)ISBN 978-952-62-2847-1 (PDF)ISSN 0355-3191 (Print)ISSN 1796-220X (Online)
U N I V E R S I TAT I S O U L U E N S I SACTAA
SCIENTIAE RERUM NATURALIUM
U N I V E R S I TAT I S O U L U E N S I SACTAA
SCIENTIAE RERUM NATURALIUM
OULU 2021
A 754
Antti Moilanen
NOVEL REGULATORY MECHANISMS AND STRUCTURAL ASPECTS OF OXIDATIVE PROTEIN FOLDING
UNIVERSITY OF OULU GRADUATE SCHOOL;UNIVERSITY OF OULU,FACULTY OF BIOCHEMISTRY AND MOLECULAR MEDICINE
A 754
AC
TAA
ntti Moilanen
ACTA UNIVERS ITAT I S OULUENS I SA S c i e n t i a e R e r u m N a t u r a l i u m 7 5 4
ANTTI MOILANEN
NOVEL REGULATORY MECHANISMS AND STRUCTURAL ASPECTS OF OXIDATIVE PROTEIN FOLDING
Academic dissertation to be presented with the assentof the Doctoral Training Committee of Health andBiosciences of the University of Oulu for publicdefence in the Leena Palotie auditorium (101A) of theFaculty of Medicine (Aapistie 5 A), on 15 February2021, at 12 noon
UNIVERSITY OF OULU, OULU 2021
Copyright © 2021Acta Univ. Oul. A 754, 2021
Supervised byProfessor Lloyd Ruddock
Reviewed byProfessor Carolyn SevierDoctor Adam Benham
ISBN 978-952-62-2846-4 (Paperback)ISBN 978-952-62-2847-1 (PDF)
ISSN 0355-3191 (Printed)ISSN 1796-220X (Online)
Cover DesignRaimo Ahonen
PUNAMUSTATAMPERE 2021
OpponentProfessor Neil Bulleid
Moilanen, Antti, Novel regulatory mechanisms and structural aspects of oxidativeprotein folding. University of Oulu Graduate School; University of Oulu, Faculty of Biochemistry andMolecular MedicineActa Univ. Oul. A 754, 2021University of Oulu, P.O. Box 8000, FI-90014 University of Oulu, Finland
Abstract
Efficient maturation of disulfide-containing proteins in the endoplasmic reticulum (ER) requiresfacilitated folding, aided by various folding catalysts and chaperones. As protein folding is acomplex process, proteins may, even in the presence of endogenous folding factors, formmisfolded states. Therefore, the cell has acquired several layers of quality control mechanisms tokeep the folding load in the ER in balance with the folding capacity. The activity of foldingcatalysts is regulated to avoid stress connected to hyperoxidation. Other quality controlmechanisms include the unfolded protein response (UPR) and ER-associated protein degradation(ERAD). The UPR consists of signaling cascades that sense unfolded protein stress and upregulatefolding factors and other components to relieve the stress. ERAD directly removes non-nativeproteins.
The main folding catalysts that introduce disulfide bonds to folding proteins in the ER aremembers of the ER oxidoreductin 1 (Ero1) family and protein disulfide isomerases (PDI). Ero1 isa sulfhydryl oxidase that reduces molecular oxygen to hydrogen peroxide to form disulfides. Thenewly generated disulfides are then transferred during the catalytic cycle to PDI which catalyzesdisulfide formation in folding proteins. The Ero1-PDI pathway has been studied extensively bycell-based techniques, and its activity is regulated in a redox dependent manner by rearrangingregulatory disulfides. However, the kinetic aspects of these regulatory functions have remainedpoorly characterized.
In this study, we report comprehensive kinetic measurements for the Ero1-PDI pathway thatreveal novel regulatory mechanisms for oxidative protein folding (OPF). These include amechanism consisting of high affinity and apparent cooperativity for substrate oxygen to allowEro1 to function efficiently at hypoxic conditions. We also show that different human Ero1isoforms have differential dependence for substrate PDI with one of the isoforms, Ero1α, showingstrong dependence for PDI for both redox activation as well as the catalytic cycle. This observationallowed us to describe a potential crosstalk between OPF and ERAD, in which OPF by the Ero1α-PDI pathway is downregulated, to potentially increase the efficiency of ERAD which requiresreduced protein substrates. This work is concluded by a structural study of a novel chaperone,pERp1, that is crucial for our immunity, as it is involved in the folding of some immunoglobulins.
Keywords: disulfide bond, endoplasmic reticulum, ERAD, Ero1, oxidative proteinfolding, PDI, UPR
Moilanen, Antti, Disulfidisidoksia sisältävien proteiinien laskostumisen uudetsäätelymekanismit ja rakenteelliset ominaisuudet. Oulun yliopiston tutkijakoulu; Oulun yliopisto, Biokemian ja molekyylilääketieteen tiedekuntaActa Univ. Oul. A 754, 2021Oulun yliopisto, PL 8000, 90014 Oulun yliopisto
Tiivistelmä
Disulfidisidoksia sisältävien proteiinien laskostumista endoplasmakalvostossa avustavat erilai-set laskostumiskatalyytit ja kaperonit. Koska laskostuminen on monimutkaista, voivat proteiinitlaskostumistekijöistä huolimatta laskostua väärin. Endoplasmakalvostoon onkin kehittynyt usei-ta laadunvarmistusmekanismeja, jotka pitävät laskostumistaakan ja -kapasiteetin tasapainossa.Laskostumiskatalyyttien aktiivisuutta voidaan säädellä, minkä avulla vältetään ylihapettumiseenliittyvä stressi. Muihin laadunvarmistuskeinoihin kuuluvat UPR (unfolded protein response) jaERAD (endoplasmic reticulum-associated degradation). UPR tunnistaa laskostumattomien pro-teiinien aiheuttaman stressin ja saa aikaan mm. laskostumistekijöiden lisätuotantoa stressinvähentämiseksi. ERAD poistaa suoraan laskostumattomia proteiineja hajottamalla niitä.
Ero1- ja PDI-proteiiniperheiden jäsenet toimivat pääasiallisina laskostumiskatalyytteinä, jot-ka muodostavat disulfideja laskostuviin proteiineihin. Ero1 on sulfhydryylioksidaasi, joka pel-kistää hapen veryperoksidiksi muodostaessaan disulfidin. Uuden disulfidin Ero1 siirtää sittenPDI:lle, joka puolestaan katalysoi disulfidien muodostuksen laskostuvissa proteiineissa. Ero1-PDI-systeemiä on tutkittu paljon solutasolla, ja sen aktiivisuutta voidaan säädellä järjestelemälläsen omia säätelydisulfideja. Tämän säätelyjärjestelmän entsyymikinetiikka tunnetaan kuitenkinhuonosti.
Tässä työssä esittelemme Ero1-PDI-systeemin laajat entsyymikinetiikan mittaukset, jotkapaljastavat uusia säätelymenetelmiä proteiinien laskostumiseen. Näihin menetelmiin kuuluvatmm. tehokkuus hypoksisissa olosuhteissa ja erilaiset kineettiset ominaisuudet ihmisen Ero1-per-heen jäsenten välillä. Tulokset osoittavat, että Ero1α vaatii korkeat pitoisuudet PDI-substraattiasekä aktivoituakseen että disulfidien siirtoon PDI:lle. Näiden tulosten perusteella havaitsimmemahdollisen laskostumisen ja ERAD:n välisen vuorovaikutuksen, jossa laskostumisen tehok-kuutta lasketaan ERAD:n toiminnan tehostamiseksi. Lisäksi esittelemme kiderakenteen uudellekaperonille, pERp1:lle, joka vasta-aineiden laskostumistekijänä on tärkeä immuniteetillemme.
Asiasanat: disulfidisidos, endoplasmakalvosto, ERAD, Ero1, PDI, proteiinienlaskostuminen, UPR
To family, to friends, to colleagues; You made this possible.
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Acknowledgements
I gratefully acknowledge the Faculty of Biochemistry and Molecular Medicine and
Biocenter Oulu for the expertise and permission to use of their facilities. I would
like to also thank the staff of both organizations who carry out excellent science in
many different fields and educate future scientists. My work was funded by the
Academy of Finland, The Finnish Cultural Foundation, and The University of Oulu
Scholarship Foundation.
Special thanks go to Prof. Lloyd Ruddock, for your excellent ideas that led me
here, for supervision, and for creating the wonderful research group and atmosphere;
to Dr. Mirva Saaranen, who made this work possible with science and help and
support; Kati Korhonen, without whom I would probably not be here, as it was you
who pulled me into this fascinating world of Ero1; to Heli and Yuko, you were also
there every day helping and guiding; Johanna and Katja, it was a pleasure to share
minds with you, and I already miss our brainstorming sessions about science and
families; and to the other group members, George, Lisette, Rosario, Angel, Aatir,
you all are part of this work.
I am thankful to the deans Prof. Kalervo Hiltunen and Prof. Peppi Karppinen,
the dean of education Adj. Prof. Tuomo Glumoff, all the professors and group
leaders, as well as the administration personnel Pia Askonen, Auli Kinnunen, Jari
Heikkinen, and Kaisa Tuominen, who managed our scientific community; you did
excellent work. It was a privilege working with you.
The final acknowledgements go to my wonderful family, who patiently
supported me through this process; to the members of my follow-up group, Dr.
Kristian Koski and Doc. Ulrich Bergmann; to the reviewers of this dissertation,
Assoc. Prof. Carolyn Sevier and Dr. Adam Benham. Creating this work was a long
and eventful adventure. Your inspiring feedback guided me here.
Oulu, 7.10.2020 Antti Moilanen
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Abbreviations
-COO- deprotonated carboxyl (-COOH) group
-NH3+ protonated amine (-NH2) group
-S-S- disulfide bond
-SH thiol group
-SOH sulfenic acid group
1SS folding intermediate with one disulfide
2SS folding intermediate with two disulfides
Ala alanine
ALR augmenter of liver regeneration
Arg arginine
ATF6 cyclic AMP-dependent transcription factor ATF-6
ATP adenosine triphosphate
BiP binding immunoglobulin protein
BLI bilayer interferometry
BPA bisphenol A
BPTI bovine pancreatic trypsin inhibitor
C-terminus carboxyl-terminus of a polypeptide
CD circular dichroism
CGHC amino acid sequence motif; C, cysteine; G, glycine; H,
histidine
CNPY canopy (protein family)
CP peroxidatic cysteine
CPXC amino acid sequence motif; C, cysteine; P, proline; X, any
amino acid
CR resolving cysteine
CRELD cysteine-rich with EGF-like domain protein
CS citrate synthase
CyDisCo cytoplasmic disulfide bond formation in Escherichia coli
Cys cysteine
CXXC amino acid sequence motif; C, cysteine; X, any amino acid
COX cytochrome c oxidase
DHA dehydroascorbate
DNA deoxyribonucleic acid
DTT dithiothreitol
Dsb thiol:disulfide interchange protein
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E2 estradiol
EDEM ER degradation-enhancing alpha-mannosidase-like protein 1
ER endoplasmic reticulum
ERAD endoplasmic reticulum-associated degradation
ERdj5 endoplasmic reticulum DNA J domain-containing protein 5
Ero1 endoplasmic reticulum oxidoreductin 1
Ero1p endoplasmic reticulum oxidoreductin 1 from Saccharomyces
cerevisiae
ERp27 endoplasmic reticulum resident protein 27
ERp44 endoplasmic reticulum resident protein 44
ERp46 thioredoxin domain-containing protein 5
ERp57 protein disulfide isomerase A3
ERp72 protein disulfide isomerase A4
Erv1p mitochondrial FAD-linked sulfhydryl oxidase ERV1, or
essential for respiration and vegetative growth protein 1, from
Saccharomyces cerevisiae
FAD flavin adenine dinucleotide
FAM20C extracellular serine/threonine protein kinase
FLAG amino acid sequence DYKDDDDK
Glu glutamate
GPx glutathione peroxidase
GR glutathione reductase
GRP94 endoplasmin, or 94 kDa glucose-regulated protein
GSH glutathione (reduced)
GSSG glutathione (oxidized)
HEK human embryo kidney cell line
HeLa human cervical cancer cell line
HSP heat shock protein
HSP70 heat shock 70 kDa protein
I folding intermediate
ID intrinsically disordered
Ig immunoglobulin
IMS intermembrane space
IRE1 serine/threonine-protein kinase/endoribonuclease IRE1
ITC isothermal titration calorimetry
k reaction rate constant
kcat catalytic rate constant
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KFWWFS peptide consisting of lysine (K), phenylalanine (F), tryptophan
(W), and serine (S)
KM Michaelis constant, or substrate concentration at which the
reaction rate is half its maximal value
LMWO low-molecular-weight oxidant
M misfolded state
MANF mesencephalic astrocyte-derived neurotrophic factor, or
ARMET
Mia40 mitochondrial intermembrane space import and assembly
protein 40
MP membrane protein
mRNA messenger ribonucleic acid
MTP microsomal transfer protein
N native state
N-terminus amino-terminus of a polypeptide
N* two-disulfide folding intermediate of the folding pathway of
bovine pancreatic trypsin inhibitor
NADPH nicotinamide adenine dinucleotide phosphate
NSS
HH the penultimate, two-disulfide folding intermediate of the
folding pathway of bovine pancreatic trypsin inhibitor
OPF oxidative protein folding
ox oxidized
P5 protein disulfide isomerase A6
PDB Protein Data Bank
PDI protein disulfide isomerase
PDI3FLAG protein disulfide isomerase with three inserted FLAG
(DYKDDDDK) sequences
PDIp protein disulfide isomerase A2, or pancreas specific protein
disulfide isomerase
PERK eukaryotic translation initiation factor 2-alpha kinase 3
pERp1 plasma cell-induced resident endoplasmic reticulum protein
Phe phenylalanine
pKa negative logarithm of the acid dissociation constant (Ka)
Prx peroxiredoxin
QC quality control
QSOX quiescin sulfhydryl oxidase
rBPTI reduced bovine pancreatic trypsin inhibitor
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red reduced
RNA ribonucleic acid
RNase ribonuclease
ROS reactive oxygen species
rpHPLC reversed phase high-performance liquid chromatography
RS- thiolate, where R denotes an alkyl or other organic substituent
RS-SR disulfide bond, where R denotes an alkyl or other organic
substituent
SAPLIP saposin-like protein
SDS-PAGE sodium dodecyl sulfate-polyacryl amide gel electrophoresis
Ser serine
SN2 bimolecular nucleophilic substitution reaction
SRP signal recognition particle
SRPR signal recognition particle receptor
SSS transition state with three sulfur atoms arranged linearly
t1/2 half-time
Trp tryptophan
U unfolded state
UPR unfolded protein response
Val valine
VKOR vitamin K epoxide reductase
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Original publications
This thesis is based on the following publications, which are referred throughout
the text by their Roman numerals:
I Moilanen, A., Korhonen, K., Saaranen, M. J., & Ruddock, L. W. (2018). Molecular analysis of human Ero1 reveals novel regulatory mechanisms for oxidative protein folding. Life Science Alliance, 1(3), e201800090. doi:10.26508/lsa.201800090
II Moilanen, A., & Ruddock, L. W. (2020). Non-native proteins inhibit the ER oxidoreductin 1 (Ero1)-protein disulfide-isomerase relay when protein folding capacity is exceeded. The Journal of Biological Chemistry, 295(26), 8647-8655. doi:10.1074/jbc.RA119.011766
III Sowa, S. T., Moilanen, A., Biterova, E., Saaranen, M. J., Lehtiö, L., & Ruddock L. W. High-resolution crystal structure of human pERp1, a saposin-like protein crucial for IgA, IgM and integrin maturation in the endoplasmic reticulum. In press.
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Contents
Abstract
Tiivistelmä
Acknowledgements 9
Abbreviations 11
Original publications 15
Contents 17
1 Introduction 19
2 Review of the literature 23
2.1 Protein folding phenomenon ................................................................... 23
2.1.1 From primary to quaternary structure ........................................... 23
2.1.2 Protein refolding in vitro and in silico .......................................... 24
2.1.3 Co-translational protein folding in vivo ....................................... 29
2.1.4 Molecular chaperones ................................................................... 31
2.2 Biochemistry of disulfide formation ....................................................... 32
2.2.1 Disulfide formation is catalyzed redox chemistry ........................ 33
2.2.2 The pKa value determines the reactivity of a thiol group ............. 36
2.2.3 Thiol-disulfide exchange reactions ............................................... 38
2.3 Oxidative protein folding in vivo ............................................................ 41
2.3.1 Disulfide formation in prokaryotes............................................... 41
2.3.2 Mitochondrial relay for disulfide bond formation ........................ 43
2.3.3 Multiple ways to form disulfides in the ER .................................. 44
2.4 Structural biology of the Ero1-PDI relay ................................................ 54
2.4.1 Structure of human Ero1 isoforms................................................ 54
2.4.2 Architecture of PDI ...................................................................... 56
2.5 Structure-function relationship of Ero1-PDI ........................................... 58
2.5.1 Substrate oxidation by PDI ........................................................... 58
2.5.2 PDI reoxidation by Ero1 ............................................................... 60
2.5.3 Ero1 reoxidation ........................................................................... 60
2.6 Quality control of oxidative protein folding ........................................... 62
2.6.1 Modulation of Ero1 activity ......................................................... 62
2.6.2 Unfolded protein response (UPR) ................................................ 64
2.6.3 Endoplasmic reticulum-associated protein degradation
(ERAD) ........................................................................................ 66
3 Aims of the present work 69
4 Methods 71
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5 Results 73
5.1 Molecular analysis of human Ero1 .......................................................... 73
5.1.1 Human Ero1 and PDI form a novel Ero1-PDI complex ............... 73
5.1.2 Generation of a non-linear regression method to analyze
Ero1 kinetics ................................................................................. 74
5.1.3 Differential substrate PDI dependence between the Ero1
isoforms ........................................................................................ 75
5.1.4 A novel regulatory mechanism allows efficient operation
in hypoxic conditions ................................................................... 77
5.2 Feedback inhibition of the Ero1α-PDI system ........................................ 78
5.2.1 The Ero1α-PDI interaction is inhibited by non-native
proteins ......................................................................................... 79
5.2.2 Quantification of the inhibition .................................................... 79
5.2.3 The inhibition is reversible ........................................................... 80
5.3 Structure-function relationship of human pERp1 ................................... 81
5.3.1 High-resolution crystal structure of pERp1 .................................. 82
5.3.2 Mapping function onto the structure ............................................. 82
5.3.3 Biochemical assays to elucidate function ..................................... 83
6 Discussion 85
7 Future prospects 93
List of references 95
Original publications 117
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1 Introduction
The smallest living unit, the cell, is a factory of a multitude of molecules, which
help the cell to carry out intra- and intercellular physiological processes - from
maintaining and translating the genetic code to signaling with neighboring cells to
ultimately organizing into tissues, organs, and organisms - to sustain life.
Macromolecules deoxyribonucleic acid (DNA), ribonucleic acid (RNA), and
proteins, which are all large polymeric molecules formed from repeating
nucleotides (DNA and RNA) or amino acids (proteins), create a cellular workflow
from blueprint (DNA) to message (RNA) to structure (proteins) (Fig. 1).
The human genome has over three billion nucleotide base pairs and contains
20,000 to 25,000 protein-coding genes (International Human Genome Sequencing
Consortium, 2004). A gene is transcribed into message RNA (mRNA) which
translates to a protein (Fig. 1). The number of proteins available for a human cell
is greater than that the number of protein-coding genes, as one gene can potentially
create several different RNA transcripts, for example, by alternative splicing,
creating different forms (isoforms) of the same protein with different functions and
attributes. The total count of proteins, including isoforms, in human cells is
therefore ~100,000 (Pan et al., 2008). The proteins are sorted during (co-) and after
translation (post-translationally) to different organelles, including the nucleus, the
cytosol, the mitochondrion, and the endoplasmic reticulum (ER). One method to
traffic proteins to specific organelles is the inclusion of a targeting signal, for
example, a signal sequence (also called signal peptide) or a transmembrane
segment that may target proteins to the ER (Fig. 1) (Blobel, 1980). A targeting
signal can also be used to retain a protein in the ER preventing its secretion
(Raykhel et al., 2007).
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Fig. 1. From genetic code to correctly folded protein. The genetic material (DNA) in the
nucleus is transcribed to mRNA and transported to the cytosol, where it binds
ribosomal subunits to create a mRNA-ribosome complex. The ribosome translates the
mRNA and produces an amino acid sequence based on the mRNA code. If a specific
amino acid sequence (a signal peptide) is translated, it will be recognized by the signal
recognition particle (SRP) which halts translation and targets the mRNA-ribosome-
nascent-polypeptide-SRP complex to an ER membrane receptor (SRPR). Otherwise,
translation continues in the cytoplasm. The SRP-SRPR interaction allows the ribosome
to contact with a membrane channel complex (translocase), SRP dissociates,
translation continues, and co-translational translocation of the growing polypeptide to
the lumen of the ER initiates (Rapoport, 2007). The signal peptide is cleaved during this
process by a signal peptidase complex (Ellgaard et al., 2016). The depicted co-
translational translocation is a common event in all cell types, but ribosome-
independent translocation is also possible, typically utilized by simpler organisms such
as bacteria and yeast (Rapoport, 2007). In the ER, proteins fold and acquire a specific
biological function defined by their native structure. As protein folding is a complicated
process, misfolded states may form that lead to aggregate formation. To avoid this, the
ER has elaborate quality control (QC) features that maintain the balance between the
folded and misfolded proteins, to allow efficient export of natively folded proteins from
the ER (dashed arrow).
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As many as 40% of all human proteins contain a signal sequence or a
transmembrane region and are translocated co-translationally to the ER (Uhlen et
al., 2015; Fass & Thorpe, 2018), the starting point of the secretory pathway that
guides new proteins to the extracellular space. There they complete translation,
acquire a three-dimensional structure (a native fold), and gain a biological function
(Fig. 1). The protein folding process is complicated, and often proteins may form
unwanted functionless misfolded states. To avoid this, the ER has evolved a
multitude of different folding factors and quality control measures that either
prevent the nascent protein from misfolding, rescue from misfolded states, or
remove terminally misfolded proteins, to promote native folding and export from
the ER (Fig. 1). Folding in the ER is governed also by many co- and post-
translational protein modifications that are important for proper folding and for
regulation of protein function (Ellgaard et al., 2016).
This work provides a basis for understanding the protein folding phenomenon,
in particular, the folding process that includes also disulfide bond formation, a
crucial modification of native folded state of most proteins of the ER and the
subsequent destinations of the secretory pathway. The key folding factors that aid
in disulfide formation are introduced, including protein disulfide isomerase (PDI),
whose molecular function will be shown by systematic review of literature to be
under the control of another ER-resident enzyme, endoplasmic reticulum
oxidoreductin 1 (Ero1). The main quality control features of the ER are reviewed
to help us understand, how cells adapt to different stress situations that are prone to
happen in the ER. Original publications will reveal novel functional, regulatory,
and structural features of the Ero1-PDI folding relay and other folding factors to
gain new insight into the important cellular decisions that influence the
folding/misfolding partitioning in the ER (Fig. 1).
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2 Review of the literature
2.1 Protein folding phenomenon
Proteins gain specific biological functions only when they acquire a native three-
dimensional fold. This review will first provide the basic physicochemical
principles of conformational protein folding, a process that governs the unfolded to
native state transition. The overall protein folding process of many proteins is
influenced by additional co- and post-translational modifications. The composite
process of conformational folding and co- and post-translational disulfide bond
formation - also termed oxidative protein folding (OPF) - is discussed next.
Disulfide bonds are the main topic of this study, and the later chapters will
concentrate on the biochemical aspects of their formation. Other co- and post-
translational modifications are out of the scope for this study, but for an overview
of the most common modifications, see Ellgaard et al. (2016).
2.1.1 From primary to quaternary structure
Protein structure is hierarchical, and it consists of four levels (Finkelstein & Ptitsyn,
2016):
1. The primary structure of a protein is defined by all covalent bonds in an amino
acid sequence. These include peptide bonds, covalent bonds of the side chains,
and any covalent co- and post-translational modifications between amino acids.
2. Secondary structure elements are then formed that define local folding of the
polypeptide chain. The secondary structure is defined by rotation around three
covalent bonds within an amino acid residue corresponding to three dihedral
angles, ω, φ, and ψ. As the peptide bond has a partial double-bond
characteristic, ω is nominally restricted to two angles, 180° (trans
configuration) or 0° (cis configuration). The trans configuration is sterically
less hindered and energetically more favored and consequently occurs more
commonly in natural proteins (the cis is limited to proline residues only, and
the trans-cis interconversion is usually actively assisted by folding factors,
which are described later). The other two main chain angles, φ and ψ, have
more degrees of freedom, and they control most of the folding processes of a
polypeptide chain. Regular secondary structures, defined by repeating dihedral
angles in a linear stretch of amino acids, are especially important for protein
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folding; for example, alpha helices and beta sheets that are stabilized by inter-
residue hydrogen bonds.
3. Compaction of the polypeptide chain and regular secondary structure elements
defines the tertiary structure of a protein. Several models have been proposed
to characterize the initial steps of this event, for example, the framework and
the hydrophobic collapse models. In the former model, compaction happens
via packing of secondary structure elements that interact, for example, via
hydrogen bonds and electrostatic interactions (Udgaonkar & Baldwin, 1988).
During hydrophobic collapse, interactions between hydrophobic residues drive
the chain to a compact state, and the hydrophobic residues become buried to
the core of the protein (Tanford, 1962; Dill, 1990). A single polypeptide chain
may utilize one or several different compaction methods and may have several
distinct tertiary structure modules, also called domains.
4. Occasionally, a quaternary structure may form when two or more polypeptide
chains (subunits) interact with each other. Such a stable protein complex is
known also as an oligomer.
Most proteins follow these fundamentals. However, there are always exceptions to
the rule: a special group of proteins called intrinsically disordered (ID) proteins (or
proteins containing ID regions) may form biologically functional entities without
regular secondary or tertiary structure elements. They are out of scope for this study,
but for a thoughtful review, see Fonin et al. (2018).
2.1.2 Protein refolding in vitro and in silico
Anfinsen and colleagues (1961) revealed more than fifty years ago that unfolded
proteins can refold in the test tube rapidly and spontaneously to their native states.
This efficient maturation suggested that a protein chain avoids a slow systematic
search of a single native state from countless possible chain configurations (a 100-
residue chain has at least 10100 potential structures, as each residue has at least ten
different combinations of backbone dihedral angles) and matures via a limited
number of folding pathways (Anfinsen et al., 1961; Finkelstein, 2018). The
conformational folding phenomenon is typically modelled by various funnel
models that conceptualize protein maturation as allowed trajectories on multi-
dimensional “rugged” energy landscapes (Fig. 2) (Leopold et al., 1992; Wolynes et
al., 1995; Dinner et al., 2000). A common feature for all these models is that
transition from the unfolded state to the native state proceeds towards intermediate
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states with higher stability and lower thermodynamic free energy, with the native
state residing at a global free energy minimum (Fig. 2). In productive folding, the
intermediate states utilize mainly native intramolecular interactions (that are found
in the native state), that on average are more stable than non-native ones (Anfinsen,
1973; Abkevich et al., 1994; Sali et al., 1994; Dobson, 2003).
Non-catalyzed protein folding is often inefficient. Productive folding
intermediate states with low energy minima, also called kinetic traps, as well as
non-native intermolecular interactions between folding intermediates that lead to
stable low-energy non-native states, for example, irreversible aggregates, may
decrease the efficiency of folding and slow down the folding process significantly.
Therefore, during the search for the native state, the folding chain needs to also
avoid and escape any traps and non-productive energy minima (Fig. 2). This is a
matter of life and death: Inefficient folding of proteins that are essential for the
development and critical functions of an organism will terminate life. Even non-
essential proteins, if folded incorrectly, may cause protein aggregation diseases that
ultimately lead to death (Dobson, 2003; Jahn & Radford, 2005). Cells have
developed special folding factors, which are described later in this review, to avoid
these potentially dangerous off-pathway reactions.
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Fig. 2. Schematic of a multidimensional protein folding funnel. The funnel includes all
possible chain configurations, the arrows depict allowed trajectories. Depicted is a
refolding process [adapted from Stigler et al. (2011)] of a two-domain protein which may
fold from unfolded (U) to native (N) state via two potential on-pathway trajectories
containing folding intermediates (I), or form one off-pathway misfolded state (M). M
must unfold to U to return to the productive pathways. The I states are defined by local
energy minima with varying energy depts and activation energy barriers to proceed to
the next state. Deep local minima may lead to very slowly folding intermediates, also
known as kinetic traps (see inset). Although the allowed trajectories are defined, the
search for N is stochastic, i.e., reversible U ↔ I, U ↔ M, I ↔ M, I ↔ N transitions are
possible. Even reversible U ↔ N transition is possible for simple one-domain proteins
with smooth energy landscapes (Finkelstein, 2018). Inset energy diagram: A free energy
diagram adapted from Finkelstein & Ptitsyn (2016) which depicts a three-state folding
process from an initial state (1) to a final state (2) via a kinetic trap (X). X is a trap, as
transition X → 2 has higher activation energy barrier (grey arrow) than the transition 1
→ X. It is also an “on-pathway” intermediate, as it is more stable (has lower free energy)
than the original state (1) but less stable than the final state (2). The conversion of
kinetic traps and the rescue of off-pathway species may be accelerated significantly by
folding factors.
27
The seminal work by Anfinsen et al. (1961) studied OPF of ribonuclease A
(RNase A), a small protein with eight cysteines that theoretically can adopt 105
different four-disulfide patterns (Anfinsen, 1973). Remarkably, only one native
disulfide pattern was observed in the experiments. This leads to a fundamental
question: are disulfides rearranged randomly until the native pattern is reached, or
does OPF utilize limited pathways – analogous to the conformational folding
described above - and favor folding intermediates with native disulfides? Two
models with stark differences have been experimentally demonstrated to answer
this question. One model is based on refolding of mature bovine pancreatic trypsin
inhibitor (BPTI) and the other on hirudin, both small proteins with three disulfides
in the native state. BPTI has an anti-parallel β-sheet core surrounded by alpha
helices that are stabilized by disulfides [Cys5-Cys55, Cys30-Cys51, Cys14-Cys38]
(Fig. 3A). A consensus view on the sequential folding events of BPTI (Fig. 3B)
(Weissman & Kim, 1991) suggests that the reduced state can first adopt any of two
different intermediates with one native disulfide, either [Cys5-Cys55] or [Cys30-
Cys51]. Subsequently, they form two-disulfide states, [Cys5-Cys55, Cys14-Cys38]
or [Cys30-Cys51, Cys14-Cys38], respectively. Attaining the final disulfide is,
counterintuitively, a complex and slow process that requires, in both pathways,
disulfide rearrangements to form a third two-disulfide intermediate, [Cys5-55,
Cys30-51]. Only this state can then efficiently fold to the three-disulfide native state.
The consensus refolding mechanism dictates that maturation of BPTI utilizes
predominantly native disulfides. In contrast, hirudin has been demonstrated to
refold with a so-called “trial and error” method, i.e., it matures with a heterogenous
mixture of folding intermediates, containing both native and non-native disulfides
(Chatrenet & Chang, 1992; Chatrenet & Chang, 1993). These two extreme in vitro
models, and other cases in between them [reviewed by Chang (2011)], suggest that
the diversity of oxidative refolding of an isolated protein molecule in the test tube
is wide and cannot be answered by a single model. Note, however, that little
evidence currently exists that non-native disulfides are required for productive
protein folding in vivo (Fass & Thorpe, 2018), and potential factors, that may
stabilize protein structure in vivo and eliminate formation of non-native disulfides,
are discussed later in Chapter 2.1.3. Therefore, this work will mainly concentrate
on the BPTI-type refolding model that utilizes native interactions.
28
Fig. 3. The BPTI fold. (A) Secondary structure representation of wild-type BPTI (Protein
Data Bank code: 1BPI). Alpha helices (α, magenta), beta strands (β, red), and the three
native disulfides ([Cys-Cys], yellow) are marked. Regions without regular secondary
structure are grey. N, N-terminus; C, C-terminus. (B) Schematic of the consensus BPTI
oxidative folding pathways (Weissman & Kim, 1991). Conversion of the reduced
unfolded state (R) via one-disulfide intermediates (1SS) to the 2SS kinetic trap states
(N* and [30-51, 14-38]) is kinetically fast. The interconversion between the 2SS kinetic
traps and conversion of the 2SS kinetic traps to the NSS
HH are slow processes, as local
structural unfolding is required to expose the free thiols for disulfide rearrangements.
Folding from NSS
HH to N is fast.
29
One fundamental question remains unanswered: Does disulfide formation
drive conformational folding or vice versa? Although the history of OPF spans
more than fifty years, the numerous “wet lab” folding studies have been unable to
answer this question explicitly. Lately, computational methods, especially
molecular simulations, have become indispensable tools for studying folding of
simple proteins. For example, Qin and coworkers (2015) demonstrated that well-
calibrated molecular simulations can refold BPTI consistent with the folding
pathways observed in the seminal biochemical studies of Weissman & Kim (1991).
Their simulations suggested that substantial compaction and formation of the
central β-sheet precedes formation of any disulfides. This hints at conformational
folding being the driving force for disulfide formation. Furthermore, BPTI has been
demonstrated to possess native subdomains, and their sequential formation as well
as reductive unfolding dictates the sequential oxidative folding pattern (Staley &
Kim, 1990; Mendoza et al., 1994). In contrast, reductive unfolding of hirudin shows
an “all-or-none” scheme, in which all three native disulfides are reduced
simultaneously, consistent with hirudin lacking any stable subdomains (Chang,
1997). The causal factor for deciding whether a protein adopts the BPTI or the
hirudin-like refolding model may therefore simply be relative structural stability of
protein subdomains in the in vitro experiments (Chang, 2011).
Collectively, the computational and in vitro biochemical evidences discussed
here suggest that conformational folding of stable (sub)domains may guide native
disulfide formation by defining the accessibility, proximity, and reactivity of thiols
(which are discussed in more detail later in this study) for efficient thiol-disulfide
chemistry.
2.1.3 Co-translational protein folding in vivo
Protein folding in vivo occurs in environments and conditions drastically different
from the in vitro refolding experiments described above. Whereas the unstructured
isolated protein at start of the refolding experiment may access all backbone
residues, a nascent polypeptide in vivo will start folding co-translationally – both
conformationally and oxidatively - while still ribosome-bound and only partially
synthesized. In other words, in vivo folding of some proteins is vectorial (N-
terminus folds first, C-terminus last) and a domain-by-domain folding process
(Bulleid & Freedman, 1988; Kolb et al., 1994; Komar et al., 1997; Holtkamp et al.,
2015; Thommen et al., 2017). The main factors that delineate in vivo folding from
test tube experiments are listed below:
30
– The reported refolding times that range from sub-seconds to tens of hours (for
example, Anfinsen et al., 1961; Schuler & Hofmann, 2013) are often
physiologically unrealistic. Minimal rates of in vivo folding are defined by
ribosomal translation rate (ktranslation), which in human cells is circa four amino
acids incorporated to a nascent chain per second. It takes two minutes to
synthesize an average-sized protein of 50 kDa (Braakman et al., 1991; Yan et
al., 2016). Also, the folding process in vivo rarely takes hours, as cells need to
be able to respond to various physiological stimuli rapidly and efficiently
(Dintzis, 1961; Braakman et al., 1991).
– ktranslation can be modulated on the mRNA and nascent chain levels (Kaiser et
al., 2011, Waudby et al., 2019) to allow controlled partial folding of
polypeptides. For example, rare codons between domain boundaries and
certain amino acid sequences that interact with the ribosome exit tunnel may
cause transient ribosomal arrests (Zhang et al., 2009; Tsai et al., 2014), favoring
the domain-by-domain folding process. Such elegant mechanisms may
significantly impact the folding energy landscape (Fig. 2) by transiently
limiting the number of allowed polypeptide configurations and preventing off-
pathway interactions that may lead to aggregation. This is especially important
for proteins (or protein domains) with high folding kinetics (kfolding). If kfolding
<< ktranslation, folding finishes post-translationally, which can be directly
observed for many proteins, as their N- and C-termini are in close proximity in
their native structures (Ellgaard et al., 2016; see also Fig. 3A).
– In vitro refolding experiments are typically carried out in mild buffer
conditions with high free volume that the folding chain can utilize during
folding. This is in stark contrast with cellular environments which are crowded
with various macromolecules, including proteins and nucleic acids.
Macromolecules may occupy up to 30% of the cytoplasmic volume and can
reach as high as 400 mg/mL concentration (Zimmerman & Trach, 1991; Fonin
et al., 2018). This is likely true also in the ER: certain folding factors in the ER
exist at very high concentrations, for instance, chaperone BiP and the folding
catalyst PDI alone reach millimolar (circa 50 mg/mL) levels (Hillson et al.,
1984; Lyles & Gilbert, 1991; Weitzmann et al., 2007). The crowding effect has
both positive and negative influences on protein folding (Rivas et al., 1999;
Ellis, 2001): on one hand, crowding may enhance early-stage folding, for
example, the intramolecular hydrophobic collapse, but also the late-stage
intermolecular associations leading to quaternary structure formation; on the
other hand, crowding increases the probability for non-native intermolecular
31
contacts that may lead to aggregation. However, the net effect is likely positive,
as cells have developed ways to minimize aggregation, specifically, molecular
chaperones (Martin & Hartl, 1997). Similarly, crowding may also stabilize
formation of the discussed protein subdomains, which may influence the
decision whether a protein adopts the scrambled hirudin-like folding scheme
or the BPTI-like model with native interaction only.
All in all, while in vitro folding studies have provided invaluable information from
thermodynamics to molecular details for protein folding in general, the co-
translational folding behavior inside the crowded cell introduces additional
complexities which are less well understood. Direct comparisons between refolding
and co-translational folding pathways of a given protein are lacking. A promising
study by Samelson and colleagues (2018) compared co-translational folding and
test tube folding of the same protein, and their results demonstrated that ribosome-
mediated folding process lacked an aggregation-prone non-native intermediate.
Thus, cellular conditions provide factors that maximize folding efficiency and
minimize non-native interactions. This review will next concentrate on two
structure-stabilizing factors that cells utilize, molecular chaperones and folding
catalysts for OPF.
2.1.4 Molecular chaperones
A general description (Hartl et al., 2011) defines a molecular chaperone as
any protein that interacts with, stabilizes, or helps another protein to acquire
its functionally active conformation, without being present in its final structure.
Molecular chaperones bind to hydrophobic regions of folding substrates and
prevent their aggregation (Hartl et al., 2011) in a way that was classically
considered not to “appreciably accelerate folding” (Puig & Gilbert, 1994). However,
the latter aspect is a matter of debate, as certain chaperones under some
circumstances seem to considerably accelerate folding (for example, Brinker et al.,
2001; Tang et al., 2006; Apetri & Horwich, 2008; Libich et al., 2015a; Libich et al.,
2015b). To avoid ambiguity, this study draws the line between “folding factor” to
describe any molecule that promotes protein folding and “folding catalyst” that
significantly accelerates the kinetics of the rate-limiting steps of peptide bond
isomerization (peptidyl-prolyl isomerases) or formation of a new covalent bond
(e.g., disulfide formation). “General chaperones” are then used in the classical way
32
to define any factors such as the heat-shock proteins (HSP) and chaperonins
(described below) that bind to hydrophobic residues, prevent aggregation, and
assist folding – regardless of whether they also accelerate folding. “Co-chaperones”
are also a class of folding factors, but they typically bind general chaperones and
enhance their activity.
General chaperones form two main classes based on their utilization of
adenosine triphosphate (ATP). The first class can bind and hydrolyze ATP to cycle
between low or high affinity states to allow controlled binding, release, and
rebinding of folding substrates (Hartl & Hayer-Hartl, 2009). ATP-dependent
chaperones include members of the HSP family (Kim et al., 2013) and chaperonins
that form a cylindrical cage for confined substrate folding (Hartl et al., 2011). The
two systems (HSP and chaperonins) are not redundant, as they likely operate in a
sequential manner: HSP family members interact with early-stage folding
intermediates and the chaperonins downstream when the substrate fails to attain
native state (Langer et al., 1992). The second class consists of ATP-independent
chaperones which mainly “buffer aggregation” and solubilize substrates by binding
and releasing hydrophobic regions in an ATP-independent manner (Hartl et al.,
2011). These proteins include many chaperones important for co- and post-
translational folding processes such as the bacterial trigger factor or the eukaryotic
nascent polypeptide-associated complex, that bind nascent polypeptides in the
ribosomal exit tunnel or as soon as they emerge from the ribosome, delaying chain
collapse until sufficient number of residues are available for productive folding
(Ferbitz et al., 2004; Kaiser et al., 2006; Gamerdinger et al., 2019). Another peculiar
molecular chaperone involved in OPF and oligomeric assembly of
immunoglobulins is introduced later in this study (Chapter 2.6.2).
2.2 Biochemistry of disulfide formation
A disulfide is a covalent bond linking the sulfur atoms of two thiol groups. A protein
disulfide connects cysteines either within the same polypeptide chain
(intramolecular), or between different chains (intermolecular or mixed disulfide).
Disulfide formation (and reduction) deals with electron flows, from an electron
donor (nucleophile) to an electron acceptor (electrophile). Spontaneous disulfide
formation by a nucleophilic attack of a free thiol on another thiol is kinetically a
slow process, with high activation energies. Disulfide formation is therefore often
the rate-limiting process for OPF (Nagy, 2013). This part of the review will
introduce the most important chemistries and mechanisms that enhance the
33
reactivity of thiols and kinetics of disulfide bond formation, to allow disulfides to
form in a physiologically relevant efficiency and time scale.
2.2.1 Disulfide formation is catalyzed redox chemistry
Disulfide bond formation is a reduction-oxidation (redox) reaction that transfers
two electrons from the dithiol to an external electron acceptor. As a result, the
dithiol is oxidized to disulfide and the electron acceptor gets reduced. The electron
acceptor can be a low-molecular-weight oxidant (LMWO) such as molecular
oxygen (O2) or hydrogen peroxide (H2O2), but often another pair of thiols acts as
the acceptor. The underlying physicochemical property that drives biological redox
reactions forward and describes the direction of the electron flux is called biological
standard reduction potential (measured in millivolts). A comprehensive review on
redox reactions and how to derive reduction potentials for a given redox pair are
reviewed by Hatahet & Ruddock (2009). The reduction potential describes only
whether a certain reaction is thermodynamically favorable to take place, but not
about kinetics of the reaction. A species with a higher reduction potential prefers to
gain electrons, i.e., oxidizes the species with a lower reduction potential (Fig. 4A).
Similarly, the redox state of an intracellular organelle can be derived by measuring
the reduction potential of a buffer component in the given compartment. Typically,
the redox potential of glutathione, which exists in various organelles as a mixture
of reduced glutathione (GSH) and its oxidized form (GSSG), is measured. In such
measurements, a cytosolic redox state is usually described to be reducing and
unfavorable for disulfide formation, and that of the ER is more oxidizing favoring
disulfide bond formation and OPF (Hwang et al., 1992; Bass et al., 2004; Dixon et
al., 2008). See Fig. 4B for examples of experimentally derived reduction potentials.
Glutathione and its buffering-capabilities are reviewed later in Chapter 2.3.3.
34
Fig. 4. Redox chemistry of disulfide formation. (A) Schematic of a reduction-oxidation
reaction with electron transfer from a redox couple with a lower reduction potential
(thiol/disulfide) to one with a higher reduction potential (O2/H2O2). -SH, thiol; -S-S-,
disulfide; O2, molecular oxygen; H2O2, hydrogen peroxide. (B) Experimentally derived
reduction potentials of low-molecular-weight oxidants and proteins as well as estimated
reduction potential ranges for selected organelles relevant for this work (Cleland, 1964;
Rost & Rapoport, 1964; Rakauskiene et al., 1989; Hwang et al., 1992; Lundstrom &
Holmgren, 1993; Huber-Wunderlich & Glockshuber, 1998; Dixon et al., 2008; Hatahet &
Ruddock, 2009; Ruddock, 2012; Birk et al., 2013; Romero et al., 2018). DTT, dithiotreitol;
DsbA, Escherichia coli thiol:disulfide interchange protein; ER, endoplasmic reticulum;
GR, glutathione reductase; NADPH, reduced nicotinamide adenine dinucleotide
phosphate.
The reduction potential of O2 is very high (Fig. 4B) which suggests that direct O2-
driven disulfide formation could form a major OPF pathway in aerobic cellular
conditions. However, this is probably not the case, as the kinetics of electron
transfer between a pair of thiols and free O2 is too slow to maintain life alone
(Ruddock, 2012). Therefore, in addition to thermodynamics, relative rates of
disulfide formation of different oxidants must be considered before their
physiological relevance can be assessed. Reported disulfide formation rates (Table
1) suggest that LMWOs oxidize thiols to disulfides rather inefficiently, and they
likely have a supporting role in disulfide formation in vivo. Catalysis can
significantly improve their oxidative effects: for example, disulfide formation by
35
O2 can be improved by transition metal ions and enzyme-bound redox cofactors
(Fass & Thorpe 2018; Romero et al., 2018). The concentration of free transition
metals is low in vivo, making them insignificant disulfide catalysts in the cell
(Ruddock, 2012). Flavin adenine dinucleotide (FAD), while inefficient free catalyst
in solution (Table 1), is a common bound cofactor in many oxidases (for example,
flavin-dependent sulfhydryl oxidases) and accelerates disulfide formation
significantly. Another important LMWO that greatly benefits from catalysis is H2O2
(Table 1). Flavin-dependent sulfhydryl oxidases and peroxidases are discussed in
more detail in Chapter 2.3.3.
Table 1. Comparison of disulfide formation rates (reported second order rate constants)
of selected oxidative systems relevant for this work.
Thiol/cofactor Electron acceptor Rate (M-1s-1) Reference
Free FAD O2 250 Massey (1994)
Enzyme-bound FAD O2 104 - 106 Mattevi (2006),
Romero et al. (2018),
and references
therein
BPTI H2O2 5 Karala et al. (2009)
Cys, DTT, GSH H2O2 18 - 26 Winterbourn &
Metodiewa (1999)
PDIa H2O2 9.2 Karala et al. (2009)
PrxIV H2O2 2.2 x 107 Wang et al. (2012)
GPx7 H2O2 2.6 x 103 Wang et al. (2014)
PDI DHA 12.5 Saaranen et al.
(2010)
BPTI DHA 163 Saaranen et al.
(2010)
BPTI Glutathione 7.3 Karala et al. (2009)
DsbA DsbB 2.7 x 105 Grauschopf et al.
(2003)
There are multiple factors influencing the reported higher enzymatic disulfide
formation rates. These include enhanced thiol nucleophilicity, optimal substrate
orientation and proximity relative to the catalytic center, and thiol-disulfide
exchange reactions, each of which are discussed next. All in all, correct disulfide
bond formation in vivo is a catalyzed redox-sensitive reaction. This sensitivity to
the redox conditions results in different organelles having low to high propensity
for disulfide formation.
36
2.2.2 The pKa value determines the reactivity of a thiol group
The rate of disulfide formation is first and foremost a function of three structural
and chemical factors: i) reactivity, ii) accessibility, iii) and proximity of thiol sulfur
atoms (Wedemeyer et al., 2000; Qin et al., 2015). Reactivity is discussed first and
accessibility and proximity later in Chapter 2.2.3. A disulfide is rarely formed by
non-charged thiol groups. A thiol can form a negatively charged state, a thiolate,
which is more efficient nucleophile (up to 1010-fold), and almost always the
preferred state during a nucleophilic attack (Fass & Thorpe, 2018). The thiol-
thiolate equilibrium is determined by the pKa value which specifies the pH of the
solution at which the thiol has a probability of 50% to exist either as a thiol or a
thiolate [see Hatahet & Ruddock (2009) for instructions on how to derive pKa
values]. If pKa is higher than pH, the equilibrium is driven towards the protonated
non-charged thiol state, while pKa less than pH will drive the group towards the
more reactive thiolate state (Fig. 5A). Thus, in general, lower pKa translates to
higher reactivity at physiological pH, but some exceptions to this rule exist, and the
nucleophilicity of each case must be carefully evaluated (Fass & Thorpe, 2018).
pKa values may vary between thiol groups. At physiological conditions (pH
circa 7), certain thiols are mainly in the reactive thiolate state (low pKa), whereas
most adopt the less-reactive thiol form (high pKa). An important modulator of pKa
is an adjacent charged group: a positive charge drives the equilibrium towards the
thiolate state, and a negative charge towards the thiol state (Fig. 5A) (Hatahet &
Ruddock, 2009). A given thiol may interconvert between the two states due to
protein dynamics, i.e., conformational changes and movement of other charged
residues can cause pKa fluctuations in proteins. This is especially important in the
catalytic mechanism of PDI (Lappi et al., 2004) which is described in more detail
in Chapter 2.5.
37
Fig. 5. The effect of pKa on the thiol:thiolate conversion. If the pKa value of the thiol
group is near the pH of the solution (the middle reaction), an equilibrium is reached
between the thiol and the thiolate states. pKa lower than pH will drive the equilibrium
towards the more reactive thiolate (upper reaction). pKa can be lowered by stabilization
of the thiolate state by a nearby positive charge. Inversely, pKa higher than pH favors
the uncharged thiol and a negatively charged group stabilizes the uncharged thiol
group to avoid repulsion between charged groups. The loss or gain of a proton during
the interconversion reactions is not depicted. (B) Thiol-disulfide exchange is a
bimolecular nucleophilic substitution (SN2) reaction. A nucleophilic sulfur atom
(cysteine residues are yellow, sulfur atoms are spheres) attacks a disulfide bond (in this
hypothetical case, a Cys53-Cys56 oxidized active site of PDI a domain) in line forming
a linear SSS transition state (Nagy et al., 2013). The product is a mixed disulfide between
Cys53 and the nucleophile and a free Cys56 thiol. The mixed disulfide is resolved by
another nucleophile with similar linear coordination chemistry (not depicted). Of note,
the chemistry of disulfide oxidation by low-molecular-weight oxidants is under debate
and may not proceed with the described mechanisms (Zeida et al., 2012). The illustration
has been drawn using PDI from PDB structure 6I7S.
38
2.2.3 Thiol-disulfide exchange reactions
A disulfide can form by a coordinated thiol-disulfide exchange reaction. Instead of
passing the electrons to an LMWO, here, electrons are transferred between two
pairs of thiols. The consensus disulfide exchange reaction proceeds with an SN2
substitution mechanism, in which the attacking sulfur atom approaches in line to
the disulfide (Fig. 5B; Fass & Thorpe, 2018). This mechanism is utilized by
oxidoreductases (but is not restricted to them) such as PDI family members, which
contain a solvent exposed thiol and a buried thiol in the active site (in the reduced
state) aligned optimally for an incoming in-line nucleophilic attack (Fig. 5B).
Therefore, efficient disulfide exchange is a series of coordinated chemistries,
including the structural orientation of the involved thiol sulfur atoms that likely
optimizes the steric requirements (orientation and proximity) of the disulfide
transfer and active modulation of pKa/nucleophilicity of the participating thiols,
that together explain the increased enzymatic disulfide transfer rates (Table 1).
The exchange occurs in two substitution steps (equations 1 and 2): in the first
step, thiolate 3 (-SR3) resolves the original disulfide between thiols 1 and 2 and a
mixed disulfide between thiols 2 and 3 is formed; the mixed disulfide is then
resolved in the second step by thiolate 4, creating a new disulfide between thiols 3
and 4.
𝑅 𝑆˗𝑆𝑅 𝑆𝑅 ↔ 𝑅 𝑆 𝑅 𝑆˗𝑆𝑅 (1) 𝑅 𝑆˗𝑆𝑅 𝑆𝑅 ↔ 𝑅 𝑆 𝑅 𝑆˗𝑆𝑅 (2)
No net disulfide formation takes places (as was described already, de novo
generation of disulfides generally requires direct or indirect participation of a
LMWO). Note also that both reactions are drawn bidirectional, as the intermediate
mixed disulfide and the transferred disulfide could theoretically be attacked by the
product thiolates. However, as already discussed in Chapter 2.2.2, the reactivity of
the participating thiols is often actively modulated to prevent the reverse reactions.
Disulfide exchange is an important mechanism for productive OPF (Fig. 6),
but exchange reactions can result in non-productive reactions which may lead to
misfolding and even aggregate formation (see the off-pathway reaction in Fig. 6).
Non-native disulfides can be corrected by reduction-oxidation cycles and by
intramolecular or intermolecular isomerization. Catalyzed intramolecular
isomerization is a physiologically important function to correct non-native
disulfides and is typically carried out by isomerases such as PDI. It is both a crucial
part of productive OPF as well as a countermeasure against disulfide-linked
39
aggregation and protein aggregation diseases (Mossuto, 2013). Intermolecular
rearrangement is not limited to isomerases, though, as it may modulate the
dynamicity of some protein assemblies, for example, disulfide-rich hair keratin and
hemostasis factors to combat mechanical stretching and shearing (Ganderton et al.,
2007). All intermolecular exchange mechanisms utilize a mixed disulfide
intermediate complex, which is an important indicator of disulfide transfer between
proteins, as these intermediates can often be trapped and identified (for example,
Molinari & Helenius, 1999; Benham et al., 2000; Bass et al., 2004).
Thiol-disulfide exchange is an important mechanism for several other
physiological processes. For example, peroxidases couple antioxidant defense
(reduction of H2O2 to water) to exchange reactions (Chapter 2.3.3). Transient
formation and resolution of disulfides may modulate enzyme activity (Chapter
2.6.1) as well as redox signaling processes (Li & Camacho, 2004; Ushioda et al.,
2016).
40
Fig. 6. Thiol-disulfide exchange mechanisms adapted from Hatahet & Ruddock (2009).
Disulfide transfer to a reduced dithiol starts with a nucleophilic attack on the disulfide
by a thiol in the reduced species, forming a mixed disulfide intermediate complex. The
productive oxidative pathway (box with an intact border) continues by resolution of the
mixed disulfide by a free thiol. A second similar reaction leads to a native state with two
disulfides. If the initial mixed disulfide is resolved by a non-native thiol, one of two non-
productive reactions may happen: A reverse reaction (not depicted) may regenerate the
starting material, or a non-native disulfide may form. The former reaction would be futile
and can be controlled by modulating the pKa of the nucleophilic thiol. The latter can be
corrected by isomerization or by reduction-oxidation cycles to first break the non-native
disulfides and then return to the productive oxidative pathway. Isomerization can either
be an intramolecular rearrangement, which requires a free thiol in the misfolded protein
or an intermolecular reaction which utilizes an external thiol. The latter can potentially
rearrange a fully oxidized misfolded system to the native state.
41
2.3 Oxidative protein folding in vivo
OPF is an enzyme-catalyzed process in vivo. At a macroscopic level, the catalytic
systems between species of the kingdoms of life seem similar (Fig. 7). Disulfides
are generated de novo by oxidoreductases, that pass electrons from their active site
dithiol ultimately to O2. The newly generated disulfides are then transferred to a
central oxidoreductase that catalyzes OPF of a nascent polypeptide. On a
microscopic level, however, the systems have stark differences, from differing
electron transfer pathways to diversity of OPF components. This chapter provides
an overview of disulfide formation in prokaryotes, eukaryotic mitochondria, and
finally the main topic of this study, the complex disulfide bond formation network
of the human ER.
2.3.1 Disulfide formation in prokaryotes
Bacteria generate disulfides mainly in the more oxidizing extracytoplasmic space,
specifically, the intermembrane space called periplasm in Gram-negative bacteria
or within the boundaries of the cell wall and the cytoplasmic membrane of Gram-
positive bacteria (Kadokura & Beckwith, 2010; Sarvas et al., 2004). After
translation in the cytoplasm and translocation to the extracytoplasmic space, OPF
of a nascent polypeptide is carried out by two potential pathways: 1) the Dsb family
in the periplasm of Gram-negative bacteria, or 2) the vitamin K system found in
both the Gram-negative and Gram-positive bacteria. DsbA, DsbB, DsbC, and DsbD
are the best characterized Dsb family members, and together they form the main
pathway for prokaryotic OPF. The DsbA-DsbB pair transfers disulfides to nascent
polypeptides, and DsbC-DsbD forms an isomerase unit to correct non-native
disulfides (Fig. 7). DsbA, which has one of the strongest reported redox potentials
of an oxidoreductase, -120 mV (Fig. 4B), is the central oxidoreductase that transfers
disulfides to folding substrates. It is re-oxidized by the membrane protein DsbB
which generates disulfides de novo, transfers them to DsbA, and regenerates its
oxidized form by passing the electrons to an electron transport chain consisting of
a quinone molecule, cytochrome c oxidase (COX), and O2 as the terminal electron
acceptor (Fig. 7). The DsbD-DsbC pair acts inversely so that DsbD reduces DsbC
which in the reduced free-thiol form may carry out reduction and isomerization of
non-native disulfides. DsbD is then reduced by cytoplasmic thioredoxin system for
a new catalytic cycle (Rietsch et al., 1997).
42
Fig. 7. The major pathways for OPF in the periplasm, the mitochondrial intermembrane
space, and the endoplasmic reticulum. Vertical and tilted arrows depict electron transfer
reactions, while horizontal arrows convert one chemical species to another. The middle
panel depicts substrate folding from unfolded to native state. This is achieved either by
direct oxidation by the oxidative pathways (lower panel) or by the corrective pathways
that rearrange (isomerization) or reduce non-native disulfides (upper panel). The lower
panels show electron transfer reactions from a central oxidoreductase (DsbA, Mia40, or
PDI) to terminal electron acceptors (O2 or H2O2). The cytoplasmic reducing pathways
that are connected to the ER are shown at the top. The reductive system of the ER is
currently under investigation with recent reports suggesting that a cytoplasmic
thioredoxin reduces an unknown ER membrane protein (MP) that may control the redox
state of the PDI reductases (Poet et al., 2017; Cao et al., 2020).
43
The vitamin K system was characterized independently by two groups (Dutton et
al., 2008; Singh et al., 2008). They showed that an atypical membrane protein was
able to catalyze disulfide formation in the extracytoplasmic space of a Dsb-
deficient prokaryotic species as well as E. coli artificially made devoid of the Dsb
pathways. The vitamin K system seems to have evolved independently of the Dsb
system (Dutton et al., 2008), but shares similarities in the catalyzed reactions: the
membrane protein vitamin K epoxide reductase (VKOR) oxidizes a DsbA-like
protein and subsequently passes the electrons to the electron transport chain via
vitamin K (a quinone derivative). VKOR has peculiar homologs that may catalyze
disulfide bond formation in the cytoplasm, e.g., in extremophiles that require
disulfides for thermal adaptation. Their catalytic components reside on the
cytoplasmic side (Hatahet & Ruddock, 2013). These rare cases of cytoplasmic
disulfide bond formation are out of scope for this study; see Saaranen & Ruddock
(2013) for more information.
2.3.2 Mitochondrial relay for disulfide bond formation
The best-characterized mitochondrial system for disulfide bond formation is found
in the intermembrane space (IMS) (Fischer et al., 2013). The central oxidoreductase
is Mia40, which introduces disulfides to imported nascent polypeptides (Fig. 7).
Mia40 is re-oxidized by essential for respiration and vegetative growth protein 1
(Erv1, in yeast) or augmenter of liver regeneration (ALR, in human), both flavin-
dependent sulfhydryl oxidases. Erv1 and ALR then pass electrons to the already
described cytochrome c/COX/O2 electron transport chain. Whereas the bacterial
Dsb system has a well-characterized pathway for disulfide isomerization, the
proofreading system of the mitochondrial IMS is still under debate. Glutathione,
which is possibly kept in the reduced state by glutathione reductase (Kojer et al.,
2012), has been shown to increase the efficiency of the Mia40-Erv1 relay,
suggesting that it may directly reduce non-native disulfides in folding intermediates
(Bien et al., 2010). On the other hand, Mia40 shows isomerase activity in vitro, and
it exists as a mixture of redox states in vivo, which together suggest that Mia40 may
carry out both oxidation and isomerization in the IMS (Kojer et al., 2012; Koch &
Schmid, 2014). Possibly, both systems contribute to and are crucial for the quality
control of disulfide formation in the IMS, similarly to the Ero1-PDI-glutathione
system found in the ER (Fig. 7; see Chapter 2.3.3.).
44
2.3.3 Multiple ways to form disulfides in the ER
40% of all human genes encode for proteins that are translocated to start folding in
the ER (Uhlen et al., 2015). This large proteome includes secreted hormones,
immunoglobulins, plasma membrane proteins, extracellular matrix proteins, and
other proteins that permanently reside within the organelles of the secretory
pathway (the ER, the Golgi apparatus, lysosomes, and the vesicles travelling
between them). A recent genome-wide analysis combined quantitative
transcriptomics to immunohistochemistry and demonstrated that as many as two-
thirds of all transcripts of the pancreas and the salivary glands may encode for
secreted proteins (Uhlen et al., 2015). The burden of protein synthesis and disulfide
formation in the ER may therefore reach extremely high levels, and it comes with
no surprise that the ER has developed an elaborate protein folding network of
multitude of folding factors.
The prokaryotic extracytoplasmic spaces and the mitochondrial IMS each had
clear major pathways for OPF. Dissecting the ER for major and minor pathways is
more complicated, however. GSSG was originally suggested to be the main source
of disulfides in the ER (Hwang et al., 1992). This view was later challenged when
a novel sulfhydryl oxidase, Ero1, was discovered that seemed to fuel PDI-driven
OPF (Frand & Kaiser, 1998; Cabibbo et al., 2000; Pagani et al., 2000). The
consensus main oxidative pathway shifted to the Ero1-PDI relay, while glutathione
was given an indirect role in disulfide formation by controlling the redox state of
other proteins (Fig. 8A). The plausibility of this new model was supported by
essentiality of the single Ero1 gene in yeast (Frand & Kaiser, 1998) and by
biochemical experiments carried out in human cells, demonstrating that Ero1 and
glutathione constitute the central element for ER redox homeostasis (Appenzeller-
Herzog et al., 2010). The story is not that straightforward though: the single gene
for Ero1 is non-essential in Drosophila melanogaster (Tien et al., 2008), and
deletion of both Ero1 isoforms creates viable double mutant mice (Zito et al.,
2010a). These observations argue that the eukaryotic ER contains an elaborate
network of Ero1-independent OPF pathways that may substitute the Ero1-PDI
pathway, at least in some organisms. The major pathways for disulfide formation
in the ER, that are formed by the PDI family members, the flavin-dependent
sulfhydryl oxidases, peroxidases, and important LMWOs, are reviewed next.
45
Fig. 8. The major pathways for disulfide formation in the human ER. (A) Schematic of
the main Ero1-PDI pathway for disulfide formation, and other major pathways that either
connect to the PDI hub or bypass it. (B) PDI family members and their domain
architecture. White box, active site or active site-like domain with active site (or -like)
motif marked; grey box, structural or substrate-binding domain; black oval,
transmembrane domain; white oval, J domain. The flexible x-linker, that is considered
to modulate the accessibility of the substrate-binding site (Nguyen et al., 2008), is
shown as a small black box between domains b' and a'.
46
OPF hub in the ER: the PDI family
PDI family members are at the center of OPF in the ER by connecting many
important oxidative pathways (Fig. 8A). The PDI family consists of more than
twenty oxidoreductases (Fig. 8B), and it is defined by localization to the ER and
similarity to the canonical PDI (from now on, the word PDI refers to this
archetypical PDI family member, and the family is discussed as PDI family). They
catalyze oxidation, reduction, and isomerization of disulfides utilizing a Cys-X-X-
Cys (where X is non-cysteine and can be the same or different amino acid) active
site motif. However, a few peculiar human PDI family members exist that have
either a Cys-X-X-Ser or Ser-X-X-Cys active site or do not have an active site at all
(Fig. 8B), and they are likely limited to isomerization or chaperone functions, only
(Cai et al., 1994; Appenzeller-Herzog & Ellgaard, 2008b; Hatahet & Ruddock,
2009; Kober et al., 2013).
Active site reduction potentials vary between PDI family members, and some
members may prefer to oxidize substrate dithiols, while others act primarily as
reductases. Although the structural aspects of PDI are reviewed in detail in Chapter
2.4.2, it is worthwhile to discuss already here the underlying determinants that
modulate the active site reduction potential and define the thiol-disulfide exchange
function of the PDI family member. Firstly, a charged residue in between the active
site cysteines has a major impact on defining the role of a PDI family member. For
example, a positively charged histidine lowers the pKa of the N-terminal active site
thiol. This may result in a destabilized active site disulfide, an increase in the
reduction potential, and a preference to oxidize other thiols (Hatahet & Ruddock,
2009). Inversely, if the positively charged histidine of the canonical CGHC motif
is mutated to a non-charged proline, the reduction potential decreases significantly
from ~-170 to ~-230 mV (cf. Fig. 4; Chambers et al., 2010). Secondly, the bulkiness
of the juxtaposed amino acids may influence the reduction potential of the active
site. A bulky proline next to the N-terminal cysteine (CPXC) may strain and
destabilize the active site disulfide, again favoring the oxidative function. Of note,
a proline in the second position does not destabilize the active site disulfide in a
similar manner, as reductases with a stable active site disulfide may have a CXPC
active site motif (Chivers et al., 1997; Hatahet & Ruddock, 2009). To summarize,
stability of the active site disulfide, which is strongly influenced by chemistries of
the juxtaposed amino acids, is the defining factor in deciding whether a PDI family
member primarily acts as an oxidase (unstable active site disulfide, prefers to
47
donate it away) or a reductase (stable active site disulfide, prefers to reduce others
by gaining a disulfide from them).
The exact physiological role of most PDI family members is still unclear. The
select few, that have been characterized, demonstrate a wide variety of functions
and tissue distributions:
1. PDI is a generalist, capable of each of the three described thiol-disulfide
reactions (reduction, oxidation, and isomerization). It exists as a mixture of
different redox states in the ER (circa 50% in fully reduced state and 50% in
semi- or fully oxidized state) (Appenzeller-Herzog & Ellgaard; 2008a).
2. ERp57 is critical for OPF of glycoproteins, and this function is strongly
influenced by interaction with the ER lectins calnexin and calreticulin
(Molinari & Helenius, 1999; Frickel et al., 2002; Pollock et al., 2004).
3. PDIp shows a peculiar tissue enrichment in the pancreas and the brain (Desilva
et al., 1996; Conn et al., 2004).
4. P5 controls the activity of some ER quality control signaling pathways by
reducing disulfides in the signaling components (Eletto et al., 2014).
5. ERp72 is a component of large multiprotein chaperone complexes involved in
biogenesis of immunoglobulins (Meunier et al., 2002).
6. ERdj5 is a dedicated reductase with three CXPC motifs involved in protein
quality control, namely endoplasmic reticulum-associated protein degradation
(ERAD) (for more information, see Chapter 2.5.4.) (Ushioda et al., 2008).
These examples demonstrate that the multitude of PDI family members has evolved
to have differential client repertoire to take part in different quality control features
of the ER - and not just to specialize in certain thiol-disulfide exchange reactions
for OPF.
Flavin-dependent sulfhydryl oxidases
Flavin-dependent sulfhydryl oxidases couple direct reduction of O2 within a FAD
redox center to de novo disulfide formation (Romero et al., 2018). This process
forms equimolar H2O2 (equation 3).
𝑂 2 ˗𝑆𝐻 → ˗𝑆˗𝑆˗ 𝐻 𝑂 (3)
They are net generators of disulfide bonds, whereas many other oxidoreductases
such as PDI family members solely exchange disulfides between thiols. Eukaryotic
flavin-dependent sulfhydryl oxidases have three major subfamilies: Erv1/ALR,
48
quiescin sulfhydryl oxidases (QSOX), and Ero1 (Fass, 2008; Kodali & Thorpe,
2010; Araki & Inaba, 2012). Whereas the Erv1/ALR and QSOX members reside
outside the ER (for example, Erv1p and ALR in the mitochondrial IMS and QSOX
mainly in the post-ER organelles of the secretory pathway or the extracellular
space), all Ero1 isoforms are ER-resident sulfhydryl oxidases. Saccharomyces
cerevisiae, Caenorhabditis elegans, and D. melanogaster express only one isoform
and most mammals, including Homo sapiens and Mus musculus, have two genes
for Ero1 that encode isoforms Ero1α and Ero1β. The main reasons for the
requirement of a second isoform have not been established explicitly. The isoforms
have differential tissue distribution under basal OPF levels: Ero1α is found from a
variety of tissues with high enrichment in esophagus, while Ero1β is enriched
highly in pancreas and other exocrine and endocrine tissues, including stomach and
testis (Pagani et al., 2000; Dias-Gunasekara et al., 2005; Uhlen et al., 2015). Under
stress conditions, both Ero1α and Ero1β have been suggested to be upregulated by
the unfolded protein response (UPR), but by different mechanisms (see Chapter
2.6.2). Importantly, Ero1α has been reported to be upregulated by hypoxia (May et
al., 2005; Gess et al., 2003), but the biochemistry of human Ero1 isoforms at low
oxygen concentrations has not been previously characterized. Despite the
differences, both isoforms seem to have a limited substrate repertoire in vivo, as
mixed disulfides have been reported to form with PDI family members only
(Benham et al., 2000; Mezghrani et al., 2001; Appenzeller-Herzog et al., 2008;
Appenzeller-Herzog et al., 2010). In vitro, PDI is the preferred substrate of all tested
PDI family members (Araki et al., 2013).
QSOX is the other flavin-dependent sulfhydryl oxidase that may exist in the
human ER. The uncertainty of its localization derives from two observations:
Firstly, QSOX contains a clear ER-targeting signal peptide, but it seems to traffic
to various intra- and extracellular places such as secreted fluids, constituents of the
extracellular matrix, plasma and nuclear membrane, and the Golgi (Kodali &
Thorpe, 2010). Secondly, it lacks an ER retention signal, and apparently only one
report by Tury and coworkers (2004) suggests that the membrane-associated form
may be retained in the ER. QSOX is expressed as several poorly characterized
isoforms and splice variants, including a membrane bound form and a cleaved
soluble form, so its localization to the ER is plausible (Kodali & Thorpe, 2010).
Why would the ER express multiple different flavin-dependent sulfhydryl
oxidases? One clear denominator that may eliminate redundancy is substrate
specificity. In contrast to Ero1 isoforms, QSOX seems to have low PDI-oxidase
activity, and it prefers to directly oxidize a variety of reduced unfolded protein
49
substrates, instead (Rancy & Thorpe, 2008). This segregation would allow targeted
oxidation of nascent polypeptides without redundant consumption of reduced PDI
which is required for isomerization – and it would also stress the role of Ero1 as
the main regulator of PDI redox state. Secondly, the structure of QSOX is peculiar:
it contains two thioredoxin-like domains connected to an Erv/ALR domain via a
helix-rich region. The first of the two thioredoxin-like domains contains the
canonical CGHC motif, found also in many PDI family members (Fig. 8B). The
Erv/ALR domain contains two additional CXXC motifs. Mechanistically, it is the
CGHC motif in the thioredoxin-like domain that transfers a disulfide to a folding
substrate. The CGHC motif is then reoxidized by a CXXC motif in the Erv/ALR
domain, and the catalytic site in the Erv/ALR domain is finally reoxidized by FAD
and O2. All in all, the differential substrate specificity, and the structural features of
QSOX would suggest that this enzyme has evolved to fulfill both roles, the central
oxidoreductase and the sulfhydryl oxidase to directly engage in substrate oxidation,
bypassing the PDI hub. This would allow the PDI pathway to remain specifically
under the control of Ero1. Whether QSOX is found in sufficient amounts in the ER
to form an Ero1-independent pathway under physiological conditions is to be
established.
Peroxidases
The sulfhydryl oxidases produce reactive oxygen species (ROS), namely H2O2 as
a side product of their catalytic cycle. In the ER, Ero1 is the main source for H2O2,
and this ROS remains in the ER without leaking to the cytosol (Harding et al., 2003;
Ramming et al., 2014; Konno et al., 2015). Accumulation of H2O2 may lead to
oxidative damage and protein misfolding that can be relieved by externally supplied
antioxidants (Malhotra et al., 2008; Karala et al., 2009). Therefore, it is feasible that
ER must have developed its own antioxidant defense to counter the unspecific
oxidative damage by H2O2.
Recently, three ER-resident peroxidases, peroxiredoxin IV (PrxIV) and
glutathione peroxidases (GPx) 7 and 8, have attracted attention as the main
scavengers of H2O2 in the ER. They very efficiently (Table 1) neutralize H2O2 to
water and form a new disulfide in the process (equation 4) (Utomo et al., 2004;
Tavender et al., 2010; Zito et al., 2010b; Nguyen et al., 2011).
𝐻 𝑂 2 ˗𝑆𝐻 → ˗𝑆˗𝑆˗ 2 𝐻 𝑂 (4)
50
Mechanistically, they generate a disulfide by the actions of a peroxidatic cysteine
(CP) and a resolving cysteine (CR) that are both reduced at the start of the reaction
cycle. CP reacts with H2O2 and is modified to contain sulfenic acid (CP-SOH). CR
then attacks CP-SOH forming a CP-CR disulfide, which can then be transferred to
other proteins (Tavender et al., 2008; Wang et al., 2014). As the peroxidase is in a
reduced state at the end of the catalytic cycle, and no other cellular resources are
consumed during the cycle, the net reaction (equation 5), that combines the partial
reactions by flavin-dependent sulfhydryl oxidases and the peroxidases, is two
disulfides formed from one molecule of O2.
𝑂 2 ˗𝑆𝐻 → ˗𝑆˗𝑆˗ 𝐻 𝑂 (3) 𝐻 𝑂 2 ˗𝑆𝐻 → ˗𝑆˗𝑆˗ 2 𝐻 𝑂 (4) 𝑂 4 ˗𝑆𝐻 → 2 ˗𝑆˗𝑆˗ 2 𝐻 𝑂 (5)
The only known efficient substrates for all the three peroxidases are PDI family
members (Tavender et al., 2010; Zito et al., 2010b; Nguyen et al., 2011; Wang et
al., 2014). Therefore, the peroxidatic disulfide formation is connected to the OPF
hub (Fig. 8A). Of note, the GPx naming convention is outdated and misleading, as
these two GPx isoforms do not have significant glutathione peroxidase activity
(Nguyen et al., 2011) - they are PDI peroxidases, instead.
The three peroxidases catalyze the same reaction in the same cellular
compartment. Do they have redundancy? Likely no, as the current knowledge
seems to place PrxIV in an Ero1-independent pathway, while both GPx are linked
to the Ero1-PDI pathway. Substrate specificity is again one clear denominator, as
GPx7/8 prefer to oxidize PDI, while PrxIV favors ERp46 and P5 over PDI (Nguyen
et al., 2011; Sato et al., 2013; Wang et al., 2014). Konno and colleagues (2015)
have even suggested that an unknown Ero1-independent lumenal H2O2 source
could fuel PrxIV, whereas the GPx isoforms interact directly with Ero1 and utilize
Ero1-derived H2O2 (Nguyen et al., 2011; Wang et al., 2014). The Ero1-
independency of PrxIV is further supported by knockdown studies. Loss of both
Ero1 isoforms in mice cells does not seem to markedly alter cell viability (Zito et
al., 2010a). However, mice cells lacking both Ero1 isoforms and PrxIV lose
viability (Zito et al., 2010b). These experiments are supported by yeast studies that
demonstrated that ER-localized PrxIV can rescue yeast cells with the lethal Ero1
knockout (Zito et al., 2010b). Collectively, these observations suggest that the GPx
isoforms contribute to the main Ero1-PDI pathway and PrvIV may bypass the PDI
51
hub and form an Ero1-PDI independent pathway for OPF in the ER. Speculatively,
this segregation, in combination with the above described PDI-independency of
QSOX, would place Ero1 in even stronger position to regulate the redox state of
PDI and the rates of PDI-driven oxidative protein folding.
Glutathione
Glutathione is a tripeptide (γ-glutamyl‐cysteinyl‐glycine) which contains a thiol
group. It was considered to form the main pathway for disulfide formation in the
ER before the discovery of Ero1 (Hwang et al., 1992). This is not surprising, as
glutathione is an abundant peptide in the ER (estimates range from 4 to 19 mM)
(Dixon et al., 2008; Birk et al., 2013) and capable of carrying out thiol-disulfide
exchange reactions either in the reduced monopeptide form (GSH) or the disulfide-
containing oxidized homodimer (GSSG). Both exchange reactions consist of two
steps that involve formation of a transient protein-glutathione mixed disulfide
intermediate species (Fig. 9). At first glance, the equations look like reverse
reactions of each other. However, the overall rate of reduction depends on GSH
concentration, as both steps of the overall scheme depend on [GSH]. This creates a
kinetic partitioning event for a potential reverse reaction and regeneration of the
starting material at low [GSH]. In contrast, the oxidation series has only one
direction, as the latter steps are independent of [GSSG] (Lappi & Ruddock, 2011).
Therefore, a careful control of redox conditions is a necessity to decide which thiol-
disulfide exchange reactions glutathione is to carry out.
Glutathione forms the main redox buffer in the ER. While the cytosolic GSH
to GSSG ratio has been reported to range from 30:1 to 100:1, values that translate
to a highly reducing environment, the ER is more oxidizing with GSH:GSSG ratios
of 3:1 to 6:1 (Hwang et al., 1992; Birk et al., 2013). This stark difference between
the organelles is affected mainly by two factors. Firstly, ER lacks glutathione
synthesis, and glutathione is transported to the ER in the reduced form from the
cytosol, potentially in a redox-regulated manner (Banhegyi et al., 1999; Ponsero et
al., 2017). Secondly, glutathione redox state dynamically both maintains the
reduced fraction of PDI family members (required for isomerization) and
reoxidizes target proteins (Chakravarthi & Bulleid, 2004; Appenzeller-Herzog et
al., 2010). Therefore, glutathione influences oxidation, reduction, and
isomerization of the ER disulfide pool at least indirectly via PDI family members
(Fig. 8A). To what extent it oxidizes nascent polypeptides directly in vivo is not
well understood. On one hand, glutathione reacts with thiols in unfolded proteins
52
with relatively slow kinetics in vitro (Table 1), arguing that it may not form a major
independent OPF pathway in vivo. On the other hand, the concentration of
glutathione in the ER is extremely high, which may lead to a high cumulative thiol-
disulfide exchange activity. Furthermore, the estimated basal GSH:GSSG ratio in
the ER (see above) was observed to be optimal for RNase A refolding in vitro (Lyles
& Gilbert, 1991). These observations suggest that glutathione has the capability for
efficient oxidation of protein thiols and could bypass the PDI hub to form a major
pathway for OPF (Fig. 8A).
Fig. 9. Reaction mechanisms for PDI reduction (upper reaction) and oxidation (bottom
reaction) by glutathione; adapted from Lappi et al. (2011). Both forward steps of the
reduction series are intermolecular reactions and depend on [GSH], which creates a
potential intramolecular reverse reaction from the mixed disulfide intermediate to the
starting material at low [GSH]. The oxidation series is unidirectional, as the later steps
are intramolecular reactions. The interconversion of the PDI C-terminal thiol to a thiolate
in the oxidation series has been suggested to be controlled by an intramolecular
conformational change that brings the positively charged side chain of Arg120 close to
the active site locale (Lappi et al., 2004; Karala et al., 2010; Lappi et al., 2011).
53
Vitamin-utilizing oxidative systems
Two vitamins and their derivatives potentially influence disulfide formation in the
ER (Fig. 8A). The first pathway utilizes vitamin K and the already described VKOR.
The main difference between the prokaryotic and the eukaryotic VKOR pathways
is that the former is an independent pathway, whereas the latter is coupled to
another metabolic pathway, γ-carboxylation of proteins (Stenflo et al., 1974; Furie
et al., 1999; Schulman et al., 2010). This process is driven by VKOR which reduces
vitamin K derivatives needed for the γ-carboxylation reaction. VKOR is then
recycled back to the reduced state by PDI family members which get oxidized in
the process (Schulman et al., 2010). VKOR pathway is therefore linked to the PDI-
driven OPF network (Fig. 8A). Of note, γ-carboxylation is a modification of a
minor subset of clients, and, as suggested by Ruddock (2012), this pathway is
unlikely to form a major pathway for disulfide formation. A prokaryotic-like
independent VKOR pathway could significantly contribute to the overall disulfide
pool, but such a pathway has not been discovered in the ER, yet.
The other vitamin that could influence disulfide formation is vitamin C
(ascorbate). It is an abundant molecule in the cell found possibly in millimolar
levels (Chaudiere & Ferrari-Iliou, 1999) and best known for its roles as an
antioxidant (Duarte & Lunec, 2005) and in collagen biosynthesis (Kivirikko et al.,
1989). The oxidized redox state of ascorbate, dehydroascorbate (DHA), can
additionally promote refolding of disulfide-containing proteins (Saaranen et al.,
2010; Szarka & Lorincz, 2014). Remarkably, it seems to favor oxidation of
unstructured proteins over the active site of PDI and oxidizes them more efficiently
than glutathione or H2O2 do (Table 1; Karala et al., 2009; Lappi & Ruddock, 2011).
A potential biochemical pathway for DHA formation in the ER has been described
(Zito et al., 2012). It combines reduction of protein cysteinyl-sulfenic acid
modifications to free thiols with concomitant ascorbate oxidation to DHA (Zito et
al., 2012). If the ascorbate to DHA interconversion rates are found to be sufficiently
high in the ER, possibly via the cellular pathway described, this vitamin could form
a major PDI-independent pathway for disulfide formation in the ER (Fig. 8A).
54
2.4 Structural biology of the Ero1-PDI relay
The human Ero1 isoforms have evolved to oxidize specifically PDI, which together
form a regulated key pathway for disulfide bond formation in the ER. This chapter
elucidates the important structural determinants for the specificity of the Ero1-PDI
interaction, and establishes the catalytic and regulatory elements of Ero1, which are
subsequently discussed in Chapters 2.5 and 2.6.
2.4.1 Structure of human Ero1 isoforms
Human Ero1α (445 amino acids, 52 kDa) and Ero1β (434 amino acids, 50 kDa)
share 65.4% amino acid identity and 74.8% similarity with conserved structural,
active site, and regulatory cysteines (Fig. 10A; Pagani et al., 2000). The isoforms
contain two active sites: a CXXC inner active site (Cys394/Cys397 in Ero1α and
Cys393/Cys396 in Eroβ) and a CX4C outer active site (Cys94/Cys99 in Ero1α and
Cys90/Cys95 in Eroβ). The remaining cysteines are either structural or regulatory
in nature. Cys131 in Ero1α forms a key regulatory switch for modulating Ero1
activity (Appenzeller-Herzog et al., 2008; Baker et al., 2008; Appenzeller-Herzog
et al., 2010). This function is discussed more in Chapter 2.6.1. The other proposed
regulatory cysteines, Cys208 and Cys241 in Ero1α, potentially control
conformational change to create a diffusion tunnel for the substrate O2 and the
product H2O2 (see Chapter 2.5.3; Ramming et al., 2015). Three cysteines are
controversial: Cys166 in Ero1α has been regarded non-functional, and many in
vitro studies mutate it to alanine (for example, Inaba et al., 2010; Araki & Nagata,
2011; Masui et al., 2011). However, it is potentially functional, as Appenzeller-
Herzog et al. (2008) suggested that Cys166 may form a disulfide to an as-yet-
unknown partner. Cysteines 85 and 391 form a disulfide that was initially
considered regulatory (Baker et al., 2008) but was later observed to be stable and
therefore likely structural (Araki & Nagata, 2011; Zhang et al., 2014). Whether this
disulfide is reduced under some conditions, like the counterpart in yeast Ero1p
(Sevier et al., 2007), remains to be established. The structural features of Ero1β are
poorly characterized and are typically correlated to those of Ero1α, based on high
conservation of the established residues. The isoform specific Cys262 of Ero1β is
buried and likely does not participate in intra- or intermolecular disulfide bond
(Hansen et al., 2014).
55
Fig. 10. Structural overview of human Ero1 isoforms and PDI. (A) Disulfide patterns of
the inactive states of human Ero1 isoforms. Circles depict cysteines (active site
cysteines are yellow, regulatory cysteines red, and structural cysteines or of unknown
function are grey) and black lines are disulfides. The functions of cysteines 166 and 165
have been unknown before this study, but Appenzeller-Herzog et al. (2008) suggested
that Cys166 may form a disulfide to an unknown partner. The amino acid numberings
are for full-length proteins. (B-C) Crystal structures of inactive Ero1α (PDB code 3AHR)
(B) and PDI (subunit of MTP, PDB code 6I7S) (C). Active site cysteines (and the
regulatory Cys131) and potential disulfides connecting them are colored yellow, FAD
blue, and other elements as per in Fig. 3.
Ero1α is a single-domain enzyme based on two existing crystal structures of mutant
forms that fix Ero1α either in an inactive state or in a state with increased activity,
also called hyperactive state (the inactive state is shown in Fig. 10B) (Inaba et al.,
2010). The FAD cofactor is juxtaposed to the inner active site cysteines and buried
in both structures within a four-helix bundle, a common structural element found
in many other flavin-dependent sulfhydryl oxidases, e.g., Erv1p, Erv2p, ALR, and
56
QSOX (Gross et al., 2002; Wu et al., 2003; Alon et al., 2010; Guo et al., 2012). The
cysteines in the outer active site and the regulatory cysteines 104 and 131 are in a
flexible loop (residues 90-131) which is missing in the structural model of the
hyperactive form and partially missing (residues 95-130) for the inactive state. The
inactive state confirms the existence of the regulatory Cys94-Cys131 disulfide (Fig.
10B). Two other flexible loops are absent in the structures: residues 166-172,
containing the controversial Cys166, and residues 212-238 flanked by the
regulatory cysteines 208 and 241. Other distinguishing features are two
hydrophobic residues mediating PDI interaction: Val101 near the outer active site,
and Trp272 at the tip of a β-hairpin structure (residues Arg265-His279) (Fig. 10B).
Val101 has been shown to be important for efficient PDI oxidation but not for
regulation of Ero1α activity (Zhang et al., 2019). While Trp272 is also crucial for
PDI oxidation (Masui et al., 2011), its roles for any regulatory functions have not
been reported before this work.
2.4.2 Architecture of PDI
Human PDI contains four domains, a, b, b', and a', with each domain having a
thioredoxin-like fold (defined by regular secondary structure elements β-α-β-α-β-
β-α) (Fig. 10C). PDI also contains an acidic C-terminal tail termed c and a linker
region of 19 amino acids called x between domains b' and a'. Together, the domains
and the extensions add up to 491 amino acids and 55 kDa (Freedman et al., 1998;
Alanen et al., 2003). The active sites for thiol-disulfide exchange are in the a and
a' domains. Other domains are non-catalytic: b is likely structural and b' forms the
principle substrate binding site (Fig. 10C). PDI has six cysteines of which four
(Cys53 and Cys56 in a, Cys397 and Cys400 in a’) are in the active site Cys-Gly-
His-Cys (CGHC) motifs, and the remaining two cysteines are in the b' domain and
have no reported functions, excepting S-nitrosylation of Cys343 which has a link
to neurodegenerative diseases (Ogura et al., 2020).
Three crystal structures of (near) full length mature human PDI have been
published: two monomers either in the oxidized or reduced redox states, both
lacking the flexible C-terminal tail (Wang et al., 2013); and a full length PDI as the
beta subunit of the microsomal transfer protein (MTP) (Biterova et al., 2019). The
two monomeric structures show slight conformational differences, mainly how the
a and a' domains are situated around the rigid b-b' axis. The reduced state is ‘closed’
with the a and a’ domains closer to each other, while the oxidized state is more
open and accessible for larger substrates. The open and closed conformations were
57
originally suggested to reflect the catalytic oxidation mechanism, so that in the open
oxidized state PDI binds substrates and releases them in the closed reduced state
(Wang et al., 2013). The real mechanism is likely more complicated though, as full-
length PDI bound to the large alpha subunit of MTP is in the reduced state with an
open conformation (Biterova et al., 2019). The open and closed conformations may
therefore only reflect conformational flexibility of PDI for binding substrates of
different sizes. Regardless, PDI in all the three crystal structures has a U-shaped
structure with the pocket between the domains forming the substrate-binding site
(Fig. 10C).
The PDI active site cysteines are located at the N-termini of the second alpha
helices of the a and a' domains (Fig. 5B, Fig. 10C). While the first cysteine of the
CGHC motif is solvent exposed and located at the N-terminus of the helix, the C-
terminal cysteine is partially buried having limited solvent exposure. The pKa
values of the active site cysteine thiol groups are different and influenced mainly
by two factors: 1) the structure of α-helix, as the N-terminus of the helix has a
permanent positive and the C-terminus permanent negative dipole moment causing
slight decrease in the pKa of the N-terminal cysteine (Kortemme & Creighton,
1995); 2) electrostatic effects from nearby residues (Fig. 5A), for example, from
the histidine residue within the CGHC motif which is partially protonated and has
a net positive charge at physiological pH. The N-terminal cysteine adopts a thiolate
form at physiological pH, as the lowest reported pKa is 4.5 (Kortemme et al., 1996).
In contrast, the pKa value of the C-terminal cysteine is high (up to 12.8; Lappi et
al., 2004), so it will be in the non-charged thiol form for the most part of the
catalytic cycle of PDI. However, the pKa of this residue may be modulated by
conformational changes in the catalytic domain resulting in a decrease of the pKa
value and formation of the thiolate group. This is important during reoxidation of
the active site when the C-terminal cysteine needs to resolve the mixed disulfide
bond between the PDI N-terminal cysteine and a substrate cysteine (discussed in
Chapter 2.5.2).
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2.5 Structure-function relationship of Ero1-PDI
Efficient substrate oxidation by the Ero1-PDI relay involves intermolecular steps
between the folding substrate and oxidized PDI as well as between Ero1 and PDI
to reoxidize PDI. Intramolecular electron transfer within Ero1 and to O2 will
reoxidize Ero1 to complete the catalytic cycle (Fig. 11). This chapter compiles
reported structure-function details of each step of the catalytic cycle.
2.5.1 Substrate oxidation by PDI
PDI must first bind the substrate for efficient thiol-disulfide exchange. The binding
is mediated by hydrophobic interactions between exposed hydrophobic substrate
residues and hydrophobic residues on the surface of the U-shaped substrate-binding
pocket of PDI (Fig. 10C). b’ forms the principal binding site and is alone sufficient
to bind small substrates such as peptides. However, both a and a’ are additionally
needed for binding larger substrates, for example, non-native proteins (Klappa et
al., 1998) or subunits in protein complexes (Koivunen et al., 2005; Biterova et al.,
2019). The U-shaped binding surface can also form a hydrophobic cavity, when
two PDI monomers form a face-to-face dimer in the oxidized state, potentially
increasing the rate of OPF (Okumura et al., 2019). The binding specificity of PDI
is poorly understood. Whereas PDIp requires a single tyrosine or a tryptophan
residue for efficient binding (Ruddock et al., 2000), PDI has several low affinity
binding sites that accept different hydrophobic residues, and, consequently, it has a
broad binding-capability for non-native proteins. Of note, thiol-disulfide exchange
is not prerequisite for substrate binding, as PDI can also bind various substrates
without any disulfides (Hatahet & Ruddock, 2007).
The thiol-disulfide exchange reaction from an oxidized PDI active site to the
reduced substrate dithiol proceeds essentially with the productive pathway of the
reaction scheme depicted in Fig. 6. This involves mixed disulfide bond formation
between the N-terminal active site cysteine of PDI and a substrate thiol followed
by a subsequent nucleophilic attack of the second substrate thiol to release oxidized
substrate (Fig. 11). To avoid a potential reverse reaction from the mixed disulfide
intermediate, the reactivity of the C-terminal active site cysteine of PDI during this
step is low (high pKa).
59
Fig. 11. Molecular mechanisms of disulfide formation by the Ero1-PDI relay. Substrate
oxidation by PDI and PDI oxidation by Ero1 are carried out by the thiol-disulfide
mechanism described earlier in this study; briefly, oxidized PDI forms mixed disulfide
with a reduced substrate, a substrate thiol resolves the mixed disulfide to form an
oxidized substrate, the reduced PDI is then reoxidized by an oxidized Ero1 outer active
site. For the catalytic cycle to continue, Ero1 outer active site must be reoxidized. This
is carried out by the inner active site that gains a disulfide by de novo oxidation by
FAD/O2. For efficient PDI catalytic cycle and PDI reoxidation, the reactions depicted
must be unidirectional and any reverse reactions (small arrows) must be inhibited. The
pKa of the C-terminal thiol of PDI during substrate oxidation is very high which
eliminates one of the reverse reactions depicted. The pKa values of Ero1 thiols are not
known, but one mechanism that may control the directions of Ero1 reactions is the
mobility of the outer active site which shuttles between the surface of the protein and
the inner active site (Gross et al., 2004).
60
2.5.2 PDI reoxidation by Ero1
Reduced PDI, once dissociated from the oxidized substrate, may bind Ero1 for
reoxidation (Fig. 11). Oxidized PDI will compete for binding oxidized Ero1, but
reduced PDI has approximately 10-fold higher affinity towards the active state of
Ero1 (Masui et al., 2011). The minimal PDI fragment that interacts with Ero1 is
b’xa’ (Wang et al., 2009). The interaction is thought to be mediated by hydrophobic
interactions: Ero1 Trp272 at the tip of the β-hairpin binds to the b’ domain of PDI,
potentially to residues Phe240 or Phe304 (Masui et al., 2011); and Ero1 Val101
recruits the PDI a’ domain to the vicinity of the Ero1 outer active site (Zhang et al.,
2019).
Current knowledge suggests that human Ero1 oxidizes specifically the a'
domain of PDI (Baker et al., 2008; Wang et al., 2009). The a domain is possibly
oxidized by intramolecular electron transfer between the PDI active sites (Araki &
Nagata, 2011) and/or intermolecularly by PDI peroxidases (Wang et al., 2014).
Mechanistically, the disulfide transfer between the Ero1 outer active site and the
PDI a' domain utilizes the described thiol-disulfide exchange mechanism. An Ero1-
PDI mixed disulfide complex can be trapped and coimmunoprecipitated from
mammalian cells or abrogated by Cys94Ser mutation (Appenzeller-Herzog et al.,
2010). It is tempting to speculate that in order to release the oxidized PDI (and
avoid the reverse reaction that reforms the oxidized Ero1), the mixed disulfide
might be resolved by the same principles that were described for the resolution of
the PDI-glutathione mixed disulfide (Fig. 9). In this cycle, the positively charged
side chain of PDI Arg120 is first brought close to the PDI active site locale by a
conformation change. The pKa of the PDI C-terminal active site cysteine is lowered
as a result, and the C-terminal thiol is converted to a thiolate. The more reactive
thiolate state can then resolve the PDI-Ero1 mixed disulfide intermediate, releasing
the oxidized PDI and reduced Ero1.
2.5.3 Ero1 reoxidation
PDI reoxidation results in a reduced Ero1 outer active site. Oxidation of Ero1
combines de novo disulfide formation in the core of Ero1 to thiol-disulfide
exchange reaction to pass the newly generated disulfide from the inner active site
to the outer active site. Therefore, reoxidation of Ero1 consists of a complex series
of chemistries, and some of the specifics are poorly understood. Therefore, for an
61
overall picture of Ero1 reoxidation, we need to review mechanisms from other
characterized flavoenzymes.
The FAD cofactor needs to be oxidized to be able to form a new disulfide to
the inner active site. A typical oxidase flavin cycle proceeds via formation of a
transient radical pair, in which the FAD is in a semi-reduced state and O2 has formed
a superoxide anion (Romero et al., 2018). Typically, the superoxide anion then
reacts to form H2O2 to oxidize FAD, but sometimes superoxide may escape. For
example, ALR releases in vitro one-third of its ROS product as superoxide
(Daithankar et al., 2012). The ratio of the side products for human Ero1 is not
known. The overall reoxidation reaction is spontaneous and irreversible as the
redox potential of the O2/H2O2 pair is significantly higher (+300 mV; see Fig. 4B)
than that of typical protein-bound flavins (+150 to -400 mV) (Romero et al., 2018).
Is O2 the only terminal electron acceptor for human Ero1? Certain sulfhydryl
oxidases are conditional oxidases switching from one electron acceptor to another
based on cellular conditions. For example, the yeast Ero1 homolog, Ero1p, is a
conditional oxidase that alternates between O2 and fumarate under aerobic and
unaerobic conditions, respectively (Kim et al., 2018). The yeast mitochondrial
sulfhydryl oxidase Erv1p consumes oxygen in the test tube (Ang & Lu, 2009; Tang
et al., 2019) but in vivo prefers to pass electrons to cytochrome c (Bihlmaier et al.,
2007; Dabir et al., 2007). Any alternative electron acceptors for human Ero1 have
not been identified, yet.
With the FAD cofactor now oxidized, the inner active site gains a disulfide by
series of chemistries that involve formation and resolution of a covalent adduct
between an inner active site thiol and FAD (Romero et al., 2018). The newly
generated disulfide can then be transferred from the inner active site to the
incoming mobile outer active site by thiol-disulfide exchange (Fig. 11).
Oxygen tunneling
O2 must reach the core of the flavin-dependent oxidase to re-oxidize FAD. Typically,
this is achieved by transient oxygen diffusion tunnels that are formed by dynamic
conformational changes (Romero et al., 2018). A diffusion tunnel has been
proposed for human Ero1α (Ramming et al., 2015). Based on molecular
simulations, the four-helix bundle surrounding the FAD moiety could undergo
conformational change to open a tunnel for substrate O2 and product H2O2 with a
gate (a disulfide between cysteines Cys208 and Cys241) at the distal side of the
enzyme. The gating mechanism could be controlled directly by PDI, which in the
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reduced form attacks the gating disulfide opening the helix bundle for oxygen entry
(Ramming et al., 2015).
2.6 Quality control of oxidative protein folding
The eukaryotic ER requires flexibility to maintain a balance between the high
folding load and the folding capacity provided by folding factors. If the proteostasis
is destabilized, misfolded proteins may accumulate causing stress and ultimately
even disease states (Mossuto, 2013). The ER can regulate its folding homeostasis
mainly by three mechanisms: i) modulate Ero1 activity, ii) upregulate folding
factors by UPR, and iii) directly remove misfolded protein by ERAD (Fig. 12). The
following chapters describe these main quality control systems in more detail.
2.6.1 Modulation of Ero1 activity
Uncontrolled disulfide formation by Ero1 can lead to accumulation of ROS and
hyperoxidation of PDI which may lead to redox and protein-folding imbalances
and diseases (Cao & Kaufman, 2014). To avoid these, Ero1 activity can be
regulated. Two mechanisms to alter human Ero1 activity have been previously
reported: intramolecular disulfide rearrangement by the inactivation switch
(Cys131 in Ero1α and Cys130 in Ero1β) and post-translational phosphorylation. Of
the two mechanisms, the inactivation switch is more important for downregulation
of ROS and redox state of the ER, as it defines both the on/off states of Ero1.
Mechanistically, to inactivate Ero1α, Cys131 pairs to the active site Cys94 (Fig.
10A) effectively blocking the outer active site of forming a disulfide for PDI
oxidation (Appenzeller-Herzog et al., 2008; Baker et al., 2008; Wang et al., 2011).
This rearrangement process is thought to include also active site Cys99 forming a
disulfide to Cys104 (Fig. 10A; Hansen et al., 2012). The regulatory switch is
controlled directly by PDI in a feedback manner, so that increased levels of reduced
PDI in the ER lead to reduction/rearrangement of the regulatory switch, activation
of Ero1, and generation of new disulfides to the free Cys94-Cys99 outer active site
to oxidize PDI. The formation of the Cys94-Cys131 link, i.e., inactivation of Ero1
may utilize both autonomous and PDI-dependent oxidation of the regulatory
disulfides, when oxidized PDI accumulates, instead (Kim et al., 2012; Shepherd et
al., 2014; Zhang et al., 2014). Kinetic details of either process (activation or
inactivation) have been unknown before this study.
63
Fig. 12. Quality control pathways of protein maturation in the ER, from the unfolded (U)
state via disulfide-containing intermediates (I) to the native (N) state. Efficient
maturation of disulfide-containing proteins to the N state combines oxidation to form
disulfides and isomerization/reduction of (non-native) disulfides by PDI family members.
Reduction is required also for efficient ERAD to avoid accumulation of non-native
proteins and aggregate formation (not depicted). UPR may increase folding capacity of
the ER. The redox states of the ER and PDI modulate activity of Ero1 in a feedback
manner: in reducing conditions, the high levels of reduced PDI (PDIred) activate Ero1,
resulting in increased levels of oxidized PDI (PDIox). In more-oxidizing conditions,
proportion of PDIred decreases, and the levels of inactive Ero1 increase. Only the N state
is efficiently exported from the ER to the organelles of the secretory pathway, including
the Golgi, and ultimately to the extracellular space.
64
The activity of Ero1 was recently suggested to be modulated by a second activity
switch, phosphorylation of Ero1α Ser145. This switch is controlled by a kinase,
FAM20C, which phosphorylates Ero1α in Ero1α-overexpressing HeLa cells in the
Golgi apparatus (with retrograde translocation to the ER by ERp44) and mouse
mammary glands (Zhang et al. 2018). Ser145Glu mutant, which mimics
phosphorylation at this site, enhanced in vitro oxygen consumption activity as well
as J-chain folding in HeLa cells (Zhang et al. 2018). The physiological importance
of this switch for maintaining the redox balance of the ER is still unclear, as the
abundance of the phosphorylated species in vivo as well as any phosphatase for
reversibility of the regulatory switch have not been established, yet.
2.6.2 Unfolded protein response (UPR)
The mammalian ER has evolved an elaborate network of signaling cascades to
battle folding stress, collectively termed the UPR. They ensure that the redox and
protein-folding homeostases are sustained, if the direct modulation of Ero1 activity
is insufficient to maintain the balance. UPR-like mechanisms are crucial also for
other functions such as plasma cell differentiation (Janssens et al., 2014).
The mammalian UPR consists of three signaling cascades mediated by three
different membrane protein families: ATF6, which contains isoforms A and B; IRE1,
with isoforms alpha and beta, and PERK. The goal of each cascade is to transduce
signals from the ER via cytosol to the nucleus to produce various transcription
factors which activate target genes to produce proteins that aim to restore the ER
homeostasis (Fig. 12). The pathways are distinct, they work in parallel, and utilize
unique molecular mechanisms for signal transduction (Walter & Ron, 2011; Hetz
& Papa, 2018).
The first and foremost method to avoid prolonged unfolded protein stress is to
increase the folding capacity to enhance client oxidation and isomerization. This is
carried out mainly by the ATF6 and IRE1 pathways which, as an early response,
upregulate folding factors such as Ero1β (but not Ero1α), PDI family members, and
molecular chaperones as well as components to regulate lipid biosynthesis, ER
expansion and ERAD (Yoshida et al., 1998; Pagani et al., 2000; Yoshida et al., 2003;
Shaffer et al., 2004; Sriburi et al., 2004; Yamamoto et al., 2007; Adachi et al., 2008;
Bommiasamy et al., 2009; Quan et al., 2018). Secondly, IRE1 and PERK reduce
the folding load directly by inhibiting further protein translation (Harding et al.,
2000; Hollien et al., 2009). In a prolonged stress, the PERK pathway may activate
65
pro-apoptotic genes, including the gene encoding for Ero1α, to start programmed
cell death, apoptosis (Marciniak et al., 2004; Li et al., 2009).
Molecular chaperones are prominent targets of the quality control systems of
the ER, specifically, UPR and ERAD (Hetz & Papa, 2018). These targets include
BiP of the HSP70 family which is involved in wide variety functions, from folding
to signaling to protein translocation across the ER membrane (Matlack et al., 1999;
Pobre et al., 2019); GRP94 which is involved in protein folding as well as immune
and antitumor responses (Luo & Lee, 2013); ERp27, a peculiar PDI family member
upregulated by the UPR which contains only the non-catalytic b-type PDI domains
(Fig. 8B) and binds hydrophobic targets as well as ERp57 in a calreticulin/calnexin-
manner (Alanen et al., 2006; Kober et al., 2013); among numerous other ER
chaperones (Fewell et al., 2001). Important for this study, a novel chaperone,
pERp1, is introduced next which is involved in an often-neglected function of
molecular chaperones, assembly of protein quaternary structure (Makhnevych &
Houry, 2012).
pERp1
pERp1 (also known as Mzb1) is a novel 18 kDa soluble ER-resident protein. It
belongs to the canopy (CNPY) protein family which consists of predicted saposin-
like proteins (SAPLIPs). SAPLIPs are defined by an alpha-helical rich fold and six
cysteines that potentially form three disulfide bonds (Shimizu et al., 2009; van
Anken et al., 2009; Schildknegt et al., 2019). pERp1 is expressed in low levels in
B and T lymphocytes, but it is strongly upregulated by a UPR-like mechanism
during plasma cell differentiation. Its link to OPF arises from observations that it
promotes assembly and secretion of immunoglobulins IgA and IgM and influences
activation and function of some integrins (Shimizu et al., 2009; van Anken et al.,
2009; Flach et al., 2010; Andreani et al., 2018; Xiong et al., 2019). The role in IgM
biosynthesis is especially noteworthy. IgM is a challenging ER client, as it can exist
as a pentamer or hexamer of Ig subunits, i.e., it contains at least twenty subunits -
10 light chains and 10 heavy chains which are stabilized by a J chain subunit - and
by more than hundred disulfide bonds (Muller et al., 2013). Knockdown of pERp1
in plasma cells leads to a severe defect in IgM assembly and secretion, suggesting
that pERp1 plays a major role in our immune system (van Anken et al., 2009).
Mechanistic details of how pERp1 promotes OPF of immunoglobulins are still
unclear. Originally, pERp1 was suggested to be an oxidoreductase, as it contains a
CXXC motif, has slight oxidoreductase activity in vitro, and possibly
66
immunoprecipitates in covalent complexes with IgM monomers (Shimizu et al.,
2009; van Anken et al., 2009). However, the covalent nature of the interaction with
IgM was challenged by van Anken and coworkers (2009) who showed that the
presence of a reducing agent DTT did not influence the results of another
immunoprecipitation experiment. A redox state analysis of cysteine mutants
supported this, as cysteines in the CXXC motif seemed to link to other cysteines
far away in the sequence and not form a PDI-like active site. Further cellular studies
suggested that another potential role could be chaperone-assisted prevention of off-
pathway IgM assembly intermediates (heavy chain oligomers) or as a cochaperone
in GRP94 or BiP complexes (Shimizu et al., 2009; Rosenbaum et al., 2014). No
structure or direct biochemical assays for chaperone activity have been reported
before this study to further elucidate its roles as a folding factor.
2.6.3 Endoplasmic reticulum-associated protein degradation (ERAD)
The other quality control mechanisms described herein do not always ensure that
protein molecules fold successfully to the native state. In fact, protein folding is
rather inefficient, and a large portion of ER clients fail to fold correctly (Smith et
al., 2011). The ER has a special pathway called ERAD to monitor and clear out any
terminally misfolded proteins. The same mechanism is also utilized to degrade
some correctly folded and active ER-resident proteins to regulate metabolic
pathways (Foresti et al., 2013; Ruggiano et al., 2014).
ERAD consists of three steps:
1. recognition and processing of ERAD substrates,
2. retrotranslocation across the ER membrane to the cytosol, and
3. cytosolic ubiquitination and proteosomal degradation.
High efficiency of recognition is required to avoid accumulation of misfolded
proteins. On the other hand, the system cannot be overactive, removing productive
folding intermediates for degradation (Ruggiano et al., 2014). One recognition
mechanism of glycosylated proteins is well characterized: a protein complex
consisting of EDEM, ERdj5, and BiP extracts misfolded glycoproteins, but not
folding-competent glycoproteins, from the calnexin quality control cycle, adds a
mannose signal to the glycan to target it for retrotranslocation, and unfolds the
protein for the retrotranslocation and the cytosolic degradation processes
(Hosokawa et al., 2001; Molinari et al., 2003; Oda et al., 2003; Quan et al., 2008;
67
Ushioda et al., 2008). In addition to ERdj5, PDI has been implicated as a potential
reductase for ERAD processing (Molinari et al., 2002; He et al., 2015).
Is disulfide reduction mandatory for ERAD? Several biochemical and
structural studies suggest that it is the reduced unfolded state of the client that is
efficiently retrotranslocated through the ER membrane. For example, disulfide
reduction has been directly observed prior to ERAD for some substrates (Mancini
et al., 2000; Fagioli et al., 2001; Okuda-Shimizu & Hendershot, 2007), redox
conditions of the ER play a major role in controlling ERAD efficiency (Tortorella
et al., 1998; Medrano-Fernandez et al., 2014), and the retrotranslocation channel is
too narrow to fit folded proteins (Schoebel et al., 2017; Ellgaard et al., 2018). As
disulfide reduction seems to be an important prerequisite for ERAD, the main
question under debate is that how are the dedicated reductases such as ERdj5 and
PDI kept in the reduced state? A possible answer is protein thiols, as they have been
suggested to form a larger thiol redox buffer than glutathione itself (Hansen et al.
2009). A likely explanation is also that the reduction of the reductases is connected
to the cytosolic thioredoxin pathways, analogous to the prokaryotic and
mitochondrial reducing pathways (Fig. 7). However, a membrane component that
would transfer electrons between the organelles has not been identified, yet.
Another puzzling problem is a potential futile cycle created by parallel client
oxidation by an active Ero1-PDI relay and the prerequisite of reduction for ERAD
(Fig. 12). Both processes have been characterized separately and are supposedly
active when the concentration of non-native proteins increase in the ER – by the
regulatory feedback and signaling mechanisms described. However, any crosstalk
between the systems, which might help to avoid the futile cycle, has not been
described. Speculatively, such regulation would require either spatial segregation
of the systems, or that they are mutually exclusive systems in the same space by
inverse modulation of their activity.
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69
3 Aims of the present work
The Ero1 isoforms and PDI form the main pathway for disulfide formation in the
human ER. The previous in vitro studies (for example, Baker et al., 2008; Araki &
Nagata, 2011; Araki et al., 2013) have consistently reported a triphasic activity trace
for Ero1 consisting of an initial redox activation lag phase, a linear maximal rate
period during the catalytic cycle, and a rapid decline of activity at low [O2].
Considering the importance of the Ero1-PDI relay for OPF in the ER, it was
surprising that the previous in vitro studies focused mainly on the kinetics of the
catalytic cycle, and only minor details have been reported for the other two phases
that likely contain information for regulation of Ero1 function. We hypothesized
that one reason for this has been a lack of proper enzyme kinetic tools to study the
complex kinetics of the Ero1 system. The first aims were therefore to generate
proper analytical tools to model the triphasic kinetics, characterize each of the three
phases, and reveal novel aspects of Ero1 function connected to the redox activation
and utilization of substrate O2. Secondly, as the Ero1-PDI relay forms the main
redox controller of the ER (Appenzeller-Herzog et al., 2010), the results obtained
may contain important information for the overall quality control homeostasis of
the ER. In other words, novel regulatory functions of the Ero1-PDI relay may reveal
new insight into how the ER maintains balance between efficient maturation of
disulfide-containing proteins and avoids hyperoxidation of clients. The latter would
have detrimental effects, if uncontrolled, for other quality control features such as
ERAD that utilizes reduced protein substrates (Ellgaard et al., 2018).
The final aim of this work was to gain insight into the non-catalytic aspects of
protein folding. Specifically, we undertook structural studies of a novel protein,
pERp1, which has been suggested to be a chaperone and shown to be a crucial
component of the biogenesis of immunoglobulins as well as other complex
disulfide-rich proteins (see Chapter 2.6.2). The main aim of this project was to
solve the crystal structure of pERp1, which might reveal new insight into the poorly
understood structure-function relationship of how pERp1 promotes
immunoglobulin maturation in the ER.
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71
4 Methods
The main methods used in this study are listed in Table 2. Further information is
provided in the referred articles.
Table 2. A list of methods and their references.
Method Original publications
Molecular biology techniques
ꞏ Construction of expression vectors I, II, III
ꞏ Generation of expression strains I, II, III
Protein expression and purification I, II, III
Protein analysis
ꞏ Sodium dodecyl sulfate-polyacrylamide gel
electrophoresis (SDS-PAGE) and sample preparation
I
ꞏ Circular dichroism (CD) and analytical reversed phase
high-performance liquid chromatography (rpHPLC)
I
ꞏ Protein concentration and determination of FAD content I
Protein functional studies
ꞏ PDI exchange assay I
ꞏ Ero1 activity assay – oxygen electrode I, II
ꞏ Ero1 activity assay – gel based I
ꞏ Binding assay – isothermal titration calorimetry (ITC) II
ꞏ Binding assay – bilayer interferometry (BLI) II, III
ꞏ Chaperone assay III
Generation and validation of in silico methods for Ero1 kinetics I
Crystallography – crystallization, data collection and
processing, structure determination
III
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73
5 Results
The main results of I-III are briefly introduced in this section. Chapter 5.1 details
kinetic characterization of the two human Ero1 isoforms, revealing novel
regulatory mechanisms. Chapter 5.2 follows these observations and suggests an
uncharacterized cross-talk between OPF and other quality control features of the
ER, namely ERAD. Chapter 5.3 describes structure-function relationship of pERp1,
a potential chaperone involved in biogenesis of IgA, IgM, and integrins.
5.1 Molecular analysis of human Ero1
5.1.1 Human Ero1 and PDI form a novel Ero1-PDI complex
Human wild type Ero1 isoforms were produced in a folded state in E. coli using the
CyDisCo system. A redox state analysis of purified Ero1 revealed stable mixed
disulfide complex formed between Ero1 and PDI (Fig. 1B in I), but not between
Ero1 and ERp57 (Fig. S3 in I). Both Ero1 isoforms in their respective complexes
were oxidized and contained long-range disulfides, as they migrated faster on non-
reducing gel (Fig. 1B in I), resembling the previously reported inactive OX states
(Benham et al., 2000; Dias-Gunasekara et al., 2005). Other biophysical
experiments (Table S1 in I and S2 in I, Fig. S1 in I, Fig. S2 in I) and FAD:complex
ratio (0.99 ± 0.07 FAD per Ero1α complex; 0.92 ± 0.13 FAD per Ero1β complex)
suggested that the Ero1 isoforms were natively folded.
PDI forms transient mixed disulfides to the Ero1 outer active site cysteines as
well as to the adjacent regulatory cysteines (see Fig. 9 and Chapter 2.6.1). The
stable complexes reported here formed for other reasons than these, as Cys to Ala
mutations in the Ero1 active site and/or the adjacent regulatory cysteines did not
abolish the complexes (Fig. 1C in I). Interestingly, the mixed disulfide formed
between a strongly conserved (see discussion in I) Cys166 in Ero1α or Cys165 in
Ero1β (Fig. 1D in I) and the N-terminal active site cysteine (Cys397) of the a'
domain in PDI (Fig. 1E in I). Cys166 has previously been considered non-
functional (Araki & Nagata, 2011) and is often mutated in in vitro functional studies
(for example, Araki & Nagata, 2011; Masui et al., 2011; Araki et al., 2013).
A redox state analysis revealed that PDI in the complex was a dynamic partner
to Ero1, as exogenous higher-molecular-weight PDI with three FLAG tag
sequences (PDI3FLAG) incorporated to the complex replacing the wild type PDI
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(Fig. 1F in I). The molar ratio of incorporated PDI to the original PDI could be
modulated by varying the concentration of the supplied PDI variant (Fig. 1F in I).
The results were similar if the redox state of all components were oxidized by
GSSG prior to the analysis. None of other tested PDI family members could replace
the complexed wild-type PDI in similar manner (Fig. S4 in I).
Collectively, these data suggest that Cys166 and 165 are functional, and that
they may interact covalently with PDI. The mixed disulfide complexes were stable
in the absence of exogenous PDI, while supplemented PDI exchanged (relatively
slowly in the tested oxidizing conditions) with the complexed PDI, suggesting that
PDI is a dynamic partner instead of a permanent subunit, when using the
Cys166/165 site.
5.1.2 Generation of a non-linear regression method to analyze Ero1
kinetics
The purified wild type Ero1-PDI complexes, with Ero1 in apparent inactive state,
were expected to undergo redox activation and show catalytic activity in the
presence of exogenous reduced PDI. To test this, we recorded a comprehensive
kinetic data set by measuring oxygen consumption in the presence of the reducing
PDI/GSH natural substrate system (Baker et al., 2008). The resulting oxygen
consumption traces consisted of the three expected phases: the initial non-linear lag
phase for redox activation, a linear maximal catalytic activity phase, and rapid loss
of activity at low [O2] (Fig. 2A in I).
No analytical tools have been reported previously to model the complex
kinetics of the Ero1 systems. We generated a non-linear regression method that
modelled the complete time course of an oxygen consumption trace. The simplest
non-linear equation that fit to the derivative of the oxygen consumption data, i.e.,
the rate of oxygen consumption at any given time point, consisted of Michaelis-
Menten kinetics with additional terms for one- or two-step exponential activation
process and Hill coefficient for potential cooperativity of oxygen binding (Fig. 2B
in I and 2C in I). The regression method could return simultaneously activation rate
constants (which can be transformed to half-time of activation, t1/2), catalytic
turnover number (kcat), half-maximal concentration for oxygen [KM(O2)], and Hill
coefficient (Table 1 in I).
75
The natural substrate assay and the regression methods were validated as
follows:
– Glutathione was kept at physiological concentrations and in high excess over
PDI to observe specifically Ero1 activity, i.e., the rate of disulfide transfer from
PDI to GSH was not limiting the Ero1 to PDI transfer rate (Fig. S8A in I).
– Other PDI-family members showed specificity (Fig. 3B in I and 3C in I)
consistent with previous in vitro studies (Araki et al., 2013): PDI was the most
active substrate and ERp57, ERp46, ERp72, and P5 showed only marginal
activity with both Ero1 isoforms. PDIp showed high activity for Ero1β but low
activity for Ero1α.
– As the main statistical tests for accuracy, residual analyses were carried out for
each fit individually (for example, Fig. 2B in I and 2C in I), and the fits would
be integrated back over the original oxygen consumption data (for example,
Fig. S5 in I).
– The obtained kinetic parameters for the Ero1 systems were cross compared to
those of another sulfhydryl oxidase, yeast Erv1p. Erv1p does not have
regulatory disulfides, and, consistent with this, it showed no lag phase (Fig.
S7A in I and S7B in I). It had stark differences in both KM(O2) and Hill
coefficient compared to those of the Ero1 systems (Table 1 in I), consistent
with Erv1p utilizing cytochrome c instead of O2 as the terminal electron
acceptor in vivo (Bihlmaier et al., 2007; Dabir et al., 2007). These data suggest
that the high affinity for oxygen and high Hill coefficients observed for the
Ero1 systems were genuine features, and not modelling errors.
5.1.3 Differential substrate PDI dependence between the Ero1
isoforms
The activation lag phase and the linear maximal rate period were first characterized
with the validated non-linear regression method. To this end, an oxygen
consumption data set with varying substrate [PDI] was recorded. When kcat values
were plotted versus [PDI] (Fig. 3A in I and S9 in I), similar maximal PDI oxidation
rates but differential midpoints for [PDI] were observed for the wild-type Ero1
isoforms with Ero1α showing clearly higher dependence for [PDI] for efficient
turnover (Table 3). Plots of activation data vs [PDI] (Fig. 4A in I and 4B in I)
suggested that the two Ero1 isoforms had differing PDI dependencies for redox
activation as well (Table 3). Whereas Ero1α showed a two-step activation process
76
with a strong dependence for [PDI] for both steps, the activation of Ero1β collapsed
to a single step [PDI]-independent process already at low [PDI]. These kinetic data
suggest that Ero1α has a high and Ero1β a low overall dependency for reduced
substrate PDI for both efficient redox activation and catalytic turnover. Importantly,
the midpoints for turnover and activation for a given isoform were of similar
magnitude (Table 3), suggesting that both processes, activation and catalytic
turnover likely utilize the same molecular determinants for the Ero1-PDI
interaction, namely the reported β-hairpin of Ero1 interacting with the substrate-
binding pocket of PDI (Masui et al., 2011). This hypothesis was followed up in II
and is described later in this work.
Table 3. Differential PDI dependency of turnover and activation between the Ero1
isoforms. All values are mean ± STD.
Enzyme Maximal turnover1
(s-1)
KM(PDI) for
turnover
(µM)
Activation
process
KM(PDI) for
activation
(µM)
Overall2
[PDI]
dependency
Ero1α-PDI 0.78 ± 0.03 8.5 ± 1.1 Two [PDI]-
dependent
steps
13.3 ± 2.1,
19.0 ± 3.3
high
Ero1β-PDI 1.06 ± 0.02 2.8 ± 0.3 One [PDI]-
dependent,
one [PDI]-
independent
step
4.6 ± 0.8 low
1at saturated [PDI] and [O2], 2accounting both redox activation and turnover.
Cys to Ala mutagenesis was carried out to dissect which disulfides are reduced
during activation, potentially explaining the differences in the activation processes
between the isoforms. Two regulatory Ero1α mutants, Cys104/131Ala and
Cys208/241Ala, that have the regulatory switch and the gating disulfide (see
Chapters 2.5.3 and 2.6.1) mutated, respectively, fit to the two-step activation
process with no major differences in activation rates compared to the wild type (Fig.
4E in I). In contrast, the Ero1α Cys166Ala mutant, which purified as a monomer,
fit to a single step activation scheme, and significantly increased the activation rate
(Fig. 4E in I). A similar study for Ero1β could not be conducted, as only the
Cys100/130Ala (a hyperactive mutant based on residue homology to Ero1α) could
be efficiently produced and analyzed. This mutant, in contrast to the hyperactive
mutant of Ero1α, showed kinetics which collapsed to the PDI-independent
77
activation process and significantly increased activation rate - further highlighting
the differences in the activation mechanisms between the two isoforms.
The activation kinetics of the Ero1α regulatory mutants were expected to
deviate more strongly from those of the wild type Ero1, as the mutants require
fewer regulatory disulfide reduction steps. As this was not the case, we next tested
the possibility that the rate-limiting steps for the two PDI-dependent activation
steps are not defined by reduction of regulatory disulfides, but by the PDI exchange
mechanism that was described earlier (Chapter 5.1.1). In such a scenario, the
regulatory disulfides may get reduced, but the reduction events are not observable
in our assays, as they get masked by the slow PDI exchange reactions. We studied
this hypothesis with a gel-based redox state analysis. We mixed Ero1α complex
with exogenous PDI3FLAG in the reducing activity assay conditions and quenched
an aliquot at constant intervals with NEM to stop thiol-disulfide exchange reactions
(Fig. S12A in I). As a result, the wild type complex disappeared with two-step
kinetics (Fig. S12B in I), resulting in a halftime of disappearance (1.5 min) similar
to the halftime of activation observed in the activity assays (cf. Table 1 in I).
Concomitant, a higher-molecular-weight complex with incorporated PDI3FLAG
appeared with a consistent halftime (1.7 min). These results suggest that
exchanging substrate PDI molecules, instead of reduction of regulatory disulfides,
forms the rate-limiting steps during redox activation of Ero1α.
5.1.4 A novel regulatory mechanism allows efficient operation in
hypoxic conditions
Another poorly characterized kinetic phase for human Ero1 is the rapid loss of
activity at low [O2]. The initial experiment (Table 1 in I) showed that both Ero1
isoforms had low KM(O2) and high Hill coefficients of >3. The high Hill
coefficients were consistent in the PDI titration assays (4.0 ± 1.2 for Ero1α and 3.2
± 0.7 for Ero1β; mean ± STD; Chapter 5.1.3), suggesting that the Ero1 systems
potentially exhibit cooperative binding for oxygen. In contrast, Erv1p had a high
KM(O2) and a Hill coefficient of 1 (Table 1 in I), implying no cooperativity of
oxygen binding.
The surprising apparent cooperativity was further characterized by modeling
[O2] thresholds of inactivation in silico. We compared the kinetics of wild type Ero1
using the experimentally derived kinetic values (Table 1 in I) to those of a
theoretical enzyme that utilized the same KM(O2) but a Hill coefficient of 1.
Whereas the theoretical enzyme (without cooperativity) had a broad [O2] range of
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inactivation (90% to 10% activity) of 44.4 µM and created hyperhypoxic conditions
by effectively consuming all oxygen from the system (Fig. 5A in I), both Ero1
isoforms retained full activity till very low [O2] and inactivated before consuming
all oxygen from the system (Fig. 5A in I). This effect was markedly apparent for
Ero1α which showed an inactivation range of only 4.3 µM (Table 4). These data
suggest that the Ero1 isoforms have been fine-tuned to operate efficiently at low
oxygen concentrations and this feature is achieved by the sigmoidal kinetics
permitted by the high Hill coefficients.
Table 4. Experimental kinetic data at low [O2] for Ero1α vs a theoretical enzyme without
a Hill coefficient.
Enzyme KM(O2)
(µM)
Hill
coefficient
[O2]90%
(µM)
[O2]10%
(µM)
inactivation
range1 (µM)
Ero1α-PDI 5.0 5.2 7.6 3.3 4.3
Theoretical enzyme 5.0 1.0 45.0 0.6 44.4
1[O2]90% - [O2]10%; [O2]90%, oxygen concentration at 90% activity; [O2]10%, oxygen concentration at 10%
activity.
5.2 Feedback inhibition of the Ero1α-PDI system
This work demonstrated that human Ero1α requires high concentrations of
exogenous reduced PDI for efficient redox activation and catalytic turnover. The
kinetic data suggested also that the activation and turnover events are likely
mediated by the same molecular determinants, specifically, the protruding β-
hairpin of Ero1α interacting with the hydrophobic substrate-binding site of PDI
(Fig. 10). These observations proposed a paradoxical hypothesis: Are the redox
activation steps and the catalytic activity of Ero1α inhibited during unfolded protein
stress, if the PDI substrate-binding site is saturated by non-native proteins? Non-
native proteins with exposed hydrophobic residues bind the b’ domain of PDI
(Klappa et al., 1998) which is also needed to bind Ero1α. We further hypothesized
that the inhibitory effect would not be paradoxical if some other quality control
feature of the ER benefited from the inhibition. One candidate is ERAD which
utilizes reduced protein substrates (Ellgaard et al., 2018). Inhibition of Ero1α
activity could therefore form a feedback inhibition loop that prevents futile redox
cycles, retaining non-native proteins in more reduced state for ERAD (Fig. 1 in II).
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5.2.1 The Ero1α-PDI interaction is inhibited by non-native proteins
To test the above hypotheses, we carried out comprehensive activity assays for
Ero1α complex using different variants of BPTI as the potential inhibitors. We
performed the assays in the presence or absence of native BPTI (N) or BPTI mutant
variants Cys30/51Ala and Cys14/38Ala that mimic the late folding intermediates
N* and NSS
HH (Fig. 3) and are incapable of forming N. When N was in clear excess
over PDI, no inhibition of Ero1α activation or catalytic cycle was observed,
irrespective of whether N was added at the start (denoted as pre-activation assay
from now on; Fig. 2B in II) or with circa 50% oxygen left in the system, when
Ero1α had completely activated (post-activation assay, Fig. 2C in II). In contrast,
N* and NSS
HH severely compromised Ero1α activity, with both intermediates causing
clearly extended lag phases (Fig. 2B in II). Whereas N* inhibited Ero1α for the
duration of the measurement, NSS
HH lost its inhibitory effect at later time points,
suggesting that late folding intermediates may not inhibit the catalytic cycle. These
results were supported by post-activation assays (Fig. 2C in II), resulting in
uninhibited activity in the presence of NSS
HH and an exponential loss of activity in the
presence of N*. The exponential activity loss could be modelled in silico (Fig. 2D
in II), and the returned halftime of inactivation of circa 128 s resembled the halftime
of activation of the uninhibited reaction (126 ± 7 s), suggesting that the exponential
loss was caused by redox inactivation instead of compromised catalytic activity.
All in all, these results suggest that natively folded proteins do not inhibit Ero1
activity, as they interact with neither Ero1α nor PDI; late folding intermediates
inhibit severely the redox activation steps but not the catalytic cycle.
5.2.2 Quantification of the inhibition
We quantified the inhibition with small redox insensitive hydrophobic molecules
that contain no cysteines. First, we tested a hydrophobic peptide, KFWWFS, which
has been reported to be bound by PDI (Byrne et al., 2009). Potent inhibition by the
peptide was observed in both the pre- and post-activation assays (Fig. 3A in II). To
quantify the inhibition, we kept [PDI] constant and varied [KFWWFS]. Both dose-
dependent extension of the lag phase and inhibition of the catalytic cycle were
observed (Fig. 3B in II). A plot of kcat vs [KFWWFS] (Fig. 3C in II) revealed
midpoint for inhibition of 4.9 ± 0.6 μM. Binding studies by ITC (Fig. 4A in II) and
BLI (Fig. 4B in II) demonstrated that PDI binds KFWWFS with affinity (Kd of 6.8
μM and 12.7 μM, respectively) of the same magnitude as the midpoint of inhibition,
80
suggesting that the inhibition is directly linked to competitive binding between the
peptide and Ero1α to the substrate binding site of PDI. We confirmed this
hypothesis by measuring the affinity of PDI to immobilized protruding β-hairpin
of Ero1α in the presence or absence of excess KFWWFS. In the absence of
KFWWFS, PDI bound to the hairpin with a Kd of 4.0 μM (Fig. 4C in II). When
KFWWFS was added in excess over PDI/hairpin, the PDI to hairpin binding signal
was decreased considerably (Fig. 4C in II).
Two other potential small-molecule inhibitors, the endogenous hormone 17β-
estradiol (17β-E2) (Fu et al., 2011) and the exogenous endocrine disruptor
bisphenol A (BPA; Hiroi et al., 2006) were tested. Both potential inhibitors, but not
a control compound 17α-estradiol (17α-E2), compromised Ero1α activity severely
in both the pre- and post-activation assays (Fig. 5A in II and 5B in II). The activity
of Erv1p was not inhibited to such degree, as it lacks the β-hairpin structure and
likely utilizes different binding mechanism (Guo et al., 2012; Fig. 5D in II). The
potent inhibition by 17β-E2 and BPA was confirmed not to be caused by the
inhibitors binding directly to Ero1α, as activity was regained in the presence of the
inhibitors and DTT (Fig. 5C in II, cf. Fig. 5A in II). To conclude, the quantification
data from this assay and the peptide assay confirm the observations from the protein
folding intermediate assays that the inhibition of Ero1 activity resulted from loss
of Ero1-PDI interaction due to saturation of the PDI substrate-binding site by the
inhibitors.
5.2.3 The inhibition is reversible
The feedback inhibition mechanism suggested (Fig. 1 in II) would have to be
reversible to have physiological significance. We tested the reversibility in vitro by
two methods:
1. the inhibition caused by non-native proteins should subside when the non-
native proteins fold to the native state;
2. increased concentration of folding factors should decrease the inhibition, as the
added factors sequester the inhibitors.
In both cases, loss of inhibition would restore the Ero1α-PDI interaction and
reinitiate oxygen consumption. To test if the inhibition subsides by folding, we
carried out oxygen consumption assays in the presence of denatured and
completely reduced wild-type BPTI (rBPTI). rBPTI behaves differently from the
late-stage folding intermediates tested earlier, as its folding pathway (Fig. 3B) is
81
not obstructed at any point, allowing it to form the early- and the late-stage folding
intermediates as well as the natively folded state. In the pre-activation assay, rBPTI
severely inhibited Ero1α activity at early time points, when no substantial folding
of rBPTI had occurred (Fig. 6A in II). At later time points, Ero1α slowly regained
activity (Fig. 6A in II), consistent with inhibition subsiding with BPTI folding and
N not inhibiting Ero1α (Fig. 2 in II). Interestingly, when rBPTI was tested in the
post-activation assay (Fig. 6B in II), severe inhibition of the catalytic activity was
observed. This suggests that early-stage folding intermediates inhibit both
activation and catalytic cycle, while the late-stage folding intermediates inhibit
mainly redox activation (cf. Fig. 2C in II).
To test the second hypothesis, we repeated the assay in the presence of rBPTI,
but, this time, before substantial amount of folded BPTI had accumulated, we
injected additional PDI to the assay. When [PDI] increased over [BPTI], Ero1α
regained activity immediately (Fig. 6A in II), supporting the second hypothesis that
folding factors sequester the non-native proteins and allow Ero1α to regain activity.
Together, these two assays demonstrate that the witnessed inhibition is reversible
and may therefore form a physiologically relevant feedback inhibition mechanism.
5.3 Structure-function relationship of human pERp1
One aim of this work was to expand the understanding of protein folding from
catalyzed folding to the non-catalytic aspect of protein maturation in the ER,
specifically, the action of molecular chaperones that are important for OPF but may
not directly promote disulfide formation. We chose pERp1 as the model protein, as
it is currently classified as either a chaperone or a co-chaperone. pERp1 promotes
the biogenesis of some complex disulfide-containing proteins, for example, IgA
and IgM, but otherwise it is poorly characterized (see Chapter 2.6.2). Cell-based
studies have demonstrated the physiological importance of pERp1 for the humoral
immune response, but the exact molecular mechanisms of its action are still poorly
understood. Here, high-resolution crystal structure of human pERp1 is presented to
address this issue. Structural and bioinformatic studies reveal relationship and
differences to other SAPLIPs. Functional studies elucidate its role as a chaperone
involved in biogenesis of immunoglobulins.
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5.3.1 High-resolution crystal structure of pERp1
We determined the crystal structure of human pERp1 to 1.4 Å resolution (Table 1
in III). Two chains were in the asymmetric unit, and all residues except for
disordered N- and C-terminal regions could be modelled. The structure consisted
of the predicted α-helical saposin-like fold with five helices linked by three
conserved disulfides (Fig. 2 in III). Four inserts were identified that are missing in
the saposin C structure (Fig. 3A in III): The disordered N- and C-terminal regions
and two disordered regions linking helices 1 and 2 (an 11 amino acid loop) and
helices 2 and 3 (a 37 amino acid region with a three stranded β-sheet).
Two potential dimers of pERp1 were identified:
1. The two chains in the asymmetric unit formed an intermolecular antiparallel β-
sheet between the 37-residue inserts. This interaction was mediated mainly by
hydrophobic interactions (Fig. 2B in III).
2. PISA analysis identified another dimerization site formed from helices 2 and 3
and N-terminus (Fig. 2C in III). The chains in this dimer would be connected
by salt bridges, with a negatively charged patch in one chain interacting with a
positively charged patch in the other.
While either of the homodimers could represent a biologically relevant form of
pERp1, analytical techniques (Fig. S3 in III) suggested that the pERp1 used in this
study was a monomer in solution. As the residues involved in dimerization are not
conserved in mammalia (see below), and saposins and other SAPLIPs (Ahn et al.,
2003; Rossman et al., 2008; Bryksa et al., 2011; Garrido-Arandia et al., 2018)
utilize different dimerization mechanisms, the observed homodimers may not be
relevant for the physiological function of pERp1.
5.3.2 Mapping function onto the structure
pERp1 is important for many physiological processes, including integrin
maturation and immune response (Shimizu et al., 2009; van Anken et al., 2009;
Flach et al., 2010; Xiong et al., 2019). However, molecular mechanisms of action
are poorly understood. We were interested if the structure reported here would
reveal new insight for the reported functions. Saposin C is known to undergo
conformational changes and possess “open” and “closed” conformations that are
related to function (Fig. 4A in III). To form the open conformation in pERp1,
helices 2, 3, and 4 as well as the 37-residue insert would be required to undergo
83
major structural rearrangements. The insert packs via hydrophobic interactions
against helices 1 and 3 (Fig. 4B in III and 4C in III), which is not present in the
saposin C fold (Ahn et al., 2006). Therefore, the structural features presented here
seem to disallow formation of a similar open conformation that has been described
for saposin C.
pERp1 contains the CXXC motif found in many oxidoreductases, and it has
been shown to have slight oxidoreductase activity (van Anken et al., 2009).
However, the structural analysis in this work suggests that pERp1 is unlikely to be
an oxidoreductase: each disulfide is buried, and the structure confirms the earlier
observations (van Anken et al., 2009) that the cysteines in the CXXC motif link to
other cysteines far away in the sequence. As pERp1 is unlikely to have the reported
dynamic open/closed states of the saposin, the three disulfides in pERp1 are likely
structural instead of functional.
As the structural features could not be directly linked to any functions, we
carried out bioinformatics analyses next. A structure-based conservation analysis
of 167 pERp1 sequences of different eukaryotic organisms or of only mammalian
species revealed 18 or 36 fully conserved amino acids, respectively (Fig. 5A in III).
We focused on the mammalian data set, as the former data set could be affected by
parallel divergent evolution of interacting proteins. A map of the 36 fully conserved
residues on tertiary structure (Fig. 5B in III) revealed that most of the conserved
residues are buried and stabilize native structure. Five or eight residues formed a
highly acidic surface exposed patch (Fig. 5B in III). The mammalian data set also
revealed that pERp1 has lower overall sequence conservation compared to other
CNPY family members (Table 2 in III).
5.3.3 Biochemical assays to elucidate function
The above structural and bioinformatics analyses revealed mainly structure-
stabilizing features, while the function of the protein remained unclear. We decided
to elucidate the function of pERp1 next by biochemical assays. pERp1 has been
suggested to be either a chaperone or a cochaperone (Shimizu et al., 2009; van
Anken et al., 2009; Rosenbaum et al., 2014). We tested if pERp1 is a general
chaperone by a classical chaperone assay based on inhibition of aggregation of
thermally denatured citrate synthase (CS). Whereas molecular chaperones BiP and
ERp27 strongly inhibited aggregation of denatured CS, pERp1, even at high 10x
molar excess over CS, showed no inhibition of aggregation (Fig. 6A in III). The
84
lack of chaperone activity was not caused by thermal denaturation of pERp1 (Fig.
6B in III).
pERp1 is clearly not a general chaperone, so we next tested its substrate
specificity. pERp1 has been reported to interact with only a relatively small set of
proteins, with IgA and IgM representing the most characterized substrates (Shimizu
et al., 2009; van Anken et al., 2009; Xiong et al., 2019). Furthermore, the C-
terminal extension of IgA has been suggested to interact with pERp1 in vivo (Xiong
et al., 2019). As the extension is predicted to contain a β-strand and is highly
conserved (Fig. 6C in III), we hypothesized that the interaction between IgA and
pERp1 is potentially mediated by formation of an intermolecular β-sheet between
the IgA extension and the 37-residue insert in pERp1. We tested binding of IgA
with and without the C-terminal extension using BLI. Neither IgA form interacted
with pERp1 (Fig. 6D in III).
85
6 Discussion
Efficient maturation of disulfide-containing proteins in the mammalian ER is a
complex process that requires the concerted effort of folding catalysts and
chaperones as well as other quality control features such as UPR and ERAD.
Although the Ero1-PDI pathway became a focus of interest as the main pathway
for disulfide bond formation in the ER already two decades ago (Frand & Kaiser,
1998; Cabibbo et al., 2000; Pagani et al., 2000), no comprehensive enzyme kinetic
studies, that characterized each of the three major phases of Ero1 kinetics (redox
activation, catalytic cycle, rapid decline of activity at low [O2]), have been reported
for the pathway. In the present study, we reported comprehensive enzyme kinetic
measurements for various forms of human Ero1 isoforms, kinetic tools that could
analyze the complex triphasic kinetics, and revealed novel structural and regulatory
features for OPF and for other quality control mechanisms of the ER. These
features included:
1. a stable mixed disulfide complex between PDI and inactive Ero1, utilizing a
cysteine that has been previously considered non-functional;
2. a mechanism that allows efficient disulfide formation in hypoxic conditions
and inactivation before hyperhypoxia;
3. differential PDI dependency for activation and catalytic cycle between the
human Ero1 isoforms;
4. feedback inhibition of Ero1α activity by non-native proteins that hints at a
previously uncharacterized crosstalk between oxidative protein folding and
ERAD.
We identified a stable heterodimeric mixed disulfide complex formed between
human PDI and Ero1α or Ero1β in the inactive state. The mixed disulfide formed
between conserved Cys166 of Ero1α or Cys165 of Ero1β and Cys397 of the a'
domain of PDI. Two technical reasons stand out why these complexes have
remained uncharacterized to date: i) Cys166 has previously been considered non-
functional, and several studies mutated it to alanine (for example, Inaba et al.,
2010; Araki & Nagata, 2011; Masui et al., 2011; Araki et al., 2013); ii) Ero1 and
PDI may form at least three other transient mixed disulfide complexes [the catalytic
complex and two regulatory interactions (Appenzeller-Herzog et al., 2008;
Appenzeller-Herzog et al., 2010; Ramming et al., 2015)], and no efficient methods
have been reported to trap and isolate a specific mixed disulfide state in vivo.
Interestingly, the complex reported in this study may have been observed but not
86
followed up by others: Benham and coworkers (2000) reported in an elegant in
vitro translation study that the levels of Ero1-PDI mixed disulfide complex
increased concomitant with Ero1α shifting from the active OX1 to the inactive OX2
state. An endogenous Ero1-PDI mixed disulfide complex was later observed at
steady state from HT and HeLa cell lysates with Ero1α primarily in the OX2 state
(Benham et al., 2013). Furthermore, an immunoprecipitation experiment of Bertoli
and coworkers (2004) demonstrated that Ero1α Cys166Ala mutant bound
significantly less PDI than an unmutated Ero1α and Appenzeller-Herzog et al.
(2008) suggested that Cys166 forms a disulfide, but the partner cysteine could not
be identified. Hence, the inactivation pathway of human Ero1 consists of
intermolecular interactions with PDI involving Cys166/165, with the reported
stable mixed disulfide complex potentially representing the final state of Ero1
inactivation (see Fig. 5B in I and the discussion below).
The analysis of the triphasic kinetic profiles of human Ero1 systems
demonstrated that the wild type complexes were active with catalytic efficiencies
(second-order rate constant Kcat / KM) at saturating [PDI] and [O2] of 9.1 x 104 M-
1s-1 (Ero1α) and 3.8 x 105 M-1s-1 (Ero1β), values that are consistent with typical
enzyme-catalyzed disulfide formation rates (cf. DsbA/DsbB couple in Table 1).
Importantly, at low [O2], we observed a novel regulatory mechanism of Ero1
activity, which was a combination of high affinity for substrate oxygen and a high
Hill coefficient (>3). The high Hill coefficient was present for all tested wild type
and redox mutants of Ero1α and Ero1β as well as at all tested substrate PDI
concentrations, suggesting that cooperative binding of substrate oxygen may
modulate the activity of both Ero1 isoforms, and that redox states of the enzyme
are unlikely to play part in the apparent cooperativity. Possible explanations for the
cooperativity include classical cooperativity (multiple oxygen-binding sites),
although we found no evidence of Ero1 forming homo-oligomeric states during the
catalytic cycle, or the monomeric glucokinase-type cooperativity which arises from
order-disorder transitions in a single substrate-binding site (Larion et al., 2012;
Whittington et al., 2015).
Physiologically, the low KM(O2) and the apparent cooperativity have important
implications for OPF. Both Ero1 isoforms retain high activity to low [O2] with a
drastic reduction of activity within an extremely narrow oxygen concentration
range (Fig. 5A in I). This effect was very strong for Ero1α which was still fully
active with ca. 5% oxygen left in the system, whereas a theoretical enzyme without
cooperativity had lost much of its activity by this point (Fig. 5A in I). Another
important observation was that the theoretical enzyme was capable of consuming
87
all oxygen from the system, while Ero1α effectively inactivated with circa 2 µM
(0.8%) oxygen left in the system (Fig. 5A in I). In other words, Ero1α has been
fine-tuned to function efficiently in hypoxic conditions and avoids formation of
hyperhypoxic conditions that might be lethal for the cell. Cell-based studies have
demonstrated that hypoxia upregulates Ero1α in vivo (May et al., 2005; Gess et al.,
2003). Our data suggests that the reported upregulation is a feasible means to
increase the folding capacity and disulfide formation rates of the ER in hypoxic
conditions when availability of substrate O2 is scarce.
We characterized for the first time the early lag phase of the Ero1 kinetics. The
data revealed differential PDI dependency for activation of the human Ero1
isoforms. Whereas Ero1α activated with two [PDI]-dependent steps, the kinetics of
Ero1β collapsed to a [PDI]-independent activation process already at low [PDI].
These differences have important consequences for in vivo function: Ero1α has a
strong dependence for exogenous reduced PDI to both activate and turn over
efficiently. Ero1β, on the other hand, will activate and show higher disulfide
formation rates even under conditions with low availability of reduced PDI. Such
a situation may occur in cells with extremely high protein folding load, for example,
professional secretory cells such as insulin-secreting pancreatic cells, where Ero1β
is enriched (Pagani et al., 2000; Dias-Gunasekara et al., 2005).
It is important to note that the differences in the activation kinetics do not
necessarily mean that the Ero1 isoforms have different activation pathways. The
simplest explanation is that the isoforms utilize similar activation pathways, but
they have different rate-limiting reactions in the individual steps. We developed a
scheme for the activation and inactivation pathways based on this view (Fig. 5B in
I) that is consistent with the activation assays utilizing redox mutants (Fig 4 in I)
and with the PDI exchange assays (Fig. 1F in I and Fig. S12 in I). In the scheme,
the oxidized Ero1-PDI mixed disulfide complex is the ultimate inactivated state of
human Ero1. The complex contains three regulatory disulfides - the mixed disulfide
and two intramolecular regulatory disulfides in Ero1 - each of which need to be
reduced for activation. The complex is relatively stable in the absence of exogenous
reduced PDI, as no net change takes place in the complex in the presence of
exogenous oxidized PDI (Fig. S13 in I). When exogenous reduced PDI is available,
the oxidized PDI in the complex is exchanged to reduced PDI, which initiates
activation. Two rounds of PDI exchange is required for full activation: the first one
leads to the reduction of the mixed disulfide and one of the intramolecular
regulatory disulfides, the second step to the reduction of the final regulatory
disulfide. For Ero1α, reductions of the intramolecular regulatory disulfides are
88
relatively fast compared to the slow PDI exchange reactions; the latter were
observed as the two [PDI]-dependent kinetic steps. Consistent with this, the Ero1α
Cys166Ala mutant activated with a single step and with significantly faster kinetics
(Fig. 4E in I), suggesting that the first slow PDI exchange step was omitted with
this monomeric mutant. In contrast, Ero1β exchanges PDI extremely fast, to such
degree that the gel-based activation assay (cf. Fig. S12 in I) could not be carried
out for this isoform. This would suggest that reduction of regulatory disulfides, of
which one was observed at low [PDI] only, form the rate limiting steps for Ero1β.
Unfortunately, we could not explicitly identify which two intramolecular disulfides
were reduced in the activation pathway and the specifics of the [PDI]-independent
activation event of Ero1β. One of the reduced disulfides is likely the well-
characterized regulatory switch (Cys94-Cys131 in Ero1α, Cys90-Cys130 in Ero1β),
supported by similar kinetics between wild-type Ero1α and the regulatory switch
mutant (activation rate limited by PDI exchange) and increased activation rate of
the Ero1β regulatory switch mutant (reduction rate-limiting). The second step is
unlikely to be reduction of the gating disulfide as the Ero1α gating mutant activated
with two [PDI]-dependent steps and without major differences in activation rates.
One possible explanation is that the second reduced intramolecular disulfide is
Cys85-Cys391 (in Ero1α) and Cys81-Cys390 (in Ero1β). This disulfide was
initially thought regulatory (Baker et al., 2008) but later reconsidered structural
(Araki & Nagata, 2011; Zhang et al., 2014). We could not produce a mutant without
this controversial disulfide in sufficient amounts to confirm this hypothesis.
Based on the strong PDI-dependency of Ero1α, we hypothesized that the
activity of Ero1α might be inhibited by molecules that compete for binding PDI.
This could potentially create a feedback inhibition mechanism to avoid futile
oxidative cycles in conditions when proteins need to be in a more-reduced state, for
example, for efficient ERAD. We observed this to be true in vitro for redox
insensitive low-molecular-weight hydrophobic compounds such as natural 17β-E2
and a peptide containing no cysteines as well as for larger non-native proteins,
including a completely unfolded protein and late-stage folding intermediates.
Whereas the activated Ero1α-PDI catalytic cycle was inhibited only by a subset of
inhibitors, the activation steps were inhibited by all the tested inhibitors. As we also
observed a kinetic partitioning in the catalytic cycle that allowed a small portion of
Ero1α (circa 1.4% at the tested conditions) to become inactivated each catalytic
cycle, with kinetics that were proportional to the activation kinetics, we propose
that the inhibition of activation may present the primary mechanism for the
feedback inhibition mechanism. We developed a scheme based on these results (Fig.
89
7 in II). During basal protein synthesis, PDI, which may reach millimolar levels in
the ER (Hillson et al., 1984; Lyles & Gilbert, 1991), is likely in excess over non-
native proteins, retaining balance with an active Ero1α between the folding load
and the folding capacity. If the concentration of non-native proteins exceeds that of
the folding capacity – that consists of PDI and other folding catalysts as well as
molecular chaperones – activation of Ero1α is inhibited. This effectively shuts
down the whole Ero1α-PDI relay, as any currently active Ero1α molecules will
inactivate, per the kinetic partitioning described above, and remain inactive due to
the inhibitors sequestering reduced PDI. This inhibition happens independent of
cysteines, suggesting that all non-native proteins that contain exposed hydrophobic
residues and bind PDI may inhibit the Ero1α-PDI pathway.
The inhibition by non-native proteins was found to be reversible by removal of
the inhibiting non-native proteins by folding as well as by increased levels of
folding factors that sequestered the inhibitors, mimicking ERAD and UPR,
respectively. In each case, the physiological impact is an increase in the level of
free PDI with an unoccupied substrate-binding site, that allows reactivation of
Ero1α and restoration of OPF. Therefore, the inhibition of the Ero1α activation
pathway by non-native proteins may present a reversible regulatory mechanism to
avoid futile oxidative cycles and to decrease the dependence on the reductive
component of the ERAD, saving cellular resources.
pERp1
This work presented a novel structure for a member of the CNPY family.
Specifically, we solved the high-resolution crystal structure of pERp1, a folding
factor that has been shown by cell-based studies to be important for efficient
maturation of some complex disulfide-containing proteins in the ER, for example,
integrins and oligomeric assembly of IgA and IgM (Shimizu et al., 2009; van Anken
et al., 2009; Flach et al., 2010; Andreani et al., 2018; Xiong et al., 2019). The
structure confirms that pERp1 is a SAPLIP, as is also predicted for other CNPY
family members (Schildknegt et al., 2019). It has large and flexible inserts at both
termini and inserted between alpha helices that are missing in the canonical saposin
fold (Ahn et al., 2006). While previous studies have clearly demonstrated the
importance of pERp1 for the humoral immune response (Shimizu et al., 2009; van
Anken et al., 2009; Flach et al., 2010; Andreani et al., 2018; Xiong et al., 2019),
the molecular mechanisms of action of pERp1 have remained elusive.
Unfortunately, the structural analyses or the bioinformatics analyses presented here
90
could not explicitly answer the question either. Overall sequence conservation of
pERP1 was found to be low compared to other CNPY family members (Table 2 in
III), which may reflect higher selective pressure of genes related to immune
function (Shultz & Sackton, 2019). We could eliminate potential functions of
pERp1 by comparing the novel structure to the structure of saposin C and by
biochemical assays:
1. The structure of pERp1 is in the ‘closed’ state of saposin and likely cannot form
the ‘open’ conformation as readily as saposin due to additional interactions
between the inserts and the SAPLIP scaffold. As saposins are involved in lipid
binding (Vaccaro et al., 1999; Bruhn, 2005), and there are no reports of CNPY
family members interacting with lipids, it is possible that the open/closed
dynamics are specific for saposins and lipid binding.
2. Instead of forming a redox active CXXC motif, the six cysteines of pERp1
likely form the three buried structural disulfides like in the saposin fold (Ahn
et al., 2006). This also suggests that previous cysteine mutations that inhibited
in vivo function (Shimizu et al., 2009) disrupted structure.
3. pERp1 does not interact with unfolded proteins, so it is not a general chaperone.
4. Binding studies showed that pERp1 does not interact directly with the C-
terminal extension of IgA.
The final result presented above seem to be at odds with the recent report by Xiong
and colleagues (2019) whose in vivo data suggested an interaction between pERp1
and the C-terminal extension of IgA. We could not reproduce this interaction in
vitro. The recently characterized function of MANF, an ER-resident protein
involved in protein folding (Yan et al., 2019) may resolve this contradiction. MANF
contains a SAPLIP domain (Fig. 7A in III) which, like pERp1, is poorly conserved
(Fig. 7D in III) but has significant structural homology to pERp1 (Fig. 7C in III).
Its action in protein folding is modulated by the molecular chaperone BiP, and this
interaction involves the C-terminal SAF-A/B, Acinus, and PIAS domain, but not
the SAPLIP domain (Fig. 7B in III). Based on these functional and structural
similarities, we propose that the CNPY family should be grouped with MANF. Two
other proteins were identified that may belong to this group: CRELD1 and
CRELD2. Both proteins are ER-resident proteins that are involved in protein
folding (D'Alessandro et al., 2018; Hartley et al., 2013). They contain six cysteines
and a CXXC motif in a domain that InterPro predicts to be similar to the SAPLIP
domains of the CNPY family members. The six cysteines of CRELD are conserved
(Fig. 7E in III) and in analogous positions to those of pERp1 (Fig. 7A in III),
91
suggesting that they form the three structural disulfides and that both CRELD
indeed contain a SAPLIP domain. All in all, the CNPY family, MANF, and CRELD
may form a functional superfamily, and the members of this superfamily would be
regulators of protein folding with their SAPLIP domains acting as scaffolds to add
extensions that regulate the action of other folding factors. This would be consistent
with pERp1 interacting indirectly with the C-terminal extension of IgA, i.e., pERp1
regulates another folding factor which interacts with IgA.
Conclusions
The activity of human Ero1 isoforms is controlled by the redox state of the ER,
turning it on and off by a regulatory disulfide in reducing and oxidizing conditions,
respectively (Appenzeller-Herzog et al., 2008; Baker et al., 2008; Appenzeller-
Herzog et al., 2010). The results of the enzyme kinetic studies in the present work
have demonstrated that the activity of Ero1 and oxidative protein folding in the ER
are modulated on multiple other previously uncharacterized levels. These layers
include another “on/off switch” at low substrate oxygen concentrations that
potentially allows Ero1 to be shut down before consuming all precious oxygen from
the cell, and, inversely, allows OPF to rapidly initiate when the oxygen levels start
to rise; as well as by the controversial-sounding inhibition of OPF by unfolded
proteins that becomes actual in the larger concept of quality control network with
one reversibly downregulated quality control pathway allowing another pathway to
act more efficiently. These quality control pathways, e.g., UPR and ERAD, have
been characterized well individually, and, to the best of our knowledge, this is the
first time that a potential direct cross-talk between OPF and ERAD have been
reported. These novel layers and interconnections of the quality control features as
well as the novel SAPLIP structure and superfamily reported demonstrate that OPF
is an intriguing phenomenon that is still poorly understood, and the results of the
present work open up new fascinating possibilities for future protein-folding work.
92
93
7 Future prospects
We carried out comprehensive in vitro enzyme kinetic studies for both human Ero1
isoforms, revealing several new regulatory mechanisms for OPF. These studies
were consistent with previous works by others, and they detailed molecular
mechanisms of actions for previously observed physiological phenomena such as
the hypoxia link. However, the background that the present work created for the
novel regulatory mechanisms would benefit from further validation by cell-based
studies. Especially, the differences in the [PDI]-dependency between the Ero1
isoforms would greatly benefit from cellular studies and potentially explicitly
answer the fundamental question of why many mammalian species have two Ero1
isoforms. Additionally, establishing the OPF-ERAD link in cells would perhaps
allow ERAD to be studied in a more efficient way.
Another interesting future prospect is elucidation of the apparent cooperativity
for oxygen binding of human Ero1 isoforms. Apparently, substrate cooperativity
has not yet been described for a sulfhydryl oxidase. Molecular details of this feature
could therefore allow us to understand sulfhydryl oxidases in a new way but would
also potentially benefit biotechnology and industry. Many important protein-based
drugs contain disulfides and one major problem in their production – in addition to
formation of disulfides – is oxygen availability during protein expression. The
present work demonstrated that Erv1p, which is an important component together
with human PDI of the CyDisCo system, utilizes molecular oxygen relatively
poorly as the terminal electron acceptor. Preliminary unpublished data has
suggested that production of disulfide-rich proteins with CyDisCo is often oxygen-
limited during shake flask production, and that Ero1 can replace Erv1p in the
CyDisCo system to some extent. Establishing the molecular details of how Ero1
utilizes oxygen and how the apparent cooperativity works could potentially benefit
production of complex disulfide-rich proteins.
Finally, as both Ero1 and pERp1 have implications in the medical field,
knowledge about their molecular functions may contribute to development of new
therapeutic and diagnostic tools. Ero1 has been demonstrated to be upregulated in
cancers, is a predictor of poor prognosis of breast cancer, and the first selective
inhibitor was published already a decade ago (Blais et al., 2010; Kutomi et al., 2013,
Shergalis et al., 2020). These works have likely relied on the assumption that human
Ero1 is a monomeric protein (when not involved in catalytic disulfide transfer to
PDI). If Ero1 is found in significant amounts in vivo in the stable heterodimeric
inactive state with PDI, this state might serve as a novel therapeutic target. pERp1
94
is crucial for immune response and, thus, it has been reported to be involved in
many immune system-related diseases and cancers (for example, Matsumura et al.,
2012; Wong et al., 2013; Kanda et al., 2016; Miyagawa-Hayashino et al., 2018;
Guzeldemir-Akcakanat et al., 2019). The structure presented in this work will be
important for establishing the potential of pERp1 as a therapeutic target.
95
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Original publications
I Moilanen, A., Korhonen, K., Saaranen, M. J., & Ruddock, L. W. (2018). Molecular analysis of human Ero1 reveals novel regulatory mechanisms for oxidative protein folding. Life Science Alliance, 1(3), e201800090. doi:10.26508/lsa.201800090
II Moilanen, A., & Ruddock, L. W. (2020). Non-native proteins inhibit the ER oxidoreductin 1 (Ero1)-protein disulfide-isomerase relay when protein folding capacity is exceeded. The Journal of Biological Chemistry, 295(26), 8647-8655. doi:10.1074/jbc.RA119.011766
III Sowa, S. T., Moilanen, A., Biterova, E., Saaranen, M. J., Lehtiö, L., & Ruddock L. W. High-resolution crystal structure of human pERp1, a saposin-like protein crucial for IgA, IgM and integrin maturation in the endoplasmic reticulum. In press.
Reprinted with permission from Life Science Alliance (I) and American Society for
Biochemistry and Molecular Biology (II).
Original publications are not included in the electronic version of the dissertation.
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Antti Moilanen
NOVEL REGULATORY MECHANISMS AND STRUCTURAL ASPECTS OF OXIDATIVE PROTEIN FOLDING
UNIVERSITY OF OULU GRADUATE SCHOOL;UNIVERSITY OF OULU,FACULTY OF BIOCHEMISTRY AND MOLECULAR MEDICINE
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