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The role of the proteasome & its regulators in adaptation to oxidative stress

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i

THE ROLE OF THE PROTEASOME & ITS REGULATORS IN ADAPTATION TO
OXIDATIVE STRESS

by

Andrew Michael Pickering

A Dissertation Presented To The
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(MOLECULAR BIOLOGY)

May 2012

Copyright 2012 Andrew Michael Pickering

ii

EPIGRAPH

“Learn from yesterday, live for today, hope for tomorrow. The important thing is not to
stop questioning.”
- Albert Einstein

iii

ACKNOWLEDGEMENTS

As I reach the end of my Ph.D I look back at the last 5 years I have spent working on it. It
has certainly been an experience. An experience which I feel has made me grow both as a
scientist and as a person. For this experience I thank my mentor Dr Kelvin J. A. Davies
who has helped to shape my scientific mind set, my analytical abilities and my bench-top
abilities. I also thank Kelvin for putting up with my eccentricities for the last 4 years, for
always making time for me and finally for his friendship and sage advice.

I thank my girlfriend Miss Carolyn E. Ziminski. who has supported me during this hectic
time and provided often much needed distractions. I thank her for putting up with my late
nights at the lab, my 1am experiments and my weekend work. Finally I cannot thank her
enough for tirelessly proof reading all of my papers and my dissertation.

I thank my parents Mrs Christine L. Pickering. and Dr Michael W. Pickering who have
made me into the person I am today and have always supported me in my work and
decisions.

I’d like to thank my undergraduate mentors Dr Mark D. Fricker, Prof Alex Kacelnik and
Dr Graham K. Taylor as well as my 6
th
form college biology teacher Mr Darrell Spear
who all inspired me and helped me to develop as a scientist.

iv

I thank my collaborators Dr John G. Tower, as well as his co-workers Dr Gary Landis
and Dr Peter Poon, who worked with me on the Drosophila parts of my research and
kindly allowed me to use space in his lab and their lab resources on the project. I would
like to thank Dr Derek S. Sieburth, and his student Miss Trisha A. Staab, who worked
with me on the C. elegans parts of my research and kindly allowed me to use space in his
lab and their lab resources on the project. I would like to thank Dr Henry J. Forman and
his co-workers Dr Honqiao Zhang and Dr Honglei Liu, who provided advice on my work
with the transcription factor Nrf2, helping especially with the CHiP assay and rtPCR
assays. I would like to thank Dr Tilman Grune and Mrs Cheryl Teoh. who performed a
number of the experiments described in chapters 2 & 3.

I would like to thank the whole of the Davies lab for the help and support over the years.
I would like to thank Dr Gennady Ermak for the help, advice and experimental expertise
he has provided me. I would like to thank Mr Robert A. Linda. who has worked with me
on the Nrf2 parts of my project. Robert performed the ChIP assay and qPCR experiments
that are described in that chapter. In addition, Robert kindly proof read a lot of my
dissertation. I would like to thank undergraduates Miss Janet Chu and Miss Lesya
Vojtovich and technician Miss Alison L. Koop. Who have helped at various stages in
these projects. Dr Jenny K. Ngo. Who helped me to get started in the lab and taught me
many of the techniques that I have used. I would also like to thank Miss Aprill Hite, Miss

v

Laura C. D. Pomatto, Miss Brenda Niu, Mr Kevin Chang and Mrs Sonal N. Patel who
have all helped me over the years.

Finally I would like to thank my dissertation committee: Dr. Caleb E. Finch, Dr Enrique
Cadenas, and Dr Steven E. Finkel for the help and advice that they have all given me
over the years.

vi

TABLE OF CONTENTS

Epigraph ii
Acknowledgements iii
List of Figures viii
Abbreviations xii
Abstract xv
Chapter 1: Introduction 1
I. Formation of oxygen radicals and toxic oxidants in metabolism 2
II. Formation of oxygen radicals and toxic oxidants by external factors 7
III. Removal of oxygen radicals and toxic oxidants by the cell 7
IV. Removal of damaged proteins by the proteasome 10
V. Changes in protein damage and the proteasome over age 14
VI. Adaptation to oxidative stress 16

Chapter 2: Oxidant Induced Adaptation to Oxidative Stress. 18
I. Hydrogen peroxide induced resistance to oxidative stress challenge 19
II. Induction of proteolytic activity 20
III. Induction through different oxidants 26
IV. Adaptation under repeated oxidative stress 30
V. Adaptation under chronic oxidative stress 35
V. Summary 37

Chapter 3: The 20S and 26S Proteasome in Stress Adaptation 39
I. Role of 20S vs 26S proteasome in degrading oxidized proteins 39
II. The role of proteasome in protein synthesis independent response 43
III. The role of proteasome in protein synthesis dependent response 45
IV. Summary 51

Chapter 4: The Nrf2 Transcription Factor a Regulator of Stress Adaptation 53
I. Induction of Nrf2 under oxidative stress 53
II. Induction of proteasome under oxidative stress is Nrf2 dependant 61
III. Adaptation through Nrf2 induction 64
IV. Other stress induced transcription factors 69
V. Summary 71

vii

Chapter 5: The Role of the Immunoproteasome 75
I. Oxidative stress induction of immunoproteasome 77
II. Immunoproteasome in oxidative stress adaptation 79
III. Capacity of immunoproteasome to degrade oxidized proteins 84
VI. Induction of immunoproteasome is Nrf2 independent. 86
V. Summary 89

Chapter 6: The Role of 20S Proteasome Regulators 91
I. Pa28αβ 92
II. PARP 106
III. Pa200 107
IV. Pa28γ 115
V. Regulation of immunoproteasome 120
VI. Binding of regulators to 20S vs hybrid proteasome 123
VII. Summary 126

Chapter 7: Oxidative Stress Adaptation in Model Organisms 131
I. H₂O₂ induced adaptation to oxidative stress in C. elegans 132
II. 20S proteasome in C. elegans oxidative stress adaptation 135
III. The role of Skn-1 in oxidative stress adaptation in C. elegans 138
IV. H₂O₂ induced increase in proteolytic capacity in D.melanogaster 143
V. H₂O₂ induced adaptation to oxidative stress in D.melanogaster 146
VI. 20S proteasome in D.melanogaster oxidative stress adaptation 147
VII. Role of Cnc-C in oxidative stress adaptation in D.melanogaster 151
VIII. Summary 156

Chapter 8: Materials and Methods 159
I. Basic techniques 159
II. C.elegans and D.melanogaster techniques 173
III. Reductive labeling of proteins with the fluorophore AMC 183

Chapter 9: Discussion 213
I. Ihe proteasome in oxidative stress adaptation. 213
II. The role of Nrf2 in oxidative stress adaptation. 217
III. The proteasome regulators in response to oxidative stress 223
IV. Summary 228

Bibliography 229

viii

LIST OF FIGURES

Figure 1.1: Formation of oxygen radicals and reactive oxidants 6
Figure 1.2: 26S vs 20S proteasome 12
Figure 2.1: H
2
O
2
induced adaptation to H₂O₂ challenge 20
Figure 2.2: Proteolytic activity increases during transient adaptation to H
2
O
2

24
Figure 2.3: Proteasome is induced during transient adaptation to H
2
O
2
25
Figure 2.4: Proteolytic activity under adaptation with protein synthesis inhibition 26
Figure 2.5: Oxidant pre-treatment increases proteolytic capacity in a
proteasome dependent manner 28
Figure 2.6: Pre-treatment with different oxidants improves tolerance
to H₂O₂ challenge 29
Figure 2.7: Capacity to degrade Suc-LLVY-AMC with repeated
H₂O₂ pre-treatment 33
Figure 2.8: Adaptive effects of repeated H₂O₂ pre-treatments at 6 h intervals 34
Figure 2.9: Adaptive effects of repeated H₂O₂ pre-treatments at 12 h intervals 35
Figure 2.10: Adaptation under chronic oxidative stress 37
Figure 3.1: Importance of 20S vs 26S proteasome in degrading
oxidized proteins 42
Figure 3.2: Disassembly and reassembly of the 26S proteasome in
response to oxidative stress 44
Figure 3.3: Dissociation of 26S proteasome with H₂O₂ treatment 45
Figure 3.4: Induction of 20S proteasome with H
2
O
2
treatment 49
Figure 3.5: Importance of 20S proteasome in adaptation 50

ix

Figure 3.6: Model for adaptive response 52
Figure 4.1: Nrf2 protein levels and nuclear translocation during oxidative
stress adaptation 58
Figure 4.2: Increased proteolytic capacity is blocked by inhibition of Nrf2 59
Figure 4.3: H
2
O
2
induced 20S proteasome induction is Nrf2 dependent 63
Figure 4.4: H₂O₂ induced adaptation is Nrf2 dependent 64
Figure 4.5: Nrf2 inducers increase oxidative stress tolerance to oxidative
stress 67
Figure 4.6: Nrf2 inducers promote proteasome dependent adaptation 68
Figure 4.7: NF-κB/Nrf2/FoxO binding sites on proteasome promoter
sequences 71
Figure 4.8: Model for adaptive response 74
Figure 5.1 Diagram of subunit composition of the 20S proteasome
vs immunoproteasome 77
Figure 5.2: Expression of 20S proteasome, 26S proteasome and
immunoproteasome subunits during H₂O₂ Adaptation 79
Figure 5.3: Blocking the induction of 20S proteasome or immunoproteasome
inhibits adaptation in H₂O₂ challenged cells 82
Figure 5.4: Ability of the immunoproteasome to degrade oxidized proteins 85
Figure 5.5: Expression of immunoproteasome is Nrf2 independent 87
Figure 5.6: ARE/EpRE binding sites upstream of 20S and immunoproteasome
subunits 88
Figure 6.1: Expression of proteasome, immunoproteasome and Pa28αβ
with H ₂O ₂ treatment 94
Figure 6.2: Proteolytic capacity of wild-type & Pa28αβγ knockout MEF 97

x

Figure 6.3: Blocking the induction of 20S proteasome or immunoproteasome
inhibits adaptation in H₂O₂ challenged cells 98
Figure 6.4: H
2
O
2
induced expression of Pa28αβ is Nrf2 dependent 101
Figure 6.5: Role of Pa28αβ in Removal of Damaged Proteins by 20S
Proteasome 105
Figure 6.6: Induction of Pa200 in response to oxidative stress 110
Figure 6.7: Pa200 regulator enhances degradation of short peptides and
non-oxidized hemoglobin 114
Figure 6.8: Induction of Pa28γ regulator in response to oxidative stress 118
Figure 6.9: Regulation of immunoproteasome by Pa28αβ, Pa28γ and Pa200 122
Figure 6.10: Pa200 and, to a lesser extent, Pa28αβ and Pa28γ can form
hybrid proteasomes 125
Figure 7.1: In C.elegans, H
2
O
2
pretreatment increases H
2
O
2
tolerance and
proteolytic capacity 134
Figure 7.2 In C. elegans, H
2
O
2
increases 20S proteasome which enhances
oxidative stress tolerance 137
Figure 7.3: H
2
O
2
exposure induces Skn-1 nuclear localization and Skn-1
dependent proteasome induction in intestines 141
Figure 7.4: In Drosophila, H
2
O
2
pretreatment increases proteolytic capacity 145
Figure 7.5: In Drosophila, H
2
O
2
pretreatment increases stress resistance
and 20S proteasome levels 149
Figure 7.6: In Drosophila, 20S Proteasome is required for H
2
O
2
induced
adaptation 150
Figure 7.7: The Drosophila Nrf2 homologue Cnc-C is required for adaptation 153
Figure 8.1: Proposed reaction between AMC and the free carboxyl groups
of a protein (R-), mediated by sodium cyanoborohydride 188
Figure 8.2: AMC can be conjugated to free carboxyl groups on proteins 192

xi

Figure 8.3: Proteolysis of AMC-labeled proteins by trypsin 195
Figure 8.4: Protease and substrate titration and particle size of proteolytic
degradation products 199
Figure 8.5: Stability of AMC-labeled hemoglobin after frozen storage
or denaturation 202
Figure 8.6: pH profile of fluorescence, stability, and proteolytic
susceptibility of free AMC and Hb–AMC 205
Figure 8.7: Proteolytic susceptibility of modified AMC-labeled proteins 208
Figure 9.1: Model of oxidative stress adaptation in mammalian cells 221
Figure 9.2: Model of oxidative stress adaptation in D. melanogaster
and C. elegans 222
Figure 9.3: Summary of the complexes and roles of proteasome regulators 227

xii

ABBREVIATIONS

Act-GS-255B: Actin linked Gene switch 255B
AMC: 7-amino-4-methylcoumarin
ARE: Anti-oxidant response element
ATP: adenosine triphosphate
BSA: bovine serum albumin
BSA–AMC: 7-amino-4-methylcoumarin-labeled bovine serum albumin
BrdU: bromodeoxyuridine
Bz-VGR-AMC: α-N-benzoyl-Val-Gly-Arg-7-amino-4-methylcoumarin
C. elegans: Caenorhabditis elegans
ChIP: Chromatin Immunoprecipitation
Cnc-C: Cap n Collar isoform C
D. melanogaster: Drosophila melanogaster
Dkeap-1: drosophila Kelch-like ECH-associated protein 1
DNA: deoxyribonucleic acid
EpRE: electrophile response element
ezrin
ox
: oxidized ezrin
FOXO: Forkhead box O
GFP: Green fluorescent protein
H
2
O
2
: hydrogen peroxide
Hb: hemoglobin

xiii

Hb–AMC: 7-amino-4-methylcoumarin-labeled hemoglobin
Hb
ox
: oxidized hemoglobin
Hsp70: Heat-shock promoter 70
Keap-1: Kelch-like ECH-associated protein 1
L1, L2, L3, L4: C.elegans larval stages 1, 2, 3, 4
MEF: murine embryonic fibroblast
M.W.C.O: Molecular weight cut off
NAD+/NADH: Nicotinamide adenine dinucleotide,
Native-PAGE: polyacrylamide gel electrophoresis lacking sodium dodecyl sulfate
NF-κB: nuclear factor kappa-light-chain-enhancer of activated B cells
Nrf2: nuclear factor (erythroid-derived 2)-like 2
Pa28αβ: proteasome activator 28 αβ
Pa28γ: proteasome activator 28 γ
Pa200: proteasome activator 200
PAS-7: proteasome α subunit 7
PBS: phosphate buffer saline
PCR: polymerase chain reaction
PrOxI: protein oxidation and immunoproteasome hypothesis of MHC class I antigen
processing

PSMA1-7: proteasome subunit α1-7
PSMB1-7: proteasome subunit β1-7
PSME1-4 : proteasome activator complex element subunit 1-4
qPCR: quantitative polymerase chain reaction

xiv

RNA: ribonucleic acid
RNAi: ribonucleic acid interference
rtPCR: reverse transcriptase polymerase chain reaction
SDS-PAGE: sodium dodecyl sulfate polyacrylamide gel electrophoresis
siRNA: short interfering RNA
Skn-1: SKiNhead-1
SKN-1::GFP: The SKiNhead-1 protein linked to green fluorescent protein
SOD: superoxide dismutase
Suc-LLVY-AMC: N-succinyl-Leu-Leu-Val-Tyr-7-amino-4-methylcoumarin
Sulfo-NHS-acetate: sulfo-N-hydroxysulfosuccinimide acetate
TCA: trichloroacetic acid
Z-LLE-AMC: benzyloxycarbonyl-Leu-Leu-Glu-7-amino-4-methylcoumarin

xv

ABSTRACT

Oxidized cytoplasmic and nuclear proteins are normally degraded by the proteasome, but
accumulate with age and disease. I demonstrate the importance of various forms of the
proteasome during transient (reversible) adaptation (hormesis), to oxidative stress in
murine embryonic fibroblasts. Adaptation was achieved by 'pre-treatment' with acute
oxidative stress exposure (e.g. H
2
O
2
, peroxynitrite, menadione, paraquat), and tested by
measuring inducible resistance to a subsequent much higher 'challenge' dose of H
2
O
2
.

Oxidative stress adaptation causes an initial direct physical activation of pre-existing
proteasomes, then a subsequent de novo synthesis of 20S proteasome,
immunoproteasome and Pa28αβ during over the next 24 h. Cellular capacity to degrade
oxidatively damaged proteins increased with 20S proteasome, immunoproteasome and
Pa28αβ synthesis, and was mostly blocked by the 20S proteasome, immunoproteasome
and Pa28 siRNA knockdown treatments. Direct comparison of purified 20S proteasome
and immunoproteasome demonstrated that the immunoproteasome can selectively
degrade oxidized proteins. Cell proliferation and DNA replication both decreased and
oxidized proteins accumulated, during high H
2
O
2
challenge. However H
2
O
2
adaptation
was protective against such H
2
O
2
challenge. Importantly, siRNA knockdown of the 20S
proteasome, immunoproteasome or Pa28αβ regulator blocked 50-100% of these adaptive
increases in cell division and DNA replication. Immunoproteasome knock-down also
largely abolished protection against protein oxidation.

xvi

I show that the adaptative increase in oxidative stress tolerance and capacity to degrade
oxidized proteins is dependent induction of the Nrf2 transcription factor. Furthermore I
show that adaptation causes an increase in cellular levels of Nrf2, and translocation of
Nrf2 from the cytoplasm to the nucleus. It also causes increased binding of Nrf2 to
antioxidant response elements (ARE) or electrophile response elements (EpRE) in the 5-
untranslated region of the Proteasome β5 subunit gene [demonstrated by chromatin
immunoprecipiation (or ChIP) assay]. I go on to show that this induction of Nrf2 is a
necessary requirement for increased Proteasome/Pa28αβ levels, and for maximal
increases in proteolytic capacity and stress resistance. The oxidative stress induced
increase in immunoproteasome however did not appear to be Nrf2 dependent.

I show that Pa28αβ and Pa28γ have increased expression under mild oxidant exposure. I
also demonstrate that both of the proteasome regulators enhance the capacity of the
proteasome to selectively degrade oxidized proteins. In conjunction with their increased
expression, there is an increase in binding of the Pa28αβ regulator to 20S proteasome. I
show that the Pa200 proteasome regulator is also induced by H₂O₂ exposure. The Pa200
regulator however does not enhance the capacity of proteasome to degrade oxidized
proteins. However, it does appear to enhance the capacity of the proteasome to degrade
histones.

I also demonstrate that this adaptive response is highly conserved. Exposure to a mild
dose of an oxidant can increase oxidative stress tolerance in both Drosophila and C.

xvii

elegans. There is also, in both of these animals, an increase in proteolytic capacity and a
corresponding increase in 20S proteasome levels. If the increase in 20S proteasome or
induction by Nrf2 homologues is blocked then the adaptive response is either blunted or
completely lost in both animals.

1

CHAPTER 1: INTRODUCTION

Protein damage is a natural and common aspect of life from multi-cellular endothermic
mammals to single celled prokaryotes. It can be caused by a wide range of mechanisms.
Such protein damage can occur as a product of external stimuli including exposure to
oxidants present in air pollution (Halliwell et al., 1992; Menzel, 1994), pesticides
(Abdollahi et al., 2004), ozone (Cross et al., 1992a; Cross et al., 1992b) and various other
chemical agents. Protein damage can also be caused through exposure to radiation such
as UV (Hu and Tappel, 1992) or various forms of ionizing radiation (Leach et al., 2001).
In addition to external factors, protein damage can also be caused through internally
generated oxygen radicals and oxidizing agents which are produced during metabolism
(Kappus, 1987) or immune response (Baeuerle et al., 1996). Most protein damage will
effect non-essential parts of the protein structure and so have limited or no effect on
protein function. However some protein damage will occur at active sites or cause a
dramatic shift in protein structure which can have highly detrimental effects on protein
function (Davies, 1987, 1995; Davies and Delsignore, 1987; Davies et al., 1987a; Davies
et al., 1987b) and organismal viability (Copeland et al., 2009). Due to these severe
outcomes, the rapid and efficient removal of damaged proteins is extremely important.
These damaged proteins are removed primarily by the proteasomal system (Davies, 2001;
Pickering et al., 2010), enabling normal cell function to resume. The degree of protein
damage, however, is not static but shifts with changes in both the internal and external

2

environment. As a product of this, the ability of the cell to remove damage ‘adapts’ to the
changes in environment (Wiese et al., 1995).

In this dissertation I characterize the role of various forms of the proteasome in oxidative
stress adaptation, the pathway by which they are regulated and the function of these
proteins in removal of damaged proteins.

I. Formation of oxygen radicals and toxic oxidants in metabolism

Oxygen is one of the greatest blessings and perhaps curses to complex life; this is what is
often referred to as the ‘Oxygen Paradox’ (Latham, 1951). The paradox is that oxygen is
essential for most organic life but is also the cause of much of the damage which limits
organic life. Oxygen possesses a vital role as the final acceptor in the electron transport
chain. In the electron transport chain, a series of alternating hydrogen and electron
carriers ensure that electrons are transported across the inner mitochondrial membrane,
such that protons are transported outwards. The resulting electrochemical gradient is then
used for the formation of ATP from ADP + P
i
. In order to maintain electron flow in the
inner mitochondrial membrane, electrons must also be removed from the electron
transport chain by a terminal electron acceptor. This role is played by the cytochrome c
oxidase complex (mitochondrial complex IV) which oxidizes electrons with oxygen to
generate water, thereby ‘bleeding’ electrons from the chain, and allowing continual ATP
production. The presence of oxygen is, thus, vital for mitochondrial function and overall

3

energy balance in eukaryotes. While there are alternative processes to generate ATP that
are independent of oxygen (such as fermentation which can be performed by some single
celled organisms), these processes are considerably less efficient than oxidative
phosphorylation. As a result of the sizable difference in ATP production rates between
oxygen-dependent and oxygen-independent pathways, it has been suggested by some
groups that the increased efficiency of oxygen-dependent respiration over fermentation
was one of the key factors that permitted the formation of complex multi-cellular life
forms (Raymond and Segre, 2006).

While the role of oxygen as a final electron acceptor permits a greatly enhanced energy
production rate, the reaction has the potential to generate a number of highly toxic by-
products. In the electron transport chain, electrons pass through a series of electron
acceptors, eventually terminating with cytochrome c oxidase, which catalyzes the
formation of water. The termination of the electron transport chain seems to occur with
minimal detectable electron leakage. However significant electron leakage has been
recorded at early stages of the electron transport chain (Halliwell and Gutteridge, 2000).
Such leaked electrons can directly react with oxygen in the cell to form superoxide
radicals (O
2
•-
). Free-radicals such as O
2
•-
possess an unpaired electron in their outer
orbital. These radicals have a strong tendency to react with other molecules to either
donate the extra electron or gain another, in either case restoring their electron pairing. In
such redox reactions, the target molecule is modified by the free-radical, sometimes

4

reversibly but often irreversibly. The molecules may then go on to react further with
surrounding molecules resulting in further, potentially severe, damage to the cell (Davies,
1995).

Superoxide radicals are capable of reacting with iron or copper ions in the cell,
converting them from Fe
3+
to Fe
2+
or Cu
2+
to Cu
+
. This is harmful in itself as it results in
the modification of metal ions which form important catalytic centers for a wide range of
cellular enzymes. In addition, the Fe
2+
or Cu
+
generated by this reaction is then capable of
reacting (by the Fenton reaction (Fenton, 1894)) with the mild oxidant hydrogen peroxide
(H
2
O
2
) to form highly toxic hydroxyl radicals (
•
OH) At variance with the Fenton
reaction, the Haber-Weiss reaction: H
2
O
2
+ O
2
.–
 O
2
+ HO
–
+ HO
.
(Haber and Weiss,
1932) occurs in the absence of transition metals, proceeds at extremely slow rates, and
mostly in the domain of chemistry but not in biological systems. In addition to this,
superoxide can instead react with two protons and another electron to form hydrogen
peroxide. Hydrogen peroxide is a relatively mild oxidant, but it can cause modification of
DNA, lipids, and proteins through reactions catalyzed by transition metals. Hydrogen
peroxide also appears to be capable of modifications to some amino acids chains in
proteins (e.g. amino acids containing thiol groups such as cysteine (Brodie and Reed,
1987) or keto-acids such as pyruvate (Holleman, 1904)) . More importantly hydrogen
peroxide is capable of reacting with additional electrons to form hydroxyl radicals which
are highly toxic and capable of a reacting with a wide range of proteins as well as lipids
and DNA (Cadet et al., 1999; Wolff et al., 1986). Superoxide can also react with nitric

5

oxide radicals (NO
•
), which are produced constantly as vasodilating agents and signaling
molecules (Garthwaite and Boulton, 1995) to form peroxynitrite (ONOO
-
). This is
detrimental both through the reduction in nitric oxide signaling (Moncada and Higgs,
1995) and through the formation of peroxynitrite which is a powerful oxidant. (The
reactions are summarized in Figure 1.1).

6

1. The four electron, univalent pathway for oxygen reduction
O
2

e-
O
2
•
− e-+2H+
H
2
O
2

e-+ H+
•OH
e-+ H+
H
2
O
H
2
O

2. Superoxide dismutase can catalyze H
2
O
2
formation from O
2
•
-

O
2
•
−
+ O
2
•
−
+ 2H
+
H
2
O
2
+ O
2

3. Superoxide can reduce oxidized transition metal ions(such as iron and copper)
Fe
3+
+ O
2
•
−
Fe
2+
+ O
2

4. Reduced transition metal ions (such as iron and copper) can react with hydrogen
peroxide in the Fenton reaction to catalyze hydroxyl radical production
Fe
2+
+ H
2
O
2
Fe
3+
+ •OH + OH
−

5. Superoxide and transition metals can react together to drive Fenton chemistry in
a doublet of steps termed the Haber-Weiss reaction.
O
2
•
−
+ Fe
3+
O
2
+ Fe
2+

Fe
2+
+ H
2
O
2
Fe
3+
+ •OH + OH
−

Figure 1.1: Formation of oxygen radicals and reactive oxidants:
A stage breakdown of the Haber-Weiss reaction

7

II. Formation of oxygen radicals and toxic oxidants by external factors

In addition to oxidative stress occurring through the formation of free radicals and
oxidizing agents in metabolism, oxidative stress can occur through contact with a range
of environmental factors. One of these environmental factors is air pollution. Air
pollution contains a range of metals which can catalyze the formation of reactive
oxidizing agents, such as iron, copper, chromium and vanadium, through the Haber-
Weiss reaction (Ghio et al., 1996; Ghio et al., 1999; Lloyd et al., 1998). Many particles in
air pollution can also produce an inflammatory response, which in part involves the
formation of oxygen radicals (Becker et al., 2005). UV radiation is another source of
oxidative stress, the main source of which is solar irradiation, but there are also a range of
lifestyle related sources of exposure. The degree of solar exposure to UV radiation is
dependent on a range of environmental factors such as cloud coverage and ozone level.
UV radiation can cause the formation of both superoxide (O
2
•-
) and hydroxyl radicals
(
•
OH). In addition it can cause direct modification of proteins within the cell. Similar
effects are also seen with infra-red and ionizing radiation (Schröder and Krutmann,
2005).

III. Removal of oxygen radicals and toxic oxidants by the cell

Various oxygen and nitrogen radicals, as well as other reactive oxygen or nitrogen
species, can (directly or indirectly) modify amino acids within proteins, damage vital

8

prosthetic groups, or oxidize key transition metal centers. Although direct modification of
amino acids or protein prosthetic groups typically increases local hydrophilicity (often
changing or introducing charges), the net effect of protein oxidative modification is
typically partial unfolding, which exposes the side chains of many hydrophobic amino
acids that are normally buried in the interior of a properly folded protein. Thus, when free
radicals, or related reactive species, modify a protein they usually cause an increase in
surface hydrophobicity and an increase in accessibility of protein hydrophobic groups.

While some mild modifications might be relatively harmless, other changes will reduce,
inhibit or modify the function of the protein (Davies, 1993, 1995, 2000a). If severely
oxidatively modified proteins are allowed to accumulate, cell function will be
progressively compromised (Davies, 1995). Thus, it is extremely important for any
aerobic cell to have a system for removing oxidatively damaged proteins. It is worth
noting that both DNA and lipid oxidation also present a serious challenge to maintenance
of cell function and aerobic cells have well-developed systems for combating these types
of damage. For the purpose of this chapter, however, I will focus solely on protein
oxidation.

The cells’ first line of defense is the removal of oxygen radicals and the detoxification of
reactive oxidants, to stop proteins becoming modified in the first place. Hydrogen
peroxide is a relatively mild oxidant, although it can be reduced to generate hydroxyl
radicals which are considerably more reactive (Figure 1.1). Because of this it is important

9

for hydrogen peroxide to be removed. This is done primarily through a glutathione
peroxidases (Mills, 1959), which couples the reduction of hydrogen peroxide into water
with the oxidation of glutathione (GSH) to glutathione disulfide (GSSG). The
glutathione disulfide is later reduced back to two glutathione molecules in a reaction
catalyzed by a range of different agents including glutathione reductase (Carlberg and
Mannervik, 1985). Peroxiredoxins also catalyze the reduction of hydrogen peroxide
coupled to the oxidation of thioredoxins; the hyperoxidation state of peroxiredoxins (as it
occurs in conditions of high oxidative stress) is prevented by sulfiredoxins (Findlay et al.,
2006). Catalase is another powerful enzyme that can remove hydrogen peroxide, but its
actions seem limited to peroxisomes in eukaryotic cells.

In addition to hydrogen peroxide, superoxide is also capable of reacting with copper, iron
or nitric oxide radicals. As a result of this, even with efficient removal of hydrogen
peroxide through glutathione peroxidases and peroxiredoxins, there are other pathways
by which protein damage can occur. In response to this cells have evolved superoxide
dismutases (SOD) (McCord and Fridovich, 1969). That catalyze the disproportionation of
superoxide anions to hydrogen peroxide, thereby decreasing the reaction of superoxide
with copper, iron, or nitric oxide. Although it may seem counter-intuitive for cells to
produce SOD that generates hydrogen peroxide, it seems that the combination of SOD
and glutathione peroxidases / peroxiredoxins ensures that most of the superoxide
produced is converted harmlessly to water without going through the hydroxyl radical
step in the univalent pathway for oxygen reduction. Minimizing the production of

10

hydroxyl radical, the most powerful oxygen radical of biological significance, seems to
be the strategy that all cells employ.

There is a range of other anti-oxidant mechanisms, largely based on antioxidant
compounds derived from fruits and vegetables in the diet, that are utilized by the cell.
Ascorbic acid (vitamin C), is one of these, it is involved in a similar reaction to
glutathione in which it functions as an electron acceptor for oxygen radicals resulting in
its conversion to dehydroascorbic acid (DHA). Dehydroascorbic acid may subsequently
be reduced back to ascorbic acid by a reaction with glutathione in the endoplasmic
reticulum (Nishikimi and Yagi, 1996). Another example is vitamin E (Halliwell and
Gutteridge, 2000) which functions as a scavenger of lipid peroxyl radicals and is
considered a chain-breaking antioxidant in the sequence of steps inherent in lipid
peroxidation. Many other phenolic or polyphenolic dietary components have also been
suggested as important biological antioxidants.

IV. Removal of damaged proteins by the proteasome

Although the first-line defenses of anti-oxidant enzymes and compounds described above
are quite effective, they are not perfect, and some oxygen radicals and other reactive
oxygen species do get through and cause protein damage. To combat this problem, the
cell has mechanisms in place to remove damaged proteins and enable normal cell
function to continue. The 20S Proteasome plays an important role in the removal of these

11

damaged proteins. The 20S proteasome is an alternative form of the 26S Proteasome in
which the 19S regulator has been removed. The 20S proteasome is a seven hundred
kilodalton barrel shaped protein composed of four rings each made up of seven subunits
ranging from twenty to thirty five kilodaltons. The first ring contains subunits α1-7, the
second ring subunits β1-7, the third ring subunits β1-7 and the fourth ring subunits α1-7.
The subunits β 1, 2 and 5 each possess’ specific proteolytic activities. In the 26S
Proteasome, the 19S regulator sits at either end of the 20S proteasome barrel, and
provides the apparatus for the selective degradation of Ubiquitin-labeled proteins in an
ATP/Mg
2+
dependent manner. In the 20S proteasome, which is free from the 19S
regulatory cap, proteins are degraded in an ATP/Ubiquitin independent manner
(Ciechanover, 1994; Coux et al., 1996) (shown in Figure 1.2).

12

Figure 1.2: 26S vs 20S proteasome
The 26S proteasome is composed of the 20S proteasome core. The core is made of four
rings each containing 7 subunits. The first ring contains subunits alpha 1-7. The second
ring contains subunits beta 1-7. The third ring contains subunits beta 1-7. The fourth ring
contains subunits alpha 1-7. Sunbunits beta 1,2 and 5 are the proteolytically active
subunits. The 19S regulator fits onto the top and bottom of the 20S core. The 19S
regulator will bind polyubiquitinated proteins and help to feed them into the core. The
20S proteasome possess the 20S core but the 19S regulator is absent.

In studies using purified 20S proteasome there is a selective preference for degradation of
oxidized over non-oxidized proteins for a wide range of proteins (Davies, 2001;
Pickering et al., 2010; Reinheckel et al., 1998). The 26S proteasome by comparison does
not appear in mammalian cells to have any selectivity for degradation of oxidized
proteins (Davies, 2001). Furthermore, while depletion of 20S proteasome from cells

13

results in an approximately eighty percent loss of capacity to degrade oxidized proteins
there is only a twenty percent reduction in capacity with depletion of 26S proteasome
from a cell (Pickering et al., 2010).

The 20S proteasome appears to be relatively resistant to oxidative stress; in fact, in
studies of its capacity to degrade oxidized proteins, under conditions of oxidative stress,
its I
50
appears to be four times higher than the level of oxidation of the target protein that
would result in optimum selective degradation (Reinheckel et al., 1998). This shows that
20S proteasome would be easily functional under the non-toxic degrees of oxidative
stress that the cell could be exposed to. In comparison the I
50
of purified 26S proteasome
under oxidative stress appears to be four to sixteen times lower than that of the 20S
proteasome (Reinheckel et al., 1998). Similarly in cell culture, levels of oxidation that
completely block ATP-stimulated 26S proteasome activity do not appear to significantly
affect 20S proteasome activity (Reinheckel et al., 2000b). As a result, while both the 20S
and the 26S Proteasome are capable of degrading oxidized proteins, the 26S proteasome
is inactivated under very mild degrees of oxidation compared to the 20S proteasome
(Reinheckel et al., 1998; Reinheckel et al., 2000b). In addition, in studies of oxidative
stress it has been seen that H₂O₂ causes a modest increase in a range of markers of
protein damage including: ditryosine, protein carbonyls, insoluble proteins, hydrophobic
proteins and protein aggregates. If, however, 20S proteasome is inhibited using NLVS,
Lactacystin, or 20S proteasome anti-sense then the H₂O₂ induced increase in all of these
is much larger (Demasi and Davies, 2003). Similarly H₂O₂ exposure slightly reduces

14

protein synthesis, DNA synthesis, and colony formation, all of which are very strongly
reduced if on top of the oxidative stress exposure, 20S proteasome activity is blocked
(Demasi and Davies, 2003). This implies that proteases, especially the 20S proteasome,
play a highly important role in response to oxidative stress.

V. Changes in protein damage and the proteasome over age

Over the course of aging there is a progressive decline in protein synthesis (Rattan,
1996). Despite this decline in synthesis there is actually an increase in protein content
with age. This discrepancy is a product of a buildup of damaged non-functioning proteins
within cells (Rattan, 1996). This buildup of damaged proteins can come from protein
oxidation as well as a range of other modifications including: glycation, methylation,
ADP-ribosylation, and deamidation (Rattan, 1996). The increase in damaged proteins in
cells is extremely detrimental and decreases the functionality of all pathways in the cell.
This forms part of the widely popular “Free Radical Theory of Aging”. This theory
suggests that many of the effects of aging are at least in part the product of accumulating
oxidative damage within a cell which results in deterioration of cell function and
eventually death (Harman, 1956).

Fortunately, much of the accumulated damage can be removed and the damaged proteins
can be degraded and replaced by non-damaged ones. In fact, some degree of modification
or damage to a protein makes it a better candidate for degradation by the 20S proteasome

15

or other proteases (Davies, 1993, 2000b, 2001; Pickering et al., 2010). However, if a
protein becomes too heavily modified it starts to become a very poor candidate for
degradation (Davies, 1993, 2001; Davies and Goldberg, 1987b). While a lot of the
accumulation of protein damage is readily degradable at least some of the buildup is a
product of damaged proteins which are so highly modified that they are difficult or
impossible to degrade by the cell machinery. It has been argued that it is the buildup of
these un-degradable damaged proteins that causes age-related effects. However, the
accumulation of oxidized proteins in cells over age is exponential rather than linear,
indicating that the rise in protein oxidation is not just a product of a buildup of un-
digestible proteins but a potentially reversible change in cell function (Stadtman, 1992).

The accumulation of oxidized or otherwise damaged proteins in cells during aging could
be a product either of a rise in damaging conditions or a fall in the rate of removal of
damaged proteins. It has been observed that over age there is a rise in mitochondrial
generation of oxidants (Sohal and Dubey, 1994). Though, in addition to that it has been
shown that there is a decline in protein turnover as well. This decline in protein turnover
is at least in part the product of a sharp decrease in 20S proteasome function over age
(which has been shown in a range of different tissues including muscle, lens, T-cells, and
lymphocytes) (Carrard et al., 2003; Chondrogianni and Gonos, 2005; Ferrington et al.,
2005; Husom et al., 2004; Ponnappan et al., 1999; Viteri et al., 2004). In addition to a
decline in 20S proteasome function there is also a drop in 20S proteasome level over the
course of age (observed in the spinal cord and epidermis) which also reduces protein

16

turnover (Bulteau et al., 2000; Keller et al., 2000; Petropoulos et al., 2000). The decline
in 20S proteasome function is partly the product of an increase in modification or damage
to the 20S proteasome over age (Bulteau et al., 2000; Carrard et al., 2003; Chondrogianni
and Gonos, 2005; Keller et al., 2000). For instance, it has been seen that 20S proteasome
isolated from old rats is 50% less proteolytically active than 20S proteasome that has
been isolated from young rats (Conconi and Friguet, 1997). As a result, not only is there a
decrease in the amount of 20S proteasome present during the aging process but there is
also a decrease in the ability of that 20S proteasome which is present to degrade the
accumulating damaged proteins so resulting in an increase of damaged proteins in the cell
over age. As a result of this, older rats are less able to remove damaged proteins than
their younger counterparts which likely contributes to the change in the level of protein
damage over age in rodents.

VI. Adaptation to oxidative stress

Cells or organisms are frequently exposed to a low level of oxidative stress or protein
damage. The level of oxidative stress exposure of an organism or a cell is not static, but
varies based on both external factors, such as environmental pollution, exposure to
ionizing radiation, and ingestion of toxins, as well as internal factors, such as
mitochondrial activity, immune response and aging. It seems reasonable then that the
capacity of a cell or organism to adapt to oxidative stress should itself be fluid. This was
confirmed using pre-treatment challenge experiments in which cells were exposed to a

17

mild non-toxic oxidative stress, then, after a period of time exposed to an oxidative stress
that would normally be toxic. It was seen from this that the cells that were pre-treated
were a lot more resistant to the challenge dose than cells which had not been pre-treated
(Pickering et al., 2010; Wiese et al., 1995).

18

CHAPTER 2: OXIDANT INDUCED ADAPTATION TO OXIDATIVE STRESS

Mammalian cells (as well as bacteria and yeast) can transiently and reversibly adapt to
changes in the level of oxidative stress exposure. This process is sometimes called
hormesis. Such adaptation can be measured using pre-treatment challenge experiments.
For such experiments, one first finds a challenge dose of the oxidant (e.g. H
2
O
2
) that
normally causes an easily measurable negative effect on cell growth and division. The
challenge dose should not be so strong, however, that it causes massive cell death from
apoptosis or necrosis, but strong enough to cause cell growth arrest. Separately, one
finds a much lower pre-treatment dose (or adaptive dose) of the same oxidant and allows
a suitable time-lag to permit adaption to occur. When the normally toxic challenge dose
is applied to pre-adapted cells, we find that they are transiently more resistant to the
stress. (Davies, 1999, 2000b; Ermak et al., 2002; Wiese et al., 1995). This chapter is
mostly composed of work which has been published in Pickering, A.M., Koop, A.L.,
Teoh, C.Y., Ermak, G., Grune, T., and Davies, K.J.A. (2010). The immunoproteasome,
the 20S proteasome and the PA28αβ proteasome regulator are oxidative-stress-adaptive
proteolytic complexes. Biochem J 432, 585-594 (Pickering et al., 2010), though it does
also contain some material from Pickering, A.M., Linder, R.A., Zhang, H., Forman, H.J.,
and Davies, K.J.A. (2012). Nrf2 dependent induction of proteasome and Pa28αβ
regulator is required for adaptation to oxidative stress. J Biol Chem. 2012 Mar 23;287,
10021-31 (Pickering et al., 2012) and some unpublished work. In most of the studies
performed we used Murine Embryonic Fibroblasts (MEF) that had been transformed

19

using an SV40 virus. This line was purchased from ATCC (Manassas, VA) catalog
#CRL-2214. This was used a general model of mammalian cells.

I. Hydrogen peroxide induced resistance to oxidative stress challenge

As a demonstration that the pre-treated cells were more resistant to oxidative stress we
performed a range of different measures of tolerance to oxidative stress challenge. We
performed cell count assays (Figure 2.1 A). In a cell count assay we pre-treated cells with
an adaptive dose of H
2
O
2.
24 h later we challenged the cells with a dose of H
2
O
2
which
was sufficient to cause almost complete growth arrest. Growth arrest was measured by
taking cell counts 24 h after challenge. We found that cells which had been pre-treated
showed much less of a growth arrest with challenge than cells which had not been pre-
treated. In addition to cell count assays we also measured adaptation using BrdU
incorporation assays (Figure 2.1B). In this pre-treatment and challenge was performed as
described in the cell count assay. After challenge, BrdU was added to cell culture, 24 h
later BrdU was washed out and BrdU incorporation measured. As in the cell count assay,
we observed a strong decline in BrdU incorporation with H
2
O
2
challenge which was
much less reduced with H
2
O
2
pre-treatment. Finally we used protein carbonyl assays, 6 h
after H
2
O
2
challenge, as a measure of oxidative stress adaptation (Figure 2.1C). Protein
carbonyl content serves as a marker of protein oxidation. With H
2
O
2
challenge there was
a sharp increase in protein carbonyl content which was reduced with H
2
O
2
pre-treatment.

20

Figure 2.1: H
2
O
2
induced adaptation to H ₂O ₂ challenge
H₂O₂ pretreatment enhances resistant to subsequent H₂O₂ challenge. Three different
measures of oxidative stress resistance were used (Cell counts, BrdU incorporation and
protein carbonyl content). In all cases, MEF wells were grown to 20% confluence then
some samples were pre-treated with a mild adaptive dose of H₂O₂ (50 μM of H
2
O
2
per
10
6
cells/ml) for 1 h. 24 h later both pre-treated and non-pretreated samples were
challenged with a toxic dose of H₂O₂ (2 mM of H
2
O
2
per 10
6
cells/ml) for 1 h. A. Cells
were permitted 24 h recovery after which cell counts were taken. B. After H₂O₂ challenge
BrdU was added to cell culture, cells were then allowed 24 h recovery after which BrdU
incorporation was measured. In C. Protein carbonyl content was measured 6 h after H₂O₂
challenge In all cases Values represent Means ± Standard Error (SE) where n = 3.

II. Induction of proteolytic activity

Since proteolysis (especially by the proteasome) plays a vital role in the removal of
damaged proteins during oxidative stress (Chondrogianni et al., 2003; Davies, 1986,

21

2001; Davies and Goldberg, 1987b; Fucci et al., 1983; Grant et al., 1993; Grune et al.,
1996; Grune et al., 1995; Keller et al., 2005; Pacifici et al., 1993; Shang and Taylor,
1995; Shringarpure et al., 2003; Ullrich et al., 1999; Whittier et al., 2004), we reasoned
that the intracellular proteolytic capacity to degrade oxidized proteins might increase
significantly during stress adaptation. To begin to test this hypothesis, we pre-treated
MEF cells with a mild (adaptive) dose of hydrogen peroxide, as previously (Davies,
1999, 2000b; Ermak et al., 2002; Wiese et al., 1995), and then allowed a suitable adaptive
period of 1-48 hours. Successful adaptation (peaking at about 24 hours and then
declining) was confirmed as described above. Having confirmed adaptation we prepared
cell lysate from cell culture 24 h after H
2
O
2
pre-treatment. We then added oxidized,
tritium-labeled, hemoglobin ([
3
H]Hb
ox
) or the fluoropeptide proteolysis substrate Suc-
LLVY-AMC (typically used as a measure of the chymotrypsin-like activity of the
proteasome) to the extracts, and measured changes in proteolytic capacity (Figure 2.2).
Both activities reached a peak of 3-4 fold increases, compared to untreated controls, 24
hours following pre-treatment and then began to decline back towards baseline levels.

We next sought to characterize the proteolytic enzyme(s) responsible for the increased
proteolytic capacity observed in Figure 2.2. Since previous work strongly suggested that
proteasome was the most likely candidate (Chondrogianni et al., 2003; Davies, 1986,
2001; Davies and Goldberg, 1987b; Fucci et al., 1983; Grant et al., 1993; Grune et al.,
1996; Grune et al., 1995; Keller et al., 2005; Pacifici et al., 1993; Shang and Taylor,
1995; Shringarpure et al., 2003; Ullrich et al., 1999; Whittier et al., 2004), we measured

22

all three proteasome-dependent proteolytic activities, the chymotrypsin-like, trypsin-like
and caspase-like activities of cell lysates, twenty four hours after hydrogen peroxide pre-
treatment (Figure 2.3A). We observed a two-fold increase in trypsin-like and
chymotrypsin-like activities and a ten-fold increase in caspase-like activity, all of which
were statistically significant increases by t-test analysis (p < 0.01). To confirm the
proteasomal identity of these activities, we repeated the experiments using the
proteasomal inhibitors lactacystin and MG132, which, indeed, blocked the majority of all
three activities in both control (Figure 2.3B) and H
2
O
2
adapted lysates (Figure 2.3C).

We have previously reported that nuclear proteasome can undergo direct activation by
poly-ADP-ribose-polymerase (Ullrich et al., 1999), and other direct mechanisms of
activating (existing) proteasome complexes, may also play a role during stress, without
need for de novo synthesis of new proteasomes. To differentiate between direct activation
of pre-existing proteasome complexes and de novo synthesis of proteasome (i.e.
induction), we first pre-incubated cells with cycloheximide to block protein synthesis,
and then exposed both cycloheximide-treated and untreated cells to an adaptive dose of
H
2
O
2
. As shown in Figure 2.4 and published in (Grune et al., 2011; Pickering et al.,
2010), cycloheximide had only a 10% inhibitory effect on increased proteasome activity
during the first hour of H
2
O
2
adaptation, indicating that a significant proportion of the
hour-one increased proteolysis reported in Figure 2.1 (using identical conditions) was
actually due to direct, physical activation of existing proteasome complexes; this could be
due to poly-ADP-ribose activation of proteasome in the nucleus (12), and/or to the effects

23

of various proteasome regulators such as 19S, PA28αβ (or 11S), PA28γ (or REGγ),
PA200, HSP90, etc. We further pursue such direct activation mechanisms in subsequent
chapters. Cycloheximide inhibited the increase in proteasome activity by 58% after three
hours of H
2
O
2
adaptation, by 82%, after 24 hours, and by 95% after 48 hours (Figure
2.4). From these results we can conclude that there is a two-stage response to hydrogen
peroxide pre-treatment- an initial translation-independent physical activation of existing
proteasomes, followed by a progressive increase in proteasome transcription/translation.

24

Figure 2.2: Proteolytic activity increases during transient adaptation to H
2
O
2

MEF cells were grown to 50% confluence and exposed, in PBS, to an adaptive pre-
treatment of 20 μM of H
2
0
2
per 10
6
cells. Successful transient adaptation (peaking at
about 24 hours and then declining) was confirmed by increased capacity to survive a
subsequent (much higher) challenge dose of H
2
O
2
that significantly decreased cell
proliferation and DNA replication, and significantly increased the accumulation of
oxidized cell proteins (confirmatory data not shown at this point, but given as part of
Figure 1.1) as previously described (Ermak et al., 2002; Wiese et al., 1995). At various
time points after exposure, the cells were harvested and lysed then suspended in 50mM
Tris, 25mM KCl, 10mM NaCl, 1mM MgCl
2
, (pH 7.5). Proteolytic activity assays for
degradation of either [
3
H]Hb
ox
or Suc-LLVY-AMC were performed (Pacifici and
Davies, 1990; Reinheckel et al., 2000a; Shringarpure et al., 2003). For [
3
H]Hb
ox

experiments, percent protein degraded was calculated after addition of 20% TCA and 3%
BSA (as carrier) to precipitate the remaining intact proteins (Fucci et al., 1983;
Shringarpure et al., 2003; Ullrich et al., 1999). Percent protein degraded was determined
by release of acid-soluble counts in TCA supernatants, using liquid scintillation, in which
% Degradation = (acid-soluble counts – background counts) x 100. Values are means ±
SE, n = 3.

25

Figure 2.3: Proteasome is induced during transient adaptation to H
2
O
2

A. MEF cells were grown to 20% confluence then transiently adapted to oxidative stress
by pre-treatment with 20 μM of H
2
0
2
per 10
6
cells in complete media and incubated at
37
o
C under 5% CO
2
. After one hour, the cells were washed twice with PBS and fresh
complete media added. Twenty four hours after exposure, the cells were harvested. Cells
were then lysed and suspended in 50mM Tris, 25mM KCl, 10mM NaCl, 1mM MgCl
2

(pH 7.5). Proteolytic activity assays for degradation of Suc-LLVY-AMC, Bz-VGR-
AMC, and Z-LLE-AMC were then performed. Values are means ± SE, n = 6. B. MEF
cells were prepared, harvested and lysed as described in Figure 2A but not pre-treated
with H
2
O
2
. Samples were then incubated with 1μM of either MG312 or lactacystin for
30 minutes after which proteolytic activity assays for degradation of Suc-LLVY-AMC,
Bz-VGR-AMC, and Z-LLE-AMC were performed. Values (means ± SE, n = 3)
represent the percent reduction in proteolytic activity following treatment with inhibitors.
C. MEF cells were prepared, transiently adapted to oxidative stress by pre-treatment with
20 μM of H
2
O
2
per 10
6
cells (as described in A), harvested and lysed. Samples were then
incubated with 1 μM of either MG312 or lactacystin for 30 minutes, after which
proteolytic activity assays for degradation of Suc-LLVY-AMC, Bz-VGR-AMC, and Z-
LLE-AMC were performed. Values (means ± SE, n = 3) in B and C are the percent
reduction in proteolytic activities, reported in A, caused by treatment with the proteolytic
inhibitors MG132 or lactacystin.

26

Figure 2.4: Proteolytic activity under adaptation with protein synthesis inhibition
A. MEF cells were incubated with 100 g/ml of cycloheximide (or not treated) in an
attempt to block H
2
O
2
induced expression of proteolytic enzymes. Cells were then grown
to 50% confluence and exposed (in PBS) to a transient adaptive pre-treatment of 20 μM
of H
2
0
2
per 10
6
cells, harvested, and lysed as described in the legend to Figure 1.
Proteolytic activity assays for degradation of Suc-LLVY-AMC were then performed, as
per Figure 2.1 and 2.2. B. The difference between cycloheximide -treated cells and
control cells from A is plotted as the percent inhibition (means ± SE, n = 3) exerted by
cycloheximide against the H
2
O
2
induced (adaptive) proteasome activity induction.

III. Induction through different oxidants

We now needed to determine if the increase in proteasome is specific to H
2
O
2
, or if it is a
more general response to oxidants. We first pre-treated MEF cells with various
concentrations of H
2
O
2
, peroxynitrite, or the redox cycling agents paraquat and
menadione for 1 h. Then, 24 h later, we harvested and lysed the cells and measured

27

proteolytic capacity by degradation of the fluorogenic peptide, Suc-LLVY-AMC which is
widely used to estimate the chymotrypsin-like activity of the proteasome (Pickering et al.,
2010; Reinheckel et al., 2000a; Ullrich et al., 1999) as we did in Figures 2.1 and 2.2. We
saw a 2-fold increase in proteolytic capacity with H
2
O
2
or paraquat pre-treatment, a 2.5-
fold increase with peroxynitrite pre-treatment and a 3-fold increase with menadione pre-
treatment (Figure 2.5A-D). In lysates of untreated cells, the selective proteasome
inhibitor lactacystin caused an 80-90% inhibition of proteolysis. In lysates of oxidant pre-
treated cells, lactacystin inhibited degradation by 90-95%, indicating that proteasome is
largely responsible for most of the oxidant-induced adaptive increase in proteolytic
capacity (Figure 2.5A-D). This experiment was repeated using another proteasome-
selective inhibitor, MG132 which blocked 50% of activity in untreated cells, and 60% of
activity following under oxidative stress adaptation (Figure 2.5E-F).

Given that pre-treatment with peroxynitrite, paraquat and menadione, like H
2
O
2
caused
anincrease in proteolytic capacity, we next wanted to test if this also corresponded to an
increase in tolerance to H
2
O
2
toxicity as we have seen with H
2
O
2
. As shown in Figure
2.6, 2 mM H
2
O
2
challenge, per 10
6
cells, caused a 65% decrease in cell counts in non-
adapted cells; this was mostly due to prolonged growth arrest, as previously shown
(Davies, 1999, 2000b; Ermak et al., 2002; Pickering et al., 2010; Wiese et al., 1995). In
contrast, cells that had been pre-treated with (low concentrations of) a range of oxidants
exhibited substantially less toxicity: only a 29% growth arrest with H
2
O
2
pre-treatment,
37% with paraquat, 42% with menadione, and 50% with peroxynitrite (Figure 2.6).

28

Figure 2.5: Oxidant pre-treatment increases proteolytic capacity in a proteasome
dependent manner
Cells treated with a mild dose of a range of oxidants exhibit increased proteolytic
capacity, the majority of which (80-95%) is blocked by the proteasome selective inhibitor
lactacystin. MEF cells were grown to 10% confluence (≈250,000 cells per ml) and treated
with A: 0μM - 10μM H
2
O
2
, B: 0μM - 1μM Peroxynitrite, C: 0 nM – 100 nM Paraquat or
D: 0 nM – 100 nM Menadione. All treatments were for 1 h in complete media, following
which the media was removed and replaced with fresh complete media. After 24 h , cells
were lysed and diluted to a protein concentration of 50μg per ml. Proteolytic capacity was
determined by cleavage of the proteasome chymotrypsin-like substrate Suc-LLVY-AMC.
In some samples 5μM lactacystin was added to samples, 30 minutes prior to incubation
with suc-LLVY-AMC. Values are Means ± SE. n = 3. E: MEF cells were prepared as
described in A and pre-treated with 100nM peroxynitrite, 1μM H
2
O
2
, 1nM menadione or
1nM paraquat for 1 h in complete media; following this, the media was removed and
replaced with fresh complete media. In some samples, 1μM MG132 was added 30
minutes prior to incubation with Suc-LLVY-AMC. Cells were incubated, harvested
lysed, diluted and analyzed for proteolytic capacity by degradation of the fluorogenic
peptide suc-LLVY-AMC. Values are Means ± SE’s, n = 3. F: Results from E were re-
plotted with the decrease in activity resulting from addition of MG132 plotted as a
percent of the proteolytic capacity of cells not treated with the inhibitor.

29

Figure 2.6: Pre-treatment with different oxidants improves tolerance to H ₂O ₂
challenge
Pretreatment of cells with a mild non-toxic dose of a range of oxidants caused increased
tolerance to a subsequent toxic H
2
O
2
challenge. MEF cells were grown to 10%
confluence and treated with 1 nM peroxynitrite, 1 nM menadione, 1pM paraquat, or 100
nM H
2
O
2
for 1 h, then washed and re-suspended. After 24 h, cells were challenged with 1
mM H
2
O
2
for 1 h, then washed and re-suspended. After another 24 h, cells were
harvested and counted. Values are Means ± SE. n = 3.

30

IV. Adaptation under repeated oxidative stress

While a mild acute oxidative stress appears to have a beneficial effect, natural oxidative
stress exposure is typically not acute but is repeated. We wanted to see how the adaptive
capacity of cells was affected by repeated oxidative stress compared to a single acute
oxidative stress.

To study changes in proteolytic capacity under repeated stress we exposed cells to the
adaptive dose of 50μM of H₂O₂ per 10
6
cells ml
-1
and 24 h later measured proteolytic
capacity through degradation of the chymotrypsin like substrate Suc-LLVY-AMC
(Figure 2.7A), which formed a measure of adaption to acute stress. To measure adaption
to repeated stress the cells were exposed to the adaptive dose of H₂O₂ a second and in
some cases a third time after the initial exposure. These subsequent exposures were at 3,
6 or 12 h intervals after the initial exposure. As before we saw that a single pre-treatment
caused an increase in proteolytic capacity. If cells were pre-treated a second time 12hr
after the first treatment, the second pre-treatment was tolerated and proteolytic capacity
was not diminished by the second pre-treatment. If however samples were pre-treated a
second or even a third time within 6hr of the initial pre-treatment then the proteolytic
capacity, while still greater than that of non-pretreated cells, was reduced from the levels
seen with a single pre-treatment. Having seen that a second pre-treatment 12 h after the
initial pre-treatment was tolerated we wished to test if further 12 h interval pre-
treatments would also be tolerated. To test this, as before cells were exposed to an

31

adaptive dose of H₂O₂, 48 h later cells were harvested and proteolytic activity measured
through degradation of Suc-LLVY-AMC. In addition to the first pre-treatment, some
samples were pre-treated a further 1-3 times at 12 h intervals (Figure 2.7B). It appeared
from this that even 4 pre-treatments at 12hr intervals were tolerated, with no loss of
proteolytic capacity.

From the above results it appeared that treatments within 6hr intervals had detrimental
effects on adaptation which we decided to study further. To better examine proteolytic
capacity we examined relative capacity to degrade an oxidized and a native protein
(hemoglobin) which we have previously considered a truer measure of proteolytic
adaptation (Davies, 2001; Pickering et al., 2010). Here we saw that capacity to degrade
oxidized proteins rose with a single treatment and was un-affected by a second treatment
but was reduced with a third successive oxidative stress treatment. Capacity to degrade
native protein by comparison continued to rise in the three successive treatments. The
result of this was that the capacity to selectively degrade oxidized proteins rose with a
single treatment but then progressively declined with subsequent treatments (Figure
2.8A). We went on to look at adaption using the pre-treatment challenge model described
earlier, again we saw an increase in oxidative stress tolerance with a single treatment
which was progressively reduced with subsequent treatments (Figure 2.8B).

Having looked at oxidative stress tolerance with repeated treatments at 6hr intervals
which we had shown to be detrimental we went on to look at the adaptive capacity of

32

cells with treatments at 12 h intervals, which in terms of proteolytic capacity appeared to
be permissible. In addition we were interested in whether the concentration of the oxidant
used in the repeated stresses would affect the response. We found that using the adaptive
dose of 50μM (per 10
6
cells/ml), which we found to be the optimum treatment, repeated
stresses did not diminish the increase in tolerance to oxidative stress. In addition we
found that when we used lower doses of H ₂O ₂ than what we would normally find
adaptive the oxidative stress tolerance of cells increased with repeated treatments;
however, if doses at the higher end of what would be found to be adaptive were
employed then the repeated stress appeared to have a detrimental effect on oxidative
stress tolerance (Figure 2.9). From this we can conclude that repeated stress can be
permissible in terms of adaptation, though the degree of permissibility is dependent on
the frequency of exposure and the size of the dose employed.

33

Figure 2.7: Capacity to degrade Suc-LLVY-AMC with repeated H ₂O ₂ pre-
treatment
A. Cell were grown to 20% confluence and pre-treated with 1 μM H₂O₂ for 1 h. Some
cells were exposed to the 1 μM dose twice at either 0 h and12 h, 0 h and 6 h or 0 h and 3 h
after initial treatment. In addition some cells were exposed to H₂O₂ three times at 0 h, 6 h
and 12 h or 0 h, 3hr and 6 h after initial treatment. Samples were harvested by scraping
24hr after initial treatment. Samples were lysed through 3 cycles of free-fractionation,
protein content was adjusted by BCA assay and capacity to degrade Suc-llvy-AMC was
measured as before. Values are Mean ± SE where n = 3. B. Cells were grown to 20%
confluence and pre-treated 0-4 times at 12hr intervals with 1 μM H₂O₂. Treatments were
performed at 0 h, 12 h, 24 h and 36 h after initial treatment. Samples were harvested 48
h after initial treatment as in A. Values are Mean ± SE where n = 3.

34

Figure 2.8: Adaptive effects of repeated H ₂O ₂ pre-treatments at 6 h intervals
A. Cells were prepared, treated, harvested and lyzed as described in Figure 2.7.
Proteolytic capacity was measured using Hb-AMC and oxidized Hb-AMC. Values are
mean ± SE where n = 6. B. Cells were prepared and treated as in A. 24 h after the initial
treatment cells were challenged with 1mM H₂O₂ for 1 h. 24 h after the challenge cell
counts were taken. Values are mean ± SE where n = 6.

35

Figure 2.9: Adaptive effects of repeated H ₂O ₂ pre-treatments at 12 h intervals
Cells were grown to 10% confluence then pre-treated with different doses of H₂O₂ at 0 h,
12 h, 24 h and 36 h after initial treatment. Samples were then challenged with 1mM H₂O₂
for 1 h, 24 h later cell counts were taken. Values are Mean ± SE where n = 4.

V. Adaptation under chronic oxidative stress

As well as studying repeated oxidative stress, a limited set of experiments were
performed under chronic oxidative stress. Models of chronic stress are difficult for a
range of reasons. Given that H₂O₂ and other oxidants are rapidly degraded by cells it is
not possible to have a continuous dose of H₂O₂ in cell culture. One solution is the use of
enzymes such as glucose oxidase (GO) which catalyze the formation of H₂O₂ from
glucose. We found that if a sufficiently low dose (10fg/ml-1000fg/ml) of GO is used then

36

GO treatment can be maintained for 24 h with no significant effects to cell division rates.
To compare the effects of acute vs chronic oxidant exposure on oxidative stress
adaptation, we treated cells with glucose oxidase for either 1 h or 24 h. 24h after this
treatment cells were challenged with a toxic dose of H₂O₂ and cell counts were taken 24 h
later, as before. We found that a 1 h treatment with GO had a progressively stronger
adaptive effect on tolerance to oxidative stress from 10fg/ml to 1000fg/ml. By
comparison in the 24 h treatment regime, 10fg/ml and 100fg/ml treatment had stronger
beneficial effects than the same dose at 1 h exposure; however 1000fg exposure at 24 h
had a much weaker adaptive effect than the same dose at 1 h exposure (Figure 2.10).

37

Figure 2.10: Adaptation under chronic oxidative stress
Chronic oxidative stress exposure of mammalian cell culture produces a stronger
adaptive response than acute exposure with weaker oxidant concentrations and produces
a blunted response compared to acute exposure with higher oxidant concentrations. MEF
cells were grown to 20% confluence then treated with 10-1000fg/ml of glucose oxidase
for either 1 h or 24 h, cells were then washed and permitted to adapt and recover for 24 h.
After recovery, cells were challenged with 2 mM of H
2
O
2
per 10
6
cells/ml for 1 h, 24 h
later cell counts were taken. Values are plotted as a percent of the cell counts of samples
which were not exposed to the challenge and are shown as mean ± SE where n = 3.

VI. Summary

During adaptation to H
2
O
2
cells undergo a two-stage response: an initial direct activation
of pre-existing proteasome during the first hour [perhaps by poly-ADP-ribose in the
nucleus (6) or other proteasome regulators in the cytoplasm], followed by a much slower
de novo synthesis response. After 24 hours, the cellular capacity to degrade oxidized

38

proteins increased by more than three-fold, and essentially all of this increase could be
blocked by proteasome inhibitors. These results demonstrate that proteasome is a highly
likely candidate in this response to oxidative stress, being both activated and induced
under stress-adaptive conditions.

This response appears not just to be inducible with H₂O₂ but broadly inducible by
oxidative stress which is seen by using peroxynitrite, and the redox-cycling agents:
menadione and paraquat instead of H₂O₂ all of these oxidants increased cellular tolerance
to a subsequent oxidative stress challenge as well as increasing proteolytic activity in a
proteasome dependent manner.

An acute exposure to oxidative stress appears to cause a clear increase in oxidative stress
tolerance. A repeated exposure, however, is less beneficial and can reduce the adaptive
response. The severity of this reduction is dependent on the frequency of oxidative stress
exposure. If, however, a sufficiently long recovery time is permitted then repeated stress
proves not to be detrimental and might be slightly beneficial.

39

CHAPTER 3: THE 20S AND 26S PROTEASOME IN STRESS ADAPTATION

This chapter discusses the relative roles of the 20S and 26S proteasome in adaptation to
oxidative stress. The chapter is mainly composed of materials published in Pickering,
A.M., Koop, A.L., Teoh, C.Y., Ermak, G., Grune, T., and Davies, K.J.A. (2010). The
immunoproteasome, the 20S proteasome and the PA28αβ proteasome regulator are
oxidative-stress-adaptive proteolytic complexes. Biochem J 432, 585-594 (Pickering et
al., 2010) but also briefly discusses material published in Grune, T., Catalgol, B., Licht,
A., Ermak, G., Pickering, A.M., Ngo, J.K., and Davies, K.J.A. (2011). HSP70 mediates
dissociation and reassociation of the 26S proteasome during adaptation to oxidative
stress. Free radical biology & medicine 51, 1355-1364.(Grune et al., 2011) of which I am
a co-author.

I. Role of 20S vs 26S proteasome in degrading oxidized proteins

Many of the experiments in chapter 2, and a large number of papers published in the
literature, point to an important role for proteasome in the adaptive response to oxidative
stress. In addition, there is moderate evidence for a greater role of 20S compared to 26S
proteasome in response to oxidative stress ((Davies, 2001; Pickering et al., 2010;
Reinheckel et al., 2000a; Reinheckel et al., 1998) and reviewed in the introduction). To
further examine the relative roles of 20S and 26S proteasome in response to oxidative
stress we employed siRNA targeted to specific subunits of each complex. As shown in

40

Figure 3.1A, two days of treatment with siRNA directed against the β5 subunit of the 20S
‘core’ proteasome blocked some 50% of the capacity to degrade an oxidized hemoglobin
substrate. In contrast, two days of treatment with siRNA directed against the S4 subunit
of the 19S regulator of the 26S proteasome blocked only 20% of the capacity to degrade
oxidized hemoglobin. After five days of β5 subunit ‘knock-down,’ capacity to degrade an
oxidized hemoglobin substrate was inhibited by 80%, whereas S4 subunit knock-down
for five days still only caused a 30% inhibition.

Since we could not be sure from Figure 3.1A that 26S proteasome had actually been
successfully knocked-down, we tested the ability of the S4 siRNA-treated cells to
conduct ATP-stimulated proteolysis – an activity that depends upon the 19S subunit.
Figure 3.1B shows that ATP-stimulated proteolysis was severely compromised, thus
demonstrating the effectiveness of the S4 knock-down procedure. Addition of ATP
produced a 4.2-fold increase in proteolysis in control samples treated with control
(scrambled) siRNA (and this is shown as 100% ATP stimulation of proteolysis on day 1
in Figure 4B). By day 2 of S4 siRNA treatment, however, ATP stimulation of proteolysis
was only 10% of control values, and by day 5, ATP completely failed to stimulate
degradation. Furthermore, we see that 2 days of S4 siRNA treatment causes a
considerable loss of the S4 subunit in cells (see western blot in inset). We suggest that
this S4 siRNA-mediated loss of ATP-stimulated 26S proteasome proteolysis (Figure
3.1B) makes it reasonable to conclude that the comparison of S4 and β5 subunit knock-
down experiments in Figure 3.1A is a valid one. The dramatic fall in oxidized substrate

41

degradation with β5 knockdown implies that the core 20S proteasome plays a highly
important role in removal of oxidized proteins from the cell, while the considerably
smaller reduction under S4 knockdown implies that the ATP/ubiquitin-dependent 26S
proteasome plays a relatively minor role.

It should also be noted that interference of 26S proteasome function, e.g. by S4 subunit
knock-down, is actually known to impede synthesis of the 20S proteasome: thus S4
knock-down would eventually be expected to limit 20S proteasome activity anyway
(Muratani and Tansey, 2003). The 19S regulator has been demonstrated to have
important and diverse roles in transcription, (reviewed in (Collins and Tansey, 2006;
Lipford and Deshaies, 2003)). While exact mechanism(s) are unclear, it has been
observed that several subunits within the 19S regulator have important roles in the
recruitment of RNA polymerase (Swaffield et al., 1992). It has also been shown that the
19S proteasome regulator is required for recruitment of RNA polymerase II to promoter
sites on many genes, and absence or insufficiency of the 19S results in decreased gene
expression (Dennis et al., 2005; Lipford et al., 2005). As a result, even this small
reduction in capacity to degrade oxidized proteins with 26S proteasome knock-down
might itself be a product of reduced 20S proteasome synthesis.

42

Figure 3.1: Importance of 20S vs 26S proteasome in degrading oxidized proteins
A. MEF cells were grown to 50% confluence and treated with β5, S4 or control
(scrambled) siRNA. Cells were grown for a further 2 or 5 days. Cells were then
harvested, lysed, and suspended in 50mM Tris, 25mM KCl, 10mM NaCl, 1mM MgCl
2
at
(pH 7.5). Proteolytic activity assays for degradation of [
3
H]Hb
ox
were performed as in
Figure 2.1 (Pacifici and Davies, 1990; Reinheckel et al., 2000a; Shringarpure et al.,
2003). Values are means ± SE, n = 3. B. Cells were prepared as described in A and
treated with either S4 or control (scrambled) siRNA. Cells were grown for a further 1 to 5
days and were then harvested, lysed, and suspended in 50mM Tris, 25mM KCl, 10mM
NaCl, 1mM MgCl
2
at (pH 7.5). Proteolytic activity assays for ATP-stimulated
degradation of Suc-LLVY-AMC was then performed in the presence and absence of
10mM ATP. Addition of ATP produced a 4.2-fold increase in proteolysis in control
samples (not treated with siRNA, or treated with control (scrambled) siRNA), and this is
shown as 100% ATP stimulation of proteolysis on day 1 in B (data are means ± SE, n =
3). By day 2 of S4 siRNA treatment, however, ATP stimulation of proteolysis was only
10% of control values, and by day 5, ATP completely failed to stimulate degradation.
Inset shows a western blot of S4 levels over a period of 2 days of siRNA treatment.

43

II. The role of proteasome in protein synthesis independent response

In chapter 3 we described a two stage adaptive response to oxidative stress, and in this we
saw an initial protein synthesis independent increase in proteolytic capacity followed by a
subsequent protein synthesis dependent increase in proteolytic capacity (Pickering et al.,
2010). In collaboration with the lab of Dr. Tilman Grune we confirmed this result then
demonstrated that this initial protein synthesis independent response was the product of
the 19S regulatory cap detaching from 26S protein to form free 20S proteasome (Grune et
al., 2011). This free 20S proteasome would facilitate a transient increase in cellular
proteolytic capacity. 3-5 h after its detachment, 26S proteasome would subsequently re-
form such that normal cell function could resume. This process was shown to be assisted
by Hsp-70, which is thought to stabilize the 19S regulator while it is detached (Grune et
al., 2011). In addition, another group has evidence that phosphorylation by the regulator
ECM29 facilitates the initial detachment of the 19S regulator (Wang et al., 2010)
(diagram in Figure 3.2). We confirmed the detachment of the 19S regulator under
oxidative stress, using our MEF cell-line. In this experiment we subjected cells to
hydrogen peroxide for 1 h. Cell lysate from these cells was pre-cleared then
immunoprecipitated using the 20S proteasome subunit β5. The eluate was run on an SDS
page gel and screened for the 19S regulator S4 subunit (Figure 3.3A). Here we saw an
H₂O₂ induced decline in 26S proteasome relative to 20S proteasome. In addition, we
confirmed that this detachment provided an adaptive response by performing a 2 day
siRNA treatment on cells using siRNA directed against either the S4 subunit or a

44

scrambled vector (Figure 3.3B). siRNA knock-down of S4 appeared to increase tolerance
to oxidative stress which is consistent with our model.

Figure 3.2: Disassembly and reassembly of the 26S proteasome in response to
oxidative stress
Under oxidant exposure Ecm29 induces disasociation of the 19S regulator from the 26S
proteasome to produce a rapid increase in the amount of free 20S proteasome. 3-5h later
the 19S regulator will re-attach so that normal cell function may resume. This re-
attachment is regulated by the Hsp70 chaperone.

45

Figure 3.3: Dissociation of 26S proteasome with H ₂O ₂ treatment
A. Under H ₂O ₂ treatment there is a decrease in S4 binding to 20S proteasome. MEF
cells were grown to 10% confluence, exposed to 1mM H ₂O ₂ for 1 h, after this
immunoprecipitation of anti-β5 was performed. Samples were run on a western blot and
screened for co-immunoprecipitation of anti-S4 and anti-β5. Values are Mean ± SE
where n = 3. B. MEF cells were grown to 10% confluence and treated with siRNA for 48
h then challenged with 1 mM H
2
O
2
; 24 h later cell counts were taken. Values are means ±
SE, n = 6.

III. The role of proteasome in protein synthesis dependent response

Because the core 20S proteasome appeared to be much more important than the
ATP/ubiquitin-dependent 26S proteasome for oxidative-stress adaptive increases in the
ability to degrade oxidized proteins, we wanted to explore the possibility of its induction
as part of the protein synthesis dependent response to oxidative stress. To do this we

46

performed a series of western blot analyses of key proteasome subunits. First, we probed
lysates from control and H
2
O
2
adapted cells with antibodies against the α3, α4, β1, and β2
core 20S proteasome subunits. The four 20S subunits examined all exhibited a
progressive rise of about two-fold during 24 hours of adaptation following mild H
2
O
2
pre-
treatment (Figure 3.4), all of which were observed to have changed significantly between
the 0 and 24 hour time points, by t-test analysis (p < 0.01). Critically, we observed no
significant change in the level of 26S proteasome (S4 subunit of the 19S proteasomal
regulator) during this analysis (Figure 3.4). In conjunction with previous results, this
suggests that the oxidative stress induced increase in capacity to degrade oxidized
proteins is at least partly due to de novo synthesis of the 20S core proteasome. This
change in proteolytic activity also appears to be independent of the ATP/ubiquitin
stimulated 26S proteasome.

To determine if the observed induction of 20S proteasome was actually relevant to
adaptation we assessed the importance of these proteins in pre-treatment induced
adaptation to oxidative stress. Cells were pre-treated with a mild (adaptive) dose of
hydrogen peroxide and then challenged 24 hours later with a more toxic dose as was
described in chapter 2. We have previously demonstrated that sub-lethal oxidative stress
challenge causes a sharp decrease in DNA synthesis, transcription, translation, and rates
of cell division, in (previously) divisionally competent cells (Davies, 1999, 2000b; Ermak
et al., 2002; Wiese et al., 1995). The doses of hydrogen peroxide challenge stress used
was fairly mild (and more biologically relevant than extreme stress), and so the main

47

effect of peroxide challenge was slow growth, rather than apoptosis, for which the Cell
count assay and BrdU assays provide the most suitable means of measurement. We
confirmed these results by performing a caspase-3 assay on both challenged and
unchallenged cells (vide infra).

We initially measured stress-tolerance by incorporation of BrdU in the 24 hours
following challenge, in both transiently H
2
O
2
adapted (pre-treated) and non-adapted cells.
This served as a measure of DNA replication and hence an indirect measure of cell
division (Figure 3.5A). We then repeated the experiment looking instead at the increase
in cell population, 24 hours after a hydrogen peroxide challenge (Figure 3.5B) in both
transiently H
2
O
2
adapted (pre-treated) and non-adapted cells. This provided a measure
not of cell death but of growth-arrest following challenge.

The experiments of Figure 3.5A show that BrdU incorporation was 60% higher in H
2
O
2

adapted (pre-treated) cells than in non-adapted cells following H
2
O
2
challenge and
control siRNA treatment. In contrast, the elevated BrdU incorporation of H
2
O
2
adapted
(and then challenged) cells was lost after treatment with β1 siRNA (20S proteasome
knock-down), which is statistically significant at p < 0.01. Similar results are seen with
cell counts as in BrdU incorporation, cell counts were 50% higher in H
2
O
2
adapted (pre-
treated) cells than in non-adapted cells following H
2
O
2
challenge and control siRNA
treatment.

48

The results of the cell survival and growth assays (cell counts) in Figure 3.5B also
revealed important roles for 20S proteasome in overall oxidative stress adaptation. We
observed a 50% increase of cell counts after H
2
O
2
challenge in cells that were first
adapted by pre-treatment with low levels of H
2
O
2
(Figure 3.5B). Treatment with β1
siRNA (20S proteasome knock-down) completely abrogated the adaptive increase in cell
counts. This was again statistically significant at p < 0.01. It is, perhaps, remarkable that
simply blocking the induction of 20S proteasome could have such a negative effect on
DNA replication and cell proliferation, especially considering the large number of genes
that are up-regulated (and those that are down-regulated) during the adaptive response to
H
2
O
2
(Davies, 1999, 2000b; Ermak et al., 2002; Wiese et al., 1995). Nevertheless,
proteasome seems to be an important component of cellular defenses against oxidative
stress, and blocking its induction appears to severely compromise overall oxidative stress
adaptation.

49

Figure 3.4: Induction of 20S proteasome with H
2
O
2
treatment
MEF cells were grown to 50% confluence and exposed (in PBS) to a transient adaptive
pre-treatment of 20 μM of H
2
0
2
per 10
6
cells, then harvested through scraping 0, 1, 6 or
24 h after treatment. Cells were then lysed and the resulting samples run through
Western blot analysis in which samples were treated with antibodies raised against the
proteasome subunits β1, β2, α3, α4 and S4. An enhanced chemiluminescence kit, from
Pierce (Rockford, IL), was used for chemiluminescent detection and membranes were
developed using the Kodak GBX developing system purchased from VWR (West
Chester, PA). 20S proteasome subunit levels were quantified in comparison with
standards, and are reported as percent of control (non H
2
O
2
adapted) levels: values are
means ± SE, n = 4.

50

Figure 3.5: Importance of 20S proteasome in adaptation
A. MEF cells were grown to 20% confluence and then treated with β1 or control
(scrambled) siRNA for 24 hours. After siRNA exposure the media was replaced with
fresh complete medium and after a further 24 hours (a total of 48 hours after initial
siRNA exposure), some cells were transiently adapted to oxidative stress by pre-
treatment with 20 μM of H
2
0
2
per 10
6
cells, while others were not adapted. Cells were
incubated at 37
o
C under 5% CO
2
for 1 hour, after which the medium was replaced.
Following a 24 hour adaptation period, both adapted and non-adapted cells were
challenged by incubation with 1 mM H
2
O
2
. Cells were then harvested and reseeded at
100,000 cells per ml on 96 well plates and the BrdU assay was then performed. BrdU
results (values are means ± SE, n = 3) represent cellular BrdU incorporation into DNA in
arbitrary units. On the X-axis, “No Pre-treatment” represents samples that were treated
with control (scrambled) siRNA and challenged with high H
2
O
2
, but were not adapted by
pre-treatment with low H
2
O
2
. All other samples were first treated with siRNA, adapted
by pre-treatment with low H
2
O
2
, and then challenged by exposure to high (1.0mM) H
2
O
2
.
B. MEF cells were prepared, treated with siRNA, transiently adapted to oxidative stress
by pre-treatment with H
2
O
2
(or not pre-treated), challenged with 1 mM H
2
O
2
, and
harvested, exactly as described in A. Samples were then seeded at a density of 100,000
cells per ml in 24 well plates. Cells were incubated for a further 24 hours, then cell counts
were taken using a cell counter. Values are means ± SE, n = 3 and represent the cell
population in challenged cells which previously were either pre-treated with an adaptive
dose of H
2
O
2
or not pre-treated.

51

IV. Summary

Our studies indicate that the 20S proteasome plays a major role in the degradation of
oxidized proteins compared to a relatively minor role for the 26S proteasome. We go on
to demonstrate that the proteasome is a highly plastic system under mild oxidative stress
and that the 20S proteasome is induced during transient adaptation to oxidative stress.
Furthermore, this was demonstrated to provide a significant contribution to adaptation
and increased tolerance to oxidative stress. We show that the initial protein synthesis
independent response is a product of detachment of the 19S regulator from the 26S
complex to form more free 20S proteasome. This is followed by a slower protein
synthesis dependent response of de novo 20S proteasome synthesis.
This model is represented in Figure 3.6.

52

Figure 3.6: Model for adaptive response
With H
2
O
2
exposure there is an initial protein synthesis independent response in which
the 19S regulator temporarily detaches from the 26S proteasome to increase the cellular
level of 20S proteasome. There is then a subsequent upregulation of 20S proteasome
protein levels by de novo synthesis.

53

CHAPTER 4: THE NRF2 TRANSCRIPTION FACTOR A REGULATOR OF
OXIDATIVE STRESS ADAPTATION

This chapter will discuss work published in Pickering, A.M., Linder, R.A., Zhang, H.,
Forman, H.J., and Davies, K.J.A. (2012). Nrf2 dependent induction of proteasome and
Pa28αβ regulator is required for adaptation to oxidative stress. J Biol Chem. 2012 Mar
23;287,10021-31 (Pickering et al., 2012).

I. Induction of Nrf2 under oxidative stress

We have demonstrated that mammalian cells, as well as bacteria and yeast, can
transiently become more resilient to oxidative stress under oxidant exposure, as described
in chapter 2 (Davies, 1993, 2000b; Ermak et al., 2002; Pickering et al., 2010; Wiese et al.,
1995). This is an adaptive or hormetic process in which cells treated with a mild dose of
an oxidant will, for a period of time (≈24-48 h), become more resistant to a higher dose
of the same (or related) oxidant that would normally be toxic. We have also shown that
this occurs by a process that is dependent upon the 20S proteasomes (described in chapter
3) (Grune et al., 2011; Pickering et al., 2010). Other groups have also reported induction
of various forms of the proteasome, and proteasome regulators, by oxidative stress
(Ferrington et al., 2005; Ferrington et al., 2008; Husom et al., 2004; Kotamraju et al.,
2006; Yamano et al., 2002).

54

The Nrf2 [nuclear factor (erythroid-derived 2)-like 2] transcription factor is an important
component of responses to oxidative stress (Itoh et al., 1997; Itoh et al., 2003; Kwak et
al., 2003; McMahon et al., 2001; Moi et al., 1994; Nguyen et al., 2003; Rushmore et al.,
1991; Venugopal and Jaiswal, 1996). Under non-stressful conditions Nrf2 is maintained
at low levels through rapid degradation via Keap1-dependent ubiquitin conjugation (Itoh
et al., 1999; Kwak et al., 2003; McMahon et al., 2003), followed by targeted degradation
by the 26S Proteasome. As a product of this rapid turnover newly translated Nrf2 is found
predominantly in the cytoplasm. Keap1 can become inactivated, as a product of factors
such as oxidative stress This will cause Nrf2 levels to increases due to decreased
degradation. In addition to this Nrf2 is phosphorylated by akt and pkcδ (Zhang and
Forman, 2008) and translocates to the nucleus. Once in the nucleus, Nrf2 binds to a cis-
acting enhancer sequence, upstream of numerous antioxidant genes, known as the
antioxidant response element (ARE) or electrophile responsive element (EpRE), and
promotes the synthesis of several antioxidants, and enzymes responsible for
repairing/removing oxidative damage and restoring cell viability (Rushmore et al., 1991).

It has been shown that Nrf2 knock-out in mice results in decreased tolerance to oxidative
stress (Enomoto et al., 2001; Ramos-Gomez et al., 2001). Additionally, results by Kwak
et al (Kwak et al., 2003) showed that 3H-1,2-dithiole-3-thione (D3T), which induces
many cellular antioxidants and phase 2 enzymes, can also enhance mammalian
proteasome expression through the Keap1-Nrf2 signaling pathway. These results led us to
hypothesize that the transient stress-adaptation, involving proteasome, that was described

55

in chapters 1 and 2 (Pickering et al., 2010), might be primarily under the control of the
Nrf2 transcription factor.

ARE/EpRE sequences are present in the upstream un-translated region of all proteasome
subunits examined. If Nrf2 is involved in our hormetic model of adaptation to oxidative
stress, we would expect to see an increase in total Nrf2 protein levels as a product of
enhanced stability following detachment from the Keap1 complex, as well as
translocation of Nrf2 from the cytosol to the nucleus, indicative of Nrf2 becoming active
as a nuclear transcription factor (Itoh et al., 1997; Itoh et al., 1999; Itoh et al., 2003). As
before we used H
2
O
2
as our adaptive oxidant and found that a mild dose of H
2
O
2
caused a
two-fold increase in cellular Nrf2 levels (Figure 4.1A); this is consistent with previous
reports of oxidative stress-related induction of Nrf2 (Itoh et al., 1999; Itoh et al., 2003;
Kwak et al., 2003; McMahon et al., 2001; Venugopal and Jaiswal, 1996). When we
blocked Nrf2 synthesis, using Nrf2 siRNA, we lost the increase in Nrf2 protein (Figure
4.1B). We next examined Nrf2 localization using immunocytochemistry, and saw a
notably stronger nuclear-localized staining of Nrf2 in H
2
O
2
treated cells compared to a
more widespread staining of all cell compartments in untreated cells (Figure 4.1C).

Having determined that Nrf2 levels were increased and that Nrf2 translocated to the
nucleus under the conditions of our cellular H
2
O
2
adaptation model, we next wanted to
determine if Nrf2 is actually required for the increased proteolytic capacity reported
previously. To examine this we blocked Nrf2 expression by two distinct methods:

56

siRNA and retinoic acid. First, we pre-treated cells with Nrf2 siRNA, as Nrf2 is
maintained at extremely low levels in unstressed cells. This served to block the oxidative
stress-induced increase in Nrf2 levels more than it served to reduce baseline levels,
however both were recorded to be reduced (Figure 4.1B and Figure 4.2A inset). Cells
pre-treated with Nrf2 siRNA and then exposed to an adaptive dose of H
2
O
2
did not
exhibit an H
2
O
2
induced increase in proteolytic capacity, but cells treated with a
scrambled siRNA vector showed a normal induction of proteolytic capacity (Figure
4.2A). As a further test of Nrf2 involvement we repeated the experiment of Figure 4.2A,
using retinoic acid treatment as a different means of blocking Nrf2, which has been
shown to prevent Nrf2 expression in cells (Wang et al., 2007). When we pre-treated cells
with retinoic acid and then attempted to adapt the cells to H
2
O
2
as in Figure 4.2A, we saw
no significant increase in Nrf2 levels and no increase in proteolytic capacity (Figure
4.2B).

While the degradation of suc-LLVY-AMC provides a good approximation of the
chymotrypsin-like activity of the proteasome, what really counts is proteasomal capacity
to degrade oxidized proteins. To examine this question, we incubated cell lysates with
tritium-labeled hemoglobin ([
3
H]Hb) and oxidized [
3
H]hemoglobin ([
3
H]Hb
ox
).
Adaptation to H
2
O
2
pre-treatment caused a two-fold increase in capacity to degrade
[
3
H]Hb, but an almost four-fold increase in selectivity for [
3
H]Hb
ox
(Figure 4.2C). In
contrast, cells pre-treated with siRNA against Nrf2, prior to H
2
O
2
treatment, exhibited no
increase in [
3
H]Hb degradation and less than a 25% increase in capacity to degrade

57

[
3
H]Hb
ox
(Figure 4.2C). The results of Figure 4.2 provide strong evidence that Nrf2 has
an important role in the increase in proteolytic capacity induced during adaptation to
oxidative stress.

58

Figure 4.1: Nrf2 protein levels and nuclear translocation during oxidative stress
adaptation
A: H
2
O
2
treatment causes an increase in whole-cell levels of Nrf2. MEF cells were grown
to 10% confluence, treated for 1 h with 0 - 100μM H
2
O
2
, and then washed and re-
suspended. Cells were harvested and lysed. Cell lysates (20μg) were run on SDS-PAGE
gels and transferred to PVDF membranes. The membranes were screened with antibodies
directed against Nrf2 and β-tubulin. All experiments were repeated in triplicate and band
intensities for Nrf2 were normalized to that of β-tubulin. Values are Means ± SE, n = 3.
B: The increase in Nrf2 band intensity is lost with Nrf2 siRNA pre-treatment. Samples
were prepared as in A except that, 4 h prior to H
2
O
2
treatment, cells were pre-treated with
siRNA against Nrf2, or with a scrambled vector, and gels were then run as in A. C:
Treatment of cells with H
2
O
2
causes Nrf2 to shift from a broad cytoplasmic distribution
to a nuclear localization. MEF cells were grown to 50% confluence and treated with
H
2
O
2
for 1 h then fixed and stained with an antibody directed against Nrf2.
Representative photographs are shown, but the experiment was repeated several times
with similar results.

59

Figure 4.2: Increased proteolytic capacity is blocked by inhibition of Nrf2

A: The increase in proteolytic capacity caused by H
2
O
2
treatment was blocked by
inhibition of Nrf2 expression through pre-treatment with Nrf2 siRNA. MEF cells were
grown to 10% confluence and treated with either control or Nrf2 siRNA for 24 h. 4 h
after initiation of siRNA treatment, half the cells were exposed to 1μM H
2
O
2
for 1 h, then
washed. Capacity to degrade the fluorogenic peptide suc-LLVY-AMC was determined
24 h after initiation of siRNA treatment. Values are Mean ± SE. n = 3 and are plotted as
nM AMC liberated per minute per mg lysate. The inset to A shows a representative
Western blot. B: Treatment of cells with the Nrf2 inhibitor retinoic acid (RA) also
blocked the H
2
O
2
induced increase in proteolytic capacity. MEF cells were seeded at 5%
confluence and treated with 3 µM Retinoic acid. When cells reached 10% confluence half
were exposed to 1μM H
2
O
2
and proteolytic capacity (Suc-LLVY-AMC lysis) was
determined 24 h after treatment as in A. Values are Mean ± SE. n = 3 and are plotted as
nM AMC liberated per minute per mg lysate. The inset to B shows a representative
western blot. C: The H
2
O
2
induced increase in selective capacity to degrade oxidized
proteins is also blocked by inhibition of Nrf2 expression. MEF cells were prepared and
lysed as described in A and B. Lysates were incubated for 4 h with [
3
H]Hb or [
3
H]Hb
ox
.
Percent protein degraded was calculated after addition of 20% TCA and 3% BSA, and
centrifugation to precipitate remaining intact proteins (Fucci et al., 1983; Pickering et al.,
2010; Shringarpure et al., 2003; Ullrich et al., 1999). Percent protein degradation was
determined by release of acid soluble counts in TCA supernatants, by liquid scintillation

60

Figure 4.2, Continued
as follows: % degradation = (acid soluble counts – background counts) / total counts x
100. Results are Means ± SE. n = 3.

61

II. Induction of proteasome under oxidative stress is Nrf2 dependent

Since oxidative stress can increase the levels of 20S Proteasome (Pickering et al., 2010),
which has been shown to have critical roles in adaptation to oxidative stress (Grune et al.,
2011; Pickering et al., 2010), we wanted to determine whether the increases in proteolytic
capacity reported in Figure 4.2 are explained by changes in proteasome, and to determine
if Nrf2 plays a critical upstream role. For these studies, we used western blot analyses of
control and H
2
O
2
adapted cells, pre-treated with either Nrf2 siRNA or a scrambled
siRNA vector. With scrambled (control) siRNA treatment we saw a two-fold H
2
O
2
induced increase in 20S Proteasome (Figure 4.3A). With Nrf2 siRNA pre-treatment,
however, the H
2
O
2
induced increase in 20S Proteasome (Figure 4.3A) was lost, indicating
that 20S Proteasome is regulated by Nrf2 during adaptation to stress. We confirmed some
of the EpRE elements upstream of all 20S proteasome subunits examined. In addition to
this we demonstrated that H
2
O
2
treatment induces binding of Nrf2 to these sequences. To
do this we performed a chromatin immunoprecipiation assay (ChIP) on an EpRE element
in the 5’-untranslated region (5’-UTR) of the proteasome subunit beta 5. This subunit has
previously been shown to have functional EpRE elements (Kwak et al., 2003). This
EpRE element showed a strong increase in NRF2 binding under H
2
O
2
exposure (Figure
4.3B). Furthermore we confirmed that there was a corresponding increase in mRNA
expression of the subunit with Nrf2 binding (Figure 4.3C).

62

As described earlier we have developed a transient oxidative stress-adaption model in
which pre-treatment of cells with a low concentration of H
2
O
2
causes changes in gene
expression that permit survival of a much higher, normally toxic, challenge dose of H
2
O
2

delivered 24 h later (Pickering et al., 2010; Wiese et al., 1995). Without pre-treatment
with a mild dose of H
2
O
2
, the challenge dose causes protein oxidation, growth arrest,
diminished DNA and protein synthesis, and some degree of apoptosis. All these measures
of toxicity are avoided or minimized if cells are adapted by pre-treatment with a mild
dose of H
2
O
2
before being exposed to the challenge dose (Davies, 1999, 2000b; Ermak et
al., 2002; Pickering et al., 2010; Wiese et al., 1995). We now wanted to test if adaptive
resistance to H
2
O
2
toxicity could be achieved by pre-treatment with a wide range of Nrf2
inducers (both oxidative and non-oxidative). As shown in Figure 4.4, 1.0 mM H
2
O
2

challenge caused a 55% decrease in cell growth. This was reduced to only a 35%
decrease with H
2
O
2
pre-treatment and adaptation (Figure 4.4). However, if cells were first
pre-treated with siRNA against Nrf2 then adaptation was severely blunted and H
2
O
2
challenge induced growth arrest returned to 67% (Figure 4.4), indicating a significant role
for Nrf2 in H
2
O
2
induced tolerance to oxidative stress.

63

Figure 4.3: H
2
O
2
induced 20S proteasome induction is Nrf2 dependent
A: H
2
O
2
induced 20S proteasome synthesis appears to depend upon Nrf2 expression.
MEF cells were prepared, treated and harvested as described in Figure 4.2. The cells were
then lysed, and samples were run on SDS-PAGE gels and transferred to PVDF
membranes as in Figure 4.1. Membranes were treated with antibodies directed against
20S Proteasome subunit β1 and β-tubulin. Graphs show the level of 20S Proteasome β1
subunit divided by β-tubulin levels for each well and then plotted as a percent of control.
Values are Means ± SE. n = 3. B: H
2
O
2
treatment causes increased binding of Nrf2 to one
of the EpRE elements upstream of the promoter of the proteasome subunit β5. Cells were
grown to 10% confluence then exposed to 1μM H
2
O
2
for 1 h. ChIP analysis was then
performed. Non-specific binding was measured through performing a ChIP assay in the
absence of the Nrf2 antibody and input was as an internal control by representing 1% of
the sample prior to immunoprecipitation. C. H
2
O
2
treatment causes increased mRNA
expression of the 20S proteasome subunit β5. Cells were grown to 10% confluence then
exposed to 1μM H
2
O
2
for 1 h. After this cells were harvested, the mRNA levels of the
20S proteasome subunit β5 and the loading control GAPDH were then determined
through reverse transcriptase PCR followed by qPCR. Values are plotted as a fold change
from control samples and are adjusted by levels of GAPDH. Values are Mean ± SE
where n = 3.

64

Figure 4.4: H₂O ₂ induced adaptation is Nrf2 dependent
The increase in tolerance to H
2
O
2
challenge induced by mild oxidant pre-treatment is lost
by blocking Nrf2 expression. MEF cells were grown, and treated with siRNA directed
against Nrf2 or a scrambled vector, then pre-treated (or not) with 1μM H
2
O
2
as described
in Figure 4.2 and 4.3. After 24 h, cells were challenged with 1 mM H
2
O
2
for 1 h, then
washed and re-suspended in fresh complete media. After another 24 h, cells were
harvested and cell counts were taken. Values are plotted as a percent of unchallenged
samples treated with scrambled siRNA, and values are Means ± SE. n = 4.

III. Adaptation through Nrf2 induction

Having gained evidence to provide a convincing argument for an important regulatory
role of Nrf2 in the adaptive response, we wanted to test if inducing Nrf2 could itself

65

induce an adaptive response. We did this using the pre-treatment challenge model (Figure
4.5) described earlier: 1.0 mM H
2
O
2
causes a 65% decrease in cell counts in non-
adapted, naïve, cells. This was mostly due to prolonged growth arrest, as previously
shown (Davies, 1999, 2000b; Ermak et al., 2002; Pickering et al., 2010; Wiese et al.,
1995). In contrast, cells that had been pre-treated with (low concentrations of) inducers of
Nrf2, including DL-sulforaphane (Kamanna et al., 1991; Mandal et al., 2009; Yang et al.,
2009), curcumin (Gan et al., 2010; Lii et al., 2010; Nair et al., 2010; Zhao et al., 2010),
and lipoic acid (Shay et al., 2009; Shenvi et al., 2009; Suh et al., 2004) had an increase in
oxidative stress tolerance. Growth-arrest, induced by H
2
O
2
challenge, was decreased
from 65% to 39% with DL-sulforaphane pre-treatment, to 31% with curcumin pre-
treatment, and to only 35% with lipoic acid pre-treatment. While it is important to note
that these agents are not exclusive inducers of Nrf2 the results provide additional support
for an important role for Nrf2 in oxidative stress adaptation.

Having observed an adaptive response with the use of multiple Nrf2 ‘inducers’, we
wanted to determine if proteasome and/or Nrf2 are involved in their adaptive response.
To test this we performed western blots on cells 24 h after pre-treatment with a range of
concentrations of the 3 Nrf2 inducers. We observed modest increases (≈40%) in 20S
Proteasome with lipoic acid and curcumin treatment, and more than a two-fold increase
with DL-sulforaphane (Figure 4.6A). To test the role of both Nrf2 and proteasome in the
adaptive response to Nrf2 ‘inducers’ we used the pre-treatment and challenge model of
Figure 4.4, with a background of scrambled siRNA, Nrf2 siRNA or 20S proteasome

66

siRNA (Figure 4.6B). With H
2
O
2
challenge of non-adapted cells there was a 68% growth-
arrest. Lipoic acid pre-treatment reduced growth arrest to ≈50%, however, growth arrest
was returned to ≈85% with either Nrf2 or 20S proteasome siRNA treatment. Similarly,
DL-sulforaphane treatment reduced growth arrest to ≈40%, which was returned to ≈85%
with either Nrf2 or 20S proteasome siRNA treatment. Curcumin treatment reduced
growth arrest to ≈40% which was restored to ≈85% with 20S Proteasome siRNA and to
70% with Nrf2 siRNA (Figure 4.6B).

67

Figure 4.5: Nrf2 inducers increase oxidative stress tolerance to oxidative stress
Pretreatment of cells with a mild non-toxic dose of a range of oxidants, and other
inducers of Nrf2, caused increased tolerance to a subsequent toxic H
2
O
2
challenge. MEF
cells were grown to 10% confluence and treated with 10pM DL-sulforaphane, 500pM
curcumin, 100 nM H
2
O
2
or 500pM lipoic acid, for 1 h, then washed and resuspended.
After 24 h, cells were challenged with 1 mM H
2
O
2
for 1 h, then washed and resuspended.
After another 24 h, cells were harvested and counted. Values are Means ± SE. n = 3.

68

Figure 4.6: Nrf2 inducers promote proteasome dependent adaptation
A: MEF cells were grown to 10% confluence and treated with 1 nM-100 nM Lipoic
acid, 1 nM-100 nM DL-sulforaphane, or 1 nM-100 nM curcumin. After 24 h, cells were
harvested, lysed, run on SDS-PAGE gels and transferred to PVDF membranes as in
Figure 4.2. Membranes were treated with antibodies directed against 20S proteasome
subunit β1. B: The adaptive response produced by Nrf2 inducers is lost or blunted by
blocking either Nrf2 or proteasome expression. MEF cells were grown, and pre-treated
with siRNA directed against Nrf2, 20S proteasome subunit β1, or a scrambled vector
then, 4 h later, treated with 500pM lipoic acid, 10pM DL-sulforaphane, or 500pM
curcumin, as described in Figure 4.5. After 24 h, cells were challenged with 1 mM H
2
O
2
for 1 h, then washed and resuspended in fresh complete media. After another 24 h, cells
were harvested and cell counts were taken. Values are plotted as a percent of
unchallenged samples treated with scrambled siRNA, and values are Means ± SE. n = 4.

69

IV. Other stress induced transcription factors

In addition to Nrf2 there are a several other oxidative stress inducible transcription
factors, including NF-κB (nuclear factor kappa-light-chain-enhancer of activated B cells).
NF-κB is induced by a range of factors including inflammatory cytokines, various
pathogenic signals, oxidative stress and UV radiation (Baeuerle and Henkel, 1994).
Under these stimuli it will translocate to the nucleus and activate a wide range of stress
response genes. NF-κB, like Nrf2, is induced by H
2
O
2
exposure (Takada et al., 2003).
Moreover, using a similar adaptive model to our own, a group demonstrated that H
2
O
2

pre-treatment was able to yield an increase in oxidative stress resistance that was partially
dependent on NF-κB (Kim et al., 2001). The group showed that this was independent of
capacity to degrade the oxidants but did not determine further how the oxidative stress
protective effect was accomplished. However, there are no reports of NF-κB having a
regulator effect on proteasome. In fact, when we performed a search for NF-κB in
binding sites upstream of 20S proteasome subunits, most subunits do not posses any
potential binding sites with only five out the fourteen subunits possessing one or two NF-
κB binding sequences. By comparison all fourteen subunits possessed at least two and in
some cases up to eight Nrf2 binding sites (Figure 4.7). As a result of this it seems
unlikely that NF-κB plays a role in the regulation of 20S proteasome in response to
oxidative stress. There is some evidence, however, that NF-κB is capable of functioning

70

in conjunction with IRF-1 in the regulation of Immunoproteasome expression
(Moschonas et al., 2008). We describe the immunoproteasome in more detail in the
subsequent chapter.

Another one of the major stress inducible transcription factors is FoxO. In mammalian
cells there are a range of different FoxO genes with semi redundant function. These are
transcription factors which activate a wide range of stress response genes in response to a
range of stresses. These genes are primarily known for their role in adaptive response to
caloric or dietary restriction, but in addition to this there have been some publications
describing the induction of one of these genes (foxo4) in response to exposure to H₂O₂
(Essers et al., 2004). Un-published experiments both by this lab and our collaborators
have failed to find any role for FoxO in oxidative stress adaptation. In addition, there are
no publications showing regulation of 20S proteasome by foxO and a search for FoxO
binding sites up-stream of 20S proteasome subunits only yielded one or two binding sites
in five of the fourteen 20S proteasome subunits, with no binding sites found in the other
nine subunits, which indicates that it is unlikely that FoxO plays a regulatory role in the
oxidative stress induced increase in 20S proteasome.

71

Figure 4.7: NF-κB/Nrf2/FoxO binding sites on proteasome promoter sequences
A search was performed for NF-κB, Nrf2, or FoxO binding sites in the 5kb upstream
region of each of the fourteen, 20S proteasome subunits.

V. Summary

Our studies reveal a mechanistic link between Nrf2, the 20S proteasome and transient
adaptation to oxidative stress. It now appears clear that the Nrf2 signal transduction
pathway plays a major role in both the increased proteasomal capacity to degrade
oxidized proteins, and the increased cellular tolerance to oxidative stress that are induced
by pre-treatment with a mild dose of oxidant.

72

Cellular levels of Nrf2 were significantly increased by adaptation to oxidative stress, and
Nrf2 was seen to translocate to the nucleus. Blocking the induction of Nrf2, with siRNA
or with retinoic acid, significantly limited the adaptive increases in cellular proteolytic
capacity. Blocking Nrf2 induction also limited the increase in oxidative stress resistance
(cell growth). When, instead of using oxidant exposure, we pre-treated cells with the Nrf2
inducers lipoic acid, curcumin, or sulforaphane, we observed increased cellular
proteolytic capacity, increased 20S proteasome, and increased cellular resistance to
oxidative stress (cell growth). Both Nrf2 siRNA and 20S Proteasome β1 subunit siRNA
effectively blocked these increases.

The Nrf2 signal transduction pathway is known to respond to stressful conditions (Itoh et
al., 1997; Itoh et al., 1999; Itoh et al., 2003; Kraft et al., 2006; Kwak et al., 2003;
McMahon et al., 2001; McMahon et al., 2003; Moi et al., 1994; Nguyen et al., 2003;
Rushmore et al., 1991; Venugopal and Jaiswal, 1996). Under non-stress conditions Nrf2
is retained in the cytoplasm through the formation of a complex with several proteins,
including Keap1. In this state it is constantly turned over through ubiquitin-dependent
26S proteasome degradation. This permits a high expression rate, enabling rapid
accumulation of Nrf2 when degradation is blocked, while ensuring low Nrf2 steady-state
levels under normal conditions. Pre-treatment with an oxidant, or other Nrf2 inducers,
liberates Nrf2 from the Keap1 complex. This also prevents further Nrf2 degradation,
resulting in a dramatic rise in Nrf2 cellular levels as well as its translocation to the

73

nucleus. Once there, it can bind to anti-oxidant response elements (ARE’s) that have also
been called electrophile response elements (EpRE’s), in a range of genes.

We find that genes encoding many 20S proteasome subunits contain at least one if not
multiple ARE/EpRE sequences in their upstream, untranslated regions and have shown
that at least some of these ARE/EpRE sequences have a strong increase in Nrf2 binding
under H
2
O
2
exposure. Nrf2 is not the only protein that can bind to ARE/EpRE sequences,
and it is certainly possible that other signal transduction proteins may bind to proteasomal
ARE/EpRE elements.

In conclusion, we find that the increase in 20S proteasome expression is largely mediated
by the Nrf2 signal transduction pathway during adaptation to oxidative stress. This Nrf2-
dependent increase in 20S proteasome is shown to be important for fully effective
hormetic increases in cellular stress resistance.
This model is represented in Figure 4.8.

74

Figure 4.8: Model for adaptive response
With H
2
O
2
exposure there is an initial protein synthesis independent response in which
the 19S regulator temporarily detaches from the 26S proteasome to increase the cellular
level of 20S proteasome. H
2
O
2
exposure also causes the Nrf2 transcription factor to
detach from the Keap-1 complex this stabilizes Nrf2 increasing its levels and enabling it
to translocate to the nucleus. Once in the nucleusNrf2 will bind to EpRE elements on 20S
proteasome subunits. This causes an increase in 20S proteasome protein synthesis.

75

CHAPTER 5: THE ROLE OF THE IMMUNOPROTEASOME

Another form of the proteasome which has been suggested to be involved in removal of
oxidized proteins is the immunoproteasome. This form of proteasome differs from 20S
proteasome by the presence of three alternative subunits: β1i, β2i, and β5i. These are
sometimes known as: lmp2, mecl-1, and lmp7 (shown in Figure 5.1). These alternative
subunits result in difference in the sites of cleavage of proteins degraded by the
immunoproteasome compared to the 20S proteasome, so producing a different range of
peptides (Belich et al., 1994; Fruh et al., 1994). The peptides generated by the
immunoproteasome, are favored as peptides for MHC class 1 antigen presentation
because of their composition and structure (Tanaka and Kasahara, 1998; Teoh and
Davies, 2004). Expression of the immunoproteasome is strongly induced by the regulator
interferon-γ. Interferon-γ is a cell signaling molecule that is important in both innate and
adaptive immune response (Fruh and Yang, 1999). This along with the localization of the
immunoproteasome to the endoplasmic reticulum has led to the belief that it functions
primarily as an enzyme for generating MHC class 1 peptides for cell surface presentation
(Tanaka and Kasahara, 1998; Teoh and Davies, 2004). However New work with the
immunoproteasome has made us re-evaluate this view and has encouraged us to more
carefully test the possible contributions of the immunoproteasome in the removal of
oxidized cellular proteins. This includes the observation that interferon-γ does itself
produce an oxidative stress response. As a result of this the interferon-γ induced increase
in immunoproteasome might actually be an oxidative stress response rather than an

76

immune response (Watanabe et al., 2003). Studies by Ferrington et al. indicate that
immunoproteasome is induced in a range of aging tissues (Ethen et al., 2007; Ferrington
et al., 2005; Ferrington et al., 2008; Husom et al., 2004). In addition a study by
Kotamraju et al shows that nitric oxide induces increased immunoproteasome expression
(Kotamraju et al., 2006). Because of these finding we decided to test the role of
immunoproteasome in oxidative stress adaptation. This chapter will primarily discuss
work published in: Pickering, A.M., Koop, A.L., Teoh, C.Y., Ermak, G., Grune, T., and
Davies, K.J.A. (2010). The immunoproteasome, the 20S proteasome and the PA28αβ
proteasome regulator are oxidative-stress-adaptive proteolytic complexes. Biochem J
432, 585-594 (Pickering et al., 2010). It will also discuss work published in: Pickering,
A.M., Linder, R.A., Zhang, H., Forman, H.J., and Davies, K.J.A. (2012). Nrf2 dependent
induction of proteasome and Pa28αβ regulator is required for adaptation to oxidative
stress. J Biol Chem. 2012 Mar 23;287,10021-31 (Pickering et al., 2012).

77

Figure 5.1 Diagram of subunit composition of the 20S proteasome vs
immunoproteasome
The 20S proteasome is composed of four rings each containing 7 subunits. The first ring
contains subunits alpha 1-7. The second ring contains subunits beta 1-7. The third ring
contains subunits beta 1-7. The fourth ring contains subunits alpha 1-7. Sunbunits β1, β2,
and β5 are the proteolytically active subunits. In the immunoproteasome the 2 proteolytic
subunits are replaced with 3 alternative subunits β1i (Lmp2), β2i (Mecl-1), and β5i
(Lmp7)

I. Oxidative stress induction of immunoproteasome

Our interest in immunoproteasome under oxidative stress conditions stems from the
PrOxI hypothesis, published previously by the Davies lab (Teoh and Davies, 2004). The
PrOxI hypothesis argues that immunoproteasome might use oxidation as a universal
marker for cell surface antigen presentation. In addition reports by Kalyanaraman et al
(Kotamraju et al., 2006; Kotamraju et al., 2003; Thomas et al., 2007) and Ding et al.

78

(Ding et al., 2003), show that immunoproteasome can be induced by both NO• and H
2
O
2
.
In addition, reports by Ferrington et al (Ding et al., 2003; Kotamraju et al., 2006) show
that injury can induce immunoproteasome expression. As a result of these studies we
hypothesized that, like 20S proteasome, the immunoproteasome might be induced in
adaptation. To test if immunoproteasome was induced during oxidative stress adaptation
we used the western blots created in Figure 3.3. In which we had seen a 2 fold increase in
4 20S proteasome subunits under H₂O₂ exposure but no change in subunits specific to the
26S proteasome. We probed these same membrane with antibodies directed against the
three unique immunoproteasome subunits β1i (or Lmp2), β2 (or Mecl-1), and β5i (or
Lmp7). Importantly, we measured a three-to-four fold increase in immunoproteasome
subunits over the 24 h after H
2
O
2
treatment (Figure 5.2, p < 0.01).

79

Figure 5.2: Expression of 20S proteasome, 26S proteasome and immunoproteasome
subunits during H ₂O ₂ adaptation
Lysate was prepared and run as described in Figure 3.3 samples were then screened with
antibodies directed against immunoproteasome subunits β1i (Lmp2), β2i (Mecl-1), β5i
(Lmp7) and plotted alongside the results from Figure 3.3. The dotted line between solid
circle symbols is the arithmetic mean of 20S proteasome α3, α4, β1, and β2 subunit level
values taken from Figure 3.3. Values are means ± SE, n = 4. These values are reported as
a percent of control (non-H
2
O
2
adapted) samples.

II. Immunoproteasome in oxidative stress adaptation

We then went on to determine if the observed inductions of immunoproteasome actually
had an effect on the oxidative stress resistance of the cells. To do this cells were pre-
treated with a mild (adaptive) dose of hydrogen peroxide and then challenged 24 h later

80

with a more severe dose. Adaptation was assessed using cell count assay and BrdU
assays as done with 20S proteasome in chapter 3.

We initially measured stress-tolerance by incorporation of BrdU in the 24 h following
challenge, in both transiently H
2
O
2
adapted (pre-treated) and non-adapted cells. This
served as a measure of DNA replication and hence an indirect measure of cell division
(Figure 5.3A). We then repeated the experiment looking instead at the increase in cell
population 24 h after a H
2
O
2
challenge (Figure 5.3B) in both transiently H
2
O
2
adapted
(pre-treated) and non-adapted cells. To test the importance of the immunoproteasome in
this adaptation in both experiments we treated cells with siRNA against the β1i subunit of
the Immunoproteasome, 48 h prior to H
2
O
2
pre-treatment. This data was then plotted
alongside the data for β1 siRNA treatment from Figure 3.5. Finally to test whether the
adaptation we observed was indeed a product of an increase in cellular capacity to
degrade oxidized proteins, we performed an oxyblot on the previously assayed samples
(Figure 5.3C).

The experiments in Figure 5.3A show that BrdU incorporation was almost 300% higher
in H
2
O
2
adapted (pre-treated) cells than in non-adapted cells following H
2
O
2
challenge.
In contrast with β1 siRNA (20S proteasome knock-down), the elevation in BrdU
incorporation (under H₂O₂ challenge) with H
2
O
2
adaption was only 80% higher. With
β1i siRNA (immunoproteasome knock-down) the elevation in BrdU incorporation (under

81

H₂O₂ challenge), with H
2
O
2
adaption, was only 60% higher. All differences quoted were
statistically significant (p < 0.01).

The results of cell survival and growth assays (cell counts) in Figure 5.3B also revealed
important roles for immunoproteasome in overall oxidative stress adaptation. We
observed a doubling of cell counts after H
2
O
2
challenge in cells that were first adapted by
pre-treatment with low levels of H
2
O
2
(Figure 5.3B). Treatment with β1 siRNA (20S
proteasome knock-down) cut the adaptive increase in cell counts down to just a 30%
increase. While treatment with β1i (immunoproteasome knock-down) actually almost
completely prevented the H
2
O
2
adaptive increase in cell counts (Figure 5.3B). All
differences quoted were statistically significant (p < 0.01). This shows that both
proteasome and immunoproteasome seem to be important components of cellular
defenses against oxidative stress and blocking their induction appears to severely
compromise overall oxidative stress adaptation.

Finally, we performed studies of protein carbonyls content using Oxyblots. These studies
demonstrated that exposure to H
2
O
2
challenge increased protein oxidation (carbonyl
content) but pre-treatment and adaptation largely protected against this increase, unless
immunoproteasome induction was blocked with β1i siRNA (Figure 5.3C). These results
were statistically significant (p < 0.01) and provide further support for the importance of
the immunoproteasome both in adaptation and in the removal of oxidatively damaged
cellular proteins.

82

Figure 5.3: Blocking the induction of 20S proteasome or immunoproteasome
inhibits adaptation in H ₂O ₂ challenged cells

A. MEF cells were grown to 20% confluence and then treated with β1, β1i (Lmp2), or
control (scrambled) siRNA for 24 h. After siRNA exposure the media was replaced with
fresh complete medium and after a further 24 h (a total of 48 h after initial siRNA
exposure), some cells were transiently adapted to oxidative stress by pre-treatment with
20 μM of H
2
O
2
per 10
6
cells, while others were not adapted. Cells were incubated at 37
o
C
under 5% CO
2
for 1 h, after which the medium was replaced. Following a 24 h adaptation
period, both adapted and non-adapted cells were challenged by incubation with 1 mM
H
2
O
2
(≈25 μmol H
2
O
2
per 10
7
cells). Cells were then harvested and reseeded at 100,000
cells per ml on 96 well plates and the BrdU assay was then performed. BrdU results
(values are means ± SE, n = 3) represent cellular BrdU incorporation into DNA in
arbitrary units. On the X-axis, “No Pre-treatment” represents samples that were treated
with control (scrambled) siRNA and challenged with high H
2
O
2
, but were not adapted by
pre-treatment with low H
2
O
2
. All other samples were first treated with siRNA’s, adapted
by pre-treatment with low H
2
O
2
, and then challenged by exposure to high (1.0mM) H
2
O
2
.
B. MEF cells were prepared, treated with siRNA’s, transiently adapted to oxidative
stress by pre-treatment with H
2
O
2
(or not pre-treated), challenged with 1 mM H
2
O
2
, and
harvested,. Samples were then seeded at a density of 100,000 cells per ml in 24 well
plates. Cells were incubated for a further 24 hours, then cell counts were taken using a
cell counter. Values (means ± SE, n = 3) represent the cell popultation in challenged cells

83

Figure 5.3, Continued
which previously were either pre-treated with an adaptive dose of H
2
O
2
or not pre-
treated. C. MEF cells were prepared, treated with siRNA, transiently adapted to
oxidative stress by pre-treatment with H
2
O
2
(or not pre-treated, challenged with 1 mM
H
2
O
2
, as in A and B. Samples were then incubated for a further six hours, harvested,
lysed, diluted based on protein content, and then assayed in a protein carbonyl assay.
Values (mean ± SE, n = 3) represent the percent increase in oxidation (overall intensity of
anti-DNP antibody staining) of H
2
O
2
challenged (1.0 mM) samples, both H
2
O
2
pre-
treated and non-pre-treated, in an oxyblot, and compares the effects of control
(scrambled) and β1i siRNA’s.

84

III. Capacity of immunoproteasome to degrade oxidized proteins

Taken together, the above results of suggest a significant role for immunoproteasome in
response to oxidative stress. It is important to note, however, that immunoproteasome
has not been shown to be able to degrade oxidized proteins; we next proceeded to test this
possibility.

First, we purified both the core 20S proteasome and the immunoproteasome from MEF
cells (after induction with interferon-gamma) and measured their ability to degrade both
the control and oxidized forms of hemoglobin and ezrin (Figure 5.4A). Additionally, to
improve our confidence in the results, we repeated the assay using 20S proteasome and
immunoproteasome purified from human erythrocytes and spleen, respectively (Figure
5.4B). The purity of erythrocyte 20S proteasome (positive for β5 subunit but not for β5i)
and of spleen immunoproteasome (positive for β5i subunit but not for β5) can be readily
seen in the Western blot insert to Figure 5.4B (in which both proteasome forms are
appropriately positive for the α3 subunit, while only 20S proteasome is positive for β5
and only immunoproteasome is positive for β5i). Oxidized hemoglobin has been widely
used as a model proteolytic substrate (Davies and Goldberg, 1987b; Pacifici et al., 1993)
and ezrin undergoes substantial oxidation and proteasome-dependent degradation
following exposure of cells to oxidants (Pacifici and Davies, 1990). Our results show
that the immunoproteasome selectively degrades the oxidized forms of proteins. In
addition we see that the immunoproteasome is at least as efficient at degrading oxidized

85

proteins as is the 20S proteasome. (Figures. 5.4A and 5.4B, in which both proteasome
and immunoproteasome degraded the oxidized proteins significantly better than the non-
oxidized proteins: p < 0.01 in all cases by students t-test).

Figure 5.4: Ability of the immunoproteasome to degrade oxidized proteins
A. 20S proteasome was isolated from MEF cells and immunoproteasome was isolated
after 2 days of cell treatment with IFNγ, using the methods previously described by
Tanaka et al. (Tanahashi et al., 2000; Tanakaa. Keiji., 2006). Purified 20S proteasome or
purified immunoproteasome was then incubated for 60 minutes with [
3
H]Hb, [
3
H]ezrin,
[
3
H]Hb
ox
, or [
3
H]ezrin
ox
. The percent of protein substrate degraded was calculated, after
addition of 20% trichloroacetic acid and 3% BSA to precipitate remaining intact proteins
(6, 8-10). Percent protein degraded was determined by release of acid soluble counts in
TCA supernatants using liquid scintilation in which % Degradation = (acid-soluble
counts – background counts) x 100. Values are means ± SE, n = 3. B. 20S proteasome
purified from human erythrocytes, and Immunoproteasome purified from human spleen
were studied in a similar way to the assay performed in Figure 6A. Values are means ±
SE, n = 3. The inset represent samples of equal quantity of either 20S proteasome or
immunoproteasome screened by Western blot, with antibodies directed against β5, β5i
(Lmp7) or α3 subunits, and demonstrates the purity of the 20S and immunoproteasome
preparations.

86

VI. Induction of immunoproteasome is Nrf2 independent

We have shown that oxidative stress adaptation can increase the levels of
immunoproteasome (Ferrington et al., 2005; Ferrington et al., 2008; Pickering et al.,
2010), which we show to have a critical role in the adaptive increase in oxidative stress
tolerance (Pickering et al., 2010). We next wanted to determine whether the increase in
immunoproteasome expression is a product of regulation by the nrf2 transcription factor.
For this study we used western blot analyses of control and H
2
O
2
adapted cells, pre-
treated with either nrf2 siRNA or a scrambled siRNA vector. With scrambled (control)
siRNA treatment we saw a two- to three-fold H
2
O
2
induced increase in
immunoproteasome (Figure 5.5A). Nrf2 siRNA treatment had only a weak effect on the
H
2
O
2
induced increase in immunoproteasome levels (Figure 5.5). Thus, we must
conclude that the immunoproteasome regulation during oxidative stress is either wholly
or partially independent of nrf2, and other factor(s) must be involved. In support of this
idea, we find that, although most 20S proteasome subunits contain a few ARE/EpRE
sequences in their promoter regions, ARE/EpRE sequences are completely absent in two
of the three immunoproteasome-specific subunits (Figure 5.6).

87

Figure 5.5: Expression of immunoproteasome is Nrf2 independent
The increase in immunoproteasome with H
2
O
2
treatment may be only partly nrf2
dependent. MEF cells were grown to 10% confluence and treated with either control or
nrf2 siRNA for 24 h. 4 h after initiation of siRNA treatment, half the cells were exposed
to 1μM H
2
O
2
for 1 h, and then washed. The cells were then lyzed, and samples were run
on SDS-PAGE gels and transferred to PVDF membranes. Membranes were treated with
antibodies directed against immunoproteasome subunit β1i (LMP2) and β-tubulin. The
graph shows the level of immunoproteasome β1i or LMP2 divided by β-tubulin levels for
each well and then plotted as a percent of control. Values are Means ± SE. n = 3.

88

20S Proteasome β2
CATGTAGGGCATGTGGATCAGGAATTTAATTCTAGCACTGATGAGTTAGAGGAAGGCAGA
TTTTTGTGAATTTGAAGTTAGTCTGGTCAACACAATTAAATTAGTTCCAGGACAACCAAGG
CTACATAGTAGTAATACCCCATCTCTCAAAAAAGGCCGGGTGGTGGTGTCACAATACATTA
ATCCCAGCACTCAGAAAGCAGAGGCAGGTGGATCTCTGAGTTCGAGGTCAGCCTAGTCTA
TAAGAGTGAGTTTCAGAGTAGCCAAGAAATCCTGTCGGGACAAATCAAAATATAAAAAAT
AAGTAAATAAAAGTGAACATTTAAAAATTTGTTGGGATGAGGAACTTTGGGGAGGGGGGC
AATGGCTGGAGTGTAAATAAATACTTTTTTTTCCCATTTTTATTAGGTATTTAGCTCATTTA
CATTAATAAATACATTTTTAAAATAATATTTTAAATGTAATGAAACAGAAGTATAGTAAAA
TATTTAAATTTAAAATTTTTTTTAAAAATTATTTGTATTGGGGATGAACGCGCTAGTGTGA
TGGTCAGAGGAAAACTTTGGGGAGTCTGTTCTTTCTACCGTGGGTTCGAGGGACAGGCTTG
ACTAACGTGCTTTCTGCCTCTGTCTATCCAGTACTAGGATTTCAAGAGGGAACCACCGCCA
AGTAGGTTTTTTGTTTGTTTGTTTGTTTTGCTTTTGAGATGTGTTGTCCTAGGCTGCTCCCCT
GGTCCTTAACTCTATGGCTCGGGGAAATTGTTCTTCAATCTGGCTTGCAGGGAAAATTTTCC
CGTTTGTTTAACGAGCAAGGAAATGAAAACCAGAAAGCTGAAGACCTCCTCTCCTCTGTAG
CACCCACACGTGCTGGTACTTCTTACACAGGGTACGGCTTGATTTGCGTGCAAACTCTAAA
ATTCGACTTGGTAGATAAGGAAACGCAACGCCCCATCACCCACAGCTACGTTTCCAAACC
ACACCTCTAGACCAGCCCACTCCGCA
20S Proteasome β3
TGACTGTTTCCATCCACCGGGACAGATTTCCTCTTTCAAGTTGCTATCGTCCACTGATGCAG
ACAATGGGAAGCTGTCCAACCTCCGCAGCAAAATTTATTCTTAATCCAAGGAAAATTGCAT
TGTTAAACATGGCAGTGAGACAGAAACTTGGCTTTGCCTTTCTGTCATCCTGATTGGGAGA
AAGAATAGTGAGAGGTGGTGAGGAGACGTTAGGGATAGATCCATTCACTGACCATAGACG
GATTTCTAAATACAGTGACCATCTGGGTTGGTGTATCACAATTATTGCTTCTGGTACCTCGG
TACCGAATGTGGCTTTGGAGCATTTGCGATTTAAAAAAAAAAAAAAAAGGCCGAGACCGT
CCCTGGGTGGGCTCGAACCACCAACCTTTCGGTTAACAGCCGAACGCGCTAACCGATTGCG
CCACAGAGACGGTGACAGTTGTTTGTGAGGCAGAGAGCCGAGTCAATAGTACTGAGCCG
ATATGGCGCAAGACCAGTCACGACGCAAGCAGTTACAGTAGCTTAGAAACCCAGCGGAAC
AAAATCTGATCTCGAAGAACAGCGAATCGAGGCTGTGGCGGAGGGAACAGTGGTTGTGCA
AGGGAAGGGAAAGCAAGGGATGCACACGTCCCACCTGGGCTGCCCTGGAGCCGTTGGCCC
AGGCTATCCTCGGTCAGTGTCTAGGATGTGGCACAGAACGTTCCTTTGTAGAGTGAACAGA
GACTTAGGGCTACGTCTGTCGGCCAGCAGGGATTGGGAGTGAGAGACTCCAAACGCTGAA
CAACCTGTAGCTCACTCCCGTAAAACAAAACAAAAGCTCCATTCTGGAACTAAACCCTCCT
AGCACGAACCCTCGGCACGATCTTCCCATTCTGCAAAGAATGGGCGCCTTCGGTATACTCG
GTGCCGATAGGTTAATTGGCGCCTGCAAAAATCCCGGTGAACAGTAAGGATTTGTGAATG
AACACTGGAAGTCGGGTAGCTTGAGTAAAATG
20S Proteasome β6
CCCTTCAGAGACCTGTCACCCAACATTAGAGTCTGCAAGGTCAACATTCCAAGCCATTAG
TGACTCGGAAACAGTTCTCCTCACCTGGCAGGACAAATGTTAAGCATCTGCCCATGTCTAC
TACTATGCTCAATTCTACACACAGAGAGACCGAGTCTGTATCTCAGCTGGGGAAGCAGGG
CATAGGTAGTGTTTGATGGACACAAGAGACTCAGCTGGGGAACATGAAAACATTCTGAAG
GATGGTGACAGAATTTATTCAATACCACAAAACTGTACTTTTAAAAATAGATAAATGTGCC
AGGCAATGATGGTGAATGCCTTTAATCCCAGTACTCAGGAAGCCAGCCTTCTCTACAAAGT
GGGTACCAGGACAGGTGAGGCTACAGAAACCCTGTTTCAAAGAGTCAAAGGTCAAAACAA
AAACAAATAAAAAAGCTAAATATAACTGCAATATTATATATTTTTTAAAGGTTTGGAGAAA
TGGCCCAGTGGTTAAGTCTGGTAGAGGGCCCAGGTTTTAGTTCCTAGCATCCACATGGAAG
CTCAGAACCACCTGTAATTCCAGTTTCAAAATCTCCAATCTCTTCTTCACACTTCTATGCAC
ATCAGGCATACATAGTACACACACATACATGCAGGCAAACACTCATACACATGATTATCGT
AGCTCATTCATAAATGTAAAATAAAAATATAAAAATAAATCTTTTTTTTTTAAGGCACAGA
CCTAGAGATGCGAACCCTGCGCTTTTGGAGGGAGAAACCATTTTCACTCTGGCTGTCTCTA
ATCCATTAAATGAGCCATACTGAAAAGCTACTTCAGACTTTACAAGTCCTGGGCAAACCT
ACACAATTCATTTGCCACAGGTCACAGAAAGTTTACACATTTGCAACTGAAACAAGCAGA
AAACTGTGTAAGGTGCTGGCCACACTGCTTTCCTGCCCTTTCCTCAGTGTCCACCCCCATAC
TCCCTGCCTCCAACCGCAGGCACTTAACA
20S Proteasome α1
TCTTCAGACTCCCAAAAAAGGGTATGCAGGGACACAGACACACACACAAAACAAATGTAA
AAACAAAAAATGTACATATATATACAGACATGAAAATATTTAGAATGTGATACTAATTACT
GTGAACATAGACATTTAAATGATGTGTTACTCCGCCATCAAAGTTCAGATCCTTAAAGATC
CTTACCTGACTCTGGGTACACGTTATTCAACAGAATTAGCAGTCTTGATTTCATTTGGGGAT
TAAACCAAAACATACTCTTTTAAAACTCGTAAGAAAACTCGCACCTTCCCCCTAGTGGCCT
AATCTACCAGATACGGTATTTGGTTGTGGACAGCAGAAGATCAAGTGCCAGGACAGAAAC
CAGGTCCTACAATAGCAACTAATGATCATAAGTATCACGATCATGCGAGACCATGGAGCC
TTTAAGGATGGGCTCTCCAAGTACTCAAGCATCTCAGGCCAGGAATATGAACACACACAC
ACACACACACACACACACACACACTTGTTTTTTTTTTTTTTTTTTCTGCAGCCTCTTTCAGAC
ACGCTGTTTCCCAAACCTCTCTGTGCACTGTGACAGATCTATGAGGGACCACCGCACCATC
CCGAGCTAGGTGGCGACGTTCAACTGAGAGGACCGAGAGTTGGCGGGAATCTGCAATCCA
CTTGGGGATTAAAAAAAAAAAAAGCAAGGAAACTCTGGGCGTGACGTGAAGCCACGCCCC
CCGCTTTCTCTGCGCATTGGCTTTCCCACCCCGGAGCTTCCTGCCAATCCCACGTTCTCAGC
TTTCGCAGGCCCCGCCCACTTAAGGCTGGAGGGCGGGGCAGCGTTAGACGCCGATCCGTG
GCTCTGGGATAGATTGCACTTCCTGCGTCGCCGCCGTCAAGACTCCGCAGGCTTCTTTGAC
GATCACCGAACCGTAGTTGGGGCCGTGTTCCTGTGTCCGCTGCTGGGTCAGGTTGTCGCTG
CGCCGGAGCTATGGTGTGTTGAGCCCCG
20S Proteasome α2
CCAAGTAGCCTTGGCTTCTGTTGCTTATGTTCTTATGCTTGCCTCTTGATATCTGTTTATCTC
TGGTGTTTGCTGGACTTGCTGTCTCAAACTGGAACTTGTCCCTCCTGTGAGACTGGGAGCC
TGGGAGGGTCAGCAGTCCTGGGGTTCAAGCTGTTAGCTTGGGTGTGTCAATAGTCCTGGGG
GACAAGCTTCCCCTGGGCAGGATTGGGGTATGGAGGGCTGTAGCATACCCCAATGCACTC
AGCTCCCATGCAATGATGAAAACTAGATTGTGTTCTTGGCTGCACTGAGGTTCTTTTGTCCA
CTGGACCCCAGGCAGGTCCAACTTTGACCAGTTATTAGGGCGGTAGTGGTGGTCTCACCTG
TGATCTTGGGTGTGTCAACACTTTTGGAGGACAAGCTTTCTCTGGGTGGGCTTGGGGTGTG
GAGGGCTGTTGTACAGGGTCAGCTCCTGGGCACAGATCAAAACTGGAAGTATTGTGTTCCT
GGCCGTATGGAGGTTCATGTGTCCACTGGGCTGGATAACTTTAAACTTTAATATATACTAT
ATATTATATAATATAACCTGTTATTCACATATGTAATGTACTGTATGTAAATATATTATATA
TTATATTATATATTATATATTATATATTATATATTATATATAAAATCACATTATATATGATA
AATATGGAAACTTTTACAGTAGTAGGCTGCACATGGATATTTCAAAAAGTCTTTATTTTTTT
TTAATCCCACAATCCTTCTCTTGCCCTCTTTTATTGCCTCTACCATTTAATTCTTCCTGATTA
TGTCCAACTATGTGGGACTATGAAAGGCAGTCTGTTTTCTGGTTGAGCTAGGTTTAAACCC
CAGAGACCCGGCAGGTGTTCCTGCAAGAAATGGAAGGTGTTTGCCATGCCCCAAGACATC
AGGCTCCTGACACAAGTCCTTCACTCTCCATAATTCTGTGACCATCACTCACATAGGGCAA
TACCCTAAGCCCCTGGTAT
20S Proteasome α3
CACGTGGTCAGTTTTCAGGAAATTTAGTAATAATTCTGATTAAGGGCATTTCTTTGGAAGA
TCATCTTAGTTCTCAGTGTTCTTATGTCAAAGACAAGGAGTAAGATTTTAATGAATTGCTA
ACTTAAGACCAGAGAGCAAGTGACAGGTAGAGTAAGTCAGGGAATCTCATTTCACACATA
TATACGTGCTAGGAGTTCATGGGGATGTACCTCAGTTGGTATGATTATTGCTAGATATGTA
TAATAAAGGCTAGATATGTACAAGATTCAACCCTAGCACTCCAAATAGGAGGTGTGGTAG
TACCTGCCTGTAATCCCAGCATATAGAAGGAGGCTAAACAGAGTTAATGGTTGGCTTGGGT
TAAGTGATACACCCTGTAAAAAATATGGCCATTTTGTGTGTGTGTGTGTGTGAATAAATAC
TATTGTATTTAGAGTTAGCCACATGGCAGAGAGTGTAGTAGCTCTTCCACAGATGGGATTT
TGAAGGAATGTTTTTCTACTATCTGTCGTGACTCATGTGTTTCTTTTGTGGCAAGACTCCTG
GTCTTTAGGTACTAGGAAATCATCTTTGGCTCTTGTGTACTAGGGTTTAGTTCTAGTTAGGT
TGACACCATTGAGCAGGGTAGAGTAGCCACTTTGACTGAGGCTCACTCAGGACATGACCT
GGAGGTTTCTCATGTGTTGTGAGGGTTTTGGTTTGTTTGTTAATTCCTGTCTGTGGTGCTGG
AGGTGGAATGCAGGCTTCACCCACACTGGGTACATGCTCTGCCACCAAGTCACATCCCAGA
GGGGAGAGATTTAATCTCAAGCTGCTTCTGAAAGGCTGGATCCAGGGAATATAGTAGGGA
AAGGTTTGTCTTCCTCAGGTCAGGAGCCCTCCCAGGGGCAGGAGCCCTTCCCTACCTGCCC
TGCCTCCTGCCTCCACCTACCTTCTATTTCACTGCCGTTTCCTGTTCTATATTGATCCTGTCC
TCAGAGTTCCTCTGCTCCATCTCC
20S Proteasome β5i (Lmp7)
GTTCAGATTCTTTCATCTTCTCCTCAGCGGCACACGCCGGGATAATCTCAATGAAATGGAT
AGGATTAGCCAGTAAATCTCTTCCATTCAGGGAGGGCCCGGTGTGTGACGTCAGCAGTTCC
TGGGCAGATAATGTCCATAAAGGTTGCAAAGAAGAGGTGGGGTTGAGGATTCAAGGAAGG
AGCGCGCAGCCCTGACTTGCGCGACGTGAGCAGCCCCGGGCAAGGGCGCGTCCTCGTGGG
TGGCTCAGGGTAGTGAGACTGTTGAACCAAGAGGGGCCCTAGGAGGGACACATGAAAAG
GCCCGACTGAGTCCTCTCCTTGGTTCGTCTTTCTCCGTTTTTTCCTGGTTGTCCCAGGACCT
AAAGACCCCTGTGCATCCATTTATCGCATTTTCCAGTTTCTATAAAATGATGCTCATTTATT
TGAGTGCTGCCCCCATCCCCAGGACGCACGCCTAGGTGGATCTGCTCAATACTCCTCTCAG
GTTCCAACGTTCTAGACAACCTCTTTAGGAGATAGGACCCTATTCAGTCCTCCGCTGTTCCC
AGAAGATGCCCCCCCCCGTCCCCCGCCTGTGGGTCTTCACGCACCCCACTGCTAGGCGGCC
TCCGAAGGCGCACCTCTTCTTCCACCCCACAGCCCATTCTCTAGCTGGTTTCACTGATGTG
GCAGCAGGAAAACTCGTCCCCAACACATTGTTTTAGTTCAGACACCCATTAACTGGGCTCT
ACTCGGAAAGGGCAGGGGATGGGGGTGGGGGACTGGTTCCCTCCCCTTGGTACTGTGGCT
TTCGCTTTCACTTCCTTGTCGCAGAGTCGACGGATCTTCGGGGCCAAGTGGTCATGGCGTT
ACTGGATCTGTGCGGTGCCGCTCGGGGGCAGCGGCCCGAGTGGGCTGCCCTGGATGCGGG
AAGCGGGGGTCGCTCGGACCCGGGACACTACAGTTTCTCCGCGCAAGCTCCGGAGCTCGC
ACTTCCCCGGGGAATGCAGGTGAGGACTG
20S Proteasome β1i (Lmp2)
CCCTTCTCGGGCCTGGGTCCTTCTCTTTCCGCCTAAGTTGCCTCCTTAGTGTCTCTGACGCTT
CCCGGGACTACTTTACTCTCCCTTTCTGCCTGGAATGCCCATTGCCCTGTCCCCTGTGGCCT
GCTCTTTCACCCAGTCTCCCTTACCAAGGCAAGAGAGAATCAAGAGAACCAGAACCAGGT
AGAGACGTCTCTTCTTAGGGCCCAGGAAGCCCAGCATCCGACACAGCATGTCTCCAGCGTC
CCTGTTGCCGCTGGGCGCCCAGAGGCTCCCCAACTTGTGCCACAGGGCGGCTGCGGGCAAT
GCTGCCACATAACTGATAGCGAAGGCATCTGGACGACTGTTCCAGTACAGTAATCCAGCG
CTGTCACCCTCCCGGAGTGTTCCCCAGGCGGCCAGCTCTCGGAACAAGGCAAGTCCAGGC
AGGGCCAAACTCAGTGCGGCCACCAGCGGCTGCAAAGCAGCCAGCCAGCCATGGGCTCCT
GCGGTGACCCCGAGGACCCCGCGGACCCCTAGTCCTAGGATGGCCCAGCGACTCAGGCCC
ACCACCCAGACCCGGAGCAGCGGCACCTCGGGAACCAACAGGGAGAAGATTCCCGGGAG
CATGGGCCGCAGCAGCAGCCAGTCCACCAGAAGGAGCAGGGCGGCCGCCAGCCAGACGT
GCGCAGCCATCGAGCGTGAGCTGTCCAGAGTCTGGTCCTAGCCTGGGACTCTCGACGTCCG
CGCTGCACAGGAGTCTCCGTGGGGAAGGAAGAAGGGCGGGTCCGAGGGCGTGCGCGTAA
AGTCCCGGGAACCCGCTCGAGCTGGTGGAGCTGACTAGAAGTGCTGGCGTTTAGAGGAAG
AAGAAACCGAAAGCCGACCTCGAATCACTAGACACGCCTCCTTCTGAGACTGGCTTGCTCA
GGCTGTTCTGGAAGCTGCCGCTGAAGCTGTAGTTGGAGTTCTGCCTAGCAGCCCTGGCCTG
TCGTGTTCTTCTCCATGAATCTCATTCTCCTCT
20S Proteasome β2i (Mecl-1)
GATAACCTGCATCACCTGGCTGATGCATGCTTGATCTGGGCTTAGGGGAACACGGGGTTCA
GTGGGCTTTGCTAAGGCCCCAGACTGGACTGACTTTGAGATATGTAAATAGATTTTCATTG
TCCTAGTTGAGCTGCTCAGGAAGCTTGCTCTGTGGCTTCCTTCTTAGGGGCCCAGGACAAT
ATTCAGATCTGGGTATATGGCTGTTAGAATGGGGACACAGAGAAATAGGATTTGAAGTCG
ACTTTTGAGTTCTGGTTGGATACAGAAATTTATAATAAAGCATGGTGGTGCACGCCTTCAA
CTCAACACTTGGGAGGCAGAGGCAGGCAGATCTCTGTAGGCCACTCTGATACACACAGTT
CCAGGACAGCCAAGACCACAGAGATTCTGTCCCCAAAACACAAACAAATAAATAGAATTT
AAGATATTGGGGCTGGGGATTGTTAATGTAAGAACTTGGATTCCATGACAGCCGGAGCTAT
ACAGAGAAAACCTGTCTCGAAAAAACAAACAAATAAATGGGGCTAGAGAGATGGCTCAG
CGTCTGTTGAAAGAAGCTGTTTTTCCAGAGGACCCTGGTTCAATACCCAGCACCCACATGG
CTGCTCACAAGTGTCTGTAACTCCAGTTTTAGGGGAGCTGACACCTCACACAGACATGCAG
GCAAAACACCAACGCACATAAAATAAGCCATAAAAAAATTAAAAAATTAGGGCCACGTG
GTGGTACACATGCCTTTAATCTTCTCATCCAGGCCAACCTTGTCTACAGAGCTTATTCCAGG
GCAGACAGGGCTACACTCAACTACACTGAAAAACAGTTTTTCAAGCGCCCCCCCCCCCCAA
AAAAAAAGTTTAGGATCTGAAGAAAAGACAGTTCATCTAAAGAAGATTTAATGTGAGACA
TTAGAGAGAGAGAGAGAGCAAGCACTCAGAAGTTAAGAGCATTTTTTTGCACTTGGAGAG
GACCTGAGTTTGGTTCCCAACGCCCACGTGGTG
Figure 5.6: ARE/EpRE binding sites upstream of 20S and immunoproteasome
subunits ARE/EpRE Consensus Sequence (TGANNNNGC / GCNNNNTCA) is present
upstream of all 20S proteasome subunits examined but only one of the
immunoproteasome specific subunits. Data represents sequences 1kb upstream of
promoter based on NCBI data base of Mus musculus. ARE/EpRE sequences are
highlighted.

89

V. Summary

The immunoproteasome has long been considered as a proteasome variant that generates
peptides for MHC Class I processing. Although we had previously suggested that
oxidation might be a common protein modification that the immunoproteasome might
recognize, and although recent data show that immunoproteasome can be induced by
oxidative stress (34-40), there has been no direct demonstration that immunoproteasome
can truly degrade oxidized proteins, until now. Our current data may even indicate
(although more detailed studies are needed) that the immunoproteasome is actually
slightly more efficient than is 20S proteasome, in recognizing the oxidatively modified
form of protein substrates such as hemoglobin and ezrin. We now suggest that the PrOxI
hypothesis (42), which proposes that some fraction of all intracellular proteins undergoes
oxidation with subsequent processing for MHC Class I by immunoproteasome, now
deserves much greater scrutiny and serious testing.

Although induction of immunoproteasome synthesis during oxidative stress adaptation is
certainly interesting, the important question is whether such induced proteolytic
capacities actually contribute to the increased oxidative stress tolerance of adapted cells.
Our data reveal that the increased capacity of adapted cells to withstand a high H
2
O
2
challenge is, at least, partly dependent upon immunoproteasome induction. This finding
demonstrates the importance of 20S immunoproteasome in adaptation to oxidative stress.
However It’s induction , unlike the 20S proteasome, appears to be primarily independent

90

of Nrf2. As suggested in the previous chapter it is potentially regulated by the irf-1
transcription factor which is also induced under oxidative stress and has binding
sequences upstream of all immunoproteasome subunits.

91

CHAPTER 6: THE ROLE OF 20S PROTEASOME REGULATORS

In addition to the 20S proteasome, the 26S proteasome, and the immunoproteasome there
are a number of regulatory complexes of proteasome which bind to the 20S proteasome
core and enhance proteolytic activity. These regulators are the Pa28 αβ regulator (Ahn et
al., 1996; Gray et al., 1994), the Pa28γ regulator (also known as ki antigen) (Wojcik,
1999), and the Pa200 regulator (Ustrell et al., 2002). The function of these regulators is
largely unknown. Given that all of these regulators have been shown to enhance
proteasomeal activity we speculated that they might have a role in adaptation to oxidative
stress. In this chapter we investigate the roles of the three proteasome regulators (Pa28αβ,
Pa28γ, and Pa200) in degradation of oxidized proteins and in the process of adaptation.
From this we provide evidence of a highly complex set of interactions within the cell.
Such interactions occur between multiple forms of proteasome with a variety of
regulators. We go on to show that these regulators control a range of different proteasome
functions in different parts of the cell. This chapter is primarily composed of un-
published work, however some of the figures are published in: Pickering, A.M., Koop,
A.L., Teoh, C.Y., Ermak, G., Grune, T., and Davies, K.J.A. (2010). The
immunoproteasome, the 20S proteasome and the PA28αβ proteasome regulator are
oxidative-stress-adaptive proteolytic complexes. Biochem J 432, 585-594 (Pickering et
al., 2010). As well as: Pickering, A.M., Linder, R.A., Zhang, H., Forman, H.J., and

92

Davies, K.J.A. (2012). Nrf2 dependent induction of proteasome and Pa28αβ regulator is
required for adaptation to oxidative stress. J Biol Chem. 2012 Mar 23;287,10021-31
(Pickering et al., 2012).

I. Pa28 αβ

The proteasome regulator Pa28αβ (also known as REGαβ, 11Sαβ or PSME1 and
PSME2), is an activator of 20S proteasome activity. Pa28αβ is localized primarily in the
cytoplasm and forms a heteroheptameric ring composed of three alpha subunits and four
beta subunits (Zhang et al., 1999). This ring can fit on either end of the core of the 20S
proteasome. When associated with the 20S proteasome it induces a large increase in the
capacity to degrade short peptides (Li and Rechsteiner, 2001). There is also a strong
suggestion for a potential role for Pa28αβ in MHC class I antigen presentation. Like the
immunoproteasome Pa28αβ is regulated by interferon-γ (Fabunmi et al., 2001). This has
led to some suggestion that the two might act together with Pa28αβ forming a cap on
immunoproteasome to enable or assist MHC class I antigen presentation of self and non-
self antigens (Teoh and Davies, 2004). Additionally, it has been shown that increases in
Pa28αβ expression results in enhanced recognition of viral antigens by cytotoxic T-cells
(Groettrup et al., 1996).

93

Induction of Pa28 αβ under oxidative stress

Because of the role of Pa28 αβ as a promoter of proteolytic activity we hypothesized that
it might be involved in the oxidative stress response. To test this we wanted to see if like
20S proteasome it was induced under oxidative stress. This was done using the western
blots created in Figure 3.3. In which we saw a two fold increase in four 20S proteasome
subunits as well as a three to four fold increase in all three immunoproteasome subunits
under H₂O₂ exposure but no change in subunits specific to the 26S proteasome. We
probed these same membranes with antibodies directed against the two Pa28 regulator
subunits (Pa28 α and Pa28β). Importantly, we measured a five-to-six fold increase in the
levels of both of these subunits (Figure 6.1, p < 0.01).

94

Figure 6.1: Expression of proteasome, immunoproteasome and Pa28 αβ with H ₂O ₂
treatment
Lysate was prepared and run as described in Figure 3.3 samples were then screened with
antibodies directed against the two Pa28 αβ subunits which were plotted alongside the
results from Figure 5.2. The dotted line between solid circle symbols is the arithmetic
mean of 20S proteasome α3, α4, β1, and β2 subunit level values taken from Figure 3.3.
Values for percent change in subunit levels are means ± SE, n = 4, values are reported as
percent of control (non-H
2
O
2
adapted samples).

95

Role of Pa28αβ in oxidative stress adaptation

Having seen that Pa28αβ appeared to be induced under oxidative stress we went on to ask
if it played a role in the process of oxidative stress adaption. So to examine the
importance of the Pa28αβ regulator, we compared degradation of oxidized ezrin
([
3
H]ezrin
ox
) in lysates prepared from wild-type MEF cells and from Pa28αβγ knockout
MEF cells. Lysates from both wild-type and Pa28αβγ knockout cells were studied with
and without H
2
O
2
pre-treatment. In wild-type MEF cell lysates we observed a 15-fold
increase in ezrin degradation after 24 h, whereas lysates from the Pa28αβγ knockout cells
exhibited only an 8-fold increase (Figure 6.2). These data (especially when considered
with the results of Figure 6.1) indicate that while the Pa28αβ regulator may not be crucial
for the H
2
O
2
induced increase in proteolytic capacity, it does seem to play an important
role.

Next we wished to determine if the observed induction in Pa28αβ was actually relevant to
the adaptive increase in oxidative stress tolerance. To do this we pre-treated cells with a
mild (adaptive) dose of H₂O₂ and then challenged the cells 24 h later with a severe
(normally lethal) dose (Pickering et al., 2010; Wiese et al., 1995) as was done in Figure
5.3 and Figure 3.5. We initially measured stress-tolerance by incorporation of BrdU in
the 24 h following challenge, in both transiently H
2
O
2
adapted (pre-treated) and non-
adapted cells. This served as a measure of DNA replication and hence an indirect
measure of cell division (Figure 6.3A). We then repeated the experiment looking instead

96

at the increase in cell population 24 h after a H₂O₂ challenge (Figure 6.3B) in both
transiently H
2
O
2
adapted (pre-treated) and non-adapted cells. To test the importance of
the Pa28 αβ regulator in this adaptation we treated the cells with siRNA treatment against
the Pa28α regulator subunit 48 h prior to H
2
O
2
pre-treatment. This served to block the
H
2
O
2
pre-treatment induced increase in Pa28αβ. These experiments showed that in
control siRNA treated samples BrdU incorporation was 140% higher in H
2
O
2
adapted
(pre-treated) cells, than in non-adapted cells, following H
2
O
2
challenge. In contrast, in
Pa28α siRNA treated cells the elevated BrdU incorporation of H
2
O
2
adapted (and then
challenged) cells was only 50% higher. The results of cell survival and growth assays
(cell counts) also revealed an important role for Pa28αβ in overall oxidative stress
adaptation. We observed a 90% increase in cell counts after H
2
O
2
challenge when cells
were first adapted by pre-treatment with low levels of H
2
O
2
. Treatment with Pa28α
siRNA actually almost completely prevented the H
2
O
2
adaptive increase in cell counts.

97

Figure 6.2: Proteolytic capacity of wild-type & Pa28αβγ knockout MEF
Wild type MEF and Pa28αβγ knock-out cells developed by Yamano et al. (Yamano et
al., 2002) were transiently adapted to oxidative stress by pre-treatment with 40 μM H
2
O
2

per 10
6
cells/ml for 1 h then washed. 24 h later control H
2
O
2
adapted wild-type MEF and
Pa28αβγ knockout MEF cells were then harvested and lysed as described in chapter 3.
Lysates was then incubated for 1 h with [
3
H]ezrin
ox
. The percent protein substrate
degraded was calculated, after addition of 20% trichloroacetic acid and 3% BSA to
precipitate remaining intact proteins (Fucci et al., 1983; Shringarpure et al., 2003; Ullrich
et al., 1999). Percent protein degraded was determined by release of acid soluble counts
in TCA supernatants using liquid scintilation in which % Degradation = (acid-soluble
counts – background counts) x 100. Values are means ± SE, n = 3.

98

Figure 6.3: Blocking the induction of 20S proteasome or immunoproteasome
inhibits adaptation in H ₂O ₂ challenged cells

A. MEF cells were grown to 20% confluence and then treated with β1, β1i (Lmp2),
Pa28 α or control (scrambled) siRNA for 24 h. After siRNA exposure the media was
replaced with fresh complete medium and after a further 24 h (a total of 48 h after initial
siRNA exposure). Some cells were transiently adapted to oxidative stress by pre-
treatment with 20 μM of H
2
O
2
per 10
6
cells/ml, while others were not adapted. Cells were
incubated at 37
o
C under 5% CO
2
for 1 h, after which the medium was replaced.
Following a 24 h adaptation period, both adapted and non-adapted cells were challenged
by incubation with 1 mM H
2
O
2
. Cells were then harvested and reseeded at 100,000 cells
per ml on 96 well plates and the BrdU assay was then performed. BrdU results (values
are means ± SE, n = 3) represent cellular BrdU incorporation into DNA in arbitrary units.
On the X-axis, “No Pre-treatment” represents samples that were treated with control
(scrambled) siRNA and challenged with high H
2
O
2
, but were not adapted by pre-
treatment with low H
2
O
2
. All other samples were first treated with siRNA’s, adapted by
pre-treatment with low H
2
O
2
, and then challenged by exposure to high (1.0mM) H
2
O
2
.
B. MEF cells were prepared, treated with siRNA’s, transiently adapted to oxidative
stress by pre-treatment with H
2
O
2
(or not pre-treated), challenged with 1 mM H
2
O
2
, and
harvested, exactly as described in A. Samples were then seeded at a density of 100,000
cells per ml in 24 well plates. Cells were incubated for a further 24 hours, then cell counts
were taken using a cell counter. Values (means ± SE, n = 3) represent the cell population

99

Figure 6.3, Continued
in challenged cells which previously were either pre-treated with an adaptive dose of
H
2
O
2
or not pre-treated.

100

Regulation of Pa28 αβ by Nrf2

Since the oxidative stress induced increase in the level Pa28αβ appears to have an
important role in adaptation to oxidative stress we wanted to determine if like 20S
proteasome this response was regulated by changes in Nrf2. For this study, we used
western blot analyses of control and H
2
O
2
adapted cells pre-treated with either Nrf2
siRNA or a scrambled siRNA vector. With scrambled (control) siRNA treatment we saw
a two- to three-fold H
2
O
2
induced increase in Pa28αβ protein levels (Figure 6.4). With
Nrf2 siRNA pre-treatment, however, the H
2
O
2
induced increase in Pa28αβ was lost. This
indicates that the induction of PA28αβ in adaptation to stress is regulated by the
transcription factor Nrf2.

101

Figure 6.4: H
2
O
2
induced expression of Pa28αβ is Nrf2 dependent
H
2
O
2
treatment causes an increase in Pa28 αβ, which appears to depend upon Nrf2
expression. MEF cells were prepared, treated and harvested as described in chapter 4.
MEF cells were grown to 10% confluence and treated with either control or Nrf2 siRNA
for 24 h. 4 h after initiation of siRNA treatment, half the cells were exposed to 1μM H
2
O
2

for 1 h, and then washed. The cells were then lyzed, and samples were run on SDS-
PAGE gels and transferred to PVDF membranes. Membranes were treated with
antibodies directed against Pa28α subunit, and β-tubulin. The Graph shows the levels of
Pa28α regulator divided by β-tubulin levels for each well and then plotted as a percent of
control. Values are Means ± SE. n = 3. Representative western blots are presented to the
right.

Role of Pa28αβ in protein synthesis independent adaptation

In the previous paragraphs we demonstrated that Pa28 αβ is important in the protein
synthesis dependent response to oxidative stress. We then went on to investigate if it
might have a role in the initial protein synthesis independent response to oxidative stress

102

(Grune et al., 2011; Pickering et al., 2010) (described in chapters 1 & 2). This response
appears to be caused by a transient decline in association between core 20S proteasome
and the 19S regulator in the first 3 h after oxidative stress (Grune et al., 2011). It has been
argued that this occurs to make more 20S proteasome available such that damaged
proteins can be more rapidly degraded. Given that the Pa28 αβ regulator has been shown
to enhance the proteolytic capacity of the 20S proteasome (Ma et al., 1992), we reasoned
that it might also bind to 20S proteasome as is released by the 19S regulator. We
confirmed the results by Grune et al (Grune et al., 2011) using co-immunoprecipitation,
during which we observed a 35% decline in the association between 20S proteasome and
the 19S regulator under conditions of H₂O₂ exposure. On top of the decline in 19S-20S
proteasome association we also observed an 80% rise in the amount of the Pa28αβ
regulator associated with 20S proteasome (Figure 6.5A). These results indicate that the
increase in capacity to degrade oxidized proteins that has been previously observed
shortly after an oxidative stress (Grune et al., 2011; Pickering et al., 2010) might not just
be a product of the 19S regulator detaching from 20S proteasome, but might also be a
product of the Pa28αβ regulator attaching in its place.

The Role of Pa28αβ in Enhancing Degradation of Oxidized Proteins by the 20S
Proteasome

In chapters 2 & 3 as well as in previous publications (Grune et al., 2011; Pickering et al.,
2010; Pickering et al., 2012), we have shown that the initial increase in proteolytic

103

capacity (1-3 h after H₂O₂ treatment) is independent of protein synthesis, whereas the
subsequent increases (3-24 h) require de novo protein synthesis; this was reaffirmed in
the present studies, but confirmatory data are not shown. We have also shown that the
initial response to H₂O₂ treatment produces a temporary (1-3 h) disassociation of 26S
proteasome to form additional free 20S proteasomes, while the liberated 19S regulators
are protected by HSP70 (Grune et al., 2011; Pickering et al., 2010). Using new co-
immunoprecipitation techniques we have now discovered a rise in association of free 20S
proteasomes (β1 subunit) with Pa28αβ regulators occurring as rapidly as the first hour
after H₂O₂ treatment (Figure 6.5A). This rise in Pa28αβ regulator association occurred
over the same time course as the dissociation of 19S regulators from 26S proteasomes, as
shown by the diminished co-precipitation of the 19S S4 subbunit by 20S proteasome
β1(Figure 6.5A). It should also be noted that, as previously (Grune et al., 2011; Pickering
et al., 2010), we observed no increase in 19S regulator levels (not shown). These results
also suggested to us that the Pa28αβ regulator might actually assist in the initial adaptive
increase in proteolytic capacity following mild oxidative stress. It has been shown that
Pa28αβ enhances 20S proteasome’s proteolytic capacity to degrade short peptide
substrates (Ahn et al., 1996; Li and Rechsteiner, 2001). Under our conditions, we found
that if we incubated purified 20S proteasome with purified Pa28αβ, there was an ≈ 50
fold increase in proteolytic capacity to degrade the short peptide substrate Suc-LLVY-
AMC (Figure 6.5B).

104

An important aspect of cellular responses to oxidative stress is the capacity of cells to
selectively degrade oxidized proteins. This represents the ability of cells to remove
damage proteins and permit normal cell function to return after an oxidative stress
(Davies, 2000b, 2001; Pickering et al., 2010). It has been shown that the capacity to
degrade oxidized proteins increases under conditions of oxidative stress. It has also been
shown that this increase is largely dependent on the synthesis of 20S proteasome (Davies
and Goldberg, 1987b; Grune et al., 2011; Pickering et al., 2010) as well as the
mitochondrial Lon protease (Bota and Davies, 2002; Ngo and Davies, 2009; Ngo et al.,
2011). If the Pa28αβ regulator plays an important role in oxidative stress adaptation, as
we suggest, we would expect it to enhance the ability of 20S proteasome to selectively
degrade oxidized proteins. Although Pa28αβ increases with oxidative stress adaptation,
and such adaptive responses were diminished in Pa28αβγ mutants (Pickering et al., 2010;
Pickering et al., 2012), the potential for Pa28αβ to stimulate the selective degradation of
oxidized proteins by 20S proteasome has not been directly tested previously. To do this
we incubated purified 20S proteasome with or without purified Pa28αβ, and measured the
capacity to degrade either oxidized or native hemoglobin. Addition of Pa28αβ caused no
change in the capacity to degrade native hemoglobin but more than doubled the capacity
to degrade oxidized hemoglobin (Figure 6.5C). This is highly supportive of the idea that
Pa28αβ functions in oxidative stress adaptation through increasing the capacity of the
20S proteasome to degrade oxidized proteins.

105

Figure 6.5: Role of Pa28αβ in Removal of Damaged Proteins by 20S Proteasome
A: Following H₂O₂ treatment there is an increase in Pa28αβ association with 20S
proteasome and a corresponding decrease in S4 (19S subunit) binding. MEF cells were
grown to 10% confluence, exposed to 1mM H₂O₂ for 1 h, after this immunoprecipitation
of anti-β1 was performed. Samples were run on Western blots and screened for co-
immunoprecipitation of anti-Pa28α, anti-S4, or anti-β1. B: Pa28αβ increases the capacity
of 20S proteasome to degrade the peptide substrate Suc-LLVY-AMC. 20S proteasome
was incubated with a 4-fold molar excess of Pa28αβ for 30 minutes. Suc-LLVY-AMC
was then added to samples and proteolytic capacity was measured over the next 4hr.
Values are means ± SE where n = 3 for Suc-LLVY-AMC assays. C. Pa28αβ increases the
capacity of 20S proteasome to selectively degrade oxidized hemoglobin. 20S proteasome
was incubated with a 4-fold molar excess of Pa28αβ for 30 minutes. Hb-AMC or Hb
OX
-
AMC was then added to samples and proteolytic capacity was measured over the next
4hr. Values are means ± SE where n = 8.

106

II. PARP

As in the cytoplasm, the nucleus appears to have a rapid response system to oxidative
stress. In fact, in just fifteen minutes following an oxidative stress there is an eighteen
fold increase in proteasome dependent proteolytic activity in the nucleus (Ullrich et al.,
1999). It has been shown that this is not due to de novo protein synthesis but due to
modifications to the 20S proteasome to make it not only more proteolytically active but
also more capable of specifically degrading oxidized histones (Ullrich et al., 1999). The
degradation of oxidized histones is very important for DNA function. There is a highly
sophisticated system in place to repair oxidized or otherwise damaged DNA. The
histones around which the DNA is wrapped do however play an important role in control
of DNA expression (Jenuwein and Allis, 2001). As a result it is very important that any
damage to the histone complexes in the chromatin be repaired as well as the DNA itself.
The agent which appears to regulate this change in proteasome activity is a protein called
poly-ADP ribose polymerase (PARP). This agent functions by transferring ADP moieties
off NAD+ onto other proteins (Banasik et al., 1992). PARP is believed to use this
mechanism heavily as a means of signaling DNA damage and inducing its repair
(Piskunova et al., 2008). It appears that the 20S proteasome can also act as a substrate for
modification by PARP. Studies using purified 20S proteasome have shown that ADP
ribosylation by PARP resulted in an increase in selectivity for degradation of oxidized
histones. This accounted for the changes in selectivity and proteolytic activity observed
under oxidative stress of the nucleus. Furthermore, the increase in proteolytic activity in

107

the nucleus under oxidative stress is lost with inhibition of PARP activity. These results
indicate that PARP is a key regulator of proteasome in the nucleus (Ullrich et al., 1999).

III. Pa200

Another regulator of nuclear 20S proteasome is the Pa200 (also known as PSME4)
proteasome regulator. This is a 200 kilodalton activator of proteasome activity (hence
Pa200). It is highly conserved throughout the eukaryotic domain (Baumeister et al., 1998;
Ustrell et al., 2002). Pa200 is a nuclear localized regulator of proteasome. Like PARP,
Pa200 appears to be involved in repair of DNA damage. Unlike PARP, which appears to
assist DNA damage repair through changes of the 20S proteasome to increase preferential
degradation of damaged histones, the role which Pa200 plays is not entirely clear (Ullrich
et al., 1999; Ustrell et al., 2002). It has been seen in studies using the yeast homologue of
Pa200 (Blm10) that mutation of blm10 results in cells becoming much less resistant to
DNA damage (Schmidt et al., 2005). Similarly in mammalian culture the depletion of
Pa200 results in greater DNA damage susceptibility to ionizing radiation (Blickwedehl et
al., 2008). Pa200 is referred to as a proteasome activator because of its ability to enhance
the 20S proteasome’s ability to degrade short peptides. Interestingly, most studies to date
have shown that it does not appear to enhance the ability of 20S proteasome to degrade
whole proteins (Ustrell et al., 2002). It has however been shown that under cell damage
through ionizing radiation or H₂O₂ treatment there is a rapid formation of foci of Pa200
on the DNA chromatin. The formation of these DNA foci correlates with an increase in

108

Pa200 binding to 20S proteasome. It is unclear if Pa200 provides the described increase
in resistance to DNA damage through enhanced degradation of damaged proteins in the
chromatin or through triggering some form of DNA repair mechanism. However, it is
clear that whatever its actual role, it is a highly conserved, regulator of proteasome in
response to oxidation or other damage in the nucleus.

Induction Pa200 under oxidative stress

As seen for Pa28αβ, we found that 1 μM H₂O₂ exposure also produced significant
increases in Pa200 levels (Figure 6.6A, inset). There are 3 isoforms of Pa200 (200kDa,
160kDa and 60kDa) and we saw a sharp increase in the levels of the 200kDa isoform,
which is thought to be the functional isoform,, but no change in the 160kDa and 60kDa
isoforms. To further investigate this we examined the levels of (200kDa) Pa200 over a
range of H₂O₂ concentrations and observed a progressive rise in Pa200 levels with H₂O₂
exposures between 1 μM and 100 μM. We observed a 3 fold increase in Pa200 levels at
our peak H₂O₂ treatment of 100 μM H₂O₂ (Figure 6.6A). We performed an H
2
O
2
pre-
treatment challenge assay, as done in chapter 3&4 and previously (Pickering et al., 2010;
Pickering et al., 2012). In this assay we found that Pa200 levels increase under H
2
O
2

exposure, and that this increase enhances oxidative stress resistance (Figure 6.6B). Pa200
is localized in the nucleus where it has been shown, in previous reports, to form
punctuate foci under either strong ionizing radiation or toxic H
2
O
2
levels of exposure (Li
and Rechsteiner, 2001; Ustrell et al., 2002). This has led to the hypothesis that Pa200

109

binds to chromatin in severe stress, and initiates some aspect of DNA repair (Ustrell et
al., 2002). To determine whether Pa200 behaved similarly under our experimental
conditions in MEF cells, we exposed cells to low adaptive doses of H
2
O
2
(1-100µM), or
to a 1.0mM toxic dose of H
2
O
2
for 1 h then staining for Pa200. Interestingly, we
observed the formation of punctuate foci of Pa200 in the nuclei both following toxic
(1.0mM) H
2
O
2
treatments and under the conditions of low 1.0 or 10.0µM adaptive H
2
O
2

concentrations used throughout this paper; indeed, the formation of punctuate nuclear
Pa200 foci seems to follow a fairly straightforward H₂O₂ concentration-curve profile
(Figure 6.6C and Figure 6.6D).

110

Figure 6.6: Induction of Pa200 regulator in response to oxidative stress.

A: H₂O₂ exposure causes an increase in Pa200 protein levels. MEF cells were grown to
10% confluence and exposed to 0, 1, 10, or 100μM H₂O₂ for 1 h. Then, 24 h later
samples were harvested and run on Western blots which were then screened with
antibodies against either Pa200 or the loading control β-tubulin. Sample band intensity
was measured and adjusted based on β-tubulin band intensity. Values are means ± SE
where n = 3. The inset shows a representative blot. B: By blocking the adaptive increase
in Pa200, the H₂O₂ induced increase in oxidative stress tolerance is blunted. MEF cells
were grown to 20 % confluence and then treated with Pa200 or (scrambled) siRNA for 24
h to block induction. After siRNA exposure, the medium was replaced with fresh
complete medium. Some cells (labeled as H₂O₂ pre-treated) were transiently adapted to
oxidative stress by pre-treatment with 1μM of H₂O₂, for 1 h, after which the medium was
replaced. Following a 24 h adaptation period, both adapted and non-adapted cells were
challenged by incubation with a high dose of 100μM H₂O₂. After 1 h the medium was
replaced with fresh medium and cells were allowed to recover for 24 h. Cell counts were
then taken using a cell counter. Results and means ± S.E. where n = 3. C: Exposure to a
toxic level of H₂O₂ causes the formation of Pa200 foci in the nucleus. MEF cells were
grown to 50% confluence and exposed to 1.0mM H₂O₂ for 1hr. Immunocytochemistry
was then performed and cells were stained with anti-Pa200. D: Exposure to low,

111

Figure 6.6, Continued
adaptive doses of H₂O₂ also results in the formation of nuclear Pa200 foci. Conditions
were identical to those of panel C., except that 0, 1.0µM, or 10.0µM H₂O₂ exposures
were used.

112

The role of Pa200 in degradation of oxidized and naive proteins by the 20S
proteasome

We speculated that Pa200 might have a role in enhancing the capacity of proteasome to
degrade oxidized proteins in the nucleus. So like Pa28αβ we measured the activity of 20S
proteasome with and without Pa200. First we confirmed that Pa200 actually interacted
with the 20S proteasome which was done by measuring capacity to degrade the short
peptide substrate Suc-LLVY-AMC (Figure 6.7A). Addition of Pa200 to 20S proteasome
produced a 4 fold increase in proteolytic activity which is consistent with previous
reports (Ustrell et al., 2002). We next measured the capacity of the 20S proteasome-
Pa200 complex to degrade native and oxidized hemoglobin. Unlike the increased
degradation of Suc-LLVY-AMC, Pa200 appeared to almost completely block the 20S
proteasome from degrading both native and oxidized hemoglobin (Figure 6.7B). The
ability of Pa200 to block degradation of a normal protein has been reported previously
(Ustrell et al., 2002) but we were surprised that Pa200 could also essentially block
degradation of oxidized Hb. We wanted to test whether Pa200 could also inhibit
degradation of other oxidized proteins and, because of the oxidative-stress-induced
association of Pa200 with chromatin, we decided to use native and oxidized histone
proteins as substrates for either 20S proteasome, or 20S proteasome plus Pa200. 20S
proteasome degraded oxidized histones, significantly better than native histones, but
when we added Pa200 to the 20S proteasome it actually increased the degradation of
native histones and decreased that of oxidized histones (Figure 6.7C). This suggests that

113

Pa200 actually enhances selectivity for degradation of normal proteins, while reducing
selectivity for oxidized histones. Thus, while Pa200 may play a role in turnover of
normal histones, it does not appear to be involved in the specific removal of oxidatively
modified histones from the chromatin.

As described earlier, PARP has been shown to enhance proteasome selective degradation
of oxidized histones. PARP also appears to play an important role in the adaptive
increase in proteolytic capacity in the nucleus (Ullrich et al., 1999). Because of this, we
speculated that perhaps PARP might work in association with Pa200. To test this we
performed an immunoprecipitation assay, using pull-down with either 20S proteasome or
Pa200 antibodies. We then screened the samples pulled-out for the presence of either 20S
proteasome or PARP (Figure 6.7D). Here we saw that while both of the samples
contained 20S proteasome subunits, only the 20S proteasome pull-down contained
PARP. These results indicate that PARP only associates with proteasome that does not
contain the Pa200 regulator and, therefore, acts to increase proteasome activity and
selectivity for oxidized histones (Ullrich et al., 1999) in a mechanism independent of
Pa200.

114

Figure: 6.7: Pa200 regulator enhances degradation of short peptides and non-
oxidized hemoglobin
A: Pa200 increases the capacity of 20S proteasome to degrade the peptide substrate Suc-
LLVY-AMC. 20S proteasome was incubated with a 4 fold molar excess of Pa200 for 30
minutes. Suc-LLVY-AMC was then added to samples and proteolytic capacity measured
over the next 4 h. Values are means ± SE where n = 3. B: Pa200 reduces the capacity of
20S proteasome to degrade both oxidized and naive hemoglobin substrates. 20S
proteasome was incubated with a 4 fold molar excess of Pa200 for 30 minutes. Hb-AMC
or Hb
OX
-AMC was then added to samples and proteolytic capacity measured over the
next 4 h. Values are means ± SE, where n = 8. C: Pa200 enhances the capacity of
proteasome to degrade normal histones but reduces proteasome’s capacity to degrade
oxidized histones. 20S Proteasome ± Pa200 was prepared as in panel A and AMC labeled
normal histones, or oxidized AMC labeled histones were used as substrates. Oxidized
histones were prepared by treatment with either 1mM or 2mM H₂O₂ for 1 h. Proteolysis
values are means ± SE, where n = 6. D: Immunoprecipitation with either anti-Pa200 or
anti-β1 both result in co- immunoprecipitation of 20S proteasome subunits, however only
anti-β1 antibodies co-precipitate PARP. MEF cells were grown to 10% confluence, then
immunoprecipitation with either anti-β1 antibody, anti-Pa200 antibody, or anti-porin
antibody (as a non-specific binding control) was performed. Samples were run on
Western blots and screened for co-immunoprecipitation of anti-β1 or anti-PARP.

115

IV. Pa28γ

The final regulator of 20S proteasome is the proteasome regulator Pa28γ (also known as
REGγ, 11sγ, or PSME3). There is no evidence at present of a role for Pa28γ in removal
of damaged proteins. The Pa28γ regulator forms a homoheptameric ring on either end of
the 20S proteasome. It is nuclear localized and is a genetic ortholog of the better studied
Pa28αβ regulator (Li and Rechsteiner, 2001). It weakly enhances the 20S proteasomes
ability to degrade short peptides. However, its main function is largely unknown. Pa28γ
has been shown to be important in cell cycle progression as well as apoptosis
(Rechsteiner and Hill, 2005). It has also been shown to be a critical regulator of p53
expression during a stress response (Zhang and Zhang, 2008).

Induction Pa28γ under oxidative stress

The other proteasome regulator is Pa28γ whose subunits are highly similar to both Pa28α
and Pa28β, but Pa28γ is primarily a nuclear protein, whereas Pa28α and Pa28β are found
in the cytoplasm. It is thought that Pa28γ forms a homoheptameric ring which can attach
to either end of the 20S proteasome cylinder. The function of this, regulator, however is
largely unknown (Li and Rechsteiner, 2001). We demonstrated that like Pa28αβ and
Pa200, H₂O₂ adaptation caused an increase in expression of Pa28γ (Figure 6.8A). To
further investigate this we examined the levels of Pa28γ over a range of doses of H₂O₂
and observed somewhat of a progressive rise in Pa28γ levels with H₂O₂ exposures

116

between 1μM and 100 μM: peaking at a 2.5 fold increase in Pa200 levels with H₂O₂
treatment of 100μM (Fig. 6.8B). Interestingly however, unlike Pa28αβ (Figure 6.5A), we
did not see an immediate increase in Pa28γ binding to 20S proteasome with H₂O₂
induced dissociation of 26S proteasome (Figure 6.8C). Dissociation of 20S and 19S
components of the 26S proteasome was again confirmed, but this time we used the S3
subunit of the 19S regulator (for independent verification) and found that less of it co-
precipitated with the 20S β1 subunit after H₂O₂ treatment (Figure 6.8C). Previous reports
have indicated that Pa28γ causes a weak stimulation of the capacity of proteasome to
degrade the peptide substrate Suc-LLVY-AMC. We tested this in our MEF cell system
and observed a 60% increase in the activity of 20S proteasome with addition of Pa28γ
(Figure 6.8D) in comparison with the 50-fold increase seen with purified Pa28αβ (Figure
6.5B).

The role of Pa28γ in degradation of oxidized and native proteins by the 20S
proteasome

As with Pa28αβ and Pa200 we were interested in testing whether Pa28γ might stimulate
the selective degradation of oxidized proteins. We saw no change in the capacity to
degrade native hemoglobin with the addition of Pa28γ to purified 20S proteasome. We
did, however, see a ≈70% rise in capacity to degrade oxidized hemoglobin with the
addition of Pa28γ to 20S proteasome (Figure 6.8E). From these results it appears that
Pa28γ might play a similar, if weaker role, to Pa28αβ through facilitating the selective

117

degradation of oxidized proteins under oxidative stress. Given that Pa28αβ is primarily
located in the cytoplasm and Pa28γ in the nucleus (Brooks et al., 2000) it is possible that
both these regulators have the function of enhancing the removal of damaged proteins, in
different cellular regions.

118

Figure 6.8: Induction of Pa28γ regulator in response to oxidative stress

A: H₂O₂ exposure causes an increase in Pa28γ protein levels. MEF cells were grown to
10% confluence and exposed to 0μM or 1μM H₂O₂ for 1 h. Then, 24 h later samples were
harvested and run on Western blots which were then screened with antibodies against
either Pa200 or the loading control β-tubulin. B: H₂O₂ exposure causes an increase in
Pa28γ protein levels. MEF cells were grown to 10% confluence and exposed to 0, 1, 10,
or 100μM H₂O₂ for 1 h. Then, 24 h later samples were harvested and run on Western
blots which were then screened with antibodies against either Pa28γ or the loading
control β-tubulin. Sample band intensity was then measured and adjusted based on β-
tubulin band intensity. Values are means ± SE, where n = 3. C: Following H₂O₂
treatment there is no change in Pa28γ association with 20S proteasome. MEF cells were
grown to 10% confluence and exposed to 1mM H₂O₂ for 1 h, after which
immunoprecipitation of anti-α5 was performed. Samples were run on Western blots and
screened for co-immunoprecipitation of anti-Pa28γ, or anti-S3 (a subunit of the 19S
regulator) as a measure of 26S proteasome disassembly. anti-α5 (not shown) was used as
a loading control to confirm similar levels of 20S proteasome. The β1 blot was taken
from a separate set of samples run under the same conditions. D: Pa28γ slightly
increases the capacity of 20S proteasome to degrade the peptide substrate Suc-LLVY-
AMC. Purified 20S proteasome was incubated with a 4 fold molar excess of Pa28γ for 30
minutes. Suc-LLVY-AMC was then added to samples and proteolytic capacity measured

119

Figure 6.8, Continured
over the next 4 h. Values are means ± SE, where n = 3. E: Pa28γ increases the capacity
of 20S proteasome to degrade oxidized hemoglobin but not native hemoglobin. Purified
20S proteasome was incubated with a 4 fold molar excess of Pa28γ for 30 minutes. Hb-
AMC or Hb
OX
-AMC was then added to samples and proteolytic capacity measured over
the next 4 h. Values are means ± SE, where n = 8.

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V. Regulation of immunoproteasome

The immunoproteasome is another important protein involved in oxidative stress
responses, whose expression is induced during oxidative stress adaptation (Pickering et
al., 2010; Pickering et al., 2012). Immunoproteasome is at least as capable as the 20S
proteasome of the selective degradation of oxidized proteins, and contributes
significantly to the adaptive response (Pickering et al., 2010; Pickering et al., 2012). Both
the immunoproteasome and Pa28αβ are induced by interferon-γ and both of them have
been suggested to participate in similar functions in the cell (Teoh and Davies, 2004).
Because of this co-association of Pa28αβ and immunoproteasome it has been speculated
that they might act together. Immunoprecipitation of MEF cell lysate with an antibody
against the 20S proteasome subunit β1 did cause co-precipitation of Pa28αβ, Pa200, and
Pa28γ (showing that all three regulators do bind to 20S proteasome), however,
immunoprecipitation with an antibody against the immunoproteasome subunit β5i caused
rather weaker co-precipitation of Pa28αβ and Pa28γ but not of Pa200 (Figure 6.9A).
These results would suggest that the immunoproteasome can bind to (and be regulated
by) Pa28αβ and Pa28γ, but that Pa200 does not share this capacity. To further test this
conclusion we incubated purified immunoproteasome with purified samples of Pa28αβ,
Pa28γ or Pa200. Addition of Pa28αβ or Pa200 to immunoproteasome enhanced the
capacity to degrade the short peptide substrate Suc-LLVY-AMC by 5- to 7-fold, but
Pa28γ actually decreased peptide degradation slightly (Figure 6.9B). These results
suggest that immunoproteasome can actually be controlled by all three regulators.

121

Addition of purified Pa28αβ to immunoproteasome increased its capacity to degrade
oxidized Hb (Figure 6.9C, left panel) but, since degradation of native Hb also increased,
the increase in selectivity for oxidized proteins was not as great as that observed (in
Figure 6.5C) with 20S proteasome. It must be said, however, that the actual specific
activity of Immunoproteasome plus Pa28αβ (Figure 6.9C) was about twice that of 20S
proteasome plus Pa28αβ (Figure 6.5C). Addition of Pa28γ to immunoproteasome did not
appear to cause a notable change in the capacity of immunoproteasome to degrade either
oxidized or native hemoglobin (Figure 6.9C, central panel). Addition of Pa200 to
immunoproteasome also did not appear to have a strong effect on the capacity of
immunoproteasome to degrade labeled hemoglobin proteins (Figure 6.9C, right panel).
This is in contrast to results with 20S proteasome (Figure 6.7B,C) where Pa200 appeared
to block the majority of the capacity of 20S proteasome to degrade either oxidized or
native Hb. From the results of Fig. 6 it would appear clear that Pa28αβ positively
regulates the selective degradation of oxidized proteins by immunoproteasome. Pa28γ
appears to bind to immunoproteasome, but has little or no effect on its activity with
peptide substrates or either normal or oxidized proteins. We have no direct evidence that
Pa200 actually binds to immunoproteasome, but it does increase peptidase activity yet
slightly decrease the degradation of an oxidized Hb.

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Figure 6.9: Regulation of immunoproteasome by Pa28αβ, Pa28γ and Pa200
A: Immunoprecipitation of MEF cell lysate with an antibody against the 20S proteasome
subunit β1 causes co-precipitation of Pa28αβ, Pa200, and Pa28γ. However
immunoprecipitation with an antibody against the immunoproteasome subunit β5i causes
weak co-precipitation of Pa28αβ, and Pa28γ but not Pa200. MEF cells were grown to
10% confluence and cells were then harvested and lysed. Immunoprecipitation was
performed using anti-β1, anti-β5i, or anti-β-tubulin (as a control for non-specific binding)
antibodies. The Immunoprecipitate was analyzed by Western blotting and samples were
screened for co-immunoprecipitation of anti-Pa28α, anti-Pa200, anti-Pa28γ, anti-β1, or
anti-β5i. B: Pa28αβ and Pa200 but not Pa28γ increase the capacity of
immunoproteasome to degrade the peptide substrate Suc-LLVY-AMC.
Immunoproteasome was incubated with a 4 fold molar excess of either Pa28γ Pa28αβ or
Pa200 for 30 minutes. Suc-LLVY-AMC was then added to samples and proteolytic
capacity measured over the next 4 h. Values are means ± SE, where n = 3. C: Binding of
Pa28αβ (left panel) but not Pa28γ (center panel) or Pa200 (right panel) significantly
increases the selective degradation of oxidized hemoglobin by immunoproteasome.
Samples were prepared as in A, then AMC labeled hemoglobin or AMC labeled oxidized
hemoglobin were added to samples as substrates. Proteolytic capacity was measured over
the next 4 h. Values are means ± SE, where n = 8.

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VI. Binding of regulators to 20S vs hybrid proteasome

The majority of proteasome in mammalian cells (≈70-80%) is thought to be free 20S
proteasome while some 15% of total proteasome is believed to be 26S proteasome
(Brooks et al., 2000; Russell et al., 1999). The 20S proteasome lacks the 19S regulatory
cap that would enable it to degrade ubiquitinylated proteins in an ATP-stimulated
manner, but both 20S and immunoproteasome are very good at selectively degrading
oxidized proteins, which 26S is not (Davies, 1995, 2000b, 2001; Grune et al., 2011;
Grune et al., 1996; Grune et al., 1995; Pacifici et al., 1993; Pickering et al., 2010;
Reinheckel et al., 1998; Shringarpure et al., 2003; Ullrich et al., 1999). It has been shown
that both of the regulators Pa28αβ, and Pa200 are capable of forming so called ‘hybrid’
proteasome (Blickwedehl et al., 2008; Tanahashi et al., 2000). These hybrid proteasomes
have been described as containing the 19S regulator on one end of the 20S proteasome
core and a different regulator (either Pa28αβ or Pa200) on the other end (i.e. 19S-20S-
regulator complexes). It is, however, unclear whether 20S proteasomes may form some
sort of ‘hybrid’ proteasomes without involvement of 19S regulators. To test this we used
co-immunoprecipitation to isolate either 20S proteasome or the regulators Pa28αβ,
Pa28γ, or Pa200 from MEF cells (Figure 6.10A). We then used Western blotting to
determine what was bound to the isolated proteins (see also the results of Figure 6.9A).
We found that each of the regulator antibodies was capable of pulling down large
quantities of 20S proteasomes (measured through anti-β1).

124

We then screened the samples with an antibody against the S4 subunit specific to the 19S
regulator of the 26S proteasome. All of the samples immunoprecipitated with Pa28αβ,
Pa28γ, or Pa200 antibodies also had the 19S subunit bound, indicating that the regulators
do indeed form some hybrid (19S-20S-regulator) proteasomes (Figure 6.9A). The Pa200
sample had a comparable amount of S4 in it to the proteasome sample, indicating that a
moderate proportion of Pa200 within the cell is hybrid. Both the Pa28α and Pa28γ
samples, however, despite containing as much if not more 20S proteasome than the β1
(20S) proteasome samples, contained a very small amount of the 19S regulator subunit
S4 (Figure 6.10A). This suggested to us that a significant proportion of the proteasome to
which Pa28αβ and Pa28γ are bound might actually be free 20S proteasomes. To test this
hypothesis we performed the reverse experiment: Using multiple immunoprecipitation
washes with the S4 antibody, we removed ≈75% of the 19S regulator from our sample
(Figure 6.10B, left panel). This resulted in a complete loss of Pa200 from the sample, but
only a ≈50% reduction in Pa28α and Pa28γ (Figure 6.10B, right panel). From the results
of Figures 6.9A, and 6.9B, we can conclude that the majority of Pa200 is bound to hybrid
26S proteasome (19S-20S-Pa200) while Pa28αβ and Pa28γ hybrid proteasomes have
much greater flexibility, and may be found in 19S-20S-Pa28αβ/Pa28γ complexes, or
Pa28αβ/Pa28γ-20S complexes or even, perhaps, Pa28αβ/Pa28γ-20S- Pa28αβ/Pa28γ
complexes.

125

Figure 6.10: Pa200 and, to a lesser extent, Pa28αβ and Pa28γ can form hybrid
proteasomes. A: Immunoprecipitation of MEF cell lysate with anti-Pa28α, anti-Pa28γ,
anti-Pa200 or anti-β1 antibodies causes co-immunoprecipitation of both 20S and 26S
proteasome subunits, however while anti-β1 and anti-Pa200 pull out a comparable
amount of S4, anti-Pa28α and anti-Pa28γ antibodies are relatively poor at co-
immunoprecipitation of S4. Immunoprecipitation protocols were as reported in the
legends to Figs. 3 and 4. Non-specific binding was measured through
immunoprecipitation of anti-Porin. B: Depletion of S4 through immunoprecipitation
causes a small loss of Pa28α and Pa28γ but a very large loss of Pa200. MEF cell lysates
were subjected to three cycles of immunoprecipitation with either anti-S4 or anti-porin.
Non-immunoprecipitated supernatants were then taken and run on Western blots.

126

VII. Summary

Perhaps surprisingly, we find that three of the proteasome regulators, Pa28αβ, Pa28γ,
and Pa200 exhibit significantly increased levels in cells transiently adapted to oxidative
stress. In contrast, we now and previously found no increase in 19S proteasome regulator
levels (or ATP-stimulated proteolysis) under identical conditions (Pickering et al., 2010).
Our previous work (Pickering et al., 2010; Pickering et al., 2012) led us to propose that
the Pa28αβ and Pa28γ regulators might actually enhance the capacity of the 20S
proteasome to degrade oxidized proteins. As we now show directly in this paper this,
indeed, appears to be correct and the finding provides support to our hypothesis that
Pa28αβ and Pa28γ may augment oxidative stress adaptive responses by enhancing the
capacity of proteasome to selectively remove damaged proteins. Given also that these
two regulator are orthologs and have very different cellular localizations (Brooks et al.,
2000), the Pa28αβ regulator is primarily localized in the cytoplasm while the Pa28γ
regulator in the nucleus, it is likely that these two regulators might play similar roles in
different cell compartments.

In further support of the potential role for Pa28αβ in response to acute oxidative stress,
we see that when cells are exposed hydrogen peroxide there is not only an increase in
expression of both Pa28αβ and Pa28γ over the subsequent 24 h but there is also an almost
immediate increase in Pa28αβ binding to 20S proteasome that is evident within one hour
of exposure to the oxidant. In previous work, using WI-38 cells, we reported that there is

127

a detachment of the 19S regulator from 26S proteasomes to form more free 20S
proteasomes for a period of 1-3 hours following H₂O₂ exposure, which we proposed
results in an increased capacity to degrade oxidized proteins (Grune et al., 2011). Similar
disassembly of 26S proteasomes following H₂O₂ treatment was also reported in yeast by
Wang et al. (Davies, 2001; Ding et al., 2006; Ding et al., 2003; Grune et al., 2011; Grune
et al., 1995; Pickering et al., 2010; Reinheckel et al., 1998; Wang et al., 2010) who have
proposed Ecm29 as the protein that actually induces disassembly. Our own work
implicated HSP70 as the chaperone that preserves 19S regulators after dissociation, until
26S proteasomes are re-assembled within 5 hours of H
2
O
2
exposure (Grune et al., 2011).
We now confirm oxidant-induced 26S disassembly in a different mammalian cell line,
using two new measures: diminished (19S) subunits S3 and S4 binding to 20S
proteasomes. The finding that many of these newly detached 20S complexes immediately
bind to Pa28αβ regulators is exciting, given that, in this paper, we also directly
demonstrate the increased ability of 20S-Pa28αβ proteasome complexes to selectively
degrade oxidatively damaged proteins. Interestingly, however, there was no
corresponding immediate increase in the formation of 20S-Pa28γ (nuclear) complexes
under oxidative stress. This may be because nuclear 20S proteasomes were bound to
PARP, whose binding increases under oxidative stress(Ullrich et al., 1999), instead
during this period, and Pa28γ may be more important later on in adaptive responses after
it is synthesized de novo.

128

The immunoproteasome has long been believed to be linked with Pa28αβ due to
induction of their co-expression by interferon-γ and their similar localization (Pickering
et al., 2010; Preckel et al., 1999; Rivett et al., 2001). In this paper we directly
demonstrate, for the first time, that Pa28αβ is capable of binding to immunoproteasome
and that this binding significantly enhances the capacity of immunoproteasome to
selectively degrade oxidized proteins. In contrast Pa28γ and Pa200 do not appear to
significantly modify immunoproteasome activity. Considering the extremely large
induction of both immunoproteasome and Pa28αβ during transient adaptation to mild
oxidative stress (Pickering et al., 2010; Pickering et al., 2012), the direct demonstration
that 20S proteasome-Pa28αβ complexes are much more capable than other proteasome
forms of rapidly removing oxidized proteins is surely of physiological significance.

Pa200, like Pa28αβ and Pa28γ, is induced by hydrogen peroxide exposure. It does not,
however, appear to possess the same ability as the Pa28αβ and Pa28γ regulators to
enhance the capacity of 20S proteasome to selectively degrade oxidized proteins. In fact
addition of Pa200 to 20S proteasome resulted in an almost complete loss of capacity to
degrade both oxidized and native hemoglobin, although it did increase the degradation of
native (but not oxidized) histones, and of the Suc-LLVY-AMC peptide. There is a
moderate body of evidence that Pa200 has a role DNA damage repair (Blickwedehl et al.,
2008; Ustrell et al., 2002). We confirmed previous reports that Pa200 will form punctuate
foci in the nucleus under toxic H
2
O
2
exposure, but now show that this also occurs (albeit
to a lesser extent) at low (µM) adaptive H
2
O
2
concentrations. Pa200 punctuate nuclear

129

foci are believed to be the product of Pa200 binding to the chromatin at sites of DNA
damage (Blickwedehl et al., 2008; Ustrell et al., 2002). Interestingly we found that
Pa200 increases 20S proteasome’s ability to degrade normal histones and the Suc-LLVY-
AMC peptide, however it significantly decreases proteasome’s ability to degrade
oxidized histones and both normal and oxidized Hb. Previous work has shown that Poly-
ADP ribose polymerase (PARP) will, among its many functions, bind to 20S proteasome
and activate its ability to selectively degrade oxidized histones (Ullrich et al., 1999). In
this paper we show that while PARP will bind to 20S proteasome it will not bind to 20S-
Pa200 proteasome complexes. From these results it seems possible that both Pa200 and
PARP regulate the 20S proteasome in similar but separate ways. We suggest that PARP-
proteasome complexes may induce the removal of damaged histones so that DNA may
be repaired, while Pa200-proteasome complexes may catalyze the proteolytic removal of
undamaged histones from damaged DNA such that the DNA may be unwound and then
repaired.

Our studies reveal that the interactions between the 20S and 26S proteasome and its
regulators are even more complex and intricate than was previously thought. We also
observe a highly dynamic quality to regulator binding in which different regulators may
be switched in and out of proteasome as a product of changing cellular conditions. We
demonstrate that the three proteasome regulators: Pa28αβ, Pa28γ and Pa200 all appear to
bind to 20S proteasome, with and without the addition of a 19S regulator, forming
multiple types of hybrid proteasomes, although the Pa200 regulator appears to prefer 20S

130

proteasome with a 19S regulator at the other end. We also show that immunoproteasome
forms a hybrid complex with Pa28αβ. In summary we see that the proteasome regulators
all appear to be important in response to oxidative stress but appear to posses very
different roles. In particular, the 20Sproteasome- Pa28αβ complex appears to have much
greater significance in successful adaptation to oxidative stress than was previously
realized.

131

CHAPTER 7: OXIDATIVE STRESS ADAPTATION IN MODEL ORGANISMS

Like in mammalian cell culture, oxidative stress induced adaptation can occur at the
organismal level in the nematode worm, Caenorhabditis elegans. In this, exposure of C.
elegans to a mild heat shock or a mild dose of an oxidant can result in increased tolerance
to a future heat or oxidant assault. This also can result in increased lifespan (Cypser and
Johnson, 2002; Lithgow et al., 1995). Similarly in the fruit fly, Drosophila
melanogaster, a mild oxidant pre-treatment or irradiation can increase the tolerance of D.
melanogaster to a future toxic oxidant challenge (Moskalev et al., 2009). Because of
these studies, we were interested in whether this adaptation occurred through the same
pathways seen in mammalian cell culture. Specifically we were interested if exposure to
H₂O₂ induced an increase in oxidative stress tolerance in D. melanogaster and C. elegans.
We were also interested in whether this adaptation was dependant on an induction of 20S
proteasome, as well as if this induction is regulated by the Nrf2 transcription factor as is
seen in mammalian cell culture. Importantly we limited our studies to proteasome and did
not study Pa28αβ and Immunoproteasome which we demonstrated to also play an
important role in mammalian cell adaptation (Pickering et al., 2012). This was because
both of these proteins are absent in C. elegans and D. melanogaster. Both organisms do
have a homologue to the Pa28γ regulator, but as its function is largely unknown we did
not investigate its role either (Rechsteiner and Hill, 2005).

132

In this chapter we examine the ability of mild hydrogen peroxide pre-treatment to
enhance resistance of both C. elegans and D. melanogaster to oxidative stress challenge.
We also investigate the role of proteasome and Nrf2 in this response. This chapter is
entirely composed of unpublished work done in collaboration with Dr Derek Sieburth and
Dr John Tower.

I. H
2
O
2
induced adaptation to oxidative stress in C. elegans

To determine if C. elegans were capable of oxidative stress adaptation, we developed a
pretreatment challenge assay, similar to the assay used in mammalian cell culture. In this
assay the N2 strain of C. elegans were age matched and cultured to the L3/L4 phase of
development. After this C. elegans were transferred into a solution containing M9
solution ± an adaptive dose of H
2
O
2
. The C. elegans were incubated for 1 h then
transferred back onto normal plates. 24 h after the H
2
O
2
treatment, C. elegans were
transferred into vials containing M9 solution with a toxic dose of H
2
O
2
dissolved in it.
The C. elegans were incubated in this solution for 1 h. C. elegans were then returned to
normal plates and survival scored based on response to prodding with a pick, as done
previously (Larsen, 1993). Doing this assay we found in non-pretreated worms a
progressive decline in survival with increased H₂O₂ challenge (Figure 7.1A). A challenge
dose of 8mM (which we considered the best challenge dose) produced 30% survival in
none pretreated worms. This survival was increased to 45-50% survival with H
2
O
2

pretreatment (Figure 7.1B). From the above results, it appeared that indeed an H
2
O
2

133

treatment caused enhanced resistance to oxidative stress. The challenge doses used in the
previous experiment however were highly toxic and produced 100% mortality 24 h after
pretreatment, even in pretreated samples. To better test oxidative stress adaptation, we
considered it reasonable to look at long-term survival following H
2
O
2
challenge. To test
this we repeated the assay in Figure 7.1A, using milder H
2
O
2
challenge doses (200μM-
3.2mM). We then permitted a 24 h recovery period after H
2
O
2
challenge, before we
scored survival. In this we saw a similar pattern of declining survival with increased
H₂O₂ challenge (Figure 7.1C). We found 1.6mM of H₂O₂ to be our optimum challenge
treatment, producing a 20% survival with challenge. This was increased to 30% with
H₂O₂ pretreatment (Figure 7.1D). In mammalian cell culture experiments, we have shown
an H
2
O
2
induced increase in proteolytic capacity (Pickering et al., 2010; Pickering et al.,
2012). We went onto demonstrate that this was an important part of adaptation and
increases in oxidative stress resistance (Grune et al., 2011; Pickering et al., 2010;
Pickering et al., 2012). We were interested in whether a similar increase in proteolytic
capacity occurred in C.elegans. To test this we pretreated worms with H
2
O
2
as done in
Figure 1B,D. 24 h after pretreatment, we lysed the worms and measured the capacity of
the lysate to degrade the short peptide substrate, Suc-LLVY-AMC. In this assay we saw a
5-fold increase in proteolytic capacity with H
2
O
2
treatment (Figure 7.1E).

134

Figure 7.1: In C.elegans, H
2
O
2
pretreatment increases H
2
O
2
tolerance and
proteolytic capacity
A. Triplicate samples of 35 - 100 (typically ≈70) age matched, adult, N2 C. elegans were
challenged with 0-16mM H
2
O
2
for 1 h. Survival was scored immediately after challenge.
Values are Means ± SE where n = 3. B. 24 h before H₂O₂ challenge, C. elegans were
incubated in M9 solution ± 0.1, 1 or 10μM of H
2
O
2
for 1 h then returned to normal plates.
24 h later C. elegans were challenged with 8mM H
2
O
2
for 1 h then returned to normal
plates. Survival was scored immediately after challenge. Values are Means ± SE where n
= 3. Survival was scored in a blinded assay through randomization of plates. C.
Triplicate samples of 40 - 195 (typically ≈100) age matched, adult, N2 C. elegans were
prepared and pretreated as in A. Except C. elegans were challenged with 0-3.2mM of
H
2
O
2
and C. elegans were returned to normal plates for 24 h after which survival was
scored. D. 24 h before H₂O₂ challenge, C. elegans were incubated in M9 solution ± 0.1, 1
or 10μM of H
2
O
2
for 1 h then returned to normal plates. 24 h later worms were
challenged with 1.6mM H
2
O
2
for 1 h then returned to normal plates. Survival was scored
24 h after challenge. Values are Means ± SE where n = 3. Survival was scored in a
blinded assay through randomization of plates. E. C. elegans were exposed to 0-10μM
H
2
O
2
for 1 h and lysed 24 h later. Protein content was normalized by BCA assay and
proteolytic capacity measured through Suc-LLVY-AMC degradation. All values are
plotted as nM of AMC released per min per mg of lysate and are means ± SE, where n=
3.

135

II. 20S proteasome in C. elegans oxidative stress adaptation

We observed that oxidative stress adaptation occurred in a similar manner in C. elegans
to that seen in mammalian cells (Grune et al., 2011; Pickering et al., 2010; Wiese et al.,
1995). We also observed that there was a corresponding increase in proteolytic activity.
We next went on to question if 20S proteasome was involved. First we tested if H
2
O
2

treatment caused a rise in 20S proteasome protein levels, as had been seen with H
2
O
2

treatment of mammalian cells (Pickering et al., 2010). To do this we age matched C.
elegans, at the L3/L4 developmental stage, then we treated the C. elegans with 0μM,
0.1μM, 1μM or 10μM of H
2
O
2
, prepared in M9 solution as before. 24 h later we lysed the
C. elegans and analysed the lysate on a Western blot. We screened the lysate with
antibodies directed against the 20S proteasome subunit Pas-7, the 19S regulator subunit
Rpn-10 and β-tubulin as a loading control (Figure 7.2A,B). Here we saw a 2-3 fold
increase in 20S proteasome levels with no significant change in the levels of the 19S
regulator. Having seen that 20S proteasome was induced by H
2
O
2
treatment we went on
to ask if its induction was important for the adaptive response. To test this we repeated
the survival assay, described in Figure 7.1B, with the addition that RNAi was used as a
treatment prior to H
2
O
2
adaptation. In this experiment L3-L4 C. elegans were placed on
plates containing a bacterial lawn which produced RNAi directed against the 20S
proteasome subunits Pas-5, Pbs-3, Pbs-5, or Pbs-6. In addition some worms were placed
on a lawn containing bacteria which expressed RNAi directed against L4440 which
served as a non-specific control treatment. Worms were cultured on this lawn for 6 h.

136

This we reasoned would block new synthesis of these genes without reducing baselines
levels. We then transferred the C. elegans into M9 solution ± 1μM H
2
O
2
for 1 h to induce
adaptation. C. elegans were then returned to their RNAi plates. 24 h later worms were
collected and adaption experiments performed as in Figure 7.1B. In this C. elegans were
transferred into M9 solution containing 8mM H
2
O
2
for 1 h, after which survival was
scored. We found that C. elegans treated with the control RNAi had comparable
adaptation to that seen in Figure 7.1B. However with treatment with RNAi against the
proteasome subunits, adaptation occurred but was severely blunted (Figure 7.2C). This
indicates that the proteasome plays a role in the adaptive increase in oxidative stress
tolerance.

137

Figure 7.2: In C. elegans, H
2
O
2
increases 20S proteasome which enhances oxidative
stress tolerance
A. H
2
O
2
pretreatment causes an increase in expression of 20S proteasome but not 26S
proteasome. Triplicate samples of N2 C. elegans were pretreated with 0 μM, 0.1 μM, 1 μM
or 10μM of H
2
O
2
for 1 h then allowed to adapt for 24 h. C. elegans were then lysed and
protein content normalized by BCA assay. Western blot analysis was then performed on
samples. In this samples were screen with antibodies against the 20S proteasome subunit
Pas-7, the 19S regulator subunit Rpn-10 or β-tubulin as a loading control. B. Samples
were prepared as in A, in triplicate. Band intensity of Pas-7 and Rpn-10 was determined
and adjusted by the band intensity of the β-tubulin. Samples were then plotted as a
percent of control. The values plotted are Mean ± SE, where n = 3. C. Bacteria
expressing RNAi against Proteasome subunits Pas-5, Pbs-3, Pbs-5 or Pbs-6, or L4444
were cultured and plated. C. elegans were grown on RNAi plates for 6 h, then pretreated
with 1μM H
2
O
2
for 1 h, after which they were returned to RNAi plates for 24 h. After the
24 h adaptation time, C. elegans were challenged with a toxic dose of H
2
O
2
, and survival
was scored 1 h after challenge. All values are means ± SE, n = 3.

138

III. The role of Skn-1 in oxidative stress adaptation in C. elegans

We have seen earlier that Nrf2 is an important regulator of oxidative stress adaptation in
mammalian cell culture (Pickering et al., 2012). In addition we have shown that the
increase in 20S proteasome as a product of oxidative stress adaptation is Nrf2 dependent
(Pickering et al., 2012). Skn-1 is a functional, though not structural homologue of Nrf2
(An and Blackwell, 2003; An et al., 2005; Kahn et al., 2008; Oliveira et al., 2009). It has
been shown that high doses of H
2
O
2
treatment (10mM) can cause Skn-1 to become
nuclear localized in the intestines during early larval stages (L1-L2) of C .elegans (An
and Blackwell, 2003; An et al., 2005). We received the kind donation of a SKN-1::GFP
line generated by An and Blackwell (An and Blackwell, 2003), we then crossed this line
with the Glo-1 mutant lines to reduce non-specific fluorescence. First we confirmed that
this same Skn-1 nuclear localization occurs in older L3/L4 C .elegans (Figure 7.3A). To
do this we first age matched the C .elegans. Next we cultured the C .elegans to the L3/L4
larval stage, then we incubated the C .elegans in M9 ±10 mM H
2
O
2
for 20 minutes. After
this the C .elegans were paralysed using 2,3, butanedione monoxime and examined under
a microscope. From this we observed a visible increase in levels of Skn-1 nuclear
localization with H
2
O
2
exposure (Figure 7.3A). We next quantified this by looking at the
number of visibly GFP stained nuclei was scored ≈20-30 L3-L4 C .elegans. This was
used as a measure of Skn-1 nuclear localization. Treated C .elegans have an average of
10.00 ± 1.36 nuclei expression GFP compared to 3.67 ± 1.16 in untreated C .elegans
(Figure 7.3B). Having seen that H
2
O
2
treatment could induced Skn-1 nuclear localization

139

we were next interested to see if our adaptive treatment could produce the same response.
To do this we repeated the experiment using the adaptive dose of 1μM of H
2
O
2
. Here we
saw less of an increase in localization of Skn-1 to the nucleus than with 10mM H
2
O
2.

However we could still see a visible increase in Skn-1 localization (Figure 7.3C). As
before we did a quantification of this, because of the smaller difference we used samples
of ≈50-60 L3-L4 C .elegans instead of ≈20-30 C .elegans. With 1μM treatment we saw
an average of 6.14 ± 1.07 GFP expressing nuclei in H
2
O
2
treated worms compared to
only 2.09 ± 0.58 in control C .elegans. These results indicate that 1μM H
2
O
2
exposure
produced an increase in Skn-1 localization (Figure 7.3D). Next we wanted to test if the
induction in Skn-1 is important for H
2
O
2
induced oxidative stress adaptation. To test this
we performed the RNAi survival assay, described in Figure 7.2C, using Skn-1 RNAi
instead. In this experiment, L3-L4 C .elegans were placed on plates containing a bacterial
lawn which produced RNAi directed against Skn-1 or L4440 as a control treatment. C
.elegans were cultured on this lawn for 6 h, then transferred into M9 solution ± 1μM
H
2
O
2
for 1 h. As before this was used as an adaptive dose. C .elegans were then returned
to RNAi plates. 24 h later C .elegans were collected and an adaption experiments
performed as in Figure 7.1B. In this C .elegans were transferred into M9 solution
containing 8mM H
2
O
2
for 1 h. We found that C .elegans treated with the control RNAi
had comparable adaptation to that seen in Figure 7.1B. However with treatment with
RNAi against Skn-1, adaptation occurred but was severely blunted (Figure 7.3E). This
confirmed that not only did the adaptive response produce an increase in Skn-1 nuclear
localization but it this localization is important for the oxidative stress adaptation.

140

As described earlier we have shown in the mammalian system that Nrf-2 plays an
important role in adaptation. We have also shown that an important part of its role in
adaptation is through inducing the up regulation of 20S proteasome (Pickering et al.,
2012). Having seen that its C. elegans homologue, Skn-1, is also induced by H
2
O
2

exposure and plays a role in oxidative stress adaptation, we speculated that it might
regulate proteasome in a similar manner. This idea is supported in finding by Li et al (Li
et al., 2011), which demonstrated that half of the proteasome subunits contain Skn-1
binding sites. Furthermore they showed that RNAi depletion of Skn-1 caused a marked
reduction in the proteolytic capacity of the worm. To determine if Skn-1 regulated the
adaptive increase in 20S proteasome we used western blotting to measure the level of 20S
proteasome under H
2
O
2
exposure using N2 C .elegans and a Skn-1 mutant line (zu67)
(Bowerman et al., 1992). As in Figure 7.2A,B, H
2
O
2
exposure caused an increase in the
expression of the 20S proteasomal subunit Pas-7. However in the Skn-1 mutant line we
observed no change in the level of Pas-7 under H
2
O
2
exposure (Figure 7.3F). These
results suggest that indeed, like in mammalian culture Skn-1 regulates the oxidative stress
induced increase in 20S proteasome.

141

Figure 7.3: H
2
O
2
exposure induces Skn-1 nuclear localization and Skn-1 dependent
proteasome induction in intestines

A. 10mM H
2
O
2
treatment causes an increase in Skn-1 nuclear localization. L3/L4, SKN-
1::GFP C. elegans, were treated for 20 minutes with 0 or 10mM H
2
O
2
in M9 solution. C.
elegans were then paralysed and visualized under a microscope. B. The degree of Skn-1
localization with 10mM H
2
O
2
treatment was quantified. In this the number of GFP
stained nuclei was counted in samples of ≈ 20-30 C. elegans treated with ± 10mM H
2
O
2
.
The graph contains both a plot of the number of visibly GFP stained nuclei in each C.
elegans counted as well as a box chart of the data. C. 1μM H
2
O
2
treatment also causes an
increase in Skn-1 nuclear localization. C. elegans were prepared as in A, except C.
elegans were treated with 1μM H
2
O
2
instead of 10mM H
2
O
2
. D. The degree of Skn-1
localization with 10mM H
2
O
2
treatment was quantified. In this the number of GFP
stained nuclei was counted in samples of ≈ 50-60 C. elegans treated with ± 1μM H
2
O
2
.
The graph contains both a plot of the number of visibly GFP stained nuclei in each C.
elegans counted as well as a box chart of the data. E. Bacteria expressing RNAi against
Skn-1 or L4444 were cultured and plated. C. elegans were grown on RNAi plates for 6 h,
then pretreated with 1μM H
2
O
2
for 1 h, after which they were returned to RNAi plates for
24 h. After the 24 h adaptation time, C. elegans were challenged with a toxic dose of
H
2
O
2
, and survival was scored 1 h after challenge. All values are means ± SE, n = 3. F.
H
2
O
2
pretreatment causes an increase in expression of 20S proteasome in wild-type (N2)
C. elegans but not Skn-1 mutant (zu67) C. elegans. L3/L4 C. elegans that were

142

Figure 7.3, Continued
homozygous for the Skn-1 mutation were picked and run in parallel with similarly aged
N2 C. elegans. C. elegans were pretreated with 0 μM, or 1 μM of H
2
O
2
for 1 h in M9
solution then returned to normal plates and allowed to adapt for 24 h. C. elegans were
then lysed and protein content normalized by BCA assay. Western blot analysis was then
performed on samples. In this samples were screen with antibodies against the 20S
proteasome subunit Pas-7, or β-tubulin as a loading control. Values are plotted as the
band intensity of Pas-7 adjusted by β-tubulin. Values are Mean ± SE where n = 3. Below
the graph are representative blots.

143

IV. H
2
O
2
induced increase in proteolytic capacity in D. melanogaster

Like C. elegans, studies have shown that pre-exposure of D. melanogaster to mild doses
of an oxidants or radiation results in a subsequent increase in tolerance to oxidative
damage (Moskalev et al., 2009). We again wished to determine if this occurred through
the same or a similar mechanism to that seen in mammalian cell culture (Pickering et al.,
2010) and in C. elegans. First, we determined if H
2
O
2
pretreatment could be used to
induce an increase in proteolytic capacity as we have seen in the other model systems.
Because of the greater size and complexity of the Drosophila compared to mammalian
cell culture or nematode worms we could not simply incubate Drosophila in H
2
O
2
.
Instead two different pretreatment regimes have been developed. Either Drosophila can
be incubated in vials containing a Kim-wipes© soaked in 5% sucrose ± an adaptive dose
of H
2
O
2
(Grover et al., 2009) which the Drosophila will then ingest as food. Alternatively
Drosophila can be injected with H
2
O
2
(Grover et al., 2009). We decided to use the regime
of feeding Drosophila 5% sucrose ± H
2
O
2
as our treatment regime. This was used for the
reason that it enabled a greater consistency of treatment and involved less damaging
manipulation of the fly. In this experiment flies were collected Drosophila 0-48 h after
emerging from a pupal state. Drosophila were then allowed to mature for 4 days, after
which the flies were transferred into vials containing Kim-wipes© soaked in 5% sucrose
± 10-1000μM of H
2
O
2
for 6 h. The Drosophila were then returned to normal vials to
permit adaptation. 24 h after the initiation of the pretreatment Drosophila were ground up
and lysed. The proteolytic capacity of the lysate was then measured through degradation

144

of Suc-LLVY-AMC. In this we saw a progressive increase in proteolytic capacity in both
males and females with H
2
O
2
pretreatment (Figure 7.4A). To determine if this was a
proteasome dependant increase in proteolytic capacity we pre-exposed some of the
samples to the proteasome selective inhibitor lactacystin, 30 minutes before proteolytic
capacity was measured, as done in mammalian culture (Pickering et al., 2010). We saw
that the increase in proteolytic capacity was lost with addition of lactacystin indicating
that the increase was due to changes in proteasome (Figure 7.4B).

145

Figure 7.4: In Drosophila, H
2
O
2
pretreatment increases proteolytic capacity
A. H
2
O
2
treatment of D. melanogaster increase proteolytic capacity. Triplicates vials of
20, 4-6 day old w[1118] flies were transferred to vials containing a Kim-wipes© soaked
in 5% sucrose and 0, 10, 100 or 1000μM of H
2
O
2
for 6 h. Drosophila were then returned
to normal vials for 18 h to permit adaptation. After which Drosophila were lysed and
protein content normalized based on BCA assays. The proteolytic substrate Suc-LLVY-
AMC was then added to samples and its degradation was measured over the following 4
h. Values are plotted as nMol of AMC released per min per mg lysate, and are means ±
SE were n = 3. B. The H
2
O
2
induced increase in proteolytic capacity is proteasome
dependent. In female samples ± 100μM H
2
O
2
, generated in A, samples were treated with
± lactasystin 30 minutes prior to addition of Suc-LLVY-AMC. Values are plotted as
nMol of AMC released per min per mg lysate, and are means ± SE were n = 3.

146

V. H
2
O
2
induced adaptation to oxidative stress in D. melanogaster

Having seen an H
2
O
2
induced increase in proteolytic capacity we then wished to
determine if there was a corresponding increase in tolerance to oxidative stress. To test
this we prepared and pretreated Drosophila as we did in Figure 7.4. Then 24 h after the
initiation of pretreatment (18 h after the termination of treatment), Drosophila were
transferred to vials containing Kim-wipes© soaked in 5% sucrose and a toxic dose of
H
2
O
2
, which we determined to be 4.4M. Instead of subjecting flies to a single challenge
as in mammalian cell culture and C.elegans, the assay we used in D.melanogaster was to
incubate the Drosophila on this toxic dose of H
2
O
2
and score survival over time (Sykiotis
and Bohmann, 2008, 2011). In this assay we monitored survival every 6-8 h, typically
over 96 h. We found that untreated female Drosophila survived an average of 48 h
following initiation of challenge. By comparison female Drosophila pretreated with
100μM H
2
O
2
(which we found to be the optimum treatment) survived an average of 60 h.
This represented a significant increase in survival time (Figure 7.5A/B). By comparison
we did not see any change in survival time following pretreatment in male flies, with both
pretreated and non-pretreated surviving ≈40hr following initiation of challenge (Figure
7.5A/B).

147

VI. 20S proteasome in D.melanogaster oxidative stress adaptation

Next we wished to test if like in mammalian cell culture, 20S proteasome expression was
induced by H
2
O
2
treatment. To do this we prepared flies as before, pretreating female
Drosophila for 6 h with 100μM H
2
O
2
which appeared to be the optimum treatment
regime. 24 h after the start of treatment flies were lysed and the lysate was run on a
Western blot. The Western blot was then screened with antibodies directed against α
subunits of the 20S proteasome, the p42e subunit of the 19S regulator or β-tubulin as a
loading control. We observed a 50-60% rise in the expression 20S proteasome subunits
but no change in the 19S regulator (Figure 7.5C). Having seen an increase in 20S
proteasome as a product of oxidative stress adaptation, we wished to test if this induction
was required for adaptation to occur. To test this we used two 20S proteasome
UAS/RNAi lines purchased from the Vienna Drosophila RNAi center. We then crossed
them with the Act-GS-255B strain (Fischer et al., 1988; Ford et al., 2007) to conditionally
block 20S proteasome expression. We chose to use two 20S proteasome RNAi lines, one
against the proteasome subunit Prosβ1 and the other against the proteasome subunit
Prosβ2. Additionally, we crossed the Act-GS-255B strain with w[1118] (Wig)
Drosophila as controls for potential effects of RU486. We then incubated the female
progeny of these three lines in ± RU486 for 7 days. RU486 activates the Act-GS-255B
switch so inducing expression of the appropriate RNAi. 24 h prior to the end of
incubation we transferred the flies to vials containing Kim-wipes© soaked in 5% sucrose
± 100μM of H
2
O
2
for 6 h then returned the flies to vials containing ± RU486 for the

148

remaining 18 h. At the start of the assay we transferred the Drosophila to vials containing
5% sucrose and a toxic dose of 4.4M H
2
O
2
. We then scored Drosophila survival every
8hrs for the next 96hrs. As a minor confounding issue we consistently saw an RU486
induced increase in tolerance to H
2
O
2
challenge. We compensated for this by using ±
RU486 control in all experiments. We saw in w[1118] x Act-GS-255B Drosophila an
H
2
O
2
pretreatment induced increase in tolerance to oxidative stress both with and without
RU486 treatment (Figure 7.6A). We then repeated the experiment using the progeny of
the two RNAi lines, which expressed RNAi against 20S proteasome subunits Prosβ2 and
Prosβ1 crossed with Act-GS-255B. We found that in the RNAi line expressing Prosβ2,
non-RU486 treated flies adapted to a similar extent to control Drosophila under H
2
O
2

treatment. In contrast, H
2
O
2
treatment in Drosophila which were treated with RU486 did
not have any increase tolerance to oxidative stress as it did in control Drosophila. In fact,
H
2
O
2
treatment actually reduced oxidative stress tolerance (Figure 7.6B). Similarly in the
Prosβ1 RNAi line, non-RU486 treated Drosophila adapted to a similar extent to control
flies under H
2
O
2
treatment. When Drosophila were treated with RU486, not only did flies
lose the ability to adapt but tolerance to oxidative stress was severely reduced with and
without H
2
O
2
treatment (Figure 7.6C).

149

Figure 7.5: In Drosophila, H
2
O
2
pretreatment increases stress resistance and 20S
proteasome levels
A. H
2
O
2
pretreatment enhances resistance to oxidative stress challenge. Triplicates vials
of 20, 4-6 day old w[1118] Drosophila were transferred to vials containing a Kim-
wipes© soaked in 5% sucrose and 0, 10, 100 or 1000μM of H
2
O
2
for 6 h. Drosophila
were then returned to normal vials for 18 h to permit adaptation. Drosophila were then
transferred to vials containing a Kim-wipes© soaked in 5% sucrose and 4.4M H
2
O
2
. %
Survival was then scored every 8 h based on complete loss of motion. The assay was
terminated 80 h after its initiation once all Drosophila were dead. Values are Mean ± SE
were n = 3. B. The results from A are re-plotted as the lifespan following initiation of
H
2
O
2
challenge. The 3 vials for each condition are pooled such that each condition is a
count of the mean lifespan of 60 flies. C. H
2
O
2
pretreatment induces increased expression
of 20S but not 26S proteasome. Triplicates vials of 20, 4-6 day old w[1118] Drosophila
were transferred to vials containing a Kim-wipes© soaked in 5% sucrose and 0, 10, 100
or 1000μM of H
2
O
2
for 6 h. Drosophila were then returned to normal vials for 18 h to
permit adaptation. After which Drosophila were lysed and protein content normalized
based on BCA assays. 40μg of the lysate was then run on an SDS page gel and analysed
by western blotting. With antibodies against the α subunits of the 20S proteasome, p42E
which is a subunit specific to the 19S regulator of the 26S proteasome and β-tubulin as a
loading control. Values are plotted as mean ± SE were n = 3, representative blots are
attached as an inset. Values marked with * were significant at p <0.05 using logrank
analysis.

150

Figure 7.6: In Drosophila, 20S Proteasome is required for H
2
O
2
induced adaptation
4-6 day old female progeny of: A,B. w[1118] x Act-GS-255B C,D. Prosβ2-UAS
RNAi
x
Act-GS-255B and E,F. Prosβ1-UAS
RNAi
x Act-GS-255B were cultured for 6 days in
normal food vials containing 100μl of either RU486 (+RU486) or ethanol (-RU486).
After this Drosophila were transferred to vials containing a Kim-wipes© soaked in 5%
sucrose and 0 or 100μM of H
2
O
2
for 6 h. Drosophila were then returned to ± RU486 vials
for 18 h to permit adaptation. Drosophila were then transferred to vials containing a Kim-
wipes© soaked in 5% sucrose and 4.4M H
2
O
2
. % Survival was then scored every 8 h
based on complete loss of motion. The assay was terminated 96 h after its initiation once
all flies were dead. A,C,E Show % survival over the 96 h time course. Values are Mean
± SE were n = 3. B,D,F show the values re-plotted as the lifespan following initiation of
H
2
O
2
challenge. The 3 vials for each condition are pooled such that each condition is a
count of the mean lifespan of 60 Drosophila. Values marked with * were significant at p
<0.05 using logrank analysis.

151

VII. Role of Cnc-C in oxidative stress adaptation in D. melanogaster

We observed in C. elegans and mammalian cell culture that Skn-1/Nrf2 is important for
oxidative stress adaptation and regulates the H
2
O
2
induced increase in 20S proteasome.
We wished to test if this was also the case in D. melanogaster. The D. melanogaster
homologue is Cnc-C. Cnc-C has also been shown to be an important regulator of
oxidative stress tolerance (Sykiotis and Bohmann, 2008, 2011). First to test if Cnc-C is
involved in adaptation we crossed a line expressing RNAi against Cnc-C, created by
Sykiotis and Bohmann (Sykiotis and Bohmann, 2008, 2011), with Act-GS-255B and
performed adaption experiments on the progeny as with the proteasome RNAi lines. As
before we saw an increase in oxidative stress tolerance with H
2
O
2
treatment in non-
RU486 treated Drosophila, while we saw no adaptive increase in oxidative stress
tolerance in RU486 treated Drosophila. In fact, H
2
O
2
treatment caused a reduction in
oxidative stress tolerance (Figure 7.7A-C). In addition we also received the kind donation
of a Dkeap-1 RNAi expressing line. Dkeap1 is a repressor of Cnc-C and has a similar
function to Keap-1 in mammalian cell culture (Sykiotis and Bohmann, 2008). As before
we crossed the Dkeap-1 RNAi line with Act-GS-255B, the progeny were then treated
with ± RU486 for 5 days. The Drosophila were subsequently challenged with a toxic
dose of H
2
O
2
and survival was measured as before. We found that knock-down of Dkeap-
1 caused a significantly larger increase in resistance to oxidative stress than the increase
in oxidative stress resistance with RU486 treatment in control Drosophila (Figure 7.7D-
G). We next were interested if like in mammalian cell culture, the induction of 20S

152

proteasome is dependent on Cnc-C/Nrf2. To test this we prepared and pretreated Cnc-
C
RNAi
x ACT-GS-255B Drosophila as in Figure 7.7A. We then lysed the flies and ran the
lysate on a western blot. Next, we screened with antibodies directed against α subunits of
the 20S proteasome, or β-tubulin as a loading control. We observed that H
2
O
2
treatment
produced an increase in the expression 20S proteasome subunits in flies not treated with
RU486 but not in Drosophila treated with RU486. These results are supportive of a role
for Cnc-C in regulating the H₂O₂ induced increase in 20S proteasome (Figure 7.7H).

153

Figure 7.7: The Drosophila Nrf2 homologue Cnc-C is required for adaptation

A. 4-6 day old female progeny of Cnc-C
RNAi
x Act-GS-255B, were cultured for 3 days in
vials containing 100μl of either RU486 (+RU486) or ethanol (-RU486). After this
Drosophila were then transferred to vials containing a Kim-wipes© soaked in 5% sucrose
and 0 or 100μM of H
2
O
2
± RU486 for 24 h. Drosophila were then returned to normal ±
RU486 vials for 24 h to permit adaptation. Drosophila were then transferred to vials
containing a Kim-wipes© soaked in 5% sucrose and 4.4M H
2
O
2
. % Survival was then
scored every 8 h based on complete loss of motion. The assay was terminated 104 h after
its initiation once all drosophila were dead. Values are Mean ± SE were n = 4. Below the
survival curve the values are re-plotted as the lifespan following initiation of H
2
O
2

challenge. The 4 vials for each condition were pooled such that each condition is a count
of the mean lifespan of 80 drosophila. B. 4-6 day old female progeny of either Dkeap-
1
RNAi
x Act-GS-255B or Wig x Act-GS-255B were cultured for 5 days in vials containing
100μl of either RU486 (+RU486) or ethanol (-RU486). Drosophila were then transferred
to vials containing a Kim-wipes© soaked in 5% sucrose and 4.4M H
2
O
2
. % Survival was
then scored every 8 h based on complete loss of motion. The assay was terminated 96 h
after its initiation once all drosophila were dead. Values are Mean ± SE were n = 4.
Below the survival curve the values are re-plotted as the lifespan following initiation of
H
2
O
2
challenge. The 4 vials for each condition were pooled such that each condition is a
count of the mean lifespan of 80 drosophila. C. 4-6 day old female progeny of Cnc-C
RNAi

x Act-GS-255B, were cultured for 3 days in vials containing 100μl of either RU486

154

Figure 7.7, Continued
(+RU486) or ethanol (-RU486). After this Drosophila were then transferred to vials
containing a Kim-wipes© soaked in 5% sucrose and 0 or 100μM of H
2
O
2
± RU486 for
24 h. Drosophila were then returned to normal ± RU486 vials for 24 h to permit
adaptation. After which drosophila were lysed and protein content normalized based on
BCA assays. The 40μg of the lysate was then run on an SDS page gel and analyzed by
western blotting. With antibodies against the α subunits of the 20S proteasome, p42E
which is a subunit specific to the 19S regulator of the 26S proteasome and β-tubulin as a
loading control. D. 4-6 day old female progeny of Cnc-C
RNAi
x Act-GS-255B, were
cultured for 3 days in vials containing 100μl of either RU486 (+RU486) or ethanol (-
RU486). After this Drosophila were then transferred to vials containing a Kim-wipes©
soaked in 5% sucrose and 0 or 100μM of H
2
O
2
± RU486 for 24 h. Drosophila were then
returned to normal ± RU486 vials for 24 h to permit adaptation. 40μg of the lysate was
then run on an SDS page gel and analyzed by western blotting. With antibodies against
the α subunits of the 20S proteasome and β-tubulin as a loading control.

155

Figure 7.7, Continued

156

VIII. Summary

In mammalian cell culture, we have shown that H
2
O
2
induced adaptation to oxidative
stress occurs through a pathway that is strongly dependent on an Nrf2 dependant increase
in 20S proteasome (Pickering et al., 2010). The model organisms Ceanorhabitis elegans
and Drosophila melanogaster have also been shown to be capable of oxidant induced
adaptation to oxidative stress (Cypser and Johnson, 2002; Lithgow et al., 1995; Moskalev
et al., 2009). We showed that both model organisms are capable of such adaptation under
H
2
O
2
treatment. Like in mammalian cells (Pickering et al., 2010) we showed that this
adaptive response also produced an increase in proteolytic activity and increased
expression of 20S but not 26S proteasome. We also demonstrated that the increase in
20S proteasome expression in both model organisms was important for the adaptive
response and that it occurs through induction by homologues of the transcription factor
Nrf2.

In C.elegans hormetic adaption has been widely reported but is typically reported in
terms of changes in lifespan with only a few studies of changes in oxidative stress
tolerance (Cypser and Johnson, 2002; Lithgow et al., 1995). We find that under hormetic
adaptation C.elegans become more resistant to oxidative stress and their cellular
proteolytic capacity increases. There is also a large increase in 20S proteasome, we go
onto show that this increase is important for the process of adaption. This was
demonstrated using RNAi, in which RNAi knock down of 20S proteasome subunits

157

reduces the capacity of the worm to adapt to oxidative stress. Skn-1 is the C.elegans
homologue of Nrf2 (An and Blackwell, 2003; Oliveira et al., 2009). Nrf2 is an oxidative
stress response transcription factor which is induced under H₂O₂ exposure and is
important in for the adaptive response (Pickering et al., 2012). It has been reported that
under high doses (10mM) of H₂O₂ exposure, Skn-1 is induced and translocates to the
nucleus (An and Blackwell, 2003; An et al., 2005; Oliveira et al., 2009). We demonstrate
that not only is it induced with 10mM of H
2
O
2
but it is also induced with the adaptive
dose of 1μM of H₂O₂. In this induction, Skn-1 translocates to the nucleus in cells
throughout the intestines of C.elegans. In addition, using Skn-1 RNAi we show that this
induction is important for adaptive response. Furthermore we show that Skn-1 regulates
that adaptive increase in 20S proteasome.

Oxidant or radiation induced adaptation to oxidative stress in Drosophila melanogaster
has been previously reported (Moskalev et al., 2009). We confirmed that with H₂O₂
pretreatment Drosophila become more resistant to oxidative stress and their cellular
proteolytic capacity increases. We also demonstrated that this increase in proteolytic
capacity was proteasome dependent which is consistent with mammalian cell culture
studies (Pickering et al., 2010). Corresponding with the increase in oxidative stress
tolerance we observed an increase in 20S proteasome levels. When we blocked this
increase in 20S proteasome, using conditionally expressing RNAi lines, we lost the
adaptive response. This indicates that 20S proteasome is important for adaptation. We
went onto show that the drosophila homologue of Nrf2, Cnc-C, is also involved in

158

adaptation. We went onto demonstrate that part of this role was through regulating the
increase in 20S proteasome expression.

In conclusion we find that oxidative stress adaption appears to be highly conserved in the
animal kingdom. Mammalian cell culture, Ceanorhabitis elegans and Drosophila
melanogaster all appear to be able to be able to undergo an adaptive increase in oxidative
stress tolerance. This increase in tolerance in all cases appears to at least partially be
dependent on an increase in the expression of 20S proteasome which is regulated by the
Nrf2/Skn-1/Cnc-C transcription factor.

159

CHAPTER 8: MATERIALS AND METHODS

This section describes the experimental methods used both in published and unpublished
work, as well as detailing the materials used. In addition it describes a novel technique
which we developed to label proteins for proteolysis assays in a manner that is free from
the use of radioactive material. This technique is published in Pickering, A.M., and
Davies, K.J.A. (2012). A simple fluorescence labeling method for studies of protein
oxidation, protein modification, and proteolysis. Free radical biology & medicine 52,
239-246 (Pickering and Davies, 2012) and for which we have filed a patent with the
University of Southern California.

I. Basic techniques

Materials

All materials were purchased from VWR unless otherwise stated. Murine Embryonic
Fibroblasts (MEF), were purchased from ATCC (Manassas, VA) catalog #CRL-2214. In
addition Wild-type MEF and Pa28αβγ
-
MEF cells which were a kind donation by Tanaka
et al (Yamano et al., 2002) were used. HT1080 Human fibroblast cells (catalog #CCL-
121) were purchased from ATCC (Manassas, VA). Cells were grown in Dulbecco’s
Modified Eagle’s Medium (DMEM), catalog #10-013-CV, purchased from Mediatech
(Manassas, VA) and supplemented with 10% Fetal Bovine Serum (catalog #SH30070.03)

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purchased from Hyclone (Logan, UT); This is henceforth be referred to as ‘complete
media.’ Cells were typically incubated at 37
o
C under 5% CO
2
and ambient oxygen.

Hydrogen peroxide adaptation

MEF cells were grown to 20% confluence (≈2,000,000 cells) in 75 cm
2
flasks in 10 ml
of complete media and pre-treated with 1-10 μM H
2
O
2
(5-50 μM H
2
O
2
per 10
6
cells/ml,
which we found to be the adaptive range) to cause adaptation to oxidative stress. Cells
were incubated for one hour at 37
o
C under 5% CO
2
. Cells were then washed once with
Phosphate buffered saline (PBS), which was finally replaced with fresh complete media.

Western blots

MEF cells were harvested from 75 cm
2
flasks by trypsinization. Cells were washed twice
with PBS to remove trypsin and then lysed in RIPA buffer (catalog #89901) purchased
from Thermo Fisher (Waltham, MA) and supplemented with protease inhibitor cocktail
(catalog #11836170001) purchased from Roche (Nutley, NJ). Protein was quantified with
the BCA Protein Assay Kit from Pierce (Rockford, IL) according to the manufacturer's
instructions. For Western analysis, 40 μg - 5 g (typically 20μg) of protein was run on
SDS–PAGE and transferred to a PVDF membrane. Using standard western blot
techniques, membranes were treated with the following proteasome/Pa28 subunit
antibodies purchased from Biomol (Plymouth Meeting, PA): ant-β1i antibody (catalog

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#PW8205-0100) at 1/10,000 dilution, anti-β2i antibody (Product code: PW8150-0100) at
1/10,000 dilution, anti-β5i antibody (catalog #PW8200-0100) at 1/10,000 dilution, anti-
β2 antibody (catalog #PW9300-0025) at 1/10,000 dilution, anti-Pa28α antibody (catalog
#PW8185-0100) at 1/10,000 dilution, anti-Pa28β antibody (catalog #PW8240-0100) at
1/10,000 dilution, anti-α4 antibody (catalog # PW8120-0025) at 1/10,000 dilution.
Other membranes were probed with anti-β1 antibody (catalog # sc-67345) at 1/2000
dilutation, or anti-α3 antibody (catalog #sc-58414) at 1/2000 dilution (both) purchased
from Santa Cruz Biotechnology (Santa Cruz, CA) or anti-Nrf2 antibody (catalog # sc-
722) from SantaCruz Biotechnology (Santa Cruz, CA, USA), anti-β-tubulin (catalog#
5661) purchased from Millipore (Billerica, MA) at 1/10,000 dilution. In addition some
membranes were treated with the C.elegans antibodies anti-Pas-7 purchased from
developmental studies hybridoma bank (Iowa City, IA, USA) at a ½ dilution and anti-
S5A (catalog # ab56851) at 1/1000 purchased from abcam (Cambridge, MA, USA).
Other membranes were treated with the drosophila antibodies anti-p42E (catalog #sc-
65757) at 1/500 and an antibody against the α subunits of the 20S protesome (catalog
#sc-65755) at 1/500 both purchased from SantaCruz Biotechnology (Santa Cruz, CA,
USA). In Western blotting experiments the blocking buffer employed was
Startingblock™ buffer (catalog #37539) purchased from Thermo Fisher (Waltham, MA)
and the Wash buffer was 1x PBS containing 0.1% Tween 20. An enhanced
chemiluminescence kit, from Pierce (Rockford, IL), was used for chemiluminescent
detection and membranes were developed onto either Kodak Biomax films (purchased
from VWR; West Chester, PA) using the Kodak GBX developing system purchased from

162

VWR (West Chester, PA) or detected using the biospectrum imaging system from UVP
(Upland, CA).

siRNA ‘knock-down’ treatments

siRNA was purchased from two different companies. Custom β5 and S4 siRNA, as well
as the relevant control (non-silencing) siRNA’s were purchased from Qiagen (Huntsville,
Al). MEF were grown to 50% confluence in 75 cm
2
flasks then treated with 3.2 ml of
EC-R buffer (Qiagen catalog # 1024492) containing 750 nM of siRNA and 5.6%
transfection reagent (Qiagen product #1024491) for which complex formation had been
permitted by gently vortexing for 10s and incubated with mild agitation for 10 minutes
prior to treatment. Three hours after treatment media was removed and replaced with
fresh complete media. Other siRNA’s were purchased from Santa Cruz biotechnology
(Santa Cruz, CA). These included: β1 (catalog #sc-62865), Lmp2 (catalog #sc-35821),
Pa28α (catalog #sc-151977), Nrf2 (catalog # sc-37049) and the relevant control siRNA
(catalog #sc-29528). For experiments with these siRNA’s, MEF were seeded at a
concentration of 100,000 cells per well in 6 well plates. One day after seeding, cells were
treated with 200 l of DMEM containing 500 μM siRNA and 2.5% transfection reagent
(catalog #sc-29528). Complex formation had been permitted by incubation with mild
agitation for 30 minutes

163

prior to treatment. Cell media was later supplemented with 800 l of complete media 5
hours after treatment and media was replaced 24 hours after treatment.

Fluorpeptide proteolytic assays

MEF were harvested by cell scraping in phosphate buffer, Cells were then re-suspended
in 50 mM Tris, 25 mM KCl, 10 mM NaCl, 1 mM MgCl
2
, (pH 7.5) and lysed by 3 freeze-
thaw cycles. Protein was quantified with the Bradford assay kit (catalog #2740) from
Merck (Whitehouse Station, NJ) according to the manufacturer's instructions. From 0.02
to 5.0 g of cell lysate was then transferred to 96 well plates. Next, 2 μM of either N-
Succinyl-Leu-Leu-Val-Tyr-AMC (catalog # 80053-860) purchased from VWR (Chester,
PA, USA); or Z-Leu-Leu-Gly-AMC (Product code: ZW9345-0005) or Bz-Val-Gly-Arg-
AMC (catalog #BW9375-0005) both purchased from Biomol (Plymouth Meeting, PA),
were added to the plates. Plates were then incubated at 37
o
C and mixed periodically at
300 rpm for 4 hours during which time, fluorescence readings were taken at 10 minute
intervals using an excitation wavelength of 355nm and an emission of 444 nm.
Fluoresence was subsequently converted from Arbitrary Units to moles of free AMC
using an AMC standard curve derived through Fluorescence readings of known amounts
of AMC (catalog #164545), purchased from Merck (Whitehouse Station, NJ).
Background fluorescence was also subtracted from the results. In some inhibition
experiments, 30 minutes prior to incubation and addition of substrates, plates were treated

164

with 1 μM of the proteasome inhibitors MG312 (catalog #474790) or lactacystin (catalog
#426100), both purchased from Merck (Whitehouse Station, NJ).

BrdU assay for DNA replication and cell division

Bromodeoxyuridine (BrdU) is a synthetic thymidine analogue, which can be incorporated
into newly synthesized DNA. The BrdU assay provides a means of determining degree of
DNA replication as an indirect measure of cell division. The assay was performed as
described in the product (catalog #2750) manual from Millipore (Billerica, MA). Briefly,
cells were seeded at 100,000 cells per ml in 96 well plates, BrdU reagent was then added
and samples were incubated for 24 hours at 37
o
C under 5% CO
2
. Samples were fixed by
addition of fixing solution and incubated at 25
o
C for 30 minutes. The plates were washed
three times with wash buffer, incubated for one hour at 25
o
C with monoclonal BrdU
antibody, and washed again three times with wash buffer. Then samples were treated
with peroxidase conjugate secondary antibody, incubated for a further hour at 25
o
C, and
then washed three times with wash buffer. BrdU incorporation was detected by addition
of peroxidase substrate. Detection was performed at a wavelength of 450 nm using a
spectrophotometer (Spectromax 250) purchased from Molecular Devices (Ramsey, MN).

165

Cell counting assay

Cells were seeded at 100,000 cells per ml in 24 well plates. Cells were harvested 24 hours
after seeding, using trypsinization and 100 μl of cell suspension was combined with 10
ml of diluent isoton (catalog #8546719) purchased from Beckman Coulter (Fullerton,
CA) in a cuvette. Cell counts were obtained using a Cell Counter purchased from
Beckman Coulter (Fullerton, CA). Three cuvettes were prepared for each sample and
resulting cell densities were calculated as means ± SE.

Induction or inhibition of Nrf2

MEF cells were grown to 5% confluence and treated with varying concentrations of Nrf2
inducers. DL-sulforaphane (catalog #S2441-5mg) or curcumin (catalog #C1386-5G)
from Sigma Aldrich (St Louis, MO, USA). Lipoic Acid (catalog #L1089) was purchased
from Spectrum Chemicals (Gardena, CA, USA), dissolved in
N,N,Dimethylformaldehyde, and combined with complete media at a final concentration
of 0.1%; and a comparable concentration of N,N,Dimethylformaldehyde was added to
control cells. Curcumin was dissolved in ethanol and combined with complete media at a
final concentration of 0.1%, and a comparable concentration of ethanol was added to
control cells. In some assays cells were treated with the Nrf2 inhibitor all-Trans-retinoic
acid (catalog #R2625-100MG) purchased from Sigma Aldrich (St Louis, MO, USA).

166

Trans-retinoic acid was dissolved in ethanol and combined with complete media at a final
concentration of 0.1%; a comparable concentration of ethanol was added to control cells.

Chromatin immunoprecipitation (ChIP) assay

Four million cells were prepared at 10% confluence, the cells were exposed to either 0 or
1μM H₂O₂ for 1h. ChIP analysis was performed using the reagents and methods provided
in a Chromatin Immunoprecipitation Assay Kit (catalogue# 17-295) purchased from
Millipore (Temecula, CA, USA). Briefly cells were cross-linked with 1% formaldehyde
for 10 minutes, washed twice with PBS, dislodged through scraping and re-suspended in
1ml of 1% SDS lysis buffer containing protease inhibitor. Samples were sonicated using
10 bursts of 5 seconds, output of 50 watts (Branson Sonifier 140, Branson Ultrasonic,
Danbury, CT), and then centrifuged at 13,000g for 10 minutes. The supernatant was
removed and diluted in a 10 fold excess of ChIP dilution buffer. (1% of samples were
removed at this point to later form the input samples). Samples were pre-cleared using a
30 minute incubation with 30μl/ml of Salmon Sperm DNA/Protein A Agarose Slurry.
Samples were then incubated for 1h with 8μg/ml of Nrf2 antibody (catalog # sc-13032)
purchased from Santacruz Biotechnology (Santa Cruz, CA, USA) then 30μl/ml of
Salmon Sperm DNA/Protein A Agarose slurry was added and samples were incubated
overnight at 4
o
C under gentle agitation. After this, the bead slurry was subjected to
sequential 10 minute washes with Low Salt Immune Complex, High Salt Immune
Complex, LiCl Immune complex and TE buffer. Samples were detached from the bead

167

slurry with two washes of 250μl of 1% SDS 0.1M NaHCO
3
, then reverse cross-linked by
incubation with 200μM of NaCl for 4h at 65
o
C. 10μM EDTA, 40µM Tris-HCl, and 20μg
of Proteinase K was then added to the samples and samples were incubated for 1h at
45°C. DNA was isolated and purified from the samples using phenol-chloroform-isoacyl
alcohol. PCR was then performed on samples as described below. 5µl of DNA from each
sample was combined with 15µl of the PCR SyBr Green Master Mix (catalog # 4367659)
purchased from Applied Biosystems (Warrington, UK), 1.5µL each of 5µM working
solutions of forward and reverse PSMB5 primers designed by Kwak et al (Kwak et al.,
2003) (catalog # 2110654) and purchased from Invitrogen (Carlsbad, CA, USA), and 7µl
of DNAse/RNAse-free ddH
2
O. The forward primer sequence used was
CAGACCGGCGCTGGTATTTAGAGG and the reverse primer sequence was
TAGCCAGCGCCATGTTTAGCAAGG. PCR was carried out in a 7500 Real Time PCR
System device from Applied Biosystems, using an annealing temperature of 61
o
C and an
extension temperature of 72
o
C, for a total of 55 cycles. PCR products were then
examined on a 1% agarose gel containing 0.001% ethidium bromide.

Immunoprecipitation

Protein A Sepharose CL-4B beads (catalogue # 17-0780-01) from GE Healthcare (Little
Chalfont, UK), were washed 3 times with pbs and incubated in PBS + 3% BSA 24 h
prior to assay. MEF cells were cultured to 10% confluence in 75-225cm
2
flasks. Cells
were washed with PBS twice and harvested through scarping in PBS. Cells were lyzed

168

with 3 freeze-thaw cycles. Cell debris was removed by centrifugation at 10,000g for 15
minutes, all subsequent steps were performed at 4oC. Samples were incubated for 30mins
with either anti-β-tubulin (catalogue # 5661) from Millipore (Billerica, Massachusetts,
USA) or anti-porin (catalogue # ab15895) from Abcam (Cambridge, MA, USA), to
remove non-specific binding. Samples were then pre-cleared with 30min incubation with
100ul of sepharose beads followed by centrifugation and removal of beads. Samples were
then incubated either with PA28α antibody (catalogue #PW8185-0100) from Enzo Life
sciences, (Plymouth Meeting, PA, USA), anti-Pa200 (catalogue # ab5620) purchased
from Abcam (Cambridge, MA, USA), anti-Pa28γ (catalogue # BML-PW8190-0025)
from Enzo Life sciences, (Plymouth Meeting, PA, USA), immunoproteasome subunit
anti-LMP2/β1i antibody (catalogue #ab3328) purchased from Abcam (Cambridge, MA,
USA), 20S proteasome anti-β1 antibody (catalogue #sc-67345) purchased from Santacruz
biotechnology (Santa Cruz, CA, USA) anti-β-tubulin (catalogue # 5661) from Millipore
(Billerica, Massachusetts, USA) or anti-porin (catalogue # ab15895) from Abcam
(Cambridge, MA, USA). For 1hr. 100ul of sepharose beads was added and samples were
incubated for 4hr. Samples were then washed 3 times with PBS then protein was
detached from beads using SDS-PAGE loading Buffer and boiling. Samples were run on
then run on western blots as described previously.

169

Immunocytochemistry

MEF cells were cultured to 50% confluence. Cells were fixed with 4% paraformaldehyde
for 30mins then washed twice for 5mins with PBS. Exogenous protienase activity was
bloched through incubation with 10% methanol; 0.3% H
2
O
2
in PBS for 20minus
followed by two 5min PBS washes. Cells were permealized by incubation with 1% NP40
for 10mins followed by two 5 min washes with PBS. Samples were incubated in
Startingblock™ buffer (catalogue #37539) from Thermo Fisher (Waltham, MA, USA)
for 30mins, then blocking buffer containing 1/1000 dilution of anti-Pa200 for 30mins.
Samples were washed three times with PBS + 0.1% tween for 5 mins. Samples were then
incubated with Biotinulated -Secondary Rabbit antibody (catalogue PK-6101) purchased
from Vector laboratories (Burlingame, CA, USA) in blocking buffer for 30mins. Samples
were washed three times with PBS + 0.1% tween for 5 mins then detected using a DAB
detection kit (catalogue #SK-4100) purchased from Vector laboratories (Burlingame, CA,
USA).

AMC labeling of protein substrates

The protein substrates we used for AMC labeling were as follows: Hemoglobin
purchased from Sigma-Aldrich (St Louis, MO,USA) catalogue #H-2500, Superoxide
Dismutase purchased from Calbiochem (San Diego, CA, USA) catalogue #574594,
Catalase purchased from Calbiochem (San Diego, CA, USA) catalogue #219001, and

170

Bovine Serum Albinum purchased from thermo-Fisher (Waltham, Massachusetts, USA)
catalogue #BP1605-100. In all cases, 5mg of protein were dissolved in 1ml of 0.1M
Hepes buffer to which was added 500μM of AMC (Calbiochem, San Diego, CA, USA,
catalogue #164545), as well as 20mM sodium cyanoborohydride (final concentration)
purchased from Sigma-Aldrich (St Louis, MO, USA, catalogue #S8628-25G). The
solution was incubated at room temperature for 2 hours, then extensively dialyzed though
a 10,000 M.W.C.O centrifugal filter (Millipore, Carrigtwohil, Ireland, catalogue #4321)
and a buffer exchange was performed with proteolysis buffer (50mM Tris/HCl pH7.8,
20mM KCl, 5mM magnesium acetate, 0.5mM DTT). Protein content was then
determined using the BCA assay kit (Thermo Scientific, Rockford, IL, USA, catalogue
#PI-23225) .

[
3
H] Labeling of protein substrates

Tritium-labeled hemoglobin ([
3
H]Hb) and BSA ([
3
H]BSA) were generated in vitro as
previously described (Davies, 2001; Grune et al., 1996; Grune et al., 1995; Shringarpure
et al., 2003; Ullrich et al., 1999) using the [
3
H]formaldehyde and sodium
cyanoborohydrate method of Jentoft and Deaborn (Jentoft and Dearborn, 1979). Proteins
were then extensively dialyzed.

171

Cell culture – murine embryonic fibroblasts

Murine embryonic fibroblasts (MEF) were purchased from ATCC (Manassas, VA, USA,
catalogue #CRL-2214). Cells were grown in Dulbecco’s Modified Eagle’s Medium
(DMEM, Mediatech, Manassas, VA, catalogue #10-013-CV), and supplemented with
10% Fetal Bovine Serum (Hyclone, Logan, UT, catalog #SH30070.03). Cells were
incubated at 37
o
C under 5% CO
2
and ambient oxygen. To generate cell lysates, MEF
were grown to confluence then washed twice with PBS, cells were then scraped using a
cell lifter, and centrifuged at 5,000g for 5 minutes. The cells were then re-suspended in
proteolysis buffer and subjected to 3 freeze-thaw cycles at -20⁰C. The lysates were then
centrifuged at 10,000g for 10 minutes, after which the supernatants were retained (the
pellets discarded) and protein content was determined by BCA assay.

Proteolysis assay – common procedures

Proteolysis was measured by incubation of 1µg of AMC-labeled substrate or [
3
H]-labeled
substrate in 100ul of proteolysis buffer containing either dissolved Trypsin (VWR, West
Chester, PA, USA, catalogue #100504-332), Chymotrypsin (Sigma-Aldrich, MO, USA,
catalogue #C-7762), Pepsin (Thermo-Fisher, Waltham, Massachusetts, USA, catalogue
#P53), Proteinase K (Oncor, Gaithersburg, MD, USA, catalogue #S4508), Purified 20S
proteasome (Biomol, Plymouth Meeting, PA, USA, catalogue #PW8720-0050), or lysate

172

generated from MEF cells as above. In each experiment, pH was adjusted appropriately
for the proteinase being studied, and samples were incubated at 37
o
C for 4 hours.

Proteolysis of AMC-labeled proteins by fluorescence assay

This procedure was used with AMC-labeled proteins. It should be noted that free AMC is
soluble in water, and that it fluoresces strongly. AMC adducted to proteins, by reductive
methylation, does not fluoresce, but when liberated by proteolysis again fluoresces
strongly. During incubations described above under “Proteolysis Assay – Common
Procedures,” fluorescence was measured every 10 minutes at an emission wavelength of
444nM, with excitation at 390nM, in a Fluoroskan Ascent Microplate Fluorometer
(Thermo Fisher, Waltham, Massachusetts, USA, catalogue #5210480). Fluorescence
emission was compared with a standard curve of the fluorescence of known
concentrations of free AMC between 5nM and 5mM to determine moles AMC released
into solution.

Proteolysis of [
3
H]-labeled proteins by radioactive liquid scintillation assay

Following incubations described above under “Proteolysis Assay – Common
Procedures,” remaining intact protein was precipitated by addition of 20% trichloroacetic
acid and 3% BSA (as carrier) as previously described (Grune et al., 2002; Pacifici and
Davies, 1990; Pickering et al., 2010; Reinheckel et al., 2000a; Shringarpure et al., 2003).

173

Percent protein degraded was estimated by release of acid soluble counts into the TCA
supernatants, measured by liquid scintilatation, in which % Protein Degraded = (acid-
soluble counts – background counts) x 100.

SDS PAGE gels

50mg of BSA with 1mM AMC and or 20mM NaCNBH
3
, in 500 μl of hepes buffer for
2hr. Following 25% Nupage loading Dye (Invitrogen, Carlsbad, CA, USA, catalogue#
NP0007) containing 5% 2-mercaptoethanol, was added. Samples were boiled for 3
minutes then added to a 12% Tris-gl;ycine SDS page gel. (VWR, West Chester, PA,
USA, catalogue# 12001-042) and run 1t 80V for 2hr. Flouresence was deretmined using
an excitation wavelength of 36nm. Silver staining was performed using silverSNAP stain
kit II (Waltham, Massachusetts, USA, catalogue# 24612). Staining was performed as
described in the product manual.

II. C.elegans and D. melanogaster techniques

C. elegans culture

Strains were maintained at 20
o
C as described in Brenner et al 1974 (Brenner, 1974). A
bacterial lawn of OP50 Escherichia coli was spread on the plates for feeding. Unless
otherwise stated the N2 Bristol strain was used for experiments.

174

Preparation of age matched C. elegans culture.

We found that egg prepation using an HCl /bleach solution; produced a stress response
itself. As a result we switched to a less aggressive form of egg preparation. In this we
placed 3-5 non-starved adult C. elegans on a plate for 24 h, over which time they would
lay eggs. The C. elegans were then removed and the progeny was used for assays 48h
later.

C. elegans H
2
O
2
adaptation

C. elegans were collected using washing with M9 solution; C. elegans were washed 3
times to remove any contamination from the bacterial lawn. C. elegans were then
transferred to epindorfs containing 1ml of M9 solution containing 0μM, 0.1μM, 1μM, or
10μM of H₂O₂. C. elegans were incubated in this with mild shaking for 1 h then returned
to normal plates for 24 h to permit adaptation to occur.

C. elegans H
2
O
2
challenge

C. elegans were collected using washing with M9 solution; C. elegans were washed 3
times to remove any contamination from the bacterial lawn. C. elegans were then
transferred to epindorfs containing 1ml of M9 solution containing 100μM – 16mM of
H₂O₂. C. elegans were incubated in this with mild shaking for 1 h then returned to normal

175

plates. Survival was then either scored instantly or 24 h later through response to
prodding with a pick. This experiment was typically performed blind which was
accomplished by another member of the group encoding and randomizing the plates
shortly before they were scored.

C. elegans RNAi treatment

C. elegans were cultured on plates containing a bacterial lawn expressing RNAi against
the proteasome subunits Pas-5, Pbs-3, Pbs-5 and Pbs-6 in addition some plates contained
bacteria expressing RNAi against skn-1 or L4444 which severed as a control. C. elegans
were incubated on these plates for 6 h. After this C. elegans were collected using washing
with M9 solution. C. elegans were washed 3 times to remove any contamination from the
bacterial lawn. C. elegans were then transferred to epindorfs containing 1ml of M9
solution ±1μM of H₂O₂. C. elegans were incubated in this with mild shaking for 1 h then
returned to RNAi plates for 18 h. C. elegans were then collected using washing with M9
solution; C. elegans were washed 3 times to remove any contamination from the bacterial
lawn. C. elegans were then transferred to a 48 well plate containing 500ul of M9 solution
+ 8mM of H₂O₂. C. elegans were incubated in this with mild shaking for 1 h then
survival was scored instantly through a score of visible motility.

176

SKN-1::GFP worm imaging

We received the kind donation of a SKN-1-GFP (007) line generated by An and
Blackwell (An and Blackwell, 2003), we then crossed this line with the glo-1 mutant
lines to reduce non-specific fluorescence. We age-matched the C. elegans through egg
preparation as described above. 48 h after egg preparation C. elegans were collected
using washing with M9 solution. C. elegans were washed 3 times to remove any
contamination from the bacterial lawn. C. elegans were then transferred to epindorfs
containing 1ml of M9 solution ±1μM or 10mM of H₂O₂ (depending on the experiment).
C. elegans were incubated in ± H
2
O
2
for 20 minutes. After this C. elegans were paralyzed
through addition of 2,3, butanedione monoxime. C. elegans were then mounted on a slide
coated in an agarose solution to prevent the C. elegans from becoming squashed. Skn-1
nuclear localization was then sored in L3 and L4 C. elegans. To do this the C. elegans
analyzed a microscopre using a 500X – 1000X magnification under GFP illumination.

Drosophila culture

Drosophila were cultured on a standard agar/dextrose/corn meal/yeast media (Ashburner,
1989) at 25
o
C. Unless otherwise stated, W
1118
drosophila were used in all assays.
Drosophila were typically collected over a 48 h period from pre-cleared bottles then
allowed 4 days to mature so that at initiation of assays drosophila were 4-6 days old.

177

Drosophila H
2
O
2
adaptation

Samples of 20 drosophila were transferred to vials containing ½ a Kim-wipe© soaked in
1ml of 5% sucrose as well as 0μM, 10μM, 100μM or 1mM H₂O₂ for 6 h. Drosophila
were then returned to normal vials for 18 h to permit adaptation to occur. After which
drosophila were challenged with a toxic dose of H
2
O
2
. In the case for experiments using
cnc-C
RNAi
drosophila we found for effective adaptation to occur drosophila needed to be
incubated on H₂O₂ Kim-wipes© for 24 h instead of 6 h. In addition because of the longer
incubation time in the experiments with cnc-C
RNAi
drosophila, 10μl of either ethanol or
RU486 was added to the Kim-wipes©.

Drosophila H
2
O
2
challenge assay

Samples of 20 drosophila were transferred to vials containing ½ a Kim-wipe© soaked in
1ml of 5% sucrose and 4.4M H₂O₂. Survival was then scored ever 6 or 8 h following
initiation of challenge. Drosophila were scored as dead once they became completely
immobile.

178

Drosophila RNAi experiments

Drosophila expressing RNAi against 2 proteasome subunits were purchased from the
Vienna Drosophila RNAi center (VDRC, Vienna, Austria). These were Prosβ1
RNAi

(Transformant ID 35923), and Prosβ2
RNAi
(Transformant ID 24749). In addition we
received the kind donation of drosophila expressing RNAi against the cnc-C and dkeap-1
form Dr Dirk Bohman (Sykiotis and Bohmann, 2008, 2011). We crossed males from
these lines (or W
[1118]
as a control) with virgin females containing the Act-GS-255B
driver (Fischer et al., 1988; Ford et al., 2007). Parents were removed 4 days after
initiation of cross. Progeny then collected over a 48 h period once occlusion had
begun.The exception to this is the case of the cnc-C line where a low birthrate required us
to collect over 72 h instead. The Act-GS-255B driver is induced by RU486, drosophila
were cultured in normal vials containing either 10μl of RU486 or ethanol which had been
added to plates and air dried 24 h prior to the assay. Drosophila were incubated on these
plates for 5-7 days. If an adaptation experiment was performed drosophila then 1 day
before the end of the assay, drosophila were transferred in containing ½ a Kim-wipe©
soaked in 1ml of 5% sucrose as well as ± 100μM of H₂O₂ for 6 h. After this drosophila
were returned to RU486/ethanol plates for the remaining 18 h.

179

Western blot analysis

In the case of C. elegans, ≈1000-2000 adult C. elegans were collected per sample using
washing with M9 solution. C. elegans were washed 3 times to remove any contamination
from the bacterial lawn. C. elegans were then transferred into a solution containing RIPA
buffer, (catalog # 89901) from Thermo Fisher (Waltham, MA, USA), supplemented with
protease inhibitor cocktail (catalog #11836170001) from Roche (Nutley, NJ, USA). C.
elegans were frozen once then subjected to 3x10S burst of sonication after which lysis
was performed by 3 cycles incubation on ice for 5 mins followed by vortexing. After this
samples were centrifuged at 10,000g to remove un-lyzed fragments. Protein content was
quantified with a BCA protein assay kit (Pierce, Rockford, IL, USA) according to the
manufacturer's instructions. For Western analysis, 40 g of protein was run on SDS–
PAGE and transferred to PVDF membranes. Using standard Western blot techniques,
membranes were treated with a 20S Proteasome subunit Pas-7 antibody purchased from
developmental studies hybridoma bank (Iowa City, IA, USA) and used at a ½ dilution, or
the 19S proteasome regulator antibody S5A (catalog # ab56851) at 1/1000 purchased
from abcam (Cambridge, MA, USA), or the loading control β-tubulin, (catalogue # 05-
661), purchased from millipore (Billerica, MA, USA) and used at a 1/10,000 dilution.
The blocking buffer employed for Western blotting was Startingblock™ buffer (catalog
# 37539) from Thermo Fisher (Waltham, MA, USA) and the wash buffer was 1x PBS
containing 0.1% Tween 20. An enhanced chemiluminescence kit (Pierce, Rockford, IL)

180

was used for chemiluminescent detection and membranes were analyzed using the
biospectrum imaging system (UVP, Upland, CA, USA).

In the case of D. melanogaster, 10 drosophila were collected per sample. Drosophila
were then transferred into a solution containing RIPA buffer, (catalog # 89901) from
Thermo Fisher (Waltham, MA, USA), supplemented with protease inhibitor cocktail
(catalog #11836170001) from Roche (Nutley, NJ, USA). Drosophila were frozen then
ground-up using an electronic pestle after which lysis was performed by 3 cycles
incubation on ice for 5 mins followed by vortexing. After this samples were centrifuged
at 10,000g to remove unlyzed fragments. Protein content was quantified with the BCA
Protein Assay Kit (Pierce, Rockford, IL, USA) according to the manufacturer's
instructions. For Western analysis, 40 g of protein was run on SDS–PAGE and
transferred to PVDF membranes. Using standard Western blot techniques, membranes
were treated with a 20S Proteasome α subunit antibody (catalog #sc-65755) at 1/500, or
the 19S proteasome regulator antibody p42E (catalog #sc-65757) at 1/500 both purchased
from SantaCruz Biotechnology (Santa Cruz, CA, USA), or the loading control β-tubulin,
(catalogue # 05-661), purchased from millipore (Billerica, MA, USA) and used at a
1/10,000 dilution. The blocking buffer employed for Western blotting was
Startingblock™ buffer (catalog # 37539) from Thermo Fisher (Waltham, MA, USA) and
the wash buffer was 1x PBS containing 0.1% Tween 20. An enhanced

181

chemiluminescence kit (Pierce, Rockford, IL) was used for chemiluminescent detection
and membranes were analyzed using the biospectrum imaging system (UVP, Upland,
CA, USA).

Fluorpeptide proteolytic assays

In the case of C. elegans, ≈100-200 adult C. elegans were collected per sample using
washing with M9 solution; C. elegans were washed 3 times to remove any contamination
from the bacterial lawn. C. elegans were then transferred into a solution containing
proteolysis buffer (50 mM Tris, 25 mM KCl, 10 mM NaCl, 1 mM MgCl
2
, 1 mM DTT
(pH 7.5). C. elegans were frozen once then subjected to 3x10S burst of sonication after
which lysis was performed by 3 freeze-thaw cycles performed through incubation in
either dry ice for 5 mins followed by a room temperature water bath for 5 mins. After
this samples were centrifuged at 10,000g to remove un-lyzed fragments. Protein content
was quantified with the BCA Protein Assay Kit (Pierce, Rockford, IL, USA) according to
the manufacturer's instructions.

In the case of D. melanogaster, 10 drosophila were collected per sample. Drosophila
were then transferred into a solution containing proteolysis buffer (50 mM Tris, 25 mM
KCl, 10 mM NaCl, 1 mM MgCl
2
, 1 mM DTT (pH 7.5). drosophila were frozen once then
ground-up using an electronic pestle after which lysis was performed by 3 freeze-thaw
cycles performed through incubation in either dry ice for 5 mins followed by a room

182

temperature water bath for 5 mins. After this samples were centrifuged at 10,000g to
remove un-lyzed fragments. Protein content was quantified with the BCA protein assay
kit (Pierce, Rockford, IL, USA) according to the manufacturer's instructions.

Once samples were pre-pared 5.0 g of lysate per sample was transferred in triplicate to
96 well plates, and 2 μM of N-succinyl-Leu-Leu-Val-Tyr-AMC (catalog # 80053-860)
purchased from VWR (Chester, PA, USA) was then added to the plates. Plates were
incubated at 37
o
C and mixed at 300 rpm for 4 h. Fluorescence readings were taken at 10
minute intervals using an excitation wavelength of 355nm and an emission of 444 nm.
Following subtraction of background fluorescence, fluoresence units were converted to
moles of free AMC, with reference to an AMC standard curve of known amounts of
AMC (catalog # 164545) purchased from Merck (Whitehouse Station, NJ, USA). In
some experiments, cells were treated with 20 µM of the Proteasome inhibitor lactacystin
(catalog #80052-806) from VWR (Chester, PA, USA), 30 minutes prior to incubation and
addition of substrates. Lactacystin was dissolved in DMSO at a 100X concentration and
combined with samples at a concentration of 0.1%.

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III. Reductive labeling of proteins with the fluorophore AMC

Introduction

The use of
3
H and
14
C labeling of proteins by in vitro reductive methylation has become
an important tool to measure the proteolytic degradation of a wide range of protein
substrates by purified proteolytic enzymes, cell lysates, and cell extracts. Such
3
H and
14
C
labeled protein substrates are also widely used to assess the effects of protein
modifications, such as denaturation, oxidation, methylation, acetylation, etc., on
proteolytic susceptibility and rates of turnover. In addition, the specificity of various
proteolytic enzymes for putative substrates has frequently been tested using
3
H and
14
C
labeled proteins (Chondrogianni et al., 2005; Cuervo and Dice, 2000; Davies, 2001; Dean
et al., 1997; Farber and Levine, 1986; Ferber and Ciechanover, 1986; Friguet et al., 1994;
Grune et al., 1996; Grune et al., 1995; Hershko et al., 1983; Hershko et al., 1984; Jentoft
and Dearborn, 1979; Kirschner and Goldberg, 1983; Netland and Dice, 1985; Pickering
et al., 2010; Scrofano et al., 1998; Shringarpure et al., 2003; Stolzing et al., 2006; Tanaka
et al., 1983; Ullrich et al., 1999). The process of in vitro reductive methylation with
3
H
and
14
C, however, does have a number of drawbacks. The use of radioactive materials,
with all the attendant exposure risks for experimenters and their colleagues, and the
difficulties and ethical considerations of radioactive waste procedures rank high on the
list of drawbacks. Additionally, the costs both of purchasing radionucleotides and of
disposing of them are extremely high. Proteolytic assays with
3
H and
14
C labeled protein

184

substrates require a labor-intensive trichloloracetic acid (TCA) precipitation step, so that
undegraded (TCA-insoluble) proteins can be separated from TCA-soluble degradation
products; This further increases the volume of radioactive waste, limits the number of
samples that may be analyzed, increases experimental error, and forces an absolute
endpoint to the assay with the result that true time courses cannot be measured.

Fluorometric peptidase assays, in which a fluorophore covalently linked to a small
peptide sequence is cleaved by a protease / protienase, provides a solution to all the above
radiolabeling problems, and fluorogenic peptides are widely used to measure peptidase
activities. Such fluorogenic peptidase measurements are based on the increase in
fluoresence as the fluorophore is released from the peptide by proteolytic cleavage. TCA
precipitation is not required, thus enabling continuous readings to be made, as well as
permitting a greater number of assays to be performed. While this technology has been
highly valuable in measuring the cleavage of short peptide sequences (Pickering et al.,
2010; Reinheckel et al., 2000a; Ullrich et al., 1999), it is only a primitive model with
which to test the activities of complete proteinases which target whole proteins rather
than short peptide sequences. Additionally many proteinases are selective for various
modified forms of their protein substrates, and such selectivity cannot be measured by
peptide hydrolysis (Davies, 1987). These problems are even more severe when trying to
measure proteolytic susceptibility in cell lysates or extracts.

185

A solution would seem to be that of adapting the fluorescent labeling technique for
peptides to work with intact proteins, but there has been limited success in modifying this
technology to measure the degradation of whole proteins. Two alternate techniques have
been described for attaching fluorophores onto proteins FITC labeling has been used to
label casein (Twining, 1984), Hemoglobin (De Lumen and Tappel, 1970) and BSA (Voss
et al., 1996). However the labeled substrate is highly unstable to compensate for this the
substrate must be precipitated and stored in 50% Ammonium Sulfate then just prior to
use is transferred out of solution and dissolved in the appropriate reaction buffer, which
presents considerable contamination risks as well as limiting the time over which assays
can be performed (Voss et al., 1996). The assay is further limited by a strong dependency
on pH for the sensitivity of the fluor making assays in of strongly acidic proteases like
pepsin or stongly alkaline proteases like protienase K impractical (Jones et al., 1997). In
addition for measuring proteolysis this technique is, like radio-labeling, limited by the
requirement of TCA precipitation making it labor intensive, error prone and extremely
limited to small experiment size (Twining, 1984). The other technique describes labeling
of either Casein or BSA with BODIPY (Jones et al., 1997). This technique provides a
number of advantages over both FITC labeling and Radio-labeling, though it also has
several issues; among these are that it has a very small separation between excitation and
emission wavelengths (503/512) when compared to other fluors such as AMC (365/444)
which makes it extremely difficult to detect the signal without highly specialized
equipment, The label is relatively large and complex (389Da-634Da, depending on type
of BODIPY label) compared to the small [
3
H]formaldehyde label (32Da) used in radio-

186

labeling, this raises some concerns about modification of the protein during BODIPY
labeling, the Fluor BODIPY is relatively expensive for very small quantities when
compared with other fluors. Finally while BODIPY has benefits for some assays, there
are only a fairly small number of assays for which it has been described for. Thus, most
studies of protein degradation continue to rely on in vitro radio-labeling (
3
H or
14
C) of
purified protein substrates, using the technique of reductive methylation developed by
Jentoft and Dearborn (Jentoft and Dearborn, 1979).

While in vitro radio-labeling of protein substrates is something we would like to avoid, it
occurred to us that reductive methylation remains an efficient and relatively mild
procedure by which to attach a label to a protein, utilizing free amino groups. In
addition, the careful experiments of Jentoft and Dearborn (Jentoft and Dearborn, 1979)
demonstrated the high stability of such adducts, and thousands of studies over the past 30
years have verified the usefulness of reductively methylated protein substrates. We,
therefore, set out to test whether we could take the fluorophore 7-amino-4-
methylcoumarin (AMC), which is a small molecule (MW 175), of the kind that has
proven effective with peptides, and adduct it to protein substrates by reductive
methylation. We describe a novel technique by which an inexpensive and stable AMC
fluorophore-protein complex can be formed both quickly and simply by reductive
methylation. We go on to demonstrate that this technique is applicable to a wide range of
protein substrates, and that it can be used to measure proteolytic susceptibility with high
sensitivity, comparable to that achieved with radio-labeled proteins. Finally, we show that

187

AMC-protein adducts are stable to oxidation and various other denaturing conditions, and
can be used to measure the increased proteolytic susceptibility of modified proteins. Our
technique offers a sensitive, inexpensive, rapid, radiation-free alternative to
3
H or
14
C
labeling that also allows truly continuous monitoring.

Proposed reaction

In this system we chose to use the fluorophore 7-amino-4-methylcoumarin (AMC) which
is a small fluorescent molecule (MW 175) that is commonly used in the substrates of
peptidase activity assays (e.g. Suc-LLVY-AMC). This fluorophore has the advantages of
being very cheaply available as well as easy to store and work with. We wished to test the
idea that the fluorophore could be used as a replacement of radio-labeled formaldehyde
as a means of labeling proteins through an adaptation of the reductive methylation
reactions described by Jentoff et al (Jentoft and Dearborn, 1979) in this the reducing
agent. Sodium cyanoborohydride (NaCNBH
3
), which is commonly used to label
proteins with either H
3
or C
14
linked formaldehyde, was used to label proteins with AMC
by promoting the formation of a carbon nitrogen bond between the exposed amine group
in the AMC molecule and free carboxyl groups the protein (Figure 8.1).

188

Figure 8.1: Proposed reaction between AMC and the free carboxyl groups of a
protein (R-), mediated by sodium cyanoborohydride

Labeling with AMC

We observed a linear correlation between concentration of AMC and fluoresence (Figure
8.2A), this enabled us to convert fluoresence readings into μM of Free AMC in a
solution. We predicted that the incubation of the fluor AMC along with the protein BSA
and the reducing agent NaCNBH
3
should result in a reductive labeling reaction, in which

189

the AMC label becomes attached to the protein. While the AMC molecule is associated
with the protein, we would expect it to quench fluoresence of the AMC molecule as is
observed in short peptide substrate. To test this we incubated AMC with increasing
concentrations of BSA in the presence of NaCNBH
3
. (Figure 8.2B) As predicted we
observed a loss of fluoresence with the addition of BSA to the solution. To be restored
with degradation of BSA as shown later Figure 8.3).

To determine the nature of the interaction between BSA and AMC, we ran an SDS page
gel of both BSA and BSA treated with AMC and NaCNBH
3
(Figure 8.2C). In this we
saw a band in both samples at ≈66kDa (BSA) and when the gel was exposed to an
excitation wavelength of 365nm the band in the sample treated with AMC and NaCNBH
3
fluoresced at ≈440nm compared with no observable fluoresence from the sample lacking
AMC and NaCNBH
3
. This Implies that the Fluor is binding to the protein and that the
binding does not cause significant modification in terms of the size of the protein. We
repeated the experiment with and without NaCNBH
3
(Fig 1d). While we observed similar
protein content based on silver staining, there was a marked reduction in fluoresence in
the absence of AMC. This indicates an important role for the NaCNBH
3
reducing agent in
the reaction. To test if AMC was indeed binding to free-carboxyl groups as we
hypothesized we incubated BSA with N-(3-Dimethylamineopropyl)-N'-
ethylcarbodiimide (Figure 8.2D) which is reported to block free carboxyl groups (Hoare
and Koshland, 1967), We saw with treatment a progressive modification of the protein as
observed by the formation of a second band forming in our gel. This correlated with a

190

progressive reduction in fluorescent label observed in the gel. These results imply that
indeed the reaction is dependent upon exposed carboxyl groups on the protein. Additional
experiments were performed to test the idea that the AMC complex was being
sequestered in hydrophobic pockets by non-covalent interaction. To test this tryptamine
was used as a competitive inhibitor. This resulted in only a small loss (30%) of
association between AMC and BSA implying only a weak role for non-covalent
interactions (data not shown).

For the labeling technique to be useful for proteolysis reactions, the fluorophore should
be mostly quenched when it is bound to the protein and then fluoresce when it is liberated
(as observed in short AMC peptide substrates) , to test this and to further confirm that
labeling does indeed occur we incubated different amounts of AMC either on its own or
with BSA and or with NaCNBH
3
(Figure 8.2A). Here we observed that following AMC
labeling incubations, the remaining free AMC in solution decreased significantly with
addition of the Protein BSA and further with the addition of the reducing agent
NaCNBH
3
though critically just the addition of NaCNBH
3
with BSA did not cause any
decrease in free AMC, as measured by diminished fluorescence, indicating that AMC
was forming adducts with protein substrates (not shown). In addition we demonstrated a
progressive quenching of fluorophore with increasing BSA (Figure 8.2B)

Our expectation that the free aldehyde moiety of AMC should undergo reductive
methylation (catalyzed by NaCNBH
3,
) with free amine groups on target proteins predicts

191

that blockage of such free amines would prevent the formation of AMC-protein adducts.
Experiments with the amine reagent sulfo-NHS-acetate (Figure 8.2C) revealed that a
maximum of 77% of free amine groups could be blocked (some are not accessible to
sulfo-NHS-acetate). Thus, we might expect that as much as 77% of AMC labeling could
blocked in sulfo-NHS-acetate-treated BSA samples. Actual experiments revealed that
some 72% of BSA labeling by reductive methylation with AMC could be blocked with
sulfo-NHS-acetate (Figure 8.2C) which, we suggest, is within reasonable experimental
error.

192

Figure 8.2: AMC can be conjugated to free carboxyl groups on proteins

A. Linear correlation between free AMC concentration, from 100 nM to 1 mM, and
fluorescence. Here variousconcentrations of AMC, dissolved in proteolysis buffer, were
incubated at 37 °C on 96-well plates. Fluorescence was analyzed at an emission
wavelength of 444 nm, with excitation wavelength of 390 nm. Values are means±SE,
n=3. B. Addition of increasing amounts of BSA to AMC in the presence of NaCNBH3
progressively quenches the fluorescence of AMC. Here 0–50 mg of BSA was added to
100 μM AMC and 20 mM NaCNBH
3
and incubated for 1 h at 37 °C. Free AMC content
was determined with reference to a standard curve of known AMC concentrations.
Values are means ± SE, n=3. C. Here 50 mg/ml BSA was incubated with 1 mM AMC in
the presence or absence of 20 mM NaCNBH3 and then run on a 12% SDS–PAGE gel. A
fluorescent BSA–AMC complex was readily observed at ≈66 kDa (the approximate size
of BSA), using an excitation wavelength of 365 nm and an emission wavelength of 444
nm, when all three reagents were present, but could be only faintly discerned in the
absence of NaCNBH3. A silver stain was later performed. D. N-(3-
dimethylaminopropyl)-N′-ethylcarbodiimide (1 ng/ml to 100 μg/ml), which blocks free
carboxyl groups [26], was incubated with 50 mg of BSA for 1 h. BSA was extensively
dialyzed and then prepared as for (c). Increasing concentrations of N-(3-
dimethylaminopropyl)-N′-ethylcarbodiimide caused a progressive decrease in BSA's
electrophoretic mobility and loss of fluorescence at 66 kDa; a representative gel is shown
to the left, and fluorescence is quantified in the graph to the right.

193

Figure 8.2, Continued

194

AMC labeling in proteolysis assays

We next incubated hemoglobin with the protease trypsin, after first incubating the Hb
with NaCNBH
4
alone, AMC alone, or AMC plus NaCNBH
3
then extensively Dialyzed
the samples to remove any Free AMC or NaCNBH
3
(the full labeling procedure). As
shown in Figure 8.3A, trypsin released an extremely large amount of AMC fluorophore
from Hb, removing any remaining doubt that the fluor had actually been successfully
adducted to the protein. Reaction of Hb with AMC alone produced a Hb-AMC
proteolytic substrate with high background release of AMC, and about a six-fold increase
in AMC liberation following incubation with trypsin. In contrast, use of the full labeling
procedure, with NaCNBH
4
to increase the strength of the adduct, produced a more stable
Hb-AMC proteolytic substrate with only one-sixth the background AMC release, but
with an 80-fold increase in AMC liberation after incubation with trypsin (Figure 8.3A).
These results demonstrate that the labeling system works well, and that it can be used to
generate at least one effective proteolytic substrate. To test the broad applicability of the
AMC labeling technique for proteins in general, we attempted to bind the AMC
fluorophore to a range of proteins. We found that AMC could be used to label Bovine
Serum Albinum, Catalase, Hemoglobin, and Superoxide Dismutase (all the proteins we
tried), and that all of the AMC-labeled proteins were effective and sensitive substrates for
proteolysis by trypsin: as measured by release of fluorescent AMC (Figure 8.3B).

195

Figure 8.3: Proteolysis of AMC-labeled proteins by trypsin
A. Incubation of 1 mg/ml hemoglobin with 100 μM AMC and 20 mM NaCNBH3
followed by extensive dialysis produced astable and sensitive substrate for measuring
protease activity, in which 10 μg/ml Hb–AMC was combined with 10 μM trypsin. Free
AMC content was determined with reference to a standard curve of known AMC
concentrations. Values are means ±SE, n=3. B. AMC labeling of BSA, catalase, Hb, or
superoxide dismutase (SOD) generates valid substrates for trypsin digestion, as measured
by liberation of fluorescent AMC. All assay conditions (including trypsin concentration)
were identical to those in A, and each substrate protein was used at a final concentration
of 10 μg/ml. Free AMC content was determined with reference to a standard curve of
known AMC concentrations. Values are means ±SE, n= 3

196

Reliability and usability of label

Effective and reliable proteolytic substrates exhibit linear increases in degradation when
exposed to linear increases in protease concentration (at least over a fairly wide and
useful range), and when substrate concentration is increased in the presence of non-
limiting protease activity. To determine the usefulness and reliability of AMC-labeled
protein substrates, we assayed the AMC release over a wide range of trypsin
concentrations and a wide range of substrate concentrations, using Hb-AMC as a model
substrate. We observed a linear relationship of proteolytic activity (AMC liberation) to
trypsin concentration between 320nM and 1mM of trypsin (Figure 8.4A) and 25ng-2.5μg
of Hb-AMC substrate (Figure 8.4B), when plotted using log-log scales. With these results
we were able to plot a linear regression curves with correlation coefficients close to unity:
indicating excellent statistical reliability.

Although it could already be seen that trypsin-mediated AMC release from AMC-labeled
proteins, reflected protein degradation, and although it was clear that free AMC is
fluorescent whereas the fluorescence of protein-bound AMC is quenched, it was unclear
what particle sizes of degradation products actually produced fluorescent signals. To
study this we partially digested a sample of Hb-AMC. It was important that the Hb-AMC
substrate was only weakly digested so we could be sure that it was not full degraded to
only the smallest size peptides (or amino acids) possible. The data of Figure 8.3C show
Hb-AMC treated with 10μM trypsin, for 24hr at 4
o
C. We confirmed that the substrate

197

was only partially digested by treating it with a trypsin concentration of 100 μM, for 24hr
at 37
o
C, which produced a 100 fold stronger increase in release of fluorescent products
(data not shown). We then suspended aliquots of the partially digested substrate (10μM
trypsin, for 24hr at 4
o
C) in three dialysis tubes with <5kDa, <1kDa and <500Da size
exclusions, and dialyzed each sample against 500 X volumes of proteolysis buffer. In
each case, dialysis caused a dramatic loss of signal. Dialysis through a 500Da filter
caused an ≈80% reduction in signal, while dialysis through a 1kDa filter caused ≈90%
reduction in signal, and dialysis through a 5kDa filter caused ≈95% reduction in signal
(Figure 8.4C). From this we can conclude that the majority (80%) of fluorescent products
are smaller than 500Da, while another 15% are particles between 500Da and 5kDa, and
only some 5% of the signal comes from peptides larger than 5kDa (Figure 8.3C). These
results seem quite consistent with proteolysis assays using radiolabeled protein
substrates, in which a TCA precipitation step is routinely used to precipitate remaining
intact protein, and peptides larger than about 5kDa, so that soluble radioactivity reflects
free amino acids and only very small peptides (Pacifici and Davies, 1990)

We also considered it important to directly compare the sensitivity of proteolytic
measurements using the AMC-labeled substrates we generated to that of traditional radio-
labeled substrates (Jentoft and Dearborn, 1979). Thus, we assessed the percent
degradation of Hb-AMC versus [
3
H]Hb following incubation with various, widely
studied proteolytic systems. The results reveal broadly comparable sensitivity for both
substrates, with trypsin, MEF cell lysates, and purified 20S proteasome (Figure 8.4D).

198

Most importantly, the pattern of degradation for each substrate, as well as the background
signals, appear very similar. Thus, AMC-labeling of proteins appears to generate labeled
substrates that are as suitable as radio-labeled proteins for measurement of proteolytic
susceptibility. Small differences between the two substrates in Figure 8.4 would not
appear to be caused by the particle sizes being measured in each technique. For AMC-
labeled substrates it appears that more than 95% of signal comes from particles smaller
than 5kDa (see Figure 8.2C) and the TCA precipitation step used in measuring
degradation of radio-labeled proteins should also capture most degradation products
smaller than 5kDa (Pacifici and Davies, 1990).

199

Figure 8.4: Protease and substrate titration and particle size of proteolytic
degradation products

A. A linear relationship between the concentration of protease and AMC release is seen
at trypsin concentrations between 320 nM and 1 mM, using an Hb–AMC protein
concentration of 10 μg/ml. B. A linear relationship between the concentration of Hb–
AMC substrate and proteolytic activity (AMC release) is seen between 25 ng and 2.5 μg
of Hb–AMC. All other conditions in both A and B were as described in the legend to
Figure 8.3 and, in both, values are means ±SE, n = 3. C. Dialysis of partially digested
Hb–AMC substrate shows that the majority of liberated fluorescent AMC products
consist of particles smaller than 500 Da. For this experiment, Hb–AMC (10 μg/ml) was
incubated with 10 μM trypsin at 4 °C for 24 h in dialysis tubing, to generate sufficient
fluorescent products to measure but also to preclude complete digestion of the substrate.
Values are means ±SE, n = 4, for which the fluorescence of controls was subtracted. D.
Hb was labeled with AMC, or with tritium, by reductive labeling in both cases, as
described under Materials and methods. Protein degradation was measured in A by AMC
fluorescence and in B by release of acid soluble 3 H counts by liquid scintillation, as
described under Materials and methods. Background fluorescence or radioactivity was
measured in the absence of protease (proteolysis buffer alone), and proteolysis was
measured by increased fluorescence or acid-soluble radioactivity after incubation with 10
μM trypsin, 1 μg/ml purified 20S proteasome, or 150 μg/ml MEF cell lysate. Percentage
degradation of Hb–AMC is reported as the percentage of total fluorescence that could be

200

Figure 8.4, Continued
released from Hb–AMC after exhaustive proteolytic digestion (not shown), whereas
percentage degradation of [
3
H]Hb is reported as the percentage of total (initial)
radioactive counts released into TCA-soluble form by proteolysis. All values are means
±SE, n = 3.

201

Stability of label

Obviously, the stability of AMC-labeled substrates, the resistance of the AMC-protein
linkage to various treatments, and the reproducibility of proteolytic assays after
prolonged storage are important in weighing the usefulness of the technique. To begin
test these matters, we stored Hb-AMC at -20
o
C and then periodically thawed samples and
analyzed both their background release of free AMC (representing undesirable
breakdown of the complex) and their proteolytic degradation during incubation with
trypsin. During repeated trials over 150 days, both the background release of AMC, and
the trypsin-induced proteolysis results varied by less than 15% over the five-month
period, indicating that the substrate was quite stable and that samples can be stored for
long period of time without significant changes in proteolytic susceptibility (Figure
8.5A). As a harsher test of substrate stability we subjected Hb-AMC to repeated freeze
thaw cycles and then measured background release of free AMC. As shown in Figure
8.5B, even repeated freeze-thaw cycles did not significantly affect the stability of the Hb-
AMC complex.

Proteolytic substrates are often subjected to various modifying or denaturing conditions
to test for effects on proteolytic susceptibility, so we considered it important to test the
stability of AMC-labeled substrates over a range of harsh conditions. Hb-AMC was
almost completely stable to incubation in dilute HCl at pH 4), 10% 2-mercaptoethanol,

202

free-thawing at -80⁰C, 50% methanol, 1mM peroxynitrite, or 1mM H
2
O
2
. Even boiling
(100⁰C) for 60 minutes only caused a 3.1% breakdown of the Hb-AMC complex (Figure
8.4C).

Figure 8.5: Stability of AMC-labeled hemoglobin after frozen storage or
denaturation
A. Hb–AMC was stored at −20 °C for up to 21 weeks. At various time points, samples
were thawed, and measurements of both background fluorescence (release of free AMC
from the Hb–AMC complex) and liberation of fluorescent AMC by proteolytic digestion
with trypsin were made, as described for Figure 8.4. B. The stability of Hb–AMC was
tested with repeated −50 °C freeze–thaw cycles, by measuring release of free AMC from
the Hb–AMC complex (background fluorescence). C. Hb–AMC was incubated for 60
min in dilute HCl at pH 4, 10% 2-mercaptoethanol, 70% methanol, 1 mM peroxynitrite,
or 1 mM H
2
O
2
or was boiled at 100 °C for 60 min or was subjected to freeze–thaw at−80
°C. Release of free AMC from the Hb–AMC complex (background fluorescence) was
then measured in comparison with control (untreated Hb–AMC). Values are means ± SE,
n = 3

203

Tolerance to pH of label

While many proteolytic enzymes have pH optima in the neutral to slightly alkaline range,
others are ‘designed’ to function under strongly acidic or alkaline conditions. We,
therefore, needed to test both the fluorescent properties of free AMC over a wide pH
range, as well as the stability of protein-AMC complexes. The fluorescence of free AMC
was unaffected by mildly acidic or alkaline conditions in a broad range from pH 3-11;
highly acidic (below pH 2) or alkaline (above pH 11) conditions, however, significantly
decreased AMC fluorescence (Fig. 8.6A). It should be noted that the fluorescence
quenching effects of strong acid or base were transient, and that AMC fluorescence
returned to normal levels when pH was neutralized (not shown).

We next wanted to determine the stability of protein-AMC adducts over the same broad
range of pH. For these experiments, Hb-AMC was incubated for 4 hr under the same pH
conditions used in Figure 8.6A, after which the pH of each sample was readjusted to pH
7.8 in order to assess the stability of the Hb-AMC complex independent of any possible
quenching effects of pH on the fluorophore. We found that the Hb-AMC complex was
highly stable over the entire range from pH 1 - 12, with less than a 0.2% decrease in
stability observed under any condition (Figure 8.6B).

Having established both the stability of AMC fluorescence, and the stability of the
protein-AMC adduct, over a wide pH range, we next wished to test the viability of

204

protein-AMC complexes as substrates for proteases with widely different pH optima. As
shown in Figure 8.6C, Hb-AMC proved to be an excellent substrate for proteolysis with
enzymes as diverse as pepsin at pH 2, proteinase K at pH 11, and trypsin or chymotrypsin
at pH 7.8.

205

Figure 8.6: pH profile of fluorescence, stability, and proteolytic susceptibility of free
AMC and Hb–AMC
A. The fluorescence of free AMC was measured in proteolysis buffer over a wide range
of pH conditions. B. Samples of Hb–AMC were incubated over a range of pH conditions
for 4 h. The pH of each sample was then adjusted to pH 7.8 and AMC fluorescence was
measured. Results are expressed as a percentage of total AMC label originally
incorporated into the Hb–AMC complex, which was (separately) assessed by exhaustive
proteolytic digestion of Hb–AMC, by incubation with 500 μM trypsin for 4 h. C. Hb–
AMC was incubated with 100 μM trypsin, 10 μM chymotrypsin, 100 μM pepsin, or 100
μM proteinase K (at the pH shown for each protease) for 4 h at 37 °C and proteolysis was
measured by AMC release, as described in the legend to Figure 8.4. Values are means
±SE, n= 3.

206

Synthesis of oxidized substrates

A highly important aspect of proteolysis research centers around the specificity of
proteolytic enzymes and the ability of intracellular proteolytic systems to selectively
degrade modified or abnormal forms of protein substrates, while not attacking normal
proteins. Thus, while digestive enzymes such as trypsin, chymotrypsin, and elastase are
very efficient at degrading both normal and modified proteins, major intracellular
proteolytic enzymes, such as the Proteasome (Davies, 2001; Pickering et al., 2010) and
the Lon protease (Bota and Davies, 2002) exhibit little activity against normal proteins
while avidly degrading their modified or damaged forms. The landmark paper of Jentoft
and Dearborn (Jentoft and Dearborn, 1979) demonstrated that reductive methylation is a
relatively mild treatment and their work, backed-up by thousands of studies by other
researchers in the past 30 years have verified that radio-labeling proteins (by reductive
methylation) generates protein substrates that are not extensively modified or denatured.
Nevertheless, the use of reductive methylation in our AMC-labeling technique has not
been previously studied and, while the AMC fluor is rather small (mw 175), we had to be
concerned that AMC labeling of proteins might causes a degree of denaturation that
would increase the proteolytic susceptibility of normal proteins, while making it harder to
determine if intentional (experimental) modifications to proteins affect their degradation.
For a labeling technique to be useful in this regard, one would hope to see only minor
degradation of the ‘normal’ labeled protein but significantly increased degradation of a
suitably modified or denatured form by intracellular proteases.

207

To test this we incubated both control and oxidized forms of Hb-AMC and BSA-AMC
with purified 20S proteasome which is well known to selectively degrade oxidized
proteins (Davies, 1987, 2001; Pacifici and Davies, 1990; Shringarpure et al., 2003). Our
results show that the unoxidized forms of BSA-AMC and Hb-AMC were rather poor
substrates for the purified proteasome, but the susceptibility to proteasomal degradation
of BSA-AMC increased some four-fold following mild oxidation with H
2
O
2
, whereas
that of Hb-AMC increased by more than 300-fold (Figure 8.7A). We additionally tested
oxidation of Hb-AMC by peroxynitrite, and a number of other protein denaturing
treatments including, boiling, freezing, low pH, methanol, and 2-mercaptoethanol. Both
untreated (control) Hb-AMC and the variously treated Hb-AMC samples were then
incubated with lysates of MEF cells for measurements of proteolysis. Cell lysates and
extracts (which contain proteasome and many other intracellular proteolytic enzymes) are
widely employed in many studies of intracellular proteolytic susceptibility (Davies, 1987;
Davies and Goldberg, 1987a; Grune et al., 1996; Pickering et al., 2010). As shown in
Figure 8.7B, the modified forms of Hb-AMC generated by mercaptoethanol,
peroxynitrite, methanol, freeze-thawing, hydrogen peroxide, HCl, or boiling, were
allsignificantly better substrates for degradation during incubation with MEF cell extracts
than was the unmodified (control) Hb-AMC.

208

Figure 8.7: Proteolytic susceptibility of modified AMC-labeled proteins
A. The capacity of the 20S proteasome to degrade both the native and the oxidized forms
of Hb–AMC and BSA–AMC was measured. For both assays, 1 μg/ml of purified 20S
proteasome was combined with 10 μg/ml Hb–AMC, Hbox–AMC, BSA–AMC, or
BSAox–AMC and incubated for 4 h at 37 °C. Protein degradation was then measured as
per Fig. 3. Hbox–AMC and BSAox–AMC were prepared by treating Hb–AMC and
BSA–AMC with 1.0 mM H
2
O
2
followed by extensive dialysis. B. The capacity of MEF
cell lysates to degrade various modified forms of Hb–AMC was measured. Hb–AMC was
modified by incubation with dilute HCl at pH 4, 10% 2-mercaptoethanol, 70% methanol,
1 mM peroxynitrite, or 1 mM H
2
O
2
or was boiled at 100 °C for 60 min or was subjected
to freeze–thaw at −80 °C. The substrates were then extensively dialyzed and incubated
with 150 μg/ml MEF cell lysates for 4 h. Values are means ±SE, n= 3.

209

Summary

We tested labeling of proteins with the fluorophore, 7-amino-4-methylcoumarin (AMC),
a small fluorescent molecule (MW 175) that is commonly used to manufacture substrates
for peptidase activity assays (e.g. Suc-LLVY-AMC). This fluorophore has the advantages
of being strongly fluorescent when free in solution, very cheap to purchase, freely
available, as well as easy to store and work with. Sodium cyanoborohydride (NaCNBH
3
)
was used to label proteins with AMC as a reducing agent to induce the formation of a
carbon nitrogen bond between the aldehyde group in the AMC molecule and epsilon
amino groups of free lysine residues (as well as the amino terminus) of the target protein
(Figure 1).

Our studies describe a novel radio-isotope free technique of in vitro protein labeling to
generate a substrate for proteolysis. The technique we describe is a modification of the
labeling technique described originally by Means and Feeney in 1968 (Means and
Feeney, 1968) which was subsequently adapted by Rice and Means in 1971 (Rice and
Means, 1971) and then Jentoft and Dearborn in 1979 (Jentoft and Dearborn, 1979) In
which they described a technique by which the carboxyl group in either [
14
C] or [
3
H]
formaldehyde would form a covalent bond with the exposed amine groups of the protein
to be labeled, in a lysine-specific manner using either the reducing agent NaBH
4
or the
milder variant NaCNBH
3
.

210

We hypothesized that the same technique could be used with the carboxyl group of 7-
Amino-4-Methylcoumarin (AMC). AMC is cheaply available, highly stable and
commonly used as the fluorescent tag of short peptide protease substrates, some
examples would be Suc-LLVY-AMC which is commonly used as a measure of cellular
chymotrypsin capacity (Lipson et al., 2008; Pacifici and Davies, 1990; Reinheckel et al.,
2000a), DEVD-AMC is used to detect caspase-3 activity as marker of cellular apoptosis
(Jones et al., 1997), Ac-Arg-Gly-Lys(Ac)-AMC is used to detect histone deacetylase
activity (Wegener et al., 2003), as well as 100’s of other peptide substrates designed to
different protease specificities.

We demonstrated here that we could indeed use reductive labeling to attach the fluor
AMC onto a range of large proteins and that this produced protein substrates which
could measure protease activity with comparable sensitivity to traditional radio-labeling
for a range of different proteases and proteinases. We then demonstrated that the labeling
technique was stable under both highly acidic and highly alkaline conditions as well as
being viable as a substrate under these conditions. We also demonstrated that the
substrate could be stored stably for a long period of time at -20
o
C and could be subjected
to freeze thaw cycles with little effect on stability.

An important consideration for a protein substrates is that many biological enzymes have
a selective preference for degrading oxidized or denatured forms of proteins (Davies,
1987). Measuring oxidized or denatured protein degradation against non-oxidized or non-

211

denatured degradation is an important tool in proteomics (Chondrogianni et al., 2005;
Davies and Delsignore, 1987; Dean et al., 1997; Farber and Levine, 1986; Ferber and
Ciechanover, 1986; Friguet et al., 1994; Scrofano et al., 1998; Stolzing et al., 2006).
While this has been used reliably with traditional radio-isotope labeling, because of the
slightly larger reporter molecule used in AMC labeling compared to [H
3
]formaldehyde
labeling there was a concern that the labeling process might cause a degree of
denaturation or distortion of the protein and as a result prevent measurement of denatured
against non-denatured proteinase specificity. As a result we tested and demonstrated that
when we oxidized or denatured the substrate there was a measurable preference for the
denatured or oxidized substrate comparable to traditional radio-labeling (Figure 8).

We demonstrate in this paper that AMC labeling appears to generate substrates which are
comparable to
3
H or
14
C labeled substrates in terms of versatility, stability and
reproducibility. On top of this we believe it to have several advantages over radiolabeling
in terms of safety, labor and cost. Foremost among the benefits is the removal of radio-
isotopes which are dangerous to use, costly to dispose of and often require permits to
store. In terms of labor requirement, traditional radio-labeled protease assays require that
at the endpoint of the assay, Trichloroacetic acid and BSA are add to each sample, then
each sample is centrifuged and the supernatant transferred to scintillation vials for
quantification which is highly work intensive so limiting assay size to ≈12-24 samples by
comparison fluoresense assays may be performed trivially on 96-well plates. In addition
the manual removal of supernatant from the TCA precipitated samples in radiolabeled

212

assays is often not complete so adding an additional error factor, Finally the fluor AMC is
relatively cheap, compared with radio-label formaldehyde. This makes the labeling
process approximately 40 times cheaper.

213

CHAPTER 9: DISCUSSION

I. The proteasome in oxidative stress adaptation

My studies published in Pickering et al 2010 (Pickering et al., 2010) indicate that the 20S
proteasome, the immunoproteasome and the Pa28αβ (11S) regulator all play major roles
in the degradation of oxidized proteins. In comparison, the 26S proteasome does not
seem to be involved. We also find that the immunoproteasome is at least as capable of
degrading oxidized proteins as is the 20S proteasome. We go on to demonstrate that the
proteasome is a highly plastic system under mild oxidative stress. The 20S proteasome,
the immunoproteasome and the Pa28αβ regulator are all induced during transient
adaptation to oxidative stress. Furthermore, all of these proteins were demonstrated to
provide significant contributions to adaptation and increased tolerance to oxidative stress.
We also provide new evidence of a highly significant role for the immunoproteasome in
stress adaptation.

During adaptation to H
2
O
2
, the proteasome undergoes a two-stage response: an initial
direct activation of pre-existing proteasomes during the first 1 h, followed by a much
slower de novo synthesis of the 20S proteasome, immunoproteasome and Pa28αβ
subunits. After 24 h, the cellular capacity to degrade oxidized proteins is increased more
than 3-fold, and essentially all of this increase can be blocked by proteasome inhibitors.

214

These results demonstrate that proteasome is highly responsive to oxidative stress, being
both activated and induced under stress-adaptive conditions.

We find in a study published in Grune et al 2011 (Grune et al., 2011), which I co-
authored, that the initial protein synthesis independent response occurs through
disassociation of existing 26S proteasome. In this H₂O₂ treatment causes the dissociation
of the 19S regulator from existing 26S proteasome. This causes a temporary increase in
the capacity of the cell to selectivity degrade oxidized proteins. 3-5 h after the initiation
of the treatment the 19S regulator will re-associate so enabling normal cell function to
resume. We demonstrated that the Hsp70 regulator plays an important role in enabling
the 19S regulator to re-associate. Another lab has also shown a similar effect, and
demonstrated that the protein Ecm29 plays an important role in inducing the initial
disassociation of the 26S proteasome (Wang et al., 2010). I also have un-published work
showing that with the disassociation of the 26S proteasome there is a rise in association
between the Pa28αβ regulator and the 20S proteasome. This could also be an important
contributing factor in the increase in the adaptive increase in the capacity of cells to
degrade oxidized proteins.

The proteasomes can only be inactivated by H
2
O
2
concentrations much higher than those
used in the present studies. Studies of purified 20S and 26S proteasomes, and intact cell
studies, show that the 20S proteasome is rather resistant to oxidation, whereas the 26S
proteasome is extremely sensitive. In fact, the IC
50
for 26S inactivation by peroxynitrite,

215

hypochlorite or H
2
O
2
is an order of magnitude lower than that of the 20S proteasome
(Reinheckel et al., 1998). Despite the relative resistance of 20S proteasome to direct
oxidative inactivation, it is interesting to note that Shang and Taylor (Shang and Taylor,
1995) reported that even relatively low levels of H
2
O
2
will inactivate both the E1 and E2
enzymes of the ubiquitylation pathway, owing to highly redox-sensitive thiol groups that
are required for activity, thus diminishing further the importance of ATP- and ubiquitin-
stimulated proteolysis (26S proteasome) in the degradation of oxidized proteins.
Recently, Midicherla and Goldberg (Medicherla and Goldberg, 2008) suggested that
yeast degrade newly synthesized oxidized proteins in an ATP- and ubiquitin-stimulated
pathway. It is possible that the degradation of newly synthesized proteins may be a
special case. It is also possible (although unlikely) that yeast may handle oxidized
proteins differently from the mammalian cells which we have studied. We, and several
other groups, have repeatedly shown that oxidized proteins are degraded, by the
proteasome in the cytoplasm and nucleus of mammalian cells, by an ATP- and ubiquitin-
independent mechanism (Chondrogianni et al., 2003; Davies, 1986, 1987, 2001; Fagan et
al., 1986; Fucci et al., 1983; Grune et al., 1996; Keller et al., 2005; Pacifici and Davies,
1990; Pacifici et al., 1993; Reinheckel et al., 2000a; Shang and Taylor, 1995;
Shringarpure et al., 2003; Ullrich et al., 1999; Whittier et al., 2004), and the present
studies strongly support this view.

The immunoproteasome has long been considered as a proteasome variant that generates
peptides for MHC class I processing. Although the Davies lab had suggested previously

216

that oxidation might be a common protein modification that the immunoproteasome
might recognize, known as the ‘PrOxI hypothesis’, and recent data show that
immunoproteasome can be induced by oxidative stress (Ferrington et al., 2005;
Ferrington et al., 2008; Husom et al., 2004; Kotamraju et al., 2006; Kotamraju et al.,
2003; Thomas et al., 2007), there has been no direct demonstration that the
immunoproteasome can truly degrade oxidized proteins until now. The present studies
may even indicate (although more detailed studies are needed) that the
immunoproteasome may actually be slightly more efficient than the 20S proteasome in
recognizing the oxidatively modified forms of protein substrates such as Hb and ezrin.
We suggest that the PrOxI hypothesis (Teoh and Davies, 2004), which proposes that
some fraction of all intracellular proteins undergoes oxidation, with subsequent
processing for MHC class I by the immunoproteasome, now deserves much greater
scrutiny and serious testing.

Although induction of the 20S proteasome, immunoproteasome and Pa28αβ regulator
synthesis during oxidative stress adaptation is certainly interesting, the important
question is whether such induced proteolytic capacities actually contribute to the
increased oxidative stress tolerance of adapted cells. My data reveals that the increased
oxidative stress tolerance (as measured by BrdU incorporation, cell proliferation and
diminished accumulation of oxidized proteins) of adapted cells to withstand a high
H
2
O
2
challenge is, at least, partly dependent upon 20S proteasome induction,
immunoproteasome induction, and Pa28αβ regulator induction. These findings

217

demonstrate the importance of the 20S proteasome, immunoproteasome and Pa28αβ in
overall adaptation to oxidative stress.

II. The role of Nrf2 in oxidative stress adaptation

Our studies, published in Pickering et al. 2012 (Pickering et al., 2012) reveal a
mechanistic link between Nrf2, the 20S proteasome, the Pa28αβ (11S) proteasome
regulator, and transient adaptation to oxidative stress. It now appears clear that the Nrf2
signal transduction pathway plays a major role in both the increased proteasomal capacity
to degrade oxidized proteins, and the increased cellular tolerance to oxidative stress that
are induced by pre-treatment with a mild dose of oxidant.

We find that cellular capacity to degrade oxidized proteins, and intracellular levels of the
20S Proteasome, Immunoproteasome, and the Pa28αβ (11S) regulator are all increased
two- to three-fold during adaptation to oxidative stress. Similar results were obtained
with the oxidants H
2
O
2
and peroxynitrite, and the redox-cycling agents menadione and
paraquat. Proteasome inhibitors, and siRNA directed against the 20S Proteasome β1
subunit, the Immunoproteasome β1i (LMP2) subunit, or the Pa28α (11S) regulator
subunit, all significantly limited the increase in cellular proteolytic capacity and partially
prevented the increased resistance to oxidative stress (cell growth).

218

Cellular levels of Nrf2 were significantly increased by adaptation to oxidative stress, and
Nrf2 was seen to translocate to the nucleus, and to bind to ARE/EpRE sequence(s)
upstream of the Proteasome β5 subunit gene. Blocking the induction of Nrf2, with siRNA
or with retinoic acid, significantly limited the adaptive increases in cellular proteolytic
capacity, 20S Proteasome and the Pa28αβ regulator. Increases in the Immunoproteasome,
however, were only partially blocked by Nrf2 siRNA. Blocking Nrf2 induction also
limited the increase in oxidative stress resistance (cell growth). When instead of using
oxidant exposure, we pre-treated cells with the Nrf2 inducers lipoic acid, curcumin, or
sulforaphane, we observed increased cellular proteolytic capacity, increased 20S
Proteasome, and increased cellular resistance to oxidative stress (cell growth). We found
that both Nrf2 siRNA and 20S Proteasome β1 subunit siRNA effectively blocked the
increase in oxidative stress tolerance and 20S proteasome levels.

These results suggest that oxidants, redox cycling agents, and other Nrf2 ‘inducers’ cause
adaptation through the upregulation of Nrf2 and its translocation to the nucleus. This, in
turn, induces expression of the 20S Proteasome and the PA28αβ regulator. In contrast,
the Imunoproteasome, whose levels were also increased by adaptation to oxidative stress,
appears to be only partially regulated by Nrf2, if at all.

The Nrf2 signal transduction pathway is known to respond to stressful conditions (Itoh et
al., 1997; Itoh et al., 1999; Itoh et al., 2003; Kraft et al., 2006; Kwak et al., 2003;
McMahon et al., 2001; McMahon et al., 2003; Moi et al., 1994; Nguyen et al., 2003;

219

Rushmore et al., 1991; Venugopal and Jaiswal, 1996). Under non-stress conditions Nrf2
is retained in the cytoplasm through the formation of a complex with several proteins,
including Keap1. In this state it is constantly turned over through ubiquitin-dependant
26S Proteasome degradation. This permits a high expression rate, enabling rapid
accumulation of Nrf2 when degradation is blocked, while ensuring low Nrf2 steady-state
levels under normal conditions. Pre-treatment with an oxidant, or other Nrf2 inducer,
liberates Nrf2 from the Keap1 complex. This also prevents further Nrf2 degradation
resulting in a dramatic rise in Nrf2 cellular levels as well as its translocation to the
nucleus. Once there, it can bind to anti-oxidant response elements (ARE’s) that have also
been called electrophile response elements (EpRE’s), in a range of genes.

We find that genes encoding many 20S Proteasome subunits contain at least one if not
multiple ARE/EpRE sequences in their upstream, untranslated regions. We have also
shown that at least some of these ARE/EpRE sequences have a strong increase in Nrf2
binding under H
2
O
2
exposure. In contrast, we find only a single subunit of the three
Immunoproteasome subunits contains the ARE/EpRE sequence. It is tempting to suggest
that this difference in density of ARE/EpRE sequences may explain the differential
sensitivity of the 20S Proteasome and the Immunoproteasome to Nrf2 siRNA and retinoic
acid, and to propose that Immunoproteasome may be regulated by another mechanism.

Nrf2 is not the only protein that can bind to ARE/EpRE sequences, and it is certainly
possible that other signal transduction proteins may bind to proteasomal and Pa28αβ

220

(11S) regulator ARE/EpRE elements, and/or to Immunoproteasome. We are also
searching for other potential pathways for Immunoproteasome induction, of which the
Interferon Regulatory Factor 1 (Foss and Prydz, 1999; Namiki et al., 2005; Seifert et al.)
appears to be a good candidate. Finally, there may well be overlapping pathways of
signal transduction that act synergistically, or antagonistically, to dynamically adjust
Proteasome/Immunoproteasome levels during adaptation to oxidative stress. From these
results we hypothesized the model of adaptation shown in Figure 9.1

In Drosophila melanogaster and Caenorhabditis elegans we see a similar adaptive
response. This response appears to be dependent on 20S proteasome and Nrf2. It also
appears that Nrf2 regulates the increase in 20S proteasome in both organisms. This led us
to hypothesize the model of adaptive response shown in Figure 9.2.

In conclusion, we find that increases in 20S Proteasome and Pa28αβ (11S) regulator
expression are largely mediated by the Nrf2 signal transduction pathway during
adaptation to oxidative stress. These Nrf2-dependent increases in 20S Proteasome and
Pa28αβ (11S) are shown to be important for fully effective adaptive increases in cellular
stress resistance. In contrast, the Immunoproteasome, which also contributes to oxidative
stress adaptation, is shown to be minimally responsive to Nrf2 control.

221

Figure 9.1: Model of oxidative stress adaptation in mammalian cells
With H
2
O
2
exposure there is an initial protein synthesis independent response in which
the 19S regulator temporarily detaches from the 26S proteasome to increase the cellular
level of 20S proteasome. In addition the Pa28αβ regulator binds to the now free 20S
proteasome further increasing its capacity to degrade oxidized proteins. H
2
O
2
exposure
also causes the Nrf2 transcription factor to detach from the Keap-1 complex this
stabilizes Nrf2 increasing its levels and enabling it to translocate to the nucleus. When in
the nucleusNrf2 will bind to EpRE elements on a range of genes causing an upregulation
of 20S proteasome and Pa28αβ, Immunoproteasome, Pa28γ and Pa200 are also up-
regulated but by mechanisms that are as yet un-characterized.

222

Figure 9.2: Model of oxidative stress adaptation in D. melanogaster and C. elegans
We proposed that in Drosophila, H
2
O
2
causes Cnc-C to detach from its complex with
Dkeap-1. This stabilizes Cnc-C, so enabling its levels to increase and enabling it to
translocate to the nucleus. When in the nucleus, Cnc-C will bind to the upstream region
of proteasome subunits causing an increase in their synthesis. Similarly in C. elegans we
propose that, H
2
O
2
causes stabilization of Skn-1 (by a mechanism that is not entirely
clear) enabling its levels to increase and enabling it to translocate to the nucleus. When in
the nucleus Skn-1 will bind to the upstream region of proteasome subunits causing an
increase in their synthesis.

223

III. The proteasome regulators in response to oxidative stress

Our studies reveal that the interactions between the 20S and 26S proteasome and its
regulators are considerably more complex and intricate than was previously thought. In
support of previous findings (Pickering et al., 2010), the Pa28αβ and Pa28γ regulators
appear to enhance the capacity of the 20S proteasome to degrade oxidized proteins. This
provides support to the hypothesis that a major role for these regulators might be in
oxidative stress response is to enhance the capacity of proteasome to remove damaged
proteins after an acute stress. Given also that these two regulator are orthologues and
have very different cellular localizations (Brooks et al., 2000), the Pa28αβ regulator is
primarily localized in the cytoplasm while the Pa28γ regulator in the nucleus, it is
possible that these two regulators might posses similar roles in different organelles.

In further support of the potential role for Pa28αβ in response to acute oxidative stress,
we see that when cells are exposed H₂O₂ there is an increase in expression of both
Pa28αβ and Pa28γ over the subsequent 24 h. Furthermore there is an increase in Pa28αβ
binding to 20S proteasome 1 h after exposure to the oxidant. In previous work we have
shown in the same time course that there is a detachment of the 19S regulator from 26S
proteasome to form more 20S proteasome which results in an increase in capacity to
degrade oxidized proteins (Grune et al., 2011). From these results it seems likely that this
increase is not just a product of the formation of newly free 20S but also the product of an

224

increase in 20S-Pa28αβ complexes. Interestingly, there is not a corresponding increase in
the formation of 20S-Pa28γ complexes under oxidative stress.

Pa200 does not appear to posses the same sort of role in removal of oxidized proteins as
the Pa28αβ and Pa28γ regulators. In fact the addition of Pa200 to 20S proteasome
resulted in an almost complete loss of capacity to degrade both oxidized and native
hemoglobin. There is a moderate body of evidence that Pa200 has a role in DNA damage
repair (Blickwedehl et al., 2008; Ustrell et al., 2002). We confirmed previous reports that
Pa200 will form punctuate foci in the nucleus under hydrogen peroxide exposure which
is believed to be a product of Pa200 binding to the chromatin at sites of DNA damage.
Interestingly, we found that when Pa200 is added to 20S protesome it becomes
considerably better at degrading histones. However, the 20S proteasome-Pa200 complex
is much worse at degrading oxidized histones. Previous work has shown that Poly-ADP
ribose polymerase (PARP) will among its many functions bind to 20S proteasome and
enhance its capacity to degrade oxidized histones (Ullrich et al., 1999). We show that
while PARP will bind to 20S proteasome it will not bind to 20S-Pa200 complexes. From
these results it is possible perhaps both PARP and Pa200 will regulate the 20S
proteasome in similar but separate ways. PARP will induce the removal of damaged
histones so that the DNA may repair while Pa200 will induce the removal of undamaged
histones from damaged DNA. This would perhaps enable the DNA to unwind and be
repaired. In further support of this both Pa200 and PARP show increased binding to 20S
proteasome 30min-1 h after exposure to an acute oxidative stress (Ullrich et al., 1999). It

225

is possible that this might also explain the decline in Pa28γ association with 20S
proteasome under oxidative stress, that perhaps it is displaced by Pa200 or PARP (Ullrich
et al., 1999).

The immunoproteasome has long been believed to be linked with Pa28αβ due to both of
their co-expression and their similar localization (Pickering et al., 2010; Preckel et al.,
1999; Rivett et al., 2001). In this paper we show that Pa28αβ and Pa28γ are capable of
binding to immunoproteasome and that this enhances the capacity of immunoproteasome
to degrade oxidized proteins. In contrast Pa200 does not appear to have significant
interactions with immunoproteasome.

In summary our studies reveal that the interactions between the 20S and 26S proteasome
and its regulators are even more complex and intricate than was previously thought. We
also observe a highly dynamic quality to regulator binding in which different regulators
may be switched in and out of proteasome as a product of changing cellular conditions.
We demonstrate that the three proteasome regulators: Pa28αβ, Pa28γ and Pa200 all
appear to bind to 20S proteasome, with and without the addition of a 19S regulator,
forming multiple types of hybrid proteasomes, although the Pa200 regulator appears to
prefer 20S proteasome with a 19S regulator at the other end. We also show that
immunoproteasome forms a hybrid complex with Pa28αβ. In summary we see that the
proteasome regulators all appear to be important in response to oxidative stress but
appear to posses very different roles. In particular, the 20Sproteasome- Pa28αβ complex

226

appears to have much greater significance in successful adaptation to oxidative stress
than was previously realized.

From these finding we present the model shown in Figure 9.3 as a summary of the role of
different proteasome regulators in the degradation of oxidized proteins.

227

Figure 9.3: Summary of the complexes and roles of proteasome regulators
Addition of Pa28αβ appears to enhance the capacity of 20S proteasome or
immunoproteasome to degrade the short peptide substrate Suc-LLVY-AMC and to
selectively degrade oxidized proteins. Pa28αβ appears to be able to bind to 20S
proteasome, hybrid proteasome and Immunoproteasome. Addition of Pa28γ appears to
enhance the capacity of 20S proteasome and perhaps immunoproteasome to degrade the
short peptide substrate Suc-LLVY-AMC and to selectively degrade oxidized proteins.
Pa28γ appears to be able to bind to 20S proteasome, hybrid 26S proteasome and
Immunoproteasome. Pa200 appears to only be able to form a complex with hybrid
proteasome. Its addition increases the capacity of proteasome to degrade the short peptide
substrate Suc-LLVY-AMC. It blocks the degradation of most oxidized and native
proteins but enhances the capacity of proteasome to degrade native histones.

228

IV. Summary

In conclusion, mild oxidant exposure increases the tolerance of cells to a subsequent toxic
oxidant assault. This adaptive response is dependent on an increase in the levels of 20S
proteasome, increasing the capacity of the cell to degrade oxidized proteins. This
response occurs through an initial protein synthesis independent response. Where the,
26S proteasome will dissociate to increase the level of free 20S proteasome. Pa28αβ will
also bind to this newly free 20S proteasome. There is then a subsequent protein synthesis
dependent response. In this, the levels of 20S proteasome, immunoproteasome, Pa28αβ,
Pa28γ and Pa200 are increased. The increase in 20S proteasome and Pa28αβ, but not
immunoproteasome, is regulated by the transcription factor Nrf2. Pa28αβ and Pa28γ can
bind to 20S proteasome, hybrid proteasome and immunoproteasome. Where they both
appear to enhance the capacity of the cell to degrade oxidized proteins. Pa200 is
important in the oxidative stress response, although it does not increase the capacity of
the proteasome to degrade oxidized proteins. We also show that both Pa28αβ and Pa28γ
are able to form hybrid proteasomes with the 19S regulator. The Pa200 regulator, in
comparison, will only form a complex with hybrid proteasome. It will not bind to either
free 20S proteasome or immunoproteasome.

229

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Abstract (if available)

Abstract Oxidized cytoplasmic and nuclear proteins are normally degraded by the proteasome, but accumulate with age and disease. I demonstrate the importance of various forms of the proteasome during transient (reversible) adaptation (hormesis), to oxidative stress in murine embryonic fibroblasts. Adaptation was achieved by 'pre-treatment' with acute oxidative stress exposure (e.g. H₂O₂, peroxynitrite, menadione, paraquat), and tested by measuring inducible resistance to a subsequent much higher 'challenge' dose of H₂O₂. ❧ Oxidative stress adaptation causes an initial direct physical activation of pre-existing proteasomes, then a subsequent de novo synthesis of 20S proteasome, immunoproteasome and Pa28αβ during over the next 24 h. Cellular capacity to degrade oxidatively damaged proteins increased with 20S proteasome, immunoproteasome and Pa28αβ synthesis, and was mostly blocked by the 20S proteasome, immunoproteasome and Pa28 siRNA knockdown treatments. Direct comparison of purified 20S proteasome and immunoproteasome demonstrated that the immunoproteasome can selectively degrade oxidized proteins. Cell proliferation and DNA replication both decreased and oxidized proteins accumulated, during high H₂O₂ challenge. However H₂O₂ adaptation was protective against such H₂O₂ challenge. Importantly, siRNA knockdown of the 20S proteasome, immunoproteasome or Pa28αβ regulator blocked 50-100% of these adaptive increases in cell division and DNA replication. Immunoproteasome knock-down also largely abolished protection against protein oxidation. ❧ I show that the adaptative increase in oxidative stress tolerance and capacity to degrade oxidized proteins is dependent induction of the Nrf2 transcription factor. Furthermore I show that adaptation causes an increase in cellular levels of Nrf2, and translocation of Nrf2 from the cytoplasm to the nucleus. It also causes increased binding of Nrf2 to antioxidant response elements (ARE) or electrophile response elements (EpRE) in the 5-untranslated region of the Proteasome β5 subunit gene [demonstrated by chromatin immunoprecipiation (or ChIP) assay]. I go on to show that this induction of Nrf2 is a necessary requirement for increased Proteasome/Pa28αβ levels, and for maximal increases in proteolytic capacity and stress resistance. The oxidative stress induced increase in immunoproteasome however did not appear to be Nrf2 dependent. ❧ I show that Pa28αβ and Pa28γ have increased expression under mild oxidant exposure. I also demonstrate that both of the proteasome regulators enhance the capacity of the proteasome to selectively degrade oxidized proteins. In conjunction with their increased expression, there is an increase in binding of the Pa28αβ regulator to 20S proteasome. I show that the Pa200 proteasome regulator is also induced by H₂O₂ exposure. The Pa200 regulator however does not enhance the capacity of proteasome to degrade oxidized proteins. However, it does appear to enhance the capacity of the proteasome to degrade histones. ❧ I also demonstrate that this adaptive response is highly conserved. Exposure to a mild dose of an oxidant can increase oxidative stress tolerance in both Drosophila and C. elegans. There is also, in both of these animals, an increase in proteolytic capacity and a corresponding increase in 20S proteasome levels. If the increase in 20S proteasome or induction by Nrf2 homologues is blocked then the adaptive response is either blunted or completely lost in both animals.

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Creator Pickering, Andrew Michael (author)

Core Title The role of the proteasome & its regulators in adaptation to oxidative stress

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Molecular Biology

Publication Date 05/02/2012

Defense Date 03/06/2012

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag adaptation,C. elegans,cell culture ❧,Cnc-C,Drosophila,free radical,hormesis,immunoproteasome,mouse,Nrf2,OAI-PMH Harvest,oxidative stress,Pa200,Pa28,proteasome,SKN-1

Language English

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Advisor Davies, Kelvin J. A. (committee chair), Cadenas, Enrique (committee member), Finch, Caleb E. (committee member), Finkel, Steven E. (committee member)

Creator Email ampicker@usc.edu,pickering.andrew@gmail.com

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