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REDOX REGULATION OF
INFLAMMATION AND
IMMUNITY
SONIA SALZANO
A thesis submitted in partial fulfilment of the
requirements of the University of Brighton and the
University of Sussex for the degree of
Doctor of Philosophy
October 2013
2
Abstract
Inflammation is a consequence of the activation of innate immunity and represents
an important component of several pathological conditions, including not only the
complication of infections but also sterile and autoimmune diseases. An early event
in inflammation is represented by the production of proinflammatory cytokines and
both their production and action have often been associated to oxidative stress. The
redox status of the cell is therefore a key regulator of inflammation and
glutathionylation (formation of mixed disulphides between cysteine residues of
proteins and glutathione) is considered an important mechanism of this regulation.
While most of the studies in the past focused on glutathionylation of intracellular
proteins and transcription factors, the main goal of this project was to verify whether
glutathionylated proteins are released by inflammatory cells and if these have a
biological role. Using redox proteomics, we identified several proteins in the
supernatants from Raw 264.7 cells (murine macrophages) stimulated with bacterial
lipopolysaccharide (LPS). Among the identified proteins, we focused our attention
on Peroxiredoxin 2 (Prx2), an antioxidant enzyme involved in cells protection
against oxidative stress by removing H2O2. Released Prx2 was also detected in
supernatant from human peripheral blood mononuclear cells (PBMC) and human
macrophages. Prx2 levels were also increased in the serum of LPS-treated mice. We
could confirm that Prx2 is released in the glutathionylated form. Moreover it was
observed that the intracellular level of glutathione affects Prx2 release suggesting a
role for glutathionylation in the mechanism of its release.
The second part of the project was to verify whether released glutathionylated
proteins may act as mediators of inflammation. To this purpose, the possible
inflammatory role of released Prx2 was studied. The results showed that extracellular
Prx2 induced an increase of TNF-α production in Raw 264.7 cells and in human
macrophages.
In conclusion, Prx2 is released during inflammation in a redox-dependent manner, in
addition to its well-known intracellular role as enzyme, Prx2, in its released form,
can also play a role in inflammatory response.
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Candidate’s declaration
I declare that the research contained in this thesis, unless otherwise formally
indicated within the text, is the original work of the author. The thesis has not been
previously submitted to these or any other university for a degree, and does not
incorporate any material already submitted for a degree.
Signed: …………………………………….
Date: ……………………………………....
4
Acknowledgements
I would like to take this opportunity to thank the following people who have
contributed to my work and supported me during my PhD: Prof. Pietro Ghezzi, my
supervisor, for giving me the opportunity to do this PhD at BSMS and for offering
guidance with this project. Dr. Manuela Mengozzi for giving me support and active
collaboration with my experiments. Dr. Lucas Bowler for the initial collaboration
and his useful identification of Prx2. Dr. Sandra Sacre, Dr. Kevin Tracey and Dr. Eva
Maria Hanschmann for their kind collaboration.
My friends and laboratory colleagues: Dr. Ilaria Cervellini for all good and bad days
that we shared during our PhD. Dr. Lucia Coppo for helping me with the “tangled
world of proteins”. Dr. Paola Checconi for always giving me positive advice about
work and life. I will remember our “two single macchiato, to have here”.
All my friends in Brighton who made the last period of my time here “amazing”. A
particular thanks to Andrej, Antonio, Cata, Flora, Prateek and Stefania. Further
thanks to my friends Andrea (d’oh), Elena, Melinda and Michelle for being close by
me throughout my PhD, despite the long distance that separated us. My “English
proof reading team” Izzie, James, Nick, Rags, and Sam for giving me very useful
comments on the thesis. Finally, last but not least, I would like to thank my family
for all the support and positive energy they have given me during this journey.
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List of publications and presentations from this thesis
Priora, R., Coppo, L., Salzano, S., Di Simplicio, P., Ghezzi,P. 2010. Measurement of
mixed disulphides including glutathionylated proteins. Methods Enzymol 473, 149-
159.
Salzano, S., Coppo, L., Bowler, L., Cervellini, I., Mengozzi, M., and Ghezzi, P.
Redox regulation of inflammation and immunity: secreted oxidoreductase acting as
inflammatory cytokines. Poster presented at BSMS, Postgraduate Research Student
Symposium (Brighton and Sussex Medical School, May 2011).
Salzano, S., Coppo, L., Bowler, L., Cervellini, I., Mengozzi, M. ,and Ghezzi, P.
A proteomic approach to identify proteins involved in Redox regulation of
inflammation and immunity. Poster presented at Experimental Biology (San Diego
Convention Center, April 2012).
Salzano, S., Coppo, L., Bowler, L., Cervellini, I., Mengozzi, M., and Ghezzi, P.
A proteomic approach to identify proteins involved in Redox regulation of
inflammation and immunity. Oral presentation at BSMS, Postgraduate Research
Student Symposium (Brighton and Sussex Medical School, May 2012).
6
Contents
Chapter 1. Introduction .....................................................................................17
1.1 Immune system .........................................................................................18
1.1.1 Adaptive immunity ..............................................................................18
1.1.2 Innate immunity ..................................................................................21
1.1.3 Toll-like receptors ...............................................................................23
1.1.4 DAMP .................................................................................................28
1.1.4.1 HMGB1......................................................................................31
1.1.4.2 HSPs ..........................................................................................34
1.2 Inflammation ............................................................................................36
1.2.1 Inflammatory cytokines .......................................................................37
1.2.2 Inflammatory diseases .........................................................................42
1.3 Oxidative Stress ........................................................................................44
1.3.1. Oxidative stress in diseases ......................................................46
1.3.2. Oxidative stress and innate immunity ......................................47
1.4 Proteins thiols-disulphides metabolism in redox regulation .......................50
1.4.1. Protein glutathionylation ......................................................53
1.5 Redoxins ...................................................................................................57
1.5.1 Thioredoxin .........................................................................................57
1.5.2 Glutaredoxin ........................................................................................61
1.5.3 Protein Disulphide Isomerase...............................................................64
1.5.4 Peroxiredoxins .....................................................................................66
1.6 Aims of the study ......................................................................................71
Chapter 2. Materials and Methods .....................................................................73
2.1 Materials ...................................................................................................74
2.1.1 Instruments ..........................................................................................74
2.1.2 Chemicals, Reagents, and Kits .............................................................75
2.1.3 List of Antibodies ................................................................................77
2.1.4 Human recombinant proteins and Biotinylated oxidized Glutathione ...78
2.1.5 Cells ....................................................................................................78
2.2 Methods ....................................................................................................79
2.2.1 Cell culture ..........................................................................................79
7
2.2.2 Cell viability assay...............................................................................82
2.2.3 LDH assay ...........................................................................................82
2.2.4 Protein glutathionylation ......................................................................84
2.2.5 Proteomics of released glutathionylated proteins ..................................84
2.2.5.1 BioGEE cellular uptake and treatments .......................................84
2.2.5.2 Pull-down of biotinylated proteins ..............................................85
2.2.5.3 SDS-PAGE and Western blot .....................................................87
2.2.5.4 SDS-PAGE staining ...................................................................90
2.2.5.5 Mass Spectrometry .....................................................................92
2.2.6 Western blot of biotinylated proteins ...................................................93
2.2.6.1 Detection of Prx2 released by macrophages ................................93
2.2.6.2 Detection of Prx2 levels in mouse serum ....................................94
2.2.7 Immunoprecipitation............................................................................96
2.2.7.1 Immunoprecipitation of HMGB1 ................................................99
2.2.7.2 Immunoprecipitation of glutathionylated Prx2 ............................99
2.2.8 Intracellular GSH assay ..................................................................... 100
2.2.8.1 Glutathione depletion with BSO ............................................... 100
2.2.8.2 Protein determination................................................................ 103
2.2.9 Activity of Prx2 with Trx1 or Grx2 .................................................... 104
2.2.10 Removal of endotoxin from hrPrx2 preparation ............................. 105
2.2.11 Limulus Amebocyte Lysate test ..................................................... 106
2.2.12 ELISA ........................................................................................... 107
2.2.12.1 Mouse TNF-α ELISA ............................................................... 107
2.2.12.2 Human TNF-α ELISA .............................................................. 108
2.2.12.3 Human Prx2 ELISA .................................................................. 110
2.2.13 Statistical Analysis......................................................................... 111
Chapter 3. Studies on the glutathionylation of HMGB1................................... 112
3.1 Introduction: non-classical secretion of proteins ...................................... 113
3.2 Aim of Chapter 3 .................................................................................... 118
3.3 Results .................................................................................................... 119
3.3.1 Glutathionylation of HMGB1 ............................................................ 122
3.3.2 HMGB1 release by LPS-stimulated Raw 264.7 cells .......................... 122
3.3.3 Immunoprecipitation of HMGB1 ....................................................... 124
3.4 Discussion .............................................................................................. 125
8
3.5 Following chapter ................................................................................... 126
Chapter 4. Proteomic analysis of conditioned medium from Raw 264.7 cells .. 127
4.1. Introduction to the identification of unknown released proteins................... 128
4.2. Aim of Chapter 4 ........................................................................................ 131
4.3. MS identification of proteins released by LPS-stimulated Raw 264.7 cells .. 132
4.4. Discussion .................................................................................................. 147
4.5. Following chapter ....................................................................................... 149
Chapter 5. Prx2 release in LPS-stimulated mouse and human macrophages .... 150
5.1. Introduction: extracellular Prx2 .................................................................. 151
5.2. Aim of Chapter 5 ........................................................................................ 153
5.3. Detection of released Prx2 from Raw 264.7 cells ........................................ 154
5.4. Passive release of Prx2 ............................................................................... 167
5.4.1. CTB assay ............................................................................................ 167
5.4.2. LDH assay ........................................................................................... 167
5.5. Detection of released Prx2 from PBMC and human macrophages ............... 170
5.5.1. PBMC .................................................................................................. 170
5.5.2. Human macrophages ............................................................................ 173
5.6. Detection of Prx2 levels in serum from LPS-treated mice ........................... 176
5.7. Discussion .................................................................................................. 181
5.8. Following chapter ....................................................................................... 182
Chapter 6. Glutathionylated Prx2 released by Raw 264.7 cells ........................ 184
6.1. Introduction ................................................................................................ 185
6.2. Aim of Chapter 6 ........................................................................................ 186
6.3. Immunoprecipitation of Prx2 ...................................................................... 188
6.4. Depletion of intracellular GSH ................................................................... 190
6.4.1. Cell viability ........................................................................................ 192
6.4.2. GSH assay............................................................................................ 194
6.4.3. Detection of glutathionylated Prx2 by Western blot .............................. 196
6.5. Discussion .................................................................................................. 199
6.6. Following chapter ....................................................................................... 201
Chapter 7. Role of extracellular Prx2 .............................................................. 202
7.1 Introduction ............................................................................................ 203
7.2 Aim of Chapter 7 .................................................................................... 205
7.3 Peroxidase activity of hrPrx2 .................................................................. 206
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7.4 Removal of LPS from hrPrx2 and Western blot analysis of LPS-free hrPrx2
208
7.5 TNF-α production in Raw 264.7 cells treated with hrPrx2 ....................... 215
7.6 TNF-α production in human macrophages treated with hrPrx2 ................ 220
7.7 Discussion .............................................................................................. 222
Chapter 8. Conclusions and discussion ............................................................ 224
8.1 Summary of thesis results ....................................................................... 225
8.2 Glutathionylation and release of HMGB1 ............................................... 228
8.3 Released Prx2 by LPS-stimulated macrophages ...................................... 229
8.4 Glutathionylated Prx2 ............................................................................. 231
8.5 Inflammatory role of released Prx2 ......................................................... 232
8.6 Prx2 in the context of danger signals and oxidative stress ....................... 233
Chapter 9. References ..................................................................................... 236
10
List of Figures
Figure 1.1: TLRs and their ligands.. .......................................................................25
Figure 1.2: TLRs and their adaptor proteins.. .........................................................27
Figure 1.3: DAMPs release.. ..................................................................................29
Figure 1.4: Redox regulation of HMGB1.. .............................................................33
Figure 1.5: Cytokines and their cells source. ..........................................................39
Figure 1.6: Jak-Stat pathways in the signaling of class 1 cytokines.. .......................41
Figure 1.7: Oxidative modifications of proteins thiol.. ............................................52
Figure 1.8: Main mechanisms of glutathionylation.. ...............................................55
Figure 1.9: Redox regulation.. ................................................................................56
Figure 1.10: The Trx system.. ................................................................................59
Figure 1.11: Post-translational modification of Trx.. ..............................................60
Figure 1.12: The Grx system.. ................................................................................62
Figure 1.13: Post-translational modifications of Grx1 and Grx2.. ...........................63
Figure 1.14: PDI domains structure. .......................................................................65
Figure 1.15: Mechanism of action of Prxs.. ............................................................69
Figure 2.1: Biotinylated proteins pull-down.. .........................................................86
Figure 2.2: Overview of Western blot. ...................................................................89
Figure 2.3: Mouse serum sample preparation. ........................................................95
Figure 2.4: Schematic immunoprecipitation using the indirect method. ..................97
Figure 2.5: Schematic immunoprecipitation using the direct method. .....................98
Figure 2.6: Sample preparation scheme for GSH assay. ....................................... 102
Figure 2.7: ELISA summary steps.. ...................................................................... 109
Figure 3.1: Western blot of glutathionylated proteins.. ......................................... 120
Figure 3.2: Western blot of glutathionylated proteins (under reducing and non-
reducing conditions).. ............................................................................................ 121
Figure 3.3: Detection of glutathionylated HMGB1 by Western blot probed with
Streptavidin-POD.................................................................................................. 123
Figure 3.4: Detection of HMGB1.. ....................................................................... 123
Figure 4.1: Summary of the experimental procedure for the identification of the
released proteins.. ................................................................................................. 134
Figure 4.2: Biotinylated glutathionylated proteins in Raw 264.7 cells................... 137
Figure 4.3: Silver stained gel of conditioned medium from Raw 264.7 cells.. ....... 138
11
Figure 4.4: Tandem mass spectra (MS/MS spectra) of HSP70 peptides obtained
during analysis of tryptic digests of samples by LC-MS.. ...................................... 142
Figure 4.5: MS/MS spectra of 2 Prx2 peptides obtained during analysis of tryptic
digests of samples by LC-MS................................................................................ 143
Figure 4.6: MS/MS spectra of 2 vimentin peptides obtained during analysis of
tryptic digests of samples by LC-MS..................................................................... 144
Figure 4.7: MS/MS spectra of 5 LDH A peptides obtained during analysis of tryptic
digests of samples by LC-MS................................................................................ 146
Figure 5.1: Time course of Prx2 release in Raw 264.7 cells... ............................... 155
Figure 5.2: Western blot showing the effect of LPS 24 and LPS 4 hours on Prx2
release by Raw 264.7 cells.. .................................................................................. 156
Figure 5.3: Experimental scheme used for LPS stimulation of Raw 264.7 cells.. .. 158
Figure 5.4: Western blot showing the effect of LPS on Prx2 release by Raw 264.7
cells (Experiment 1).. ............................................................................................ 159
Figure 5.5: Western blot showing the effect of LPS on Prx2 release by Raw 264.7
cells (Experiment 2).. ............................................................................................ 160
Figure 5.6: Effect of LPS on released and intracellular Prx2 levels in Raw 264.7
cells (Experiment 3).. ............................................................................................ 162
Figure 5.7: Effect of LPS on released and intracellular Prx2 levels in Raw 264.7
cells (Experiment 4).. ............................................................................................ 163
Figure 5.8: Effect of LPS on released and intracellular Prx2 levels in Raw 264.7
cells (Experiment 5).. ............................................................................................ 164
Figure 5.9: Prx2 detection by Western blot in non-reducing conditions.. .............. 166
Figure 5.10: Effect of LPS on Raw 264.7 cell viability.. ....................................... 168
Figure 5.11: LDH release by Raw 264.7 cells....................................................... 169
Figure 5.12: Effect of LPS on Prx2 release in PBMC.. ......................................... 172
Figure 5.13: Effect of LPS on Prx2 release in human macrophages.. .................... 174
Figure 5.14: Western blot analysis of serum from LPS-treated mice (90 min or
24 hours) and vehicle-treated mice (Experiment 1).. .............................................. 177
Figure 5.15: Western blot analysis of serum from LPS-treated mice (90 min) and
vehicle-treated mice (Experiment 2).. .................................................................... 178
Figure 5.16: Serum Prx2 levels after vehicle (saline) or LPS (24 hours) injection in
mice (Experiment 3). . ........................................................................................... 179
Figure 5.17: Serum Prx2 levels after LPS (24 hours) injection in mice
(Experiment 4).. .................................................................................................... 180
12
Figure 6.1: Released glutathionylated Prx2 by Raw 264.7 cells treated with LPS..
............................................................................................................................. 189
Figure 6.2: Experimental scheme of BSO and LPS treatments of Raw 264.7 cells.
............................................................................................................................. 191
Figure 6.3: Effect of BSO and LPS on Raw 264.7 cell viability............................ 193
Figure 6.4: Effect of BSO on GSHtot (GSH + GSSG) levels in Raw 264.7 cells.. .. 195
Figure 6.5: Effect of GSH depletion by BSO 250 µM on Raw 264.7 cells.. .......... 197
Figure 6.6: Effect of GSH depletion by BSO 125 µM on Raw 264.7 cells.. .......... 198
Figure 6.7: The different oxidation states of Prx2 and the effect of H2O2 treatment..
............................................................................................................................. 200
Figure 7.1: Peroxidase activity of hrPrx2.. ........................................................... 207
Figure 7.2: Scheme for the removal and checking of LPS contamination from
hrPrx2. .................................................................................................................. 210
Figure 7.3: Protein determination of hrPrx2 after Detoxi-Gel filtration.. ............... 211
Figure 7.4: Western blot analysis of hrPrx2 after Detoxi-Gel filtration.. ............... 212
Figure 7.5: Scheme of the experiment for TNF-α production in Raw 264.7 cells
evaluated by ELISA.. ............................................................................................ 217
Figure 7.6: Effect of hrPrx2 on TNF-α production by Raw 264.7 cells using boiled
hrPrx2 as control for Prx2 contamination by endotoxin.. ....................................... 218
Figure 7.7: Effect of hrPrx2 on TNF-α production by Raw 264.7 cells using PMB to
investigate endotoxin contamination.. ................................................................... 219
Figure 7.8: Effect of hrPrx2 on TNF-α production by human macrophages.. ........ 221
13
List of Tables
Table 1.1: TLRs and their ligands...........................................................................26
Table 1.2: Some diseases associated with DAMPs. ................................................30
Table 1.3: Cytokines and their major function. .......................................................40
Table 1.4: Infection/inflammation-associated diseases with evidence for a role of
oxidative stress. ......................................................................................................49
Table 2.1: Dilutions recommended in the product datasheet for Western blot
analysis. ..................................................................................................................77
Table 2.2: (A) Plating densities for Raw 264.7 cell culture. (B) Plating densities for
primary human macrophages and PBMC. ...............................................................81
Table 4.1: Proteins identified by MS in the experiment without separation with SDS-
PAGE (Experiment 3).. ......................................................................................... 140
Table 4.2: MS results of more interesting proteins identified in three independent
experiments........................................................................................................... 141
Table 5.1: TNF-α levels in supernatants from untreated (control) and LPS-stimulated
PBMC and human macrophages.. ......................................................................... 175
Table 7.1: LAL test for endotoxin contamination.. .............................................. 214
Table 8.1: Aims and results from each chapter of this thesis. ................................ 226
14
List of Abbreviations
Abbreviation Full wording
˚C Degree Celsius
2-ME 2-Mercaptoethanol
APS Ammonium Persulphate
ARDS Acute Respiratory Distress Syndrome
AU Arbitrary Units
BCR B Cell Receptor
BioGEE Biotinylated Glutathione Ethyl Ester
BioGSSG Biotinylated Glutathione (oxidized)
BSA Bovine Serum Albumin
BSO Buthionine Sulfoximine
CGD Chronic Granulomatous Disease
CNS Central Nervous System
CSE Control Standard Endotoxin
CTB CellTiter-Blue
Cys Cysteine
DAMPs Damage-Associated Molecular Patterns
DMSO Dimethyl Sulfoxide
DTNB 5,5’-Dithiobis (2-Nitrobenzoic Acid)
DTT Dithiothreitol
ELISA Enzyme-Linked Immunosorbent Assay
ER Endoplasmic Reticulum
EU Endotoxin Unit
FBS Fetal Bovine Serum
GR Glutathione Reductase
Grx Glutaredoxin
GSH Glutathione (reduced)
GSSG Glutathione (oxidized)
HI Heat-inactivated
HMGB1 High-Mobility Group B1
HRP Horseradish Peroxidase
HrPrx2 Human recombinant Prx2
HSPs Heat Shock Proteins
15
IFN Interferon
IL Interleukin
IP Immunoprecipitation
LAL Limulus Amebocyte Lysate
LDH Lactate dehydrogenase
LPS Lipopolysaccharide
M Molar (mM, millimolar; µM, micromolar; nM, nanomolar)
MHC Major Histocompatibility Complex
Min Minute
MOF Multiple Organ Failure
MS Mass Spectrometry
MS Multiple Sclerosis
MW Molecular Weight
MyD88 Myeloid Differentiation factor 88
NADPH Nicotinamide Adenine Dinucleotide Phosphate
NEM N-Ethylmaleimide
NF-kB Nuclear factor-kappaB
NK Natural Killer
Nm Nanometer
ON Overnight
PAGE Polyacrylamide Gel Electrophoresis
PAMPs Pathogens-associated molecular patterns
PBMC Peripheral Blood Mononuclear Cells
PBS Phosphate Buffered Saline
PDI Protein Disulphide Isomerase
PMB Polymyxin B
PMN Polymorphonuclear Neutrophils
POD Peroxidase
PRRs Pattern Recognition Receptors
Prxs Peroxiredoxins
PSHs Protein thiol groups
RNS Reactive Nitrogen Species
ROIs Reactive Oxygen Intermediates
ROS Reactive Oxygen Species
RPMI 1640 Roswell Park Memorial Institute
RT Room Temperature
SA-POD Streptavidin-Peroxidase
16
SB (Laemmli) Sample Buffer
SBD Substrate Binding Domain
SDS Sodium Dodecyl Sulphate
SH Sulfhydryl group
SIRS Systemic Inflammatory Response Syndrome
SN Supernatant
SNO Nitrosothiol
SOD Superoxide Dismutase
TCA Trichloroacetic acid
TCR T cell receptor
Th T helper cells
TLRs Toll-like receptors
TMB 3,3',5,5'-Tetramethylbenzidine
TNB 5-Thio (2-Nitrobenzoic Acid)
TNF-α Tumor necrosis factor-α
Trx Thioredoxin
TrxR Trx Reductase
17
Chapter 1. Introduction
18
1.1 Immune system
The most important function of the immune system is to protect the body/host from
microbes such as bacteria, fungi, protozoa and viruses that can be dangerous (1). The
protection from the pathogens requires recognition, attack and final elimination to
avoid replication of the foreign organism. This defence against microbes is divided
into two major branches with a different function and role: the innate immune system
and the adaptive immune system. The innate immunity represents an immediate and
non-specific response against an attack by a pathogen (2). All multi-cellular
organisms use the innate immunity as a potent and rapid weapon against infection.
Furthermore, only in vertebrates is the innate immune response followed in 4-7 days
if it is necessary by adaptive/specific immune response. However, different
experimental studies suggested that the adaptive response is not only a consequence
of the innate response failure and then a second option to fight against pathogens but
that there is also a proper collaboration between the two defence systems (3) (4).
Leukocytes (or white blood cells) which include granulocytes (neutrophils, basophils
and eosinophils), macrophages (and their precursor monocytes) and lymphocytes
(B cells and T cells) are an effective weapon in immune system. In particular,
macrophages and neutrophils have phagocytic activity and are involved in innate
immunity while lymphocytes in adaptive immunity.
1.1.1 Adaptive immunity
Compared to innate immunity, adaptive immunity is most complex due to the
activity of different cell populations. Adaptive immunity can be classified as two
main types: cell-mediated immunity and humoral immunity. The main problem with
adaptive immunity is that it requires time to have its effect. Normally, at least a week
is required for a T-cell-mediated response, even longer for the antibody (humoral)
response.
Cell-mediated immunity mainly involves T lymphocytes and antigen-presenting
cells. Basically, the principle function of antigen-presenting cells such as
19
macrophages dendritic cells and B lymphocytes is to recognize and capture the
antigen, followed by its migration and T cell stimulation. For instance, dendritic cells
interact and control lymphocytes functions, in particular immature dendritic cells
also contain specific cell surface receptors that recognize pathogens antigen.
Macrophages and dendritic cells utilize phagocytosis for the antigen uptake, whereas
B lymphocytes utilize specific B cell receptor (BCR). Antigenic proteins are
processed into short peptide fragments by different enzymatic mechanism. After this
process, antigen fragments are transported to the surface of the antigen-presenting
cells and followed by formation of peptides bound to MHC (major histocompatibility
complex) class I or class II molecule. Therefore, the T cell receptor (TCR) can
recognize the antigen peptides associated to MCH (5). T lymphocytes that express
CD8+ have a cytotoxic phenotype and are activated by MCH class I, whereas MCH
class II-presented antigen activate CD4+ helper T lymphocytes (6).
Perforin/granzyme-induced apoptosis is the main pathway used by cytotoxic T cells
to kill infected or transformed cells. The perforin molecules form a pore on the
membrane surface to allow the passage of granzymes (serine proteases contained in
cytoplasmic granules of cytotoxic T lymphocytes and natural killer cells (NK)) into
the target cell cytoplasm. Inside the cell, granzymes induce the activation of caspase
(cysteine proteases) and then the apoptosis (programmed cell death) (7). The
evidence that perforin has an important role in killing cells was shown in perforin-
deficient mouse infected with viruses, where the lack of perforin molecules induce
reduction or elimination of capacity to kill cells (8).
T helper cells (Th) have no cytotoxic or phagocytic activity but they have the role of
help the other cells of the immune system (macrophages, B cells and cytotoxic cells).
T helper cells can be divided in Th1 and Th2 characterized by a different pathway
and cytokines production.
Humoral immunity has B lymphocytes as the major players responsible for antibody
production which are specific for each invading pathogens. These B cells possess a
BCR which basically resembles an antibody molecule. Antigen binding to the BCR
is followed by internalization of antigen-BCR complex and antigen fragmentation
with presentation to the MCH II. Th cells have a central role for the antibody
response. In particular, the T cell binding to the antigen presented on the BCR
20
through the T cell MHC II causes the release of different cytokines from the T cell
and then differentiation of B cells in plasma cells that produce antibodies (IgD, IgA,
IgM, IgE and IgG). Antibodies fight against pathogens through different killing
strategies such as the opsonisation for phagocytosis, the neutralization of toxins
which block the attachment of pathogens to cells or tissue, and the activation of the
complement. During the opsonization process, the antigen binds the antigen binding
area (Fab) of the antibodies that is used as a tag (or opsonin). The binding antigen-
antibody stimulates the phagocytic cells that recognize and destroy the pathogen (9).
Neutralization of toxin is another mechanism of defence against the attack from toxic
molecules secreted by pathogens. The antibody binds the toxin neutralizing its
dangerous effect. Agglutination is a reaction of antibodies that immobilize and form
a large aggregate (clumps) of infectious agents by binding to their surface antigens.
Therefore, the immobilized antigens formation prevents the harmful effect of
pathogens.
Both innate and adaptive immunity can activate the complement system (a group of
more than 30 proteins) that allows host defense during infection or tissue damage.
Complement can be activated producing a cascade of reactions through three
pathways (classical, lectin or alternative) which terminate in inflammation, lysis and
opsonization (10). In the classical pathway antibody forms antigen-antibody complex
on the surface of a pathogen and the antibody complex also binds the first component
of Complement (C1) producing an enzyme cascade. Activated C1s (subcomponent of
C1) cleaves C4 in C4a and C4b and C2 into C2a and C2b. The complex C4bC2b
forms C3 convertase that cleaves C3 into anaphylatoxins (proinflammatory
molecules) C3a and opsonin C3b (11). C3b also binds the C3 convertase to form a
C5 convertase that produces C5a with a chemotaxis role and C5b. The lectin
pathway is independent by antibody but activated directly by microbial component.
The alternative pathway starts with the spontaneous hydrolysis of C3.
21
1.1.2 Innate immunity
Innate immunity is also called non-specific immunity because its mechanisms are not
specific to a particular pathogen. Mechanism of host defences is composed of
anatomic barriers that make it difficult for the pathogen to enter the body. Anatomic
barriers include the skin and mucosal membranes such as the mucociliary escalator.
In the respiratory system, the cilia not only provide a physical barrier but also expel
bacteria trapped in the mucus (12).
Low pH, high temperature and chemical mediators represent the physiologic barriers.
The low pH of the skin and in the stomach is also an inhibitor of bacterial growth.
Chemical mediators such as interferon (IFN), complement, and lysozymes also
contribute to innate immunity defence. The normal body temperature and the fever
response are not optimal for many pathogens making the environment even more
difficult for their growth. The body’s temperature is regulated in the hypothalamic
area of the brain and high temperature inhibits pathogens growth, and therefore fever
represents an important mechanism of defence against pathogens. Cytokines such as
TNF-α (Tumor necrosis factor-α) interleukin-1 (IL-1) and interleukin-6 (IL-6) can
act as endogenous pyrogens inducing fever (13). Cytokines pyrogens initially named
leucocytic pyrogen were purified in vitro in supernatants of human peripheral
leukocytes stimulated by phagocytosis of killed staphylococci (14). IL-1 can induce
fever through the prostaglandin E2 release in different cells (monocytes, fibroblasts,
brain tissue and homogenates and muscle strips) (15). For instance, Dinarello et al.
showed (16) that human monocytes incubated with IL-1 increases prostaglandin
levels and this effect can be blocked by ibuprofen.
More complex reactions involve the inflammatory barriers, characterized by leakage
of vascular fluid containing serum proteins with antibacterial activity. Components
of immunity also include phagocytic cells (neutrophils, monocytes and macrophages)
and other cells (dendritic cells and NK cells) that defence the host through the release
of inflammatory mediators such as chemotactic factors, cytokines, Reactive Oxygen
Intermediates (ROIs) and reactive nitrogen intermediates. ROIs such as O2.-
(superoxide anion), OH.(hydroxyl radicals) and H2O2 (hydrogen peroxide) and NO
(nitric oxide) have antimicrobial activity (17) and can regulate the expression of
22
cytokines (18). These mediators also activate a number of physiological responses to
the infection, some desirable and some damaging to the host. The systemic reaction
(acute phase reaction) is characterized by fever, increase in secretion of
glucocorticoids, increased leucocytosis, decreases in serum levels of iron and zinc
and changes in the concentration of plasma proteins knows as acute-phase proteins.
In order to control the bacterial growth, the body has developed a nutritional
immunity to deprive bacteria of iron. Iron is an essential element necessary for the
metabolism and replication of bacterial pathogens, for instance it is required for RNA
and DNA synthesis and different enzyme activity. IL-1 produced during fever
induces iron deficiency (hypoferremia) which has an antimicrobial effect (19).
Siderophores, the small molecules produced by bacteria to acquire iron are decreased
by fever as showed by Garibaldi in Salmonella typhimurium (20). The body also
uses a mechanism to withhold iron, increasing synthesis of human iron-binding
proteins such as lactoferrin, transferrin, and ferritin (21). The cachexia-anorexia
syndrome characterized by weight loss is a consequence of inflammation with
cytokines production. For instance, IL-1 and TNF-α can reduce the appetite and have
a synergic effect in rats as showed by Yang et al. (22).
Inflammatory cytokines produced during an inflammatory reaction can modify the
levels of acute phase proteins. Acute phase proteins are classified in positive-acute-
phase proteins (proteins that increase during inflammatory disorders) and negative-
acute-phase proteins (proteins that decrease during inflammatory disorders) (23)
(24). For instance, IL-6 is the major regulator of acute-phase proteins synthesis in
adult human hepatocytes (25).
23
1.1.3 Toll-like receptors
The underlying concept of innate immunity is based on the fact that one cell,
particularly macrophages, can recognize potentially pathogenic microorganisms
(both viruses and bacteria) even if these had not been encountered before. This is
done by recognizing of some molecular patterns that are common to many pathogens
and whose presence of a bacterium or virus flags it as a potential pathogen. Immune
cells express pattern recognition receptors (PRRs) which include Toll-like receptors
(TLRs) that play a key role in the innate immune response. Probably the main
breakthrough in this field was the discovery that the mammalian homologs of the
Drosophila gene Toll are responsible for the response of the immune system. LPS,
the component of cell wall of Gram-negative bacteria is widely used to induce
inflammation in vitro and in vivo. In a pioneering study that gained Bruce Beutler the
Nobel Prize in Physiology or Medicine, he found that genetically LPS-resistant mice
have a mutation in TLR4 that became the prototypic innate immunity “recognition
receptor” (26) (27). This important discovery increased the interest of scientists on
activation of innate immunity and different Toll-like receptors (TLRs) that are now
known (ten TLR expressed in humans and thirteen expressed in mice). They can
recognize specific microbial components known as pathogen-associated molecular
patterns (PAMPs), and initiate intracellular signalling pathways with production of
inflammatory cytokines. In humans some TLRs (TLR2 with TLR1 or TLR6, TLR4,
TLR5 and TLR10) are distributed at the cell membrane and are particularly
important in the recognition of molecules expressed by pathogenic bacteria, parasites
and fungi. Others are located in the endosomal compartments and recognize viral,
bacterial or protozoal molecules (TLR3, TLR7, TLR8 and TLR9).
As shown in Figure 1.1, flagellin (constituent of bacterial flagella) is a ligand for
TLR5, while lipopolysaccharide (LPS) is specific for TLR4. TLR2 is able to form
heteromers with TLR1 or TLR6 and recognizes lipoproteins, lipopeptides,
peptidoglycans and lipoteichoic acid of Gram-positive bacteria (28). TLR7 and
TLR8 recognize RNA from viruses while TLR9 is specific for bacterial DNA with
unmethylated CpG motif (29). A list of TLRs with their location, PAMPs recognition
and pathogens that express these PAMPs are listed in Table 1.1.
24
Activation of the various TLRs induces transcription factors and then gene
expression for several key mediators in innate immunity. TLRs have an extracellular
domain to recognize the pathogen and a cytoplasmic domain (TIR domain). TIR is
associated with different adaptor proteins such as myeloid differentiation factor 88
(MyD88), TIR-domain-containing adaptor-inducing interferon-β (TRIF) and TRIF-
related adaptor molecule (TRAM). In particular, as shown in Figure 1.2, TLR1,
TLR2, and TLR6 utilize MyD88 and TIRAP as adaptors while TLR5, TLR7, TLR9
and TLR11 utilize MyD88. TLR4 uses four adaptors, MyD88, TIRAP, TRIF and
TRAM. TLR3 uses only TRIF adaptor (30). The end-result of the TLRs signalling
cascade mediated by the adaptor proteins is the activation of the nuclear factor-
kappaB (NF-KB) pathways with cytokine production and IRF3 pathways responsible
for the IFN production. Furthermore, the mitogen-activated protein kinase (MAPK)
pathway can also be activated by TLRs with production of inflammatory cytokines
(31).
25
Figure 1.1: TLRs and their ligands. Figure adapted from Takeda and Akira (32).
26
TLR Location Main ligand Recognize Reference
TLR1 & TLR2 cell surface triacyl
lipopeptides bacteria
(33)
TLR2 & TLR6 cell surface
diacyl
lipopeptides
zymosan
lipoteichoic acid
bacteria
fungi
bacteria
(34)
(35)
(36)
TLR3 endosomes double-stranded
RNA viruses (37)
TLR4 cell surface LPS bacteria (26)
TLR5 cell surface flagellin bacteria (38)
TLR7/8 endosomes single-stranded
RNA viruses (39)
TLR9 endosomes DNA various
pathogens
(40)
(41)
Table 1.1: TLRs and their ligands.
27
Figure 1.2: TLRs and their adaptor proteins. Figure adapted from Kaway and Akira
(30).
28
1.1.4 DAMP
There is now an agreement that tissue damage induces the release of a variety of
“danger signals” that can activate inflammation. The term damage-associated
molecular pattern (DAMPs) was proposed by analogy with PAMPs expressed by
microorganisms.
As shown in Figure 1.3, DAMPs include endogenous molecules such as high
mobility group box chromosomal protein 1 (HMGB1), heat shock proteins (HSPs),
and non-protein molecules such as DNA, uric acid, IL-1 and ATP released from cells
in response to trauma, ischemia and tissue damage (42). In addition, cytokines and
cells activated by pathogens also induce secretion of DAMPs (43). S100 proteins
(group of 20 calcium binding proteins) are another example of DAMPs as
demonstrated by high levels of proteins in inflamed tissue (44). DAMPs are
implicated in different diseases as summarized in Table 1.2. For instance, uric acid
crystals (monosodium urate) are implicated in gout disease as demonstrated by
Garrod (45). The table below lists some DAMPs such as HMGB1, S100, and HSPs,
that have more in common with the proteins identified in our proteomics
experiments.
29
Figure 1.3: DAMPs release. Figure adapted from Tang et al. (42).
30
DAMP Disease Reference
HMGB1 Sepsis
Rheumatoid arthritis
Cancer
(46)
(47)
(48)
HSPs Rheumatoid arthritis
Sepsis
Multiple sclerosis
(49)
(50)
(51)
S100 proteins Psoriasis
Rheumatoid arthritis
Systemic sclerosis
(52)
(53)
(54)
Uric acid Gout (45)
Table 1.2: Some diseases associated with DAMPs.
31
1.1.4.1 HMGB1
HMGB1 is a non-histone nuclear protein composed by two DNA-binding domains
(HMGB boxes A with Cys23 and Cys45 and boxes B with Cys106) and an acidic tail
that contains glutamic and aspartic acid (55). HMGB1 can have intracellular or
extracellular localization with different functions. In particular, inside the nucleus it
binds DNA structure and regulates gene transcription. In addition to its nuclear role,
extracellular HMGB1 is implicated in sterile and infectious inflammation, cell
differentiation, cell migration and tumor metastasis.
It is probably that act as proinflammatory cytokine by binding TLR2 and TLR4 (56)
while the binding between receptor for advanced glycation end (RAGE) and
HMGB1 mediates tumour growth and metastases as suggested by in vivo
experiments performed by Taguchi et al. (48). Both sterile injury activated by DAMP
and infection activated by molecular pathogens induce release of HMGB1.
HMGB1 can undergo various forms of post-translational modifications, including
thiol oxidation. Changes in the redox state of the three conserved cysteines (Cys) of
HMGB1 (Cys23, Cys45 and Cys106) affect the extracellular protein function.
Experiments performed in CHO (Chinese hamster ovarian) cells showed the
formation of an intramolecular disulphide bond between Cys23 and Cys45 while the
conservative Cys106 was not oxidized. Experiments with mutated forms of HMGB1
(e.g. Ser23 and/or Ser45, Ser106) demonstrated that Cys106 is involved in the
nuclear localization of HMGB1 (57). Figure 1.4 shows that only HMGB1 containing
Cys106 reduced and a disulphide bond between Cys23-Cys45 is able to induce
cytokines release and then have an inflammatory role. Therefore, as demonstrated by
Yang et al. the reduced state of Cys106 is important for the binding with TLR4 and
then release of TNF by macrophages (58). The HMGB1 with all cytokines in a
reduced state binds the chemokine CXCL12 inducing recruitment of inflammatory
cells to the site of injury (59). During apoptosis all cysteines are not reduced and
HMGB1 have not chemotactic or inflammatory activity.
Glutathionylation of HMGB1 (to form HMGB1-SSG) has a dual role representing a
mechanism of regulation of protein function and protection from irreversible
32
oxidation. Glutathionylated HMGB1 at least in its nuclear form has been described
by Hoppe suggesting that the nuclear localization of HMGB1 is mediated by Cys106
(57). Furthermore, during the EMBO meeting workshop in 2006 the link between
HMGB1 secretion and glutathionylation was discussed. Mrp1 is involved in the
transport of glutathionylated HMGB1. A study on peritoneal macrophages Mrp1 -/-
did not show HMGB1 secretion (60).
33
Figure 1.4: Redox regulation of HMGB1. Figure from Yang et al. (55).
34
1.1.4.2 HSPs
HSPs are a group of proteins whose expression is markedly increased in response to
high temperature with intracellular or extracellular localization. Following a
modified version of the HUGO Gene Nomenclature Committee, HSPs can be
divided in HSP70 family, HSP110 family, HSP40 family, small HSP family, HSP90
family and human chaperonin family (61).
HSPs can have different functions depending on their localization. A major function
of intracellular HSPs is theirs chaperone activity, providing help for the formation of
a correct protein structure (folding of non-native proteins, protein refolding and
preventing protein aggregation) (62) while extracellular HSPs are implicated in
immunological functions. Secreted HSPs can be associated with antigenic peptides
and bind the receptor on dendritic cells. The following antigen interaction with MCH
I induces the T cells activation (63).
HSPs can be passively secreted by necrotic cells but not apoptotic cells, and actively
by different cells (64). In particular HSP70, a signal peptide-less protein, is actively
released from cells in both basal and stressed conditions through alternative
pathways. Lancaster et al. (65) showed that HSP70 can be released from peripheral
blood mononuclear cells using an exocytotic pathway (exosomes-dependent) instead
of the classical secretory pathway (ER/Golgi-dependent).
HSP70 is composed by two major domains: N-terminal nucleotide binding domain
(NBD) and a C-terminal protein substrate binding domain (SBD). The SBD has a
SBDβ subdomain which contains the peptide binding pocket and a lid region (SBDα)
that controls the access of the peptide to the substrate binding cavity. Therefore, the
chaperone function is due to the switch between the open conformation (lid open)
and the closed conformation (lid closed). Binding of ATP in the NBD domain
induces the open conformation (66) (67).
HSPs can undergo various forms of post-translational modifications, including thiol
oxidation. The change in the oxidation state of conserved cysteines in HSPs can
regulate their chaperone function. For instance, Jakob et al. showed that the
35
chaperone activity of HSP33 is inactivated by treatment with the reducing agent
Dithiothreitol (DTT). This inactivation of HSP33 is reversible and the formation of a
disulphide bond after treatment with GSSG (oxidized glutathione, GSH) or H2O2
causes reactivation of the chaperone activity (68). Another example of redox
regulation of HSPs was shown by Callahan et al. in experiments on HSP70 under
oxidative stress conditions. In the oxidized form, HSP70 has chaperone activity due
to a better accessibility of the peptide to the substrate binding cavity (69).
36
1.2 Inflammation
Inflammation is an important component of the innate immunity but an exaggerated
inflammatory response is at the basis of several inflammatory diseases. The four
cardinal signs of the inflammatory response are rubor (redness), tumor (swelling)
calor (heat) and dolor (pain) as described 2000 years ago by Celsus. Therefore, the
redness phase is caused by an increase of the blood flow (vasodilatation) and
vascular permeability. Histamine, prostaglandin and nitric oxide are chemical
mediators of inflammation, inducing vasodilation and increased permeability. The
tissue swelling is caused by recruitment of inflammatory cells at the site of infection
and accumulation of the exudate (fluid with high proteins content and antibacterial
properties). The release of cytokines (IL-1 and TNF) increases the levels of
leukocyte adhesion molecules on endothelial cells (70). Increased permeability of the
blood vessels allows the passage of cells from the vessel into site of inflammation.
Among the inflammatory cells, polymorphonuclear neutrophils (PMN) are the first to
be recruited and the migration of these cells required different steps which include
rolling, adhesion and transmigration (diapedesis). PMN bind the endothelium of the
blood cells through cell adhesion molecules (selectins) causing rolling motion along
the vessel wall, followed by a tight bind between PMN and endothelium using a
different receptor. The final step of PMN migration is the passage of PMN outside of
the blood vessel (71). Neutrophils can destroy the pathogens through phagocytosis
which include recognition and engulf of the pathogens in vacuole (phagosome)
provided by enzymes that produce components with cytotoxic activity. For instance,
NADPH oxidase complex produces superoxide and hydrogen peroxide while
myeloperoxidase enzyme produces hypochlorous acid (72) (73). The key role of
PMN in defense against bacteria is exemplified by the increased susceptibility to
infection in patients whose PMN lack the ability to generate ROS (chronic
granulomatous disease, CGD).
Activated neutrophils produce chemotactic factors that attract monocytes and
dendritic cells at site of infection (74) (75). Macrophages (derived from monocytes)
are another important component of inflammation for their role in antigen
presentation, phagocytosis of invading pathogens and production of inflammatory
cytokines. There are two possible ways of macrophage activation: the classical and
37
the alternative. In the classical activation, cytokines produced by Th1 cells (IFN-γ
and TNF) and LPS activate macrophages to produce nitric oxide as killing agent. In
the alternative activation, cytokines (IL-4, IL-10, and IL-13) produced by Th2 cells
activate macrophages to produce proliferative polyamines and proline for the
collagen production (76).
1.2.1 Inflammatory cytokines
Cytokines are a group of proteins that can be divided in proinflammatory or
antinflammatory cytokines based on their activity. An important role of cytokines is
to communicate with other cells to initiate the inflammatory response and defence
the host from pathogens. They are classified with different name according to their
origin, function or binding receptor. They can be classified in type 1 and type 2
cytokines when produced by Th1 and Th2 respectively. Th1 cells produce
proinflammatory cytokines such as IFN-γ and IL-2 in response to antigen +
presenting cells and stimulate activation of macrophages and then destruction of
bacteria and parasites through phagocytosis. Th2 cells are macrophages-independent
and stimulate the production of antibodies. Th2 cytokines include IL-4, IL-5, IL-6,
IL-10 and IL-13 (77) (78) (79). As summarized by Lucey et al. in Figure 1.5, type 1
and type 2 cytokines are not only produced by CD4+ T cells but also by other cells
such as macrophages, monocytes, B cells, and dendritic cells (80).
“Cytokine” is a general term that includes lymphokines (cytokines produced by
lymphocytes (81), monokines (cytokines produced by macrophages and monocytes).
They also include chemokines, cytokines with chemotactic activity (function to
attract neutrophils and monocytes to the site of inflammation). IFN (α, β and γ)
indicate antiviral cytokines (that directly interfere with virus replication) (82). From
the point of view of the nomenclature, there is an agreement to assign cytokines and
IL number (IL-1, IL-2 etc.) where the term “interleukin” generically points to
proteins acting as signals between different leukocyte populations).
Cytokines have different effects such as the production of antibodies, cellular
proliferation, cellular differentiation, chemotaxis, inflammation and phagocytosis.
For instance, TNF is involved in killing of intracellular pathogens. Bermudez et al.
38
demonstrated that TNF stimulate macrophages to kill Mycobacterium avium (83).
The role of TNF was also shown in human neutrophils where TNF induced the
enhanced of neutrophils activity (84). IL-1 is implicated in proliferation of T cells
while IL-6 induces B cells proliferation (85). Other cytokines functions are
summarized by Tayal and Kalra in Table 1.3 (86). Therefore, a stimulus can induce
the production of cytokines from different cells and the interaction between
cytokines and specific receptors is responsible for different biological effects. The
receptors can be expressed on the cytokine-producing cell (autocrine action) or
presents on a different target cells localized close (paracrine action) or distant
(endocrine action) to the cytokine-producing cell (87).
Cytokines receptors include: class I cytokine receptor family (or hematopoietin
receptor family), class II cytokine receptor family (or IFN receptor family), TNF
receptor family, chemokine receptor family and immunoglobulin superfamily
receptor. Binding of a cytokine to its receptor normally causes activation of a
signaling class 1 cytokines associated to Jaks (a family of kinase proteins). As shown
in Figure 1.6, the binding of the cytokine to the cytokine receptor activate the Jak-
Stat signaling pathway. Jaks phosphorylate the tyrosine residue of the receptor
activating the Stat proteins resulting in activation of transcription of target genes
(88).
39
Figure 1.5: Cytokines and their cells source. Figure adapted from Lucey et al. (80).
40
Cytokine Major function
IL-1 Proliferation and differentiation, pyrogenic, bone marrow cell
proliferation
IL-2 Proliferation and activation
IL-3 Hematopoietic precursor proliferation and differentiation
IL-4 Proliferation of B and cytotoxic T-cells, enhances MCH II
expression, stimulates IgG and IgE production
IL-5 Proliferation and maturation , stimulates IgA and IgM production
IL-6 Differentation into plasma cells, IgG production
IL-7 B and T-cell growth factor
IL-8 Chemotaxis, proinflammatory
IL-9 Growth and proliferation
IL-10 Inhibits cytokines and mononuclear cell function, antinflammatory
IL-11 Differentiation, induces acute phase proteins
IL-12 Activates NK cells
IL-18 Proinflammatory, induction of IFN-γ
IFN-α Anti-viral, anti-proliferative
IFN-β Anti-viral, anti-proliferative
IFN-γ Macrophage activation, increases neutrophil and monocyte function,
MHC I and II expression on cells
TNF-α Phagocyte cell activation, endotoxin shock, tumor cytotoxicity,
cachexia
TNF-β Chemotactic, phagocytosis, oncostatic, induces other cytokines
G-CSF Granulocyte production
GM-CSF Granulocyte, monocyte, eosinophil production
M-CSF Monocyte production and activation
EPO Red blood cell production
Table 1.3: Cytokines and their major function. Table adapted from Tayal and Kalra
(86).
41
Figure 1.6: Jak-Stat pathways in the signaling of class 1 cytokines. Figure from
O’Sullivan et al. (88).
42
1.2.2 Inflammatory diseases
Inflammation can be classified, according to the time course and the tissue damage in
acute (short term) and chronic (long term) inflammation. Acute inflammation is an
aggravating component of infections. Overproduction of inflammatory cytokines
such as IL-1 (89) and TNF (90) causes systemic inflammatory response syndrome
(SIRS; called “septic shock” in the past). SIRS represent the response to a stimulus
that can be an infection condition or a non-infectious condition (such as burn,
pancreatitis, surgery and trauma). SIRS associated with an infection is called sepsis
(91). SIRS induce symptoms such as fever or hypothermia, tachycardia and change
in blood leucocyte count. Severe and untreated sepsis can lead to septic shock with
vasodilatation, hypotension, hypoperfusion, and ultimately death (92).
Other pathological conditions that can result from infections-associated inflammation
are acute respiratory distress syndrome (ARDS) and multiple organ failure (MOF).
MOF and particularly ARDS are due to the action of inflammatory cytokines on
various tissues. For instance, elevated inflammatory cytokines levels were observed
in bronchoalveolar lavage fluid of patients with ARDS. Furthermore, ARDS is also
associated with accumulation of PMN and other inflammatory cells, edema and
tissue damage (the classical signs of inflammation) in the lung (93) (94).
Chronic inflammatory diseases include, for instance, atherosclerosis, inflammatory
bowel diseases, and pulmonary fibrosis (95). Activation of TLRs by self-component
(DNA or RNA) can contribute the symptoms of autoimmune diseases such as
systemic lupus erythematosus, rheumatoid arthritis (RA) and Sjögren's syndrome.
RA is a chronic inflammation ultimately resulting in joint damage. At the molecular
level, RA is associated with high levels of TNF that can have direct effects or induce
inflammatory cascade through the production of other inflammatory cytokines (IL-1,
IL-6 and IL-8) (96) (97). Feldmann et al. demonstrated that injection of anti-TNF
monoclonal antibody into mice with collagen-induced arthritis decreases
inflammatory damage of joints (98). As a result from this knowledge, anti-TNF
molecules (anti-TNF antibodies; soluble TNF receptors) are now approved therapies
for this disease and top selling biological drugs. In addition, inhibition of specific
TLR can also be useful to treat autoimmune diseases. Barrat et al. showed the
43
protective activity of new TLR7 and TLR9 inhibitors in autoimmune diseases (99).
Other new anti-inflammatory and anti-cytokine strategies are being evaluated in
these pathologies, such as the recently introduced anti-IL-R receptor antibody
(tocilizumab) or anti-IL-1beta (canakinumab).
Injury, infection or disease can induce inflammation in central nervous system
(CNS). Inflammation in the CNS, mediated by proinflammatory cytokines (e.g. IL-1,
IL-6 or TNF) production is also a pathogenic component of neurological diseases
such as multiple sclerosis (MS) or stroke. Inflammation is an aggravating mechanism
in ischemia/reperfusion injury in several tissues (cerebral ischemia, myocardial
infarction, transplantation-associated I/R injury). Experiments show that damage
associated with cerebral ischemia is decreased blocking the IL-1 or TNF-α effect
(100). Finally, inflammation represents an important component not only in acute but
also in chronic CNS disease such as MS producing demyelination and loss of
neuronal function (101).
The occurrence of inflammation in conditions, such as ischemia, where tissue injury
is not associated with infection or autoimmunity has raised the question of what
triggers sterile inflammation. In 1994, Matzinger first introduced the concept that the
immune system can be activated not only by foreign invaders but also by substance
made or modified by stressed or injured cells (102) (103). In this context, we could
define two different types of inflammation: inflammation induced by pathogens and
sterile inflammation. Sterile inflammation indicates an inflammation not induced by
microorganisms but caused by ischemia, trauma or chemically-induced injury. The
current theory is that during sterile inflammation, DAMPs activate the inflammatory
cascade by interacting with TLRs inducing the production of proinflammatory
cytokines as described for infections (104) (105).
Lukens et al. discussed the role of IL-1 in sterile inflammation. In particular, IL-1
released from necrotic cells can activate macrophages and neutrophils with cytokines
production and then induce sterile inflammation (106). In the case of IL-1,
inflammation is triggered through a different pathway involving the inflammasome
rather than TLRs (107).
44
1.3 Oxidative Stress
Molecular oxygen (O2) is an important component for all living cells that use it in
aerobic respiration to convert fat, proteins and carbohydrates to CO2, water and
generate energy as ATP. On the other hand, O2 can be reduced to reactive oxygen
species (ROS), highly reactive and harmful agents. ROS include superoxide anion
radical (O2.-), hydrogen peroxide (H2O2) and hydroxyl radical (HO
.). Oxidants can be
produced by phagocytic cells, with an antibacterial function, for instance neutrophils
use an NADPH oxidase system to generate superoxide, followed by superoxide
dismutation with production of H2O2 (weakly microbicidal). The hydroxyl radical is
extremely reactive and can be produced by superoxide and H2O2 by the Haber-Weiss
reaction (108).
In iron-catalyzed Haber-Weiss reaction below, the step 1 and step 2 describe the
Fenton reaction while the step 3 represents the net reaction (109).
(1) Fe 3+
+ O2.- Fe
2+ + O2
(2) Fe 2+
+ H2O2 Fe 3+
+ OH- + OH
.
(3) O2.- + H2O2 O2 + OH
- + OH
.
ROS are also produced by the mitochondrial respiratory chain during ATP synthesis
(110) in peroxisomes by the oxidation of fatty acids and during the reaction cycle of
cytochrome P-450 that directly reduces O2 to O2.-. However, ROS are unstable
intermediates and the constituents of living organism are a possible target of their
toxicity. In particular, ROS modify the nucleotide bases in DNA (formation of
double bond) and induce a strand breaks (111). By inducing peroxidation of
membrane lipid ROS induce changes in the permeability and fluidity of the
membrane ultimately leading to its rupture (112), while oxidation of proteins causes,
in the case of enzymes, their inactivation (113).
The term oxidative stress was first used by Halliwell (114) to define the imbalance
between the production of chemically reactive species (ROS and Reactive Nitrogen
Species, RNS) and antioxidants. Human antioxidant defence is represented by
enzymes such as Superoxide Dismutase (SOD), catalase, GSH peroxidase,
Glutathione Reductase (GR) and non-enzymatic antioxidants such as GSH vitamins
45
(A, C and E), carotenoid and uric acid. Furthermore, Selenium and Selenium-
compounds (selenoproteins and selenoenzymes containing Selenium as
selenocysteine) have an important antioxidant role (115). For instance, Selenium is
part of the catalytic site of GSH Peroxidase and a Selenium deficient diet induces a
decrease of its enzymatic activity and protein levels (116).
There are different forms of SOD depending on the metal associated (Copper, Iron,
Manganese and Zinc) and the intracellular distribution. The metal (M) is reduced-
oxidised while the superoxide radical is oxidised-reduced (117). SOD catalyzes the
reduction of two superoxide radical molecules to H2O2 and molecular oxygen:
M (n+1)
+O2.-
M n+
+ O2
M n+
+ O2.- (+2H
+) M
(n+1) + H2O2
Overall 2O2.-
+ 2H+ O2 + H2O2
Catalase catalyzes H2O2 degradation according to the reaction:
2H2O2 2H2O + O2
Peroxidases are also important in the detoxification of H2O2. The generic reaction of
a peroxidase is the following: H2O2 + DH2 2H2O + D (where D is a generic
reductant). Particularly important, in the context of this thesis, are Glutathione
Peroxidases that use GSH to protect cells reducing H2O2 to H2O while two GSH are
oxidized to GSSG, according to the following reaction (118).
H2O2 + 2 GSH H2O + GSSG
GSH (the main thiol antioxidant in the cell) is a small tripeptide of glutamic acid,
cysteine and glycine. The first step of GSH synthesis is the production of γ-
glutamylcysteine from L-glutamate and cysteine via the enzyme γ-glutamylcysteine
synthetase. The second step is the introduction of glycine amino acid via the enzyme
glutathione synthetase. Buthionine Sulfoximine (BSO) is used as GSH synthesis
inhibitor (as it is an irreversible inhibitor of γ-glutamylcysteine synthetase).
Glutathione can exist either in a reduced form with a free thiol group (GSH) or in an
oxidized form with a disulphide located between two identical molecules (GSSG).
GSSG can be reduced to GSH by GR. The intracellular environment is in a more
reduced condition because of the relatively high GSH concentrations, while the
extracellular environment is oxidizing one because of the absence of thiolic
46
antioxidants. The reduced/oxidized GSH ratio defines the redox state of the cell,
along with the redox state of thioredoxin (reduced/oxidized Trx), which is regulated
independently of the GSH/GSSG ratio (119). The concentration of GSH in cells is
~1-10 mM and the GSH/GSSG ratio is about 100:1 in normal conditions but under
oxidative stress the GSH/GSSG ratio decreases (120). In particular, Hwang et al.
calculated the GSH/GSSG ratio 100:1 as representative of entire cell, whereas in the
highly oxidizing ER environment the GSH/GSSG ratio is about 1:1-3:1 (121).
However, Morgan et al. suggested that GSH/GSSG ratio can be only representative
of the whole-cell without to consider the individual compartments (122).
As discussed above, the main function of glutathione in its reduced form (GSH) is to
act as an antioxidant and free radical scavenger. This has been extensively
demonstrated by many studies showing that GSH depletion increases the
susceptibility to the toxic action of various oxidants, and likewise, GSH repletion by
administration of its precursors such as N-acetyl cysteine (NAC) has a protective
effect in those models (123) (124).
1.3.1. Oxidative stress in diseases
Early from the development of the concept of ROS it was clear that when their
generation exceeds the antioxidant potential of the organism, oxidative damage
occurs. This has been well demonstrated with toxicants such as paraquat or carbon
tetrachloride, as well as with radiation toxicity and “oxygen toxicity” (the toxicity
observed upon exposure to high O2 levels) (125).
Several studies using in vivo or in vitro models of diseases, or based on the
measurement of markers of oxidative damage, have suggested that oxidative stress is
a component of various human diseases (126) (127). These include, for instance,
chronic obstructive pulmonary disease (COPD) (128), atherosclerosis (129), diabetic
complications (130), Parkinson’s disease (131), and MS (132).
In the context of this thesis, oxidative stress has been implicated as a component of
the pathogenesis of infective and inflammatory diseases. In this respect, it is
important to note that ROS are not only by products of the interaction of O2 with
cellular oxidoreductases (such as the mitochondrial respiratory chain) but are also an
47
important component of the antibacterial and possibly antitumoral armamentaria of
the immune system. In particular, PMN exposed to bacteria or phagocytic stimuli
respond with rapid oxygen consumption and ROS generation, called “oxidative
burst” as also described above (133) (134).
1.3.2. Oxidative stress and innate immunity
As mentioned above, phagocyte activation is accompanied by an oxidative burst with
production of ROS. The importance of ROS as bactericidal agents is demonstrated
by the fact that deficiency in the key enzyme, a NADPH oxidase (implicated in ROS
production by PMN) is at the basis of CGD (a pathological condition associated with
high susceptibility to infection and impaired bactericidal activity of PMN) (135).
Production of ROS has also been implicated in the cytolytic activity of NK cells,
whose main biological function is the killing of tumor cells (136).
Not only ROS are used as weapons by cells of the immune system, as discussed
above. Droge et al., in 1989 reported that people affected by ADS had low levels of
cysteine, the precursor of GSH (137). Herzenberg et al. showed that increasing the
GSH levels by oral administration of NAC can improve the survival of HIV patients
(138). In 1991 Patrick Baeuerle discovered that ROS activate the transcription factor
NF-kB while thiol antioxidants, including GSH and NAC inhibit its activation. NF-
kB, along with IRFs, is a key transcription factor activated by different stimuli
(bacterial products, viral products, inflammatory cytokines and oxidative stress)
(139) and responsible for the induction of different transcriptional genes (including
inflammatory cytokines, acute phase proteins and cell adhesion molecules). It is
composed by a DNA binding protein (P50), DNA binding protein (RelA formerly
P65) and an inhibitory subunit (IkBα) which is bound to RelA (140). Activation of
IkB kinase induces phosphorylation of IkBα and its dissociation from the DNA
binding proteins. DNA binding proteins translocate from the cytosol into the nucleus.
ROS can induce the degradation of IkBα and therefore activate NF-kB translocation
and transcription of target genes.
48
Various studies have shown that oxidants, particularly H2O2, augment the production
of inflammatory cytokines and/or that cytokines may promote further ROS release.
In particular, hydrogen peroxide induces:
IL-8 production by various types of cells including hepatoma cells,
pulmonary epithelial cells, and fibroblasts (141).
TNF-alpha in MCF-7 cells (142).
IL-32 expression in pulmonary epithelial cells (143).
MIF production by fibroblasts (144).
Chemokines MIP-1 alpha and MIP-2 mRNA in alveolar macrophages (145)
(146).
IFN-γ production by NK cells (147).
MCP-1 mRNA in endothelial cells (148).
IL-6 production by retinal pigment epithelial cells (149).
In vivo, ROS and oxidative stress in general have been implicated in several
inflammatory diseases as suggested by various evidences, including protective effect
of antioxidants in animal models or evidences of oxidative stress in animal models or
in patients. A growing number of papers suggest a connection between the role of
GSH and the inflammatory response (150). A non-exhaustive list is given in Table
1.4 below.
49
Disease Evidence/end-point Reference
Influenza infection Protective effect of antioxidants in a
mouse model
(151)
ARDS Protective effect of antioxidants in animal
model
(152)
HIV infection Glutathione depletion in the blood of
patients; effect of GSH repletion
(137)
Sepsis/septic shock Protective effect of antioxidants and
worsening effect of GSH depletion in
peritoneal sepsis in mice
(153)
Atherosclerosis antiatherogenic effect of the antioxidants
in mice
(154)
Alzheimer early increase in protein oxidation in
patients/protective effect of antioxidants
in mouse model
(155)
(156)
Pulmonary fibrosis elevated markers of oxidative stress in
patients with interstitial lung diseases
/protective effect of antioxidants in animal
models
(157)
(158)
Rheumatoid arthritis Altered redox state in patients’ T cells and
normalization by antioxidants; elevated
markers of oxidative stress in RA patients
(159)
(160)
Multiple sclerosis Protective effect of SOD in a mouse
model of optic neuritis
(161)
Ischemia/reperfusion Evidence of oxidative stress/protection by
antioxidants
(162)
Table 1.4: Infection/inflammation-associated diseases with evidence for a role of
oxidative stress.
50
1.4 Proteins thiols-disulphides metabolism in redox
regulation
High levels of ROS, diminished antioxidant defence enzymes and depletion of
antioxidants may result in oxidative stress and proteins represent a possible target of
oxidative damage. Proteins can undergo different types of oxidation, depending on
the amino acid targeted and their oxidation can be reversible or irreversible.
Irreversible oxidative modifications of proteins (e.g. carbonylation, nitration, protein-
protein cross-linking) are responsible for protein degradation and aggregation, while
reversible oxidative modifications (e.g. S-nitrosylation or glutathionylation) can have
a dual role, protecting cysteine from irreversible oxidation and modulating protein
functions (redox regulation) (163).
Protein modifications include sulfoxidation, carbonylation and nitration.
Sulfoxidation is a reversible oxidation of the amino acid methionine in proteins to
methionine sulfoxide which is reduced by the enzyme methionine sulfoxide
reductase (164). One example of a protein whose activity is inhibited by methionine
sulfoxidation is the alpha-1 proteinase inhibitor (165). Moskovitz et al. showed that
mice lacking the methionine sulfoxide reductase gene are more sensitivity to
oxidative stress compared to the wild type (166). Carbonylation is an irreversible
oxidative modification and different diseases such as Parkinson’s disease,
Alzheimer’s disease, cancer and sepsis are associated with high levels of
carbonylated proteins (167). Primary protein carbonylation is the direct oxidation of
the amino acid side chains of proline, arginine and lysine with introduction of a
carbonyl group. In the secondary protein carbonylation a reactive carbonyl
compound (oxidized carbohydrates and lipids) induce the protein modification. For
instance, oxidants act on lipid producing a lipid-derived aldehydes and ketones
which induces the modification of proteins (168). Finally, tyrosine can react with a
RNS to form 3-nitrotyrosine, in a reaction called nitration (169) and has been found
elevated in various diseases including atherosclerosis (170), and Alzheimer’s disease
(171).
Cys is the most chemically reactive natural amino acid in proteins and Cys-
containing proteins are particularly susceptible to various reversible or irreversible
51
oxidative modifications. As shown in Figure 1.7, a thiol group can undergo different
post-translational modifications. Typically, it can react with another Cys to form a
disulphide bond (-S-S-). A disulphide bond can be formed between two Cys residues
within the same protein (intramolecular cross-linking) or between two Cys residues
from different proteins (intermolecular cross-linking). Mixed disulphides can also be
formed between a protein Cys and the Cys of GSH (a process also termed protein
glutathionylation or glutathionylation) or with a Cys (protein cysteinylation). S-
nitrosylation represents the reaction between a thiol group and RNS to form a
nitrosothiol (SNO). Protein thiol can also be oxidized by ROS and RNS to sulfenic
acid (Cys-SOH), eventually followed by a second oxidation with formation of
sulfinic acid (Cys-SO2H). Sulfinic acids can be further oxidized to sulfonic acids
(Cys-SO3H). While all the other forms of oxidation are reversible (that is, the
protein’s Cys can be reduced back to the free thiol), sulfonylation is an irreversible
form of oxidation.
52
Figure 1.7: Oxidative modifications of proteins thiol. Figure adapted from Dalle-
Donne et al. (172).
53
1.4.1. Protein glutathionylation
An important mechanism of protein modification is glutathionylation (protein-SSG),
a reaction between GSH and the SH (sulfhydryl group) of proteins with formation of
a mixed disulphide. The two main reactions describing (in a simplified way and
without stoichiometric values) their formation from proteins and GSH or GSSG are
the following:
Direct oxidation
PSH + GSH PSSG
Thiol-disulphide exchange
PSH + GSSG PSSG + GSH
Figure 1.8 shows the glutathionylation of proteins through a thiol-disulphide
exchange (-SH S-S) between a protein thiol and GSSG (172).
Other mechanisms for the formation of glutathionylated proteins have been
postulated, for instance via intermediates such as sulfenic acids and via thyil radicals
(173).
Sulfenic acid intermediates
PSH + H2O2 PSOH + H2O
PSOH + GSH PSSG + H2O
Formation via thiyl radicals
Reaction between GSH or PSH and hydroxyl radical forms a thiyl radical (GS. and
PS. respectively). The thiyl radical can react with a radical (radical combination
reaction) or with a thiolate (150). The reactions are described below:
PSH + OH. PS
. + H2O
PS. + GS
. PSSG (radical combination)
PS. + GSH PSSG
. (reaction with thiolate followed by reaction with O2)
PSSG. +O2 PSSG + O2
.-
The fact that GSH can form mixed disulphides with proteins has been first described
for hemoglobin by Allen et al. (174). Several studies from the laboratory of Helmut
Sies investigated protein glutathionylation describing its increase in organisms
exposed to oxidative stress. Experiments performed on isolated rat hepatocytes
54
incubated with oxidants showed a specific thiolation of a 30 kDa cytosolic protein. In
particular, the protein thiolation was slow with menadione incubation while rapid
with diamide treatments (175). More experiments were also performed to quantify
the amount of GSH in hepatic protein mixed disulphides. These studies pointed out
that a significant amount of GSH is present as protein glutathionylation in the liver,
up to 30 nmoles/gram of tissue, under normal conditions (176).
In fact, many proteins can also be glutathionylated under basal conditions, supporting
a possible function of glutathionylation in redox regulation (177). Protein
glutathionylation/deglutathionylation may represent a general mechanism similar to
protein phosphorylation/dephosphorylation that can induce changes in the activity of
proteins (activation or inactivation) in a reversible manner (Figure 1.9).
Glutathionylation can have important effects on proteins, modifying function and
physiological processes. For instance, glutathionylated glyceraldehyde-3-phosphate
dehydrogenase (178) IkB kinase, creatine kinase (179) are inactivated, while
glutathionylation stabilizes HIV-1 protease (180), and activates matrix
metalloproteinase (1-8-9) (181).
55
Figure 1.8: Main mechanisms of glutathionylation. Figure adapted from Dalle-
Donne (172).
56
Figure 1.9: Redox regulation. Glutathionylation/deglutathionylation and analogy to
phosphorylation/dephosphorylation.
57
1.5 Redoxins
The redox balance of the cells is controlled by the redoxins family which include
several enzymes such as thioredoxins (Trxs), glutaredoxins (Grxs), Protein
Disulphide Isomerase (PDI) and Peroxiredoxins (Prxs). Many redoxins act as
antioxidant by reducing disulphide, while the main role of PDI is to act as oxidant,
facilitating the formation of structural disulphides and therefore the proper folding of
newly synthetized proteins. Trxs, Grxs, and PDI are characterized by having a redox
active motif -C-X-X-C- (two Cys separated by two other amino acids) in the active
site.
1.5.1 Thioredoxin
The main function of Trxs is the reduction of disulphide bonds in proteins and was
originally described as the essential cofactor for the enzymatic activity of
ribonucleotide reductase, where the main function of Trxs is to re-generate reduced
form of the enzyme during the catalytic cycle (182). The Trx system is composed by
Trx, Trx Reductase (TrxR) and NADPH. Trxs (e.g. mammalian cytosolic Trx1,
mitochondrial Trx2 and the sperm-specific Trx located in spermatozoa) are 12 kDa
proteins present in bacteria, yeast, plant and animal cells. They have antioxidant
activity, reducing protein disulphides and act as electron donor for Prxs, methionine
sulfoxide reductases and ribonucleotide reductases (183). Trxs can also regulate the
function of transcription factor such as NF-kB (184). Furthermore, the Trx system
has anti-apoptotic and pro-survival functions. Therefore, Trx system inhibitors such
as inhibitor of TrxR, inducing apoptosis in cancer cells, are being as potential
anticancer drugs (185).
As shown in Figure 1.10, Trx can switch from the oxidized form (Trx-S2), (where the
two Cys of the active site -C-X-X-C- for a disulphide) to the reduced form (Trx-
(SH)2). Trx-(SH)2 reduces the disulphide bond of a target protein and resulting in the
formation of Trx-S2 that is then subsequently reduced by the selenoprotein TrxR
using electrons from NADPH (186). Human Trx1 contains the Cys32 and Cys35 in
the -C-X-X-C- dithiol/disulphide active site and additional cysteines Cys62, Cys69
58
and Cys73. Casagrande et al. showed the glutathionylation of Trx (mixed disulphide
binding between GSH and Cys73) treating human recombinant Trx with GSSG
(thiol-disulphide exchange reaction) (187).
As shown in Figure 1.11, other post-translational modifications described for Trxs
are thiol oxidation, nitrosylation and nitration. Thiol oxidations include
intermolecular disulphide bond between two Cys73 of Trx molecules to form a dimer
(followed by inhibition of the Trx activity due to the inaccessible TrxR on the active
site) (188) and intramolecular disulphide bond formation between two Cys of the
same Trx (e.g. Cys 32 and Cy35 in the active site, as part of the catalytic cycle, or
Cys62 and Cys69) (189). Haendeler et al. showed nitrosylation of Trx on Cys69 and
its importance for the redox activity of Trx as demonstrated using a non-
nitrosylatable Trx (C69S) (190). The nitration of Tyrosine49 induces an irreversible
conformational change of Trx and then inhibition of its redox activity (191).
59
Figure 1.10: The Trx system. Figure adapted from Yamawaki et al. (186).
Glutathionylated protein is reduced by Trx(SH)2, which is reduced by TrxR and
NADPH.
60
Figure 1.11: Post-translational modification of Trx. Figure from Ago and Sadoshima
(189).
61
1.5.2 Glutaredoxin
The Grx system comprises Grx, GSH and GR. While Trx reduces various types of
disulphides bonds in proteins, Grx is rather specific for reducing the mixed
disulphide of glutathionylated proteins. Mammalian cells contain Grx1 (cytosolic),
Grx2 (of which the Grx2a isoform is located in the mitochondria and the Grx2b
isoform in the nucleus), Grx3 (cytosolic and nuclear localization) and Grx5. (192).
Grxs contain a -C-X-X-C- motif in the active site (Grx1: Cys-Pro-Tyr-Cys; Grx2:
Cys-Ser-Tyr-Cys) where the two Cys are oxidized while reducing the disulphide
bond of the target protein (193). The mechanism of the enzymatic catalysis of the
deglutathionylation of a glutathionylated protein by Grx can be described in two
steps: 1) formation of a Grx-SSG intermediate; 2) reduction of a Grx-SG mixed
disulphide by GSH (194) (Figure 1.12). The oxidized GSH is then recycled to GSH
by NADPH dependent the enzyme GR. In addition, Grx2 can use both GSH and
TrxR as electron donor.
As shown in Figure 1.13, Grx1 contains the Cys23 and Cys26 in the active site and
additional cysteines Cys83, Cys79 and the Cys8 presents in human Grx1 but not in
the Grx1 from bovine, rabbit and pig. Grx2 contains Cys37 and Cys40 in the active
site and, in man, mouse and rat, two additional cysteines, Cys28 and Cys113. Post-
translational modifications of cytosolic Grx1 include oxidation (disulphide bond
between Cys8 and Cys79, Cys8 and Cys83 or Cys79 and Cys83) and nitrosylation of
Cys83. Hashemy et al. showed that the activity of Grx1 is decreased by treatment
with GSSG and H2O2, and can be restored after reduction with DTT.
Glutathionylation of Grx1 by reaction of GSH with the cysteines outside the active
site (Cys8, Cys79 or Cys83) can modify the structure and then the activity of Grx1.
(195).
62
Figure 1.12: The Grx system. Figure from Pillay et al. Glutathionylated protein is
reduced by Grx(SH)2, which is reduced by GSH. GSG can be regenerated from
GSSG using GR and NADPH (196).
63
Figure 1.13: Post-translational modifications of Grx1 and Grx2. Figure from
Hashemy et al. (195).
64
1.5.3 Protein Disulphide Isomerase
The endoplasmic reticulum (ER) represents the organelle where protein folding takes
place, so that only proteins with a correct conformation can transit to the Golgi. The
ER contains enzymes of the PDI family, which include several proteins (19 PDI
members have been described in humans). These proteins have an oxidase activity,
catalysing the formation of structural disulphide bonds. In addition, some PDI
members have isomerase activity (capable of structural rearrangement of incorrectly-
formed disulphide bonds) (197). Finally, PDI also act as molecular chaperones that
assist in the no covalent folding for nascent proteins (198).
As shown is Figure 1.14, PDIs has five domains (a, b, b’, a’, and c) and the domain a
and a’ have an active site motif -C-X-X-C- with two cysteine residues (-Cys-Gly-
His-Cys) (199). The active site motif is essential for the isomerase activity of PDI,
whereas modification of the active site does not influence the chaperone activity
(200). PDIs can exist in oxidized form (PDI-S2) and reduced form (PDI-(SH)2) and
its redox state regulate reaction of oxidation, reduction or isomerization as shown
below (201).
Disulphide formation:
Protein-(SH)2 + PDI-S2 Protein-S2 + PDI-(SH)2
Disulphide isomerization:
Protein-(S1-S
2)-(S
3-S
4) + PDI-(SH)2 Protein-(S
1-S
3)-(S
2-S
4) + PDI-(SH)2
Post-translational modifications of PDIs include nitrosylation (PDI-SNO) and
glutathionylation (PDI-SSG) and can reduce their activity and consequently cause an
accumulation of unfolded proteins, thus leading to the activation of an unfolded
protein response (199).
65
Figure 1.14: PDI domains structure.
66
1.5.4 Peroxiredoxins
Prxs or Trx peroxidases are antioxidant enzymes involved in protection of cells
against oxidative stress by reducing H2O2 to H2O (peroxidase activity) (202). The
schematic reaction of a peroxidase in the elimination of H2O2 is as follows:
H2O2 + Dred +2H+
H2O + Dox
where D is any electron donor. While GSH peroxidases use GSH as the electron
donor, producing GSSG, Prxs use TRX-(SH)2 as the electron donor, thus generating
TrxS2. Oxidised Trx is then regenerated by TrxR.
However, Prxs can catalyse other reactions. Peroxynitrite, a product between
superoxide and nitric oxide, can also be a substrate of Prxs (in a reaction where
ONOO- is reduced to NO2
-). Mammalian cells express six distinct Prxs, which differ
in structure, mechanism of action and mode of regulation. They are intracellular
enzymes with a different localization within the cell, for instance Prx1 is in both
cytosol and nucleus, Prx2 is predominant in the cytosol, while Prx3 is present in
mitochondria. Prx4, the only one to have a signal peptide for secretion, is present in
the plasma and identified also in the Golgi apparatus. Prx5 has only a monomeric
form and is localized in mitochondria, peroxisome and cytosol, while Prx6 only in
cytosol (203) (204).
Prxs are divided into three major groups:
1) Typical 2-Cys Prxs (Prx1, 2, 3 and 4) that contain two conserved redox active
cysteines (peroxidatic cysteine; Cys-SpH) at the N-terminal region and the
resolving cysteine (Cys-SrH) at the C-terminal region.
2) Atypical 2-Cys Prx (Prx5) which has a monomeric structure with one
conserved peroxidatic cysteine at N-terminal region and its resolving cysteine
present within the same polypeptide.
3) 1-Cys Prx (Prx6) which contains only one cysteine residue involved in
catalysis.
Their mechanism of action is described in Figure 1.15 by Fujii and Ikeda (203). In
the first step, common to all Prxs, Cys-SpH is oxidised to a sulfenic acid (Cys-
SpOH). In the second step the Cys-SpOH of 2-Cys Prxs can react with the Cys-SrH
67
of another monomer to form an intermolecular disulphide bond (homodimeric
structure). In contrast, the second step for the atypical 2-Cys Prx is slightly different.
In this case, in fact the Cys-SpOH is attacked from Cys-SrH present in the same
monomer and forms an intramolecular disulphide. In particular, the peroxidatic
cysteine of Prx2 is Cys51 that interacts with Cys172 (resolving cysteine) of another
subunit after its oxidation (205).
In addition, in 1-Cys Prx (Prx6) the peroxidase catalysis is performed using a
different reaction mechanism. In fact, as described in Figure 1.15, the only Cys
involved in the catalytic action of Prx6 reacts with an electron donor and is oxidized
to sulfenic acid. Following this reaction, there is a formation of a disulphide bond
due to interaction of the oxidized group with a thiol compound (e.g. Trx). The Prxs
can be recycled by thiol donor molecules (e.g. GSH and thioredoxin) as shown in
Figure 1.15. In particular, typical 2-Cys Prxs and atypical 2-Cys Prx use Trx to
reduce the disulphide bond of Prxs while GSH and cyclophilin A have been proposed
as electron donors for 1-Cys Prx (204).
While Prx5, as mentioned above, can be found as a monomer while the oligomeric
structure of the other Prxs is more complex. As shown in Figure 1.15, Prx5 has a
different mechanism compared to typical 2-Cys Prxs. In particular, Prx5 can only
form intramolecular disulphide bond between cysteines in the same monomer while
typical 2-Cys can form intermolecular disulphide bond and then dimers. Prx6
contains only one Cys at the N-terminal position and can form an intermolecular
disulphide bond with GSH during the catalytic cycle (203). Modification in the
structure of Prxs subunit (presence of monomer, dimer or decamer), under oxidative
stress or during the catalytic cycle, are analysed by non-reducing or reducing Sodium
dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE gel). Several
articles show the switching of typical 2-Cys Prxs from monomer to dimer by
treatment with H2O2. Peskin et al. demonstrated that oxidation of Prx2 by H2O2 forms
a disulphide-bonded homodimer. In particular, Western blot in non-reducing
conditions using an antibody against Prx2 showed a band corresponding to the dimer,
while in reducing conditions Prx2 was only migrating as a monomer (206). Cox et al.
showed the redox transformation of Prxs with formation of disulphide bond by H2O2.
In contrast, in reducing conditions, the disulphide bond was reduced and only the
monomer form was visible. Under overoxidation conditions, typical 2-Cys Prxs are
68
monomeric due to the presence of cysteines in the sulfinic acid state and then unable
to form a disulphide bonds (207).
Under normal conditions, Prx1, Prx2 and Prx3 exist as monomer and dimer but in a
different ratio (Prx1 predominant as monomer, Prx2 predominantly as dimer and
Prx3 half monomers and half dimers). A study by Schröder et al. also confirmed the
redox modification of Prxs. In particular, analysis of Prxs from rat hearts perfused
with H2O2 showed that the oxidant induces the formation of a disulphide bond and
then Prx1 and Prx2 run as dimer in non-reducing SDS-PAGE gel. Overoxidation by
H2O2 of Prx1 and Prx2 induces the formation of a decameric structure that migrates
as MW > 250 kDa band. Prx5 runs in a gel as a monomer both under basal
conditions and after exposure to H2O2 but the formation of an intramolecular
disulphide bond forms a small monomer that runs faster. Prx6 runs in a gel as a
monomer under both basal and oxidative condition (208).
69
Figure 1.15: Mechanism of action of Prxs. Figure adapted from Fujii and Ikeda
(203).
70
Prxs have dual functions, firstly the peroxidatic activity as described above, and
second they can also behave as molecular chaperones (e.g. assisting the folding of
newly synthesized proteins and preventing proteins aggregation), switching from one
activity to another. They can undergo various forms of post-translational
modifications, not limited to thiol oxidation. Overoxidation of the sulfenic
intermediate to the sulfinic acid derivate (Cys-SO2H) inactivates Prxs (209). In
addition, a decrease of Prxs activity can be induced by phosphorylation (210) and
proteolysis. In a similar way to phosphorylation, glutathionylation is another possible
mechanism for Prxs regulation. In particular Fratelli et al. (177) have shown
glutathionylation of Prx1 in human T lymphocytes treated with diamide, while a
recent study (211) found glutathionylation of Prx1 and Prx2 after treatment with
H2O2 in Hela cells (cervical carcinoma cells). Sulfinylated Prxs can then be reduced
and thus reactivated by sulfiredoxin (212). Glutathionylation induces structural
modification involving predominant formation of dimers and loss of chaperone
activity. Experiments by Park et al. (213) have shown that deglutathionylation of 2-
Cys Prxs is catalysed by sulfiredoxin. Structural modification of Prxs, changing from
small oligomers to decameric structure (pentamers of dimers), confers chaperone
activity (204).
Prx2, initially identified as a cytoplasmic protein named calpromotin, is the third
most abundant protein in erythrocytes (0.25 mM; 5 mg/ml) (214) Numerous studies
on Prx2 knockout mice highlight the importance of its protective role against the
oxidative stress. In particular, Prx2 knockout mice develop haemolytic anemia (215)
as a consequences of hemoglobin endogenous oxidation. Lee et al. demonstrated
using Prx2 knockdown that Prx2 is also involved protecting cancer cells from cell
death (216). Furthermore, high levels of Prx2 causes resistance to ionizing radiation
in MCF-7 breast cancer cells (217) and resistance to cisplatin and anticancer agents
in SNU638 gastric cancer cell line (218).
71
1.6 Aims of the study
The overarching hypothesis for this study was that protein glutathionylation can
target proteins that are released extracellularly by inflammatory cells. Thus the
primary aim of the study was to identify glutathionylated proteins eventually released
by LPS-stimulated macrophages. This required setting up methods for labelling
glutathionylated proteins and to identify them in proteomics experiments. Once these
proteins have been identified, a secondary aim was to characterize their biological
activity in the context of inflammation. For this purpose, the recombinant
preparations of the proteins identified were tested to study their effect on TNF-α
production by macrophages. All methods used to detect and study glutathionylated
released proteins are described in Chapter 2. The specific aims of each chapter are
detailed below.
Chapter 3
Experiments of Chapter 3 focused on HMGB1, a nuclear protein also known to be
released and involved in inflammation.
The main aim was:
To investigate whether extracellular HMGB1 can be glutathionylated upon
exposure to LPS.
The labelling of the intracellular GSH pool with BioGEE followed by gel
electrophoresis and immunoprecipitation were used to test this hypothesis. A
Western blot with Streptavidin Peroxidase (Streptavidin–POD) conjugate was also
performed to confirm the presence of proteins labelled with biotin using the BioGEE
method.
Chapter 4
The aim described in Chapter 4 was:
To identify proteins released in glutathionylated form from LPS-stimulated
macrophages (Raw 264.7 cells).
Different released proteins were identified and Prx2 was selected as the protein of
interest for further studies. The BioGEE labelling method and SDS-PAGE followed
72
by tryptic digestion and MS were used to identify the glutathionylated proteins
released.
Chapter 5
The three main aims of the experiments described in Chapter 5 were:
To confirm the release of Prx2 by Raw 264.7 cells using Western blot.
To extend the experiments to human cells (human macrophages and PBMC)
measuring the Prx2 released by Enzyme-Linked Immunosorbent Assay
(ELISA).
To extend the experiments to an in vivo model of inflammation, measuring
Prx2 levels in serum of mice after LPS injection.
Chapter 6
The two main aims described in Chapter 6 were:
To confirm the glutathionylation of Prx2 by immunoprecipitation with anti-
Prx2 antibodies followed by Western blot with anti-GSH antibodies.
To determine whether modification of intracellular GSH levels affected the
release of Prx2.
To this purpose, various methods of GSH depletion, GSH assay and Western blot
were used.
Chapter 7
The main aim in Chapter 7 was:
To study the possible inflammatory role of released Prx2.
This was done by evaluating TNF-α production in Raw 264.7 cells and human
macrophages stimulated with the human recombinant Prx2 (hrPrx2). TNF production
was measured by ELISA.
73
Chapter 2. Materials and Methods
74
2.1 Materials
2.1.1 Instruments
Product Name Supplier
Bioworks v3.3.1 Thermo Scientific (Bremen, Germany)
Block heater SBH130 Stuart (Chelmford, Essex, UK)
Heraeus fresco 17 centrifuge Thermo Scientific (Waltham, MA, US)
Heraeus multicentrifuge 3SR+ centrifuge Thermo Scientific (Waltham, MA, US)
Incubator Heracell Thermo Scientific (Waltham, MA, US)
Inverted Microscope CKX41 Olympus (Southend-on-Sea, Essex, UK)
LTQ Orbitrap hybrid FTMS Thermo Scientific (Bremen, Germany)
Magnetic stirrer C-MAG MS 7 IKAMAG Ika (Staufen, Germany)
Microplate reader Synergy HT Biotek (Winooski, VT, US)
Mini-PROTEAN Tetra Cell electrophoresis
apparatus
Bio-Rad (Hemel Hempstead, UK)
Mini-Trans-Blot electrophoretic transfer cell
apparatus
Bio-Rad (Hemel Hempstead, UK)
pH Meter 3510 Jenway (Staffordshire, UK)
Platform rocker STR6 Stuart (Chelmford, Essex, UK)
Reverse phase column (Acclaim PepMap100,
C18)
Dionex, Thermo Scientific (Bremen,
(Germany)
Ultimate U3000 nano-LC system Dionex, Thermo Scientific (Bremen,
(Germany)
X-ray film cassette (Hypercassette) GE Healthcare (Chalfont St. Giles, UK)
75
2.1.2 Chemicals, Reagents, and Kits
Product Name Supplier
2-Mercaptoethanol Sigma
30% Acrylamide/Bis Solution 29:1 Bio-Rad
5,5’-Dithiobis (2-nitrobenzoic acid) Sigma
Acetic Acid Acros organics
Albumin Sigma
Ammonium Persulphate Sigma
BioGEE Invitrogen
Bovine Serum Albumin Sigma
Bromophenol blue Sigma
Buthionine Sulfoximine Sigma
CellTiter-Blue Cell viability assay Promega
Coomassie (GelCode Blue Stain Reagent) Thermo Scientific
CytoTox 96® Non-Radio Promega
DC Protein Assay kit Bio-Rad
Detoxi-Gel Endotoxin Removing columns Thermo Scientific
Dimethyl Sulfoxide Fisher
Disodium phosphate Fisher
Dithiothreitol Sigma
ECL Blotting Detection Reagents GE Healthcare
Ethanol Sigma
Fetal Bovine Serum Invitrogen
Ficoll (Lympholite-H) Cedarlane
Full range molecular weight marker rainbow GE Healthcare
GBX developer and replenisher Kodak
Glutamine Invitrogen
Glutathione (oxidized) Sigma
Glutathione (reduced) Sigma
Glutathione Reductase (GR) from yeast Roche
Glycerol Sigma
Glycine for electrophoresis Sigma
Human Prx2 ELISA kit Antibodies-online
Human TNF-alpha DuoSet ELISA kit R&D Sistems
Hybond ECL nitrocellulose membrane GE Healthcare
76
Hyperfilm ECL GE Healthcare
Limulus Amebocyte Lysate, Pyrogent Plus test Lonza
Lipopolysaccharides from Escherichia coli 055:B5 Sigma
Methanol Fisher
Mouse TNF-alpha DuoSet ELISA kit R&D Sistems
NADPH Tetrasodium salt Roche
N-Ethylmaleimide Fluka
Nitrocellulose membrane (Hybond ECL) Amersham Biosciences
Opti-MEM I medium Life Technologies
PBS 1X PAA laboratories
Penicillin-Streptomicin Gibco
Polymyxin B Sulfate Salt Sigma
Ponceau S solution Sigma
Protein G agarose Pierce
RPMI 1640 medium PAA laboratories
Silver Quest (Silver Staining Kit) Invitrogen
Sodium Deoxycholate Sigma
Sodium dihydrogen orthophosphate dehydrate Fisher
Sodium Dodecyl Sulphate Sigma
Streptavidin immobilized on Agarose CL-4B Fluka
Temed Sigma
Trichloroacetic Acid Sigma
Trizma base Sigma
Trizma hydrochloride Sigma
Tween 20 Fisher
Vivaspin 500, 6, 20 (centrifugal concentrator) Sartorius
Western Blot Stripping buffer Thermo Scientific
77
2.1.3 List of Antibodies
Primary antibodies
Anti-GSH (Virogen): IgG2a mouse monoclonal antibody for GSH-protein
complexes. Protein A purified.
Origin: Mouse.
Anti-HMGB1 (2G7): mouse anti-HMGB1 monoclonal antibody was kindly
provided by Dr. Kevin Tracey (The Feinstein Institute for Medical Research, NY).
Anti-Prx2 (Sigma): produced in rabbit, affinity isolated antibody. Polyclonal
antibody.
Streptavidin-POD conjugate (Roche).
Secondary antibodies
Anti-mouse (Stressgen): goat anti-mouse IgG (Fab), HRP-conjugated.
Immunoaffinity purified.
Anti-rabbit (Sigma): anti-rabbit IgG (whole molecule)-peroxidase antibody
produced in goat.
Antibody Working dilution
Anti-GSH 1:1000
Anti-HMGB1 2 µg/ml
Anti-mouse 1:5000
Anti-Prx2 1:1000
Anti-rabbit 1:160000
Streptavidin-POD 1:25000
Table 2.1: Dilutions recommended in the product datasheet for Western blot
analysis.
78
2.1.4 Human recombinant proteins and Biotinylated oxidized
Glutathione
The hrPrx2 used for the experiments to measure the TNF-α production, was a gift of
Dr. C.H. Lillig (Philipps University Marburg, Germany).
The human recombinant HMGB1 (hrHMGB1) was a gift of Kevin Tracey’s
laboratory.
Biotinylated oxidized GSH (BioGSSG) used for glutathionylation of HMGB1 was a
kind gift from Professor Philip Eaton (King’s college, London. UK).
2.1.5 Cells
Raw 264.7 cells are a macrophage-like, Abelson leukemia virus transformed cell line
derived from BALB/c mice and was kindly provided by Dr. Jon Mabley (University
of Brighton, UK).
Peripheral Blood Mononuclear Cells (PBMCs) were isolated from healthy blood
donors as described in Methods (2.2.1). The study was approved by the local ethics
committee and all donors gave written informed consent.
Primary human macrophages were a gift of Dr. Sandra Sacre (Brighton and Sussex
Medical School, UK). They were obtained as described in Methods (2.2.1). The
study was approved by the local ethics committee and all donors gave written
informed consent.
79
2.2 Methods
All the techniques to investigate released glutathionylated proteins and its role have
been described in this chapter. These included cell culture, proteomic techniques
(e.g. Western blot and Mass Spectrometry), ELISA, immunoprecipitation, and
specific assay to detect GSH levels.
2.2.1 Cell culture
Macrophages and monocytes play an important role in inflammation, so all
experiments were conducted using macrophages known to respond to LPS (agonist
of TLR4) with cytokine and ROS production (219). Therefore, experiments were
performed treating Raw 264.7 cells, a murine macrophages cell line with LPS.
Furthermore, the study was also extended to human macrophages and PMBC.
Raw 264.7 cells
Raw 264.7 cells were cultured under standard incubation conditions (37˚C and
5% CO2) in complete RPMI 1640 medium (Roswell Park Memorial Institute
medium) containing 100 U/ml penicillin, 100 µg/ml streptomycin, 2 mM L-
glutamine and 10% heat-inactivated Fetal Bovine Serum (HI FBS). Cells were
splitted (1:3 ratio) every 3 days with a cell scraper and plated as indicated in Table
2.2A.
In particular, all experimental treatments of cells followed by Western blot analysis
(or MS, GSH assay and immunoprecipitation) were carried out in serum-free Opti-
MEM I medium which contains growth factor instead of HI FBS. For the
experiments to measure the TNF-α production, cells treatments were performed in
complete RPMI 1640 medium.
80
Peripheral blood mononuclear cells
Peripheral Blood Mononuclear Cells (PBMCs) were isolated from heparinized
venous blood of healthy volunteers with informed consent. Briefly, blood was diluted
in equal volume with PBS and then Ficoll (Lympholite-H) was added to allow the
cell separation after centrifugation at 2000 rpm for 20 min. From the resultant layers
(from top to bottom: plasma-platelets; PBMC; Ficoll and red blood cells with
granulocytes), the PBMC was separated and diluted with PBS before centrifugation
at 1500 rpm for 10 min. The pellet (mononuclear cells) was washed twice with PBS
by centrifugation at 1000 rpm for 10 min and then resuspended in RPMI 1640
complete with 5% HI FBS. Cells were plated in 96-well plates at 0.5 x 106
for the
experiments to measure the TNF-α production as indicated in Table 2.2B.
Primary human macrophages
Primary human macrophages were differentiated from peripheral blood monocytes in
presence of 100 ng/ml human recombinant macrophage colony-stimulating factor
(M-CSF) for 4 days. Briefly, PBMC collected by Lympholite-H separation were
washed, resuspended in Hanks’ balanced salt solution (HBSS), layered with
isosmotic Percoll plus and centrifuged at 2200 rpm for 30 min. Monocytes were
recovered on the top of the gradient, washed with HBSS and resuspended in RPMI
1640 medium complete with 5% HI FBS. Cells were plated for the experiments to
measure the TNF-α production as indicated in Table 2.2 B. Human macrophages
were seeded in 96-well plates at 0.2 x 106 in 100 µl RPMI 1640 complete. After ON,
100 µl medium containing cell treatments were added and then the supernatant was
analysed by ELISA.
81
Plate Cells/well Growing
medium
Working
medium Method
96-well
(Raw 264.7)
3-5 x 10
4
RPMI 1640
+10% HI FBS
(100 µl) ON
Added 100 µl RPMI 1640
+10% HI FBS
with treatments (tot. 200 µl)
ELISA
Cell viability
24-well
(Raw 264.7)
0.3 x 106
RPMI 1640
+10% HI FBS (1 ml)
ON
Washes and
changed
medium with Opti-MEM I
(200-300 µl)
Western blot
6-well (Raw 264.7)
1.0-1.4 x 106
RPMI 1640 +10% HI FBS
(3 ml)
ON
Washes and
changed medium with
Opti-MEM I
(1 ml)
MS GSH assay
IP
(A)
Plate Cells/well Growing
medium
Working
medium Method
96-well (human
macrophages)
0.2 x 106
RPMI 1640 +5% HI FBS
(100 µl)
ON
Added 100 µl
RPMI 1640 +5% HI FBS
with treatments
(tot. 200 µl)
ELISA
96-well (PBMC)
0.5 x 10
6
RPMI 1640
+5% HI FBS
(100 µl) ON
Added 100 µl RPMI 1640
+5% HI FBS
with treatments (tot. 200 µl)
ELISA
(B)
Table 2.2: (A) Plating densities for Raw 264.7 cell culture. (B) Plating densities for
primary human macrophages and PBMC.
Abbreviations word: ELISA (Enzyme-Linked Immunosorbent Assay), FBS (Fetal
Bovine Serum), GSH (Glutathione reduced), HI (Heat-inactivated), IP
(Immunoprecipitation), MS (Mass Spectrometry), ON (Overnight), PBMC
(Peripheral Blood Mononuclear Cells) and RPMI 1640 (Roswell Park Memorial
Institute).
82
2.2.2 Cell viability assay
Cell viability was measured by CellTiter-Blue (CTB) assay following the
manufacturer’s instructions. This method is based on the ability of mitochondrial
dehydrogenases in viable cells to convert resazurin (a blue non fluorescent dye) into
resofurin (pink fluorescent dye). The fluorescence produced is proportional to the
number of viable cells.
In order to detect cell viability with this method, Raw 264.7 cells were cultured in
96-well plates at density of 5 x 104/well in complete RPMI 1640 following the
experimental scheme. CTB reagent was added directly to the cells (20 μl/well) and
incubated for 2-4 hours at 37°C. Fluorescence was measured (530/25 nm excitation
filter, 590/35 nm emission filter) using a microplate reader.
2.2.3 LDH assay
Quantification of lactate dehydrogenase (LDH) in the supernatant is a method to
investigate the cytotoxicity of a substance based on the fact that LDH is a cytosolic
enzyme, realised only upon cell lysis. LDH release from cells was measured by the
CytoTox96 colorimetric assay. The LDH released converts a tetrazolium salt (INT)
into a red formazan product.
LDH was measured in conditioned medium of Raw 264.7 cells with and without
incubation of 100 ng/ml LPS. Cells were plated in 24-well plates at 0.3 x 106 in
RMPI 1640 complete and treatments were carried out in Opti-MEM I medium for 24
hours.
96 well-plate reaction:
An assay was performed in 96-well plates in three main steps. In the first step of the
assay, medium with substrate mix was taken as blank and LDH plus substrate mix as
a positive control. Sample was prepared using the supernatant (SN) from treated or
untreated cells with substrate mix. Finally the 100% LDH released from the cells was
83
obtained by adding 20 µl/well of lysis buffer for 1 hour at 37°C and then mixed with
substrate mix. Note: in the above reaction, 50 µl substrate mix (provided in the
CytoTox96 kit) and 50 µl compounds were used. In the second step, the reaction was
stopped by adding 50 µl Stop Solution (provided in the CytoTox96 kit). In the final
step, the optical density reading at 490 nm was obtained using a microplate reader.
Reagent preparation
Reconstitute substrate mix: 12 ml of assay buffer to a bottle of substrate mix
LDH (bovine heart LDH): 2 µl of LDH stock into 10 ml of PBS (1:5000 dilution).
Five two-fold serial dilutions in PBS starting from LDH diluted 1:5000.
Calculations
The average value obtained measuring LDH release in medium without cells (blank)
was used to remove the background from the others samples whilst LDH was added
to separate wells as positive control to verify the performance of the assay.
Furthermore to obtain the maximum release of LDH (Total LDH, 100%) to compare
with LDH released in the supernatant, cells were lysed by adding lysis buffer for
1 hour at 37°C. The final mean value is expressed as percentage (%).
84
2.2.4 Protein glutathionylation
Proteins can be glutathionylated with different approaches, for instance using GSH in
presence of oxidant (e.g. H2O2 or diamide) or with a thiol-disulphide exchange
mechanism (glutathionylation of protein thiols using GSSG). In order to obtain
glutathionylated HMGB1, it was treated with biotinylated oxidized GSH (BioGSSG)
using all the advantages of biotinylated proteins detection.
Briefly, HMGB1 (30 µM) was incubated in presence of 5 mM BioGSSG at RT for
2 hours in H2O. BioGSSG-sample treatment was followed by incubation with thiol-
specific alkylating reagent N-Ethylmalemide (NEM) to a final concentration of
5 mM. In order to detect glutathionylated HMGB1 by Western blot, the protein
previously treated with BioGSSG was separated by 12% SDS-PAGE under non-
reducing conditions or with 10% 2-Mercaptoethanol (2-ME) (reducing conditions)
and biotinylated protein was detected by a Streptavidin-POD Western blot.
2.2.5 Proteomics of released glutathionylated proteins
Released glutathionylated proteins were analysed by a two steps procedure: the first
step (2.2.5.1) involved the cell labelling with BioGEE, a membrane-permeable
analogue of GSH that is transported into cells and is hydrolysed to form intracellular
GSH. In the second step (2.2.5.2, 2.2.5.3, 2.2.5.4 and 2.2.5.5) after the incorporation
of BioGEE, biotinylated proteins were purified from the protein complex using
streptavidin-agarose beads and then identified by a combination of one-dimensional
gel electrophoresis and MS. Western blot with Streptavidin–POD was also
performed to confirm the biotinylation of proteins.
2.2.5.1 BioGEE cellular uptake and treatments
All experiments to detect glutathionylated proteins using the BioGEE method were
performed in Raw 264.7 cells. Raw 264.7 cells were plated in 6-well plates as
described in Table 2.2 and treatments were carried out in 1 ml Opti-MEM I medium.
Cell were pre-incubated with BioGEE (200 µM) for 1 hour and then treated with
85
LPS 100 ng/ml for 24 hours to allow an inflammatory response. A stock solution of
BioGEE (100X concentrated) was prepared in Dimethyl Sulfoxide (DMSO),
immediately before use. NEM was added to the supernatant to a final concentration
of 40 mM to avoid artificial disulphide bond formation. Collected supernatant from
LPS stimulated macrophages was centrifuged at 10000 x g for 3 min to remove cell
debris. Samples prior to proteomic analysis were subjected to a desalting and
concentration process using Vivaspin 20 (ultrafiltration spin columns) with a
molecular weight cut-off of 5 kDa following the manufacturer’s instructions, then
low molecular weight (low-Mr) solutes were filtrate while high molecular weight
(high-Mr) solutes were retained. The first centrifugation with Vivaspin 20 was
followed by a second dilution adding PBS for desalting. To concentrate less volume
of medium are also available Vivaspin 500 (volume range 100-500 µl) and
Vivaspin 6 (volume range 2-6 ml).
2.2.5.2 Pull-down of biotinylated proteins
Biotinylated proteins were purified from the protein complex using streptavidin-
agarose beads (streptavidin immobilized on Agarose CL-4B). This method is
described in three steps: sample incubation with streptavidin-agarose beads; washes
to remove unbound protein and elution.
In the first step the supernatant prepared as described above was incubated for 30
min at 4˚C with 70 µl of streptavidin-agarose beads with rocking. The beads were
pelleted by centrifugation at 8000 x g for 30 sec. In the second step unbound proteins
were removed by washing four times in PBS and once in water. In the third step, the
captured biotinylated proteins were eluted from beads by incubation for 30 min with
¼ vol. of Laemmli Sample Buffer (SB) 4X with 10 mM DTT at RT. The elution step
was repeated for the second time by adding equal volume of SB with DTT and
samples were boiled for 10 min and the collected eluates were loaded on Sodium
Dodecyl Sulphate-Polyacrylamide Gel Electrophoresis (SDS-PAGE). The method
described previously is a modification of Sullivan method (220) (Figure 2.1).
86
Figure 2.1: Biotinylated proteins pull-down. Figure adapted from Sullivan (220).
Interaction between biotinylated proteins and streptavidin-agarose beads was
followed by washes to remove unbound proteins. The beads were eluted with SB
containing DTT.
87
2.2.5.3 SDS-PAGE and Western blot
Proteins were separated by size using a SDS-PAGE. Protein size expressed in kDa
and gel percentage are inversely proportional (high percentage of bis acrylamide is
used to detect small proteins).
The different steps in a Western blot procedure are summarized in Figure 2.2.
Sample preparation and electrophoresis step: samples were mixed with
¼ vol. of SB 4X, boiled for 8 min and then subjected to 12% SDS-PAGE
under reducing condition (10% 2-ME) or non-reducing conditions in
Running buffer for 1.5-2 hours at 100V.
Transfer step: after electrophoresis, the proteins were blotted onto
nitrocellulose membrane (Hybond ECL) in Transfer buffer for 1 hour at
100V.
Blocking step: non-specific binding sites were blocked for 1 hour in
Blocking buffer and membrane was washed three times in PBS-Tween 20
(PBS-T).
Antibodies incubation step: membrane was incubated with a primary
antibody for 1 hour and washed in PBS-T before incubation with a secondary
antibody. All washes and incubations were carried out at RT on a rocker
shaker.
Development step: in a dark room the membrane was covered by ECL
reagent (equal amount of ECL1 and ECL2) for 1 min and then exposed to
autoradiography film (Hyperfilm ECL) using an X-ray film cassette
(Hypercassette). The bands were visualized following the manual
development with developer and fixer solution.
Densitometric quantification of bands was measured with image analysis
software (GeneTools, Syngene).
88
Western blot buffers
Gel preparation: 12% Resolving Gel (bottom phase) and 5% Stacking Gel (top
phase).
12% Resolving gel solution: 4 ml Acrylamide/bis solution , 2.5 ml Running
buffer 4X, 100 µl SDS 10%, 100 µl APS 10%, 10 µl temed and 3.4 ml H2O.
Running gel buffer -4X stock (1.5 M Tris-HCl, pH 8.8): Tris base
(18.17 g/100 ml H2O).
5% Stacking gel solution: 835 µl Acrylamide/bis solution, 1.25 ml Stacking
buffer 4X, 50 µl SDS 10%, 75 µl APS 10%, 5 µl temed and 2.86 ml H2O.
Stacking gel buffer -4X stock (0.5 M Tris-HCl, pH 6.8): Tris base (6 g/100
ml H2O).
Laemmli Sample Buffer -4X stock (10 ml): 250 mM Tris-HCl pH 6.8 (2.5 ml Tris-
HCl 1M); 8% (w/v) SDS; 40% (v/v) glycerol; 0.04% (w/v) bromophenol blue.
10% 2-ME was added to sample immediately prior to heating if required for reducing
conditions.
Tris-HCl 1 M, pH 6.8: 12 g/100 ml H2O
Glycine Running buffer -1X stock: 3.03 g Tris base, 14.4 g glycine and 1 g
SDS/1L H2O.
Transfer buffer -1X stock: 3.03 g Tris base, 14.4 g glycine and 200 ml methanol/1
L H2O.
Wash buffer: 0.1% (v/v) Tween 20 in PBS -1X stock (PBS-T).
Blocking buffer: 5% (w/v) skim milk (or BSA) in PBS-T.
For instance to detect biotinylated or phosphorylated proteins by Western blot is
suggested to use BSA instead of milk because milk components (biotin and
phosphoproteins like casein) can interfere with proteins detection.
89
Figure 2.2: Overview of Western blot.
90
2.2.5.4 SDS-PAGE staining
Following the BioGEE incorporation, the proteins separated by streptavidin beads
(pull down) were identified by MS. We used three different strategies to identify
these biotinylated released proteins:
1) MS from Coomassie stain
Proteins eluted from streptavidin beads were separated by 12% SDS-PAGE. After
electrophoresis as previously described, proteins were visualized by staining gels
with Coomassie and bands of interest were excised from the gel.
2) MS from Silver stain
In order to improve bands visualization, Silver Staining, which is about 10-100 times
more sensitive (in the low nanogram range) than Coomassie was used.
3) MS without protein separation on SDS-PAGE and staining procedures
(shotgun proteomics)
Proteins were identified by MS without separation on SDS-PAGE. Following the
biotinylated protein pull-down the elution step was performed in 50 mM ammonium
bicarbonate.
Coomassie Stain
Following 12% SDS-PAGE, the gel was stained with Coomassie stain according to
protocol. Briefly, the gel was incubated with the fixation solution 45% (v/v)
methanol, 5% (v/v) acetic acid and 50% (v/v) H2O) for 15 min, washed 3 times for
10 min in H2O and then stained with Coomassie stain (GelCode Blue Stain Reagent)
at RT. Protein bands were visualized within 30-60 min and then destained with H2O.
Silver Stain
Silver stain is a colorimetric method for total proteins detection where the brown
colour depends from the reduction of the silver ions (silver nitrate) binding proteins
to metallic silver. Proteins were identified by Silver staining, according to the
manufacturer’s instructions. Briefly, after electrophoresis gel was fixed with fixation
solution for 20 min, followed by 1 wash with 30% (v/v) ethanol in H2O for 10 min
and then incubated with Sensitizing solution for 10 min. After 1 wash with 30% (v/v)
ethanol in H2O and 2 washes with H2O for 10 min each, the gel was stained with
91
Staining solution for 15 min and then washed again with H2O for 1 min before
incubation with Developer solution for 4-8 min until the bands appeared. The
Stopping solution was added and incubated for 10 min. Then, Destaining solution
was added and incubated for 15 min. Finally the gel was washed twice with H2O and
used for MS analysis.
Silver Solutions
All solutions were prepared from reagents provided in the kit, immediately before
starting the Silver Stain protocol.
Fixation solution: 40% (v/v) Methanol, 10% (v/v) acetic acid and 50% (v/v) H2O.
Sensitizing solution: 30 ml Ethanol and 10 ml Sensitizer (Sodium
Thiosulphate)/ 100 ml H2O.
Staining solution: 1 ml Stainer (Silver Nitrate)/100 ml H2O.
Developing solution: 10 ml Developer (Sodium carbonate) and 1 drop Developer
enhancer (formaldehyde)/100 ml H2O.
Destaining solution: 50 µl Destainer A and 50 µl Destainer B.
92
2.2.5.5 Mass Spectrometry
MS is described in several articles as the most common method for protein
identification (221) (222). MS experiments for this study were performed after gel
staining (Silver and Coomassie) or directly after streptavidin pull-down as described
in Methods (2.2.5.2).
Protein bands of interest were excised from gels and then In-gel digestion and MS
analysis were carried out by Dr. Lucas Bowler (Sussex Proteomics Centre,
University of Sussex). Briefly, bands were washed 15 min in acetonitrile, reduced
with 10 mM DTT/25 mM NH4HCO3 at 56°C for 1 hour and then alkylated with
55 mM Iodoacetamide (IAA)/25 mM NH4HCO3 for 45 min at RT in the dark. Bands
were washed with acetonitrile for 20 min and with 25 mM NH4HCO3 for 5 min and
then again with acetonitrile. The bands were dried in a speedvac for approximately
30 min. Finally the tryptic digestion was performed with 5 µl of 25 ng/ml trypsin and
45 µl of 25 mM NH4HCO3 in ice for 45 min, incubated ON at 37°C and then stopped
with trifluoroacetic acid. The resulting tryptic peptides were extracted by 5% formic
acid /50% acetonitrile and concentrated using speedvac. The tryptic peptide extract
was analysed by Liquid Chromatography-Mass Spectrometry (LC-MS) using reverse
phase chromatography coupled to a tandem mass spectrometer.
Specifically, resulting peptides were fractionated on a 250 mm x 0.075 mm reverse
phase column using an Ultimate U3000 nano-LC system and a 2 hour linear gradient
from 95% solvent A (0.1% formic acid in water) and 5 % B (0.1% formic acid in
95% acetonitrile) to 50% B at a flow rate of 250 nL/min. Eluting peptides were
directly analysed by tandem MS using a LTQ Orbitrap XL hybrid FTMS and the
derived MS/MS data searched against the ipi. MOUSE.v3.72 database (56957
entries) using Sequest version SRF v.5 as implemented in Bioworks v3.3.1, assuming
carboxyamidomethylation (Cys), deamidation (Asn) and oxidation (Mat) as variable
modifications and using a peptide tolerance of 10ppm and a fragment ion tolerance
of 1.0 Da. Filtering criteria used for positive protein identifications are Xcorr values
greater than 1.9 for +1 spectra, 2.2 for +2 spectra and 3.75 for +3 spectra and a delta
correlation (DCn) cut-off of 0.1.
93
2.2.6 Western blot of biotinylated proteins
Biotinylated proteins were detected using Streptavidin-POD following the standard
protocol of Western blot for the running and transfer steps as described above
(2.2.5.3). Then, non-specific binding sites were blocked for 1 hour in blocking buffer
(BSA) containing 5 mM NEM to prevent reduction of disulphides. Membrane was
then washed three times in PBS-T, incubated with Streptavidin-POD for 30 min and
washed in PBS-T before development.
2.2.6.1 Detection of Prx2 released by macrophages
Conditioned medium obtained by Raw 264.7 cells plated in 24-well plates, as
indicated in Table 2.2 and treated with BioGEE and LPS as described above, was
analysed by Western blot for the detection of Prx2. Time course experiment was
performed following the same procedure and cells were incubated with LPS and
collected at different time points (0, 2, 4, 8 hours and ON). All subsequent
experiments were conducted with 100 ng/ml LPS for 24 hours on the basis of the
results obtained in the previous experiments.
Equal volumes of supernatants (20 µl/lane) were mixed with ¼ vol. SB 4X, boiled
for 8 min and separated by 12 % SDS-PAGE as described above and analysed by
Western blot using a rabbit anti-Prx2 polyclonal antibody (primary antibody). Briefly
proteins transferred to the nitrocellulose membrane were incubated for 1 hour with
primary antibody solution (rabbit anti-Prx2 antibody in PBS-T with 1% BSA) and
then washed before the incubation for 1 hour with a goat anti-rabbit HRP-labelled
secondary antibody diluted in PBS-T with 1% BSA. Concentration of antibodies is
indicated in the Material section (2.1.3). Cell lysate directly in SB 1.5 X (50 µl/well)
with 10% 2-ME and boiled for 8 min was also used for Prx2 detection. Total cell
lysates (5-10 µl of each sample) were analysed by Western blot using an anti-Prx2
antibody (as described for supernatants).
94
2.2.6.2 Detection of Prx2 levels in mouse serum
All animal studies were carried out in accordance with the European Communities
Council Directive #86/609 for care of laboratory animals and in agreement with
national regulations on animal research in UK and working under an appropriate
project license by Dr. Ilaria Cervellini (Brighton and Sussex Medical School) with a
personal license.
First of all, for the in vivo experiments it was considered the choice between to
analyse proteins in mouse serum or plasma. The main difference between plasma and
serum is that plasma is obtained by adding anticoagulant (e.g. heparin, sodium citrate
or EDTA) and contains fibrinogen (coagulant factor). In contrast, serum is prepared
without anticoagulant and the fibrinogen is converted in fibrin during blood
coagulation. Detection of proteins in serum by Western blot was preferred over
plasma analysis considering that the literature recommended for the most part of
studies to use serum for different reasons. For instance, higher concentration of
proteins in plasma (e.g. fibrinogen or other proteins not present in serum) can cover
proteins present in the lowest concentrations that are for this reason not detected.
Furthermore, anticoagulant present in plasma can modify proteins values. Only
serum samples that were not haemolysed were analysed by Western blot.
Experiments were performed using male CD-1 mice (4-6 weeks) that were injected
intraperitoneally (IP) with saline (vehicle) or with 400 µg/mouse LPS (LPS-treated
mice). Blood was collected 90 min or 24 hours later in tube without anticoagulant
and incubated for 2 hours at RT to allow clotting. The sample was centrifuged at
5000 rpm for 5 min. The supernatant (serum) was transferred in a new tube and
centrifuged for the second time to remove all potentially remaining cells (Figure 2.3).
Serum was diluted 1:10 in H2O and then equal volumes were separated by a 12%
SDS-PAGE under reducing conditions. Western blot for Prx2 was performed as
described previously.
95
Figure 2.3: Mouse serum sample preparation.
96
2.2.7 Immunoprecipitation
Immunoprecipitation (IP) is used to purify a protein from a complex sample by
interaction with a specific antibody and Protein G beads. All experiments for this
thesis were performed using the indirect method or the direct method.
Indirect Method: interaction of antibody and the sample followed by incubation with
Protein G (Figure 2.4). IP procedure used for this thesis is based on the method
described in the instructions of Pierce Protein G agarose. Briefly, sample
(conditioned medium) was precleared with Protein G agarose for 1-2 hours at 4˚C,
followed by immunoprecipitation with primary antibody ON at 4˚C.
Immunocomplex (Ag-Ab complex) was precipitated with Protein G agarose beads
for 2 hours at RT. The immunoprecipitate was washed three times with IP buffer
(25 mM Tris, 150 mM NaCl, 1% (v/v) Triton; pH 7.2). The immunocomplex was
eluted with glycine elution buffer (0.2 M glycine-HCl buffer, pH 2.5) or with SB
elution. Dissociated immunocomplex was then subjected to Western blot using a
specific antibody against the proteins of interest.
Direct Method: binding of antibody to Protein G followed by incubation with sample
(Figure 2.5). Briefly, primary antibody and Protein G were incubated for 2 hours at
4°C and then washed to remove the unbound antibody. The precleared sample
prepared as described above was added to the antibody-beads for 4 hours at 4°C. The
antibody-beads protein complex was washed 4 times with PBS and once with water
before elution with non-reducing SB 2X for 10 min at 50˚C. The elution step was
followed by analysis of immunoprecipitated proteins by SDS-PAGE.
97
Figure 2.4: Schematic immunoprecipitation using the indirect method.
98
Figure 2.5: Schematic immunoprecipitation using the direct method.
99
2.2.7.1 Immunoprecipitation of HMGB1
Following the direct method, 4 µg anti-HMGB1 (antibody) was incubated with
Protein G beads (20 µl) and then with the precleared sample (supernatant pool from
two 6-well plates incubated with 20 µl of Protein G beads for 1 hour at 4˚C). The
sample was eluted from the Protein G beads with 1.5 X SB and loaded onto 12%
SDS-PAGE and then blotted with Streptavidin-POD.
Immunoprecipitation of HMGB1 was also performed using the indirect method.
Supernatant was precleared as described above for the direct method and then the
precleared samples were incubated with 4 µg anti-HMGB1 overnight at 4˚C.
Immunocomplexes were precipitated with Protein G beads for 2 hours at room
temperature and then the beads were washed three times with 0.5 ml of IP buffer.
The immunocomplexes were eluted with 50 µl of elution buffer (0.2 M glycine-HCl
buffer, pH 2.5). The dissociated immunocomplexes were then subjected to Western
blot using anti-HMGB1.
2.2.7.2 Immunoprecipitation of glutathionylated Prx2
Raw 264.7 cells were plated in 6-well plates at 1 x 106 in 3 ml RPMI 1640 complete
and treated in 1 ml Opti-MEM I medium with 100 ng/ml LPS for 24 hours, before
adding 40 mM NEM to the supernatant. Prx2 contained into supernatant was
immunoprecipitated following the indirect method (Ab-sample and then incubated
with Protein G). Briefly, supernatant from 3 wells were pooled together and
centrifuged with Vivaspin 6. The concentrated sample was precleared using Protein
G and then incubated with 5 µl anti-Prx2 (5 µg) at 4ºC for 4 hours. The
immunocomplex (Ab-sample) was incubated with 40 µl Protein G ON at 4ºC. The
sample was eluted from the beads with 25 µl SB 2X for 4 min at 100ºC.
100
2.2.8 Intracellular GSH assay
Cellular GSH was measured according to the enzymatic cycling method of Tietze
(223) with modifications as described by Rahman (224). DTNB (5, 5’-Dithiobis-2-
Nitrobenzoic acid, Ellman’s reagent) reacts with GSH present in the sample,
producing TNB (5-Thio-2-Nitrobenzoic acid) and mixed disulphide GSTNB. The
rate of formation of TNB, measured at 412 nm, is proportional to the concentration
of GSH. GSTNB is then reduced by GR in presence of NADPH to GSH and TNB.
GSSG is also reduced by GR, the amount of glutathione measured represent the total
glutathione (GSH and GSSG) in the sample. Preincubation of the sample with the
thiol masking agent 2-vinylpyridine, prevents the measurement of GSH, resulting in
measurement of GSSG only. However results were obtained measuring only total
glutathione.
2.2.8.1 Glutathione depletion with BSO
Raw 264.7 cells were plated in 6-well plates at a density of 1 x 106
in 2 ml of
complete RPMI-1640 medium. Intracellular GSH levels were depleted in Raw 264.7
cells using BSO. The medium was replaced with 1 ml Opti-MEM I after ON
incubation. A stock solution of BSO 50 mM was prepared in PBS, sterilized and
stored at –20˚C and diluted at 100X concentrations, immediately before use. Various
amounts of BSO (final concentration: 0, 125 µM, 250 µM) were added to Raw 264.7
cells for 24 hours and then stimulated with 100 ng/ml LPS for an additional 24 hours.
The sample preparation method is illustrated in Figure 2.6, and adherent cells treated
as described above were washed twice in PBS. Cells were deproteinized with 6%
trichloroacetic acid and left 1 hour on ice, centrifuged at 2100 rpm for 10 min at 4˚C.
The supernatants were used for the measurement of total glutathione (GSH + GSSG).
The assay was performed in a 96-well plate and each well contained reagents
prepared as described in the reagent preparation section. A standard GSH
concentration curve was prepared in duplicate and were used the concentrations
starting from 6.5 µM of GSH.
101
96 well-plate reaction:
20 µl Sample (GSH standard or PB) +120 µl DTNB: GR + 60 µl NADPH 0.8 mM.
Concentration of total GSH was calculated from a standard curve generated using
GSH prepared in 0.2 M phosphate buffer pH 7.4 (PB). The pellets were dissolved in
0.5 ml of NaOH 0.05 M and used for protein determination to normalize GSH
content. The protein concentrations were determined using the DC Protein Assay kit
from Bio-Rad, using BSA as a standard.
Reagent preparation
Phosphate buffer (Buffer A + Buffer B), pH 7.4
Buffer A: 0.2 M NaH2PO4.
Buffer B: 0.2 M Na2HPO4.
0.2 M Buffer at pH 7.4: 19 ml Buffer A + 81 ml Buffer B.
1.68 mM DTNB: 2 mg in 3 ml PB.
3 U/ml GR: 15 µl of GR 600 U/ml in 3 ml PB.
DTNB + GR (1:1): 3 ml DTNB + 3 ml GR.
NADPH (0.8 mM): 2 mg in 3 ml PB.
GSH standards
GSH 10 µg/ml (32.5 µM): dilution 1:100 from GSH 1 mg/ml.
GSH 26 32.5 µM: 800 µl GSH 32.5 µM + 200 µl PB.
GSH from 13 µM to 0.102 µM: two-fold serial dilutions from 13 µM to
0.102 µM.
102
Figure 2.6: Sample preparation scheme for GSH assay.
103
2.2.8.2 Protein determination
The protein concentration was determined using the colorimetric assay, DC Protein
Assay kit according to manufacturer’s instructions. This assay was performed at RT
in 96-well plate by adding 5 µl of sample (or BSA as a standard), 25 µl of reagent A
(alkaline copper tartrate solution) and 200 µl of reagent B (Folin reagent) into each
well.
A standard protein concentration curve was prepared in duplicate (two-fold dilution
starting from 1.5 mg/ml of BSA in the same buffer as the sample). The absorbance
was measured at 690 nm using a microplate reader. Total protein concentration was
calculated following linear regression analysis using Microsoft Excel (version Office
2010). Linear Regression (y=mx+b, where x=absorbance value, m=slope and b=y-
intercept). Total protein (mg/ml) =sample absorbance – (y-intercept)/ slope).
104
2.2.9 Activity of Prx2 with Trx1 or Grx2
The peroxidatic activity of hrPrx2 was evaluated by Dr. Eva Maria Hanschmann
following the protocol described by Hanschmann et al. (225) using the Trx and Grx
systems (Trx1, TrxR and NADPH; Grx, GSH, GR and NADPH, respectively) as
electron donor. As reported in the article, the activity of Prx2 was measured by a
spectrophotometric assays in 96-well plates (200 µl/well) containing 100 mM Tris-
HCl, pH 7.0, 1 mM EDTA, 120 µM H2O2, ≥1 µg TrxR, 200 µM NADPH, and the
cytosolic form of Grx2 or Trx2 (1–80 µM). In some experiments TrxR was replaced
by 1 mM GSH and yeast GR for GSH-dependent Grx reactions. The reaction was
started by the addition of the Prx2 (0.05–1 µM) and followed as a decrease in
absorbance at 340 nm. The linear part of the reaction was used to determine the rates
of NADPH consumption. Kinetic constants were determined by nonlinear curve
fitting of the Michaelis-Menten equation using Grace (225).
105
2.2.10 Removal of endotoxin from hrPrx2 preparation
A Detoxi-Gel column, immobilized with Polymyxin B (PMB) was used to remove
LPS contamination from hrPrx2 following the manufacturer’s instructions. All steps
(regeneration column, washes, equilibration, sample incubation and elution) were
performed at RT.
Regeneration: 5 ml of 1% (w/v) Sodium Deoxycholate in H2O.
Washes: 9 washes (5 ml each) with PBS.
Equilibration: 5 ml of PBS.
Sample incubation: 200 µl hrPrx2 at 9.7 mg/ml for 1 hour incubation.
Protein elution: 2 ml fractions eluted in PBS (only the first eluted fraction
was used for all experiments).
hrPrx2 free LPS was sterilized by a filter membrane (0.22 µm) and stored at -20 °C.
An aliquot (~10 µl) was used for protein determination with DC Protein Assay, as
described in Methods (2.2.8.2).
106
2.2.11 Limulus Amebocyte Lysate test
The Limulus Amebocyte Lysate (LAL) test is specific for the detection of endotoxin
using a gel-clot method. The Pyrogent Plus kit was used following the
manufacturer’s instructions. In the test tube containing the LAL reagent (clotting
agent from the blood cells of the horseshoe crab) was added 250 µl of sample or the
Control Standard Endotoxin (CSE). CSE and protein were prepared with pyrogen-
free water and incubated for 1 hour at 37° C. The formation of a stable solid clot
evaluated by 180 °C inversion of the tubes confirms the presence of endotoxin
contamination.
Reagents preparation
CSE stock solution: one vial of lyophilized preparation purified endotoxin from
E. coli strain 0111:B4 was reconstituted with 1 ml of LAL reagent water to obtain a
stock solution of 17 EU/ml. From the stock solution, CSE serial dilutions were made
to obtain CSE at 0.5 EU/ml, 0.125 EU/ml, 0.06 EU/ml and 0.03 EU/ml.
hrPrx2 was diluted with pyrogen-free water at different concentrations (40 µg/ml,
20 µg/ml, 10 µg/ml, 2.5 µg/ml) from hrPrx2 (concentration of the stock solution
determined with DC Protein assay).
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2.2.12 ELISA
2.2.12.1 Mouse TNF-α ELISA
A sandwich ELISA was used to detect and quantify TNF-α using specific antibody
(primary antibody and biotinylated secondary antibody). The different ELISA steps
are described in Figure 2.7.
TNF-α concentration in the supernatant from murine cells was measured by the
mouse TNF-α Duoset (ELISA kit) according to manufacturer’s instructions. Briefly,
a 96-well plate was coated with 50 µl/well of the capture antibody ON at 4°C. After
3 washes with wash buffer, the plate was blocked with 200 µl/well of reagent diluent
at RT for a minimum of 1 hour. After 3 washes with wash buffer, 50 µl of samples or
TNF-α (standard) were added to the plate for 2 hours at RT followed by 4 washes.
Then, 50 µl/well of detection antibody was added for 2 hours at RT followed by
6 washes. 50 µl/well Streptavidin-POD was added for 30 min at RT followed by
8 washes, then 100 µl/well of substrate solution (TMB) for 20-30 min at RT. The
reaction was blocked with 50 µl/well of stop solution and absorbance was measured
(value at 570 nm was subtracted from 450 nm value) using a microplate reader.
ELISA solutions
Capture antibody (monoclonal anti-mouse TNF-α): 0.8 µg/ml in PBS (dilution
1:180 of stock 144 µg/ml).
Wash buffer: 0.05% (v/v) Tween 20 in PBS.
Reagent diluent: 1% (w/v) BSA in PBS.
Recombinant mouse TNF-α (standard): seven two-fold serial dilutions in reagent
diluent starting from 1 ng/ml (dilution 1:270 of stock 270 ng/ml).
Detection antibody (biotinylated anti-mouse TNF-α): 200 ng/ml in reagent diluent
(dilution 1:180 of stock 36 µg/ml).
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SA-POD: dilution 1:200 of stock in reagent diluent.
Substrate solution: color reagent A (TMB) and color reagent B (H2O2).
Stop solution: H2SO4 1M.
2.2.12.2 Human TNF-α ELISA
TNF-α concentration in the supernatant of human macrophages and PBMC was
measured by the human TNF-α Duoset (ELISA kit) according to manufacturer’s
instructions. Following the same detection steps as described for mouse TNF-α, a 96-
well plate was coated with 50 µl/well of monoclonal anti-hu TNF-α ON at 4°C. After
3 washes with wash buffer, the plate was blocked with 100 µl/well of reagent diluent
at RT for a minimum of 1 hour. After 3 washes with wash buffer, 50 µl of samples or
TNF-α (standard) were added to the plate for 2 hours at RT followed by 3 washes.
Then, 50 µl/well of detection antibody was added for 2 hours at RT followed by
6 washes. 50 µl/well SA-POD was added for 30 min at RT followed by 6 washes,
then 50 µl/well of substrate solution (TMB) for 20-30 min at RT. The reaction was
blocked with 25 µl/well of stop solution.
ELISA solutions
Capture antibody (monoclonal anti-hu TNF-α): 4 µg/ml in PBS (dilution 1:180 of
stock 720 µg/ml).
Wash buffer: 0.05% (v/v) Tween 20 in PBS.
Reagent diluent: 1% (w/v) BSA in PBS.
Recombinant human TNF-α (standard): seven two-fold serial dilutions in reagent
diluent starting from 1 ng/ml (dilution 1:320 of stock 320 ng/ml).
109
Figure 2.7: ELISA summary steps. (1) Plate is coated with a capture antibody
(primary antibody); (2) standard or sample is added; (3) incubation with detection
antibody (secondary biotinylated antibody); (4) SA-POD and substrate solution are
added; (5) the reaction is stopped with stop solution.
110
2.2.12.3 Human Prx2 ELISA
Prx2 levels from supernatant of human macrophages and PBMC were measured by
ELISA kit according to manufacturer’s instructions. Briefly, sample or standard
(from 50 ng/ml to 0.78 ng/ml) was incubated for 2 hours at 37 °C in a 96-well plate
pre-coated with a monoclonal antibody specific to human Prx2.
100 µl of Detection Reagent A were added to each well for 1 hour at 37 °C followed
by three washes with Wash solution. After the last wash, 100 µl of Detection
Reagent B were added for 30 min at 37 °C followed by 5 washes. Substrate solution
(90 µl) was incubated for 15 min at 37 °C then stopped with 50 µl of stop solution.
Absorbance was measured at 450 nm.
ELISA solutions
Wash buffer: 1 volume of wash buffer concentrate (30X) with 29 volume of H2O.
Assay diluent A: equal volume of diluent A concentrate (2X) and H2O.
Assay diluent B: equal volume of diluent B concentrate (2X) and H2O.
Recombinant human Prx2 (standard): dilution 1:5 of stock 200 ng/ml to obtain
50 ng/ml and then six two-fold dilutions (25 ng/ml, 12.5 ng/ml, 6.25 ng/ml,
3.13 ng/ml, 1.56 ng/ml, 0.78 ng/ml) in standard diluent.
Detection reagent A (biotin conjugated polyclonal antibody): dilution 1:100 of
stock in assay diluent A.
Detection reagent B (avidin conjugated to HRP): dilution 1:100 of stock in assay
diluent B.
Substrate solution: TMB substrate.
Stop solution: 1M H2SO4.
111
2.2.13 Statistical Analysis
In this thesis, all the results are expressed as mean and minimum (min)-maximum
(max) range values. Standard deviation (SD) values were not calculated for
experiments performed using duplicate samples. When experiments were performed
using triplicate or quadruplicate samples, SD was calculated. When experiments
were performed using triplicate or quadruplicate samples, the statistical significance
of differences between experimental groups was assessed by Student’s t-test. In
figures, asterisks (*p<0.05); **p<0.01; *** p<0.001) and hash sign (#p<0.05;
##p<0.01); ###p<0.001) indicate statistically significant differences. We thank Dr.
Liz Cheek, Senior Lecturer at University of Brighton, School of Computing,
Mathematical and Information Sciences for suggestions and advice.
112
Chapter 3. Studies on the glutathionylation of
HMGB1
113
3.1 Introduction: non-classical secretion of proteins
There are different proteins that can be present in both intracellular compartments, or
actively secreted in an extracellular environment (e.g. plasma and culture medium)
(226). The study of secreted proteins can give important new information, for
instance the discovery of different biological functions depending on protein
localization.
Many proteins are secreted with a process known as the classical or ER/Golgi-
dependent secretory pathway (227). Proteins that contain a signal peptide (amino
acid motif within the protein) are directed from the ER to the Golgi apparatus. After
eventual glycosylation in the Golgi, proteins are distributed to secretory vesicles that
can release their contents into the extracellular space (228). Both brefeldin A and
monensin block the classical secretory pathway but with a different mechanism. In
particular, brefeldin A blocks the exit from the ER while monensin has effects on
Golgi apparatus (229) (230).
Although most cytokines are efficiently and rapidly released through the classical
secretory pathway, there are some cytokines lacking a signal peptide that are secreted
using non-classical secretory/release pathways. For instance, leaderless cytokines
such as IL-1 β, Trx and HMGB1 utilize other mechanisms for their secretion (231).
Rubartelli et al. demonstrated that IL-1β is released by human monocytes activated
with LPS through an alternative secretory pathway. In fact, 1L-1β secretion was not
blocked but increased by incubation with monensin and brefeldin A (inhibitory drugs
of the ER/Golgi pathway). Furthermore, IL-1 β secretion was increased by high
temperature (42°C) but blocked by low temperature (18°C), and it was also inhibited
by serum free medium or Methylamine (inhibitor of endocytosis). The alternative
secretory pathway of 1L-1β may use endosome and lysosome (232). Rubartelli et al.
showed the secretion of 1L-1β through endolysosomal-related organelles and its
correlation with extracellular ATP concentration, whereby a high concentration of
ATP was necessary to increase the secretion of 1L-1β (233). MacKenzie et al.
studied the rapid (after 2-5 min) secretion of 1L-1β by microvesicle shedding and
suggested this as a possible previously unrecognized pathway for 1L-1β secretion
(234). As summarized by Dubyak, all the possible secretory pathway of IL-1 β
114
known in literature include the exosome pathway, the microvesicle pathway, the
secretory lysosome pathway, autophagy pathway and the necrotic pathway (235).
Rubartelli et al. also studied the secretion of Trx from normal and neoplastic cells
suggesting an alternative pathway similar to the secretory pathway found for IL-1β
(e.g. inhibition of secretion by methylamine or increasing with brefeldin A and
monensin) (236). Tanudji et al. showed the secretion of Trx from breast cancer,
colon carcinoma, and multiple myeloma via a non-classical pathway was not blocked
by brefeldin A. Furthermore, the secretion of Trx was not regulated by the redox
state as demonstrated using mutated forms of Trx (Trx C35S and Trx C73S) (237).
HMGB1 also lacks of a signal peptide, and is not secreted through the classical
ER/Golgi pathway. HMGB1 is a nuclear protein and the first step of the secretion
includes the exit from the nucleus into the cytoplasm, followed by translocation from
the cytosol into cytoplasmic organelles and finally secretion into extracellular
environment by exocytosis (231).
Wang et al. demonstrated that HMGB1 can be secreted actively by macrophages
stimulated with endotoxin, TNF or IL-1 and can act as proinflammatory cytokine
(238). HMGB1 is also actively secreted by NK cells and represents a potential
defence in HIV-1 infections (239). In addition, as demonstrated by Scaffidi et al.,
HMGB1 can be also released passively by necrotic cells (but not by apoptotic cells)
where the interaction between protein and chromatin is weak and the protein can be
easily released (240). IL-1 and Trx show a kinetics secretion that decreases after 6
hours (236), while HMGB1 is secreted by LPS-activated monocytes with a slower
kinetics, from 6 hours until 18-30 hours (231).
Secreted proteins may undergo post-translational modifications such as methylation,
acetylation, phosphorylation or glutathionylation. Several articles have highlighted
the importance of post-transcriptional modification for the secretion of HMGB1. As
summarized by Pisetsky et al., the acetylation, phosphorylation or methylation of
HMGB1 can modify the interaction between the protein and the chromatin,
facilitating HMGB1 secretion (241). The secretion of HMGB1 from activated
macrophages and monocytes is regulated by acetylation of target lysine contained in
HMGB1. In particular, acetylated HMGB1 is transferred from the nucleus into
115
secretory lysosomes in the cytoplasm (242). During the EMBO meeting workshop in
2006 the link between HMGB1 secretion and glutathionylation was also discussed
(60). In order to understand the mechanism of HMGB1 release, this study examined
the possible link between glutathionylation and the release of HMGB1 in
macrophages treated with LPS.
It was important for this research to select the most appropriate method to detect
glutathionylated proteins. Gao et al. (243) described the main techniques comparing
advantages and drawbacks. For instance, Fratelli et al. identified glutathionylated
proteins in primary rat hepatocytes and human HepG62 using 35
S-GSH
(radiolabelled GSH with 35
S-Cys) (244). 35
S-GSH was also used to study
glutathionylation of complex I in bovine heart mitochondria (245). This method is
very sensitive and quantitative but there are a number of drawbacks. One drawback
of this method is that it requires the presence of a protein synthesis inhibitor
(cycloheximide) to avoid the incorporation of 35
S-Cys into protein in the process of
protein synthesis, independent of glutathionylation. Clearly, the addition of
cycloheximide alters the cell physiology significantly. Furthermore, the use of
radioactivity makes this methodology particularly problematic because radioactivity
is being used less and less in research laboratories. In addition, this method can only
be used for cell culture.
Another possible method is based on the use of commercial anti-GSH antibodies,
available from Chemicon or Virogen. For instance Dalle-Donne et al. showed the
detection of glutathionylated actin by Western blot using an anti-GSH (167).
Furthermore, glutathionylation of IkB was also studied using anti-GSH antibody
(246). However, reviewing the literature, it was not possible to find this method used
in the identification of unknown glutathionylated proteins but rather to either
evaluate overall cellular glutathionylation (247) or to demonstrate glutathionylation
of specific proteins, using purified protein (248). The anti-GSH antibody has the
advantage of not requiring pre-treatments with cycloheximide but has the drawback
of allowing the detection of only very abundant proteins due to the low sensitivity of
these anti-GSH antibodies.
A very sensitive detection method for glutathionylated proteins employs BioGSSG,
Biotinylated GSH (BioGSH) or BioGEE (a BioGSH with an added ethyl ester group
116
which increases the permeability into cells). Biotinylated GSSG that was also used to
identify glutathionylated proteins by redox proteomics has been developed in the
laboratory of Phil Eaton (249) (250). Biotinylated GSH has been used in the past to
study glutathionylation of specific proteins and allow their visualization in gel
electrophoresis (251). A further refinement of this approach has been achieved with
the use of BioGEE originally described by Sullivan et al in 2000 (220). In fact, it is
well known that GSH is poorly transported across the cell membrane and the use of
its esters developed by Alton Meister has been a successful way to circumvent this
problem (252). The ester would then be cleaved by cellular esterases to release GSH.
BioGSH and BioGEE require incubation with an oxidant and are glutathionylate by
either direct oxidation but also by thiol-disulphide exchange following oxidation of
GSH to GSSG while BioGSSG uses the mechanism of thiol-disulphide exchange. It
should be pointed out that it is unclear, which is the most important reaction in the
formation of glutathionylated proteins, although one study on human platelets
suggested that thiol-disulphide exchange is not the main mechanism (253). Overall,
due to its cell penetration as well as its commercial availability, BioGEE is probably
the most widely-used reagent for the study of glutathionylation. The work by
Sullivan validates the use of BioGEE by comparing the results with those obtained
with 35
S (220). Since then, BioGEE is commercially available and has been used in
several studies of protein glutathionylation following visualization in Western blots
(254) (255).
Streptavidin has a high affinity for biotin and proteins glutathionylated with
biotinylated molecules (BioGSSG, BioGSH and BioGEE) can be enriched using
streptavidin (by affinity purification on streptavidin-agarose beads) and/or visualized
in Western blot with Streptavidin-POD or anti-biotin antibodies. On the other hand,
the presence of the biotin group represents a possible drawback. In fact a non-
specific background can derive by the biotin/streptavidin system (256). Furthermore,
the biotin group on the GSH could interact with the proteins modifying their
function. Zaffagnini and Lamaire (unpublished results reported in reference (243))
observed that protein glutathionylated with BioGSH could not be deglutathionylated
by Grxs.
Importantly, all these approaches require the use of non-reducing conditions during
the processing of the sample, including electrophoresis when present, because the
117
biotin tag will be attached through a disulphide bond between the protein and GSH.
Another aspect that is often overlooked is that the mixed disulphide is labile. Even in
the absence of an added reducing agent (such as DTT or 2-ME) protein thiols can
reduce mixed disulphides (257). It is therefore important to block any free thiol in the
sample by alkylation.
Another means of affinity-purifying glutathionylated proteins was developed in the
laboratory of Ian Cotgreave and consists of blocking the free thiols in the sample by
alkylation with NEM, followed by specific reduction of glutathionylated proteins
with E. coli Grx3 and re-alkylation of these newly exposed thiols with biotinylated
NEM. As a result, the proteins that originally were glutathionylated will have a
Biotin tag thus allowing affinity purification and visualization on Western blots
(258). However, this method has never been used outside the laboratory that
developed it and the several steps required introduce a further degree of complexity.
To study the released glutathionylated proteins, the BioGEE method was preferred to
the other methods described above because this could allow us to first evaluate if
there was a release of glutathionylated proteins in the supernatant, and if this
preliminary experiment was positive, pursue their identification following affinity
purification of streptavidin beads. The samples also were alkylated with NEM to
block all free SH groups and thus stabilize the mixed disulphides as described above.
Following the selection of the method to detect glutathionylated proteins, the next
step in the research has been to face the problems related to the analysis of the
released proteins as described by Chevallet et al. (259). One of the most important
problems is the relative concentration of the secreted proteins compared to
intracellular proteins. In addition, the secreted proteins are in low concentration and
can be contaminated by serum proteins present in the medium (260). The approaches
I used to tackle the problem of relative concentration have been to carry out the
experiments in an Opti-MEM I medium without FBS, to concentrate the samples
with Vivaspin (as described in Methods) and to culture the cells in a reduced volume
of medium.
118
3.2 Aim of Chapter 3
One aim of this project was to study proteins released from macrophages activated
by LPS and particular attention focused on proteins that were not secreted with the
classical pathway. This chapter describes the investigation of one specific protein
already known to be involved in inflammation, HMGB1. This was chosen as the
protein of interest since it is released with an unknown secretion mechanism, and is
involved in the activation of inflammation (238). In particular, I investigated whether
extracellular HMGB1 can be glutathionylated upon exposure to LPS, and thus be
involved in the redox regulation of inflammation and immunity.
119
3.3 Results
The results of the preliminary experiments which were performed to define the role
of redox regulation of released HMGB1 upon exposure to inflammatory stimuli
(LPS) are showed below. Released HMGB1 was analysed from Raw 264.7 cells
labelled with BioGEE followed by immunoprecipitation and Western blot.
Preliminary experiments were performed to demonstrate that the proteins were
efficiently biotinylated with the method using BioGEE. For this purpose, Raw 264.7
cells were preincubated with BioGEE at a final concentration of approximately
200 µM (modification of Sullivan’s method) (220) 1 hour before the incubation with
100 ng/ml LPS to label the cells as described in Methods. The supernatant containing
the biotinylated proteins was subjected to 12% SDS-PAGE and proteins were
visualized by Western blot using Streptavidin-POD. The protein bands were present
only in the sample preincubated with BioGEE. No bands were observed with
Streptavidin-POD in the sample without prepared BioGEE showing the absence of
non-specific bands (Figure 3.1).
In a similar experiment, Raw 264.7 cells preincubated with BioGEE and treated with
LPS were analysed by Streptavidin-POD Western blot in reducing or non-reducing
conditions. Signals were decreased by the addition of the reducing agent 2-ME
providing evidence that these proteins were indeed glutathionylated (Figure 3.2).
120
Figure 3.1: Western blot of glutathionylated proteins. Supernatant from Raw 264.7
was analysed by SDS-PAGE (12% gel) followed by Western blot probed with
Streptavidin-POD. Sample from supernatant loaded with BioGEE and incubated with
LPS (lane 1) and sample without BioGEE and incubated with LPS (lane 2).
121
Figure 3.2: Western blot of glutathionylated proteins (under reducing and non-
reducing conditions). Supernatant from Raw 264.7 cells loaded with BioGEE and
incubated with LPS was analysed by SDS-PAGE (12% gel) under non-reducing
(lane 1) and reducing conditions (lane 2) followed by Western blot probed with
Streptavidin-POD.
122
3.3.1 Glutathionylation of HMGB1
Before embarking in experiments using cell culture, we first investigated whether
HMGB1 can be glutathionylated using the purified protein in a cell-free system.
BioGSSG was used to show that HMGB1 could be glutathionylated and visualized
using the methods listed in the previous section. BioGSSG and HMGB1 were
incubated (as described in Methods, 2.2.4) to allow HMGB1 glutathionylation. In
fact, incubation of susceptible proteins with GSSG causes their glutathionylation by
thiol-disulphide exchange (187) (261).
The experiment in Figure 3.3 shows that there is a band corresponding to HMGB1
detected by Streptavidin-POD Western blot in the sample incubated with 5 mM
BioGSSG. The fact that the band was not observed when the sample was reduced by
2-ME indicates that the labelling is actually due to a disulphide bond. Furthermore
no bands were detected for samples where HMGB1 was incubated only with H2O, in
the absence of BioGSSG, showing the absence of non-specific bindings.
3.3.2 HMGB1 release by LPS-stimulated Raw 264.7 cells
To determine whether HMGB1 was released into the extracellular milieu in
glutathionylated form, Raw 264.7 cells were cultured in 6-well plates and incubated
with BioGEE and LPS as described in Methods. After centrifugation to remove cell
debris, supernatant was incubated without (lane 1) or with streptavidin beads (lane 2)
to detect only the released biotinylated HMGB1. An aliquot of supernatant was
analysed by Western blot using a monoclonal antibody for HMGB1.
As seen by Western blot analysis (Figure 3.4), the anti-HMGB1 recognizes only one
band of the expected mass size (24 kDa) confirming the presence of HMGB1 in the
medium. However, no band appears in proteins purified using streptavidin beads
(lane 2).
123
Figure 3.3: Detection of glutathionylated HMGB1 by Western blot probed with
Streptavidin-POD. Samples were prepared as described in Methods and separated on
12% SDS-PAGE under non-reducing conditions (lanes 1-2) and under reducing
conditions (lanes 3-4). There is no detectable HMGB1 glutathionylation in samples
(lanes 2-4) without BioGSSG treatment. Signals were abolished by the addition of 2-
ME.
____________________________________________________________________
Figure 3.4: Detection of HMGB1. HMGB1 was detected as a ~24 kDa protein
(arrow) in supernatant from LPS-stimulated macrophages (lane 1). The 24 kDa
protein was absent in supernatant purified with Streptavidin beads (lane 2).
124
3.3.3 Immunoprecipitation of HMGB1
In the experiment whose results are reported in Figure 3.3, I have shown that
HMGB1 can be glutathionylated with BioGSSG. To study whether the HMGB1
released by LPS-stimulated macrophages is glutathionylated, immunoprecipitated
proteins were subject to Western blot using anti-HMGB1, and then Streptavidin–
POD or anti-HMGB1 was used to identify the biotinylated proteins or HMGB1
respectively.
Briefly, supernatants from LPS-stimulated Raw 264.7 cells, prepared as described in
Methods (Table 2.2) were immunoprecipitated with the direct method or the indirect
method (Figure 2.4 and Figure 2.5). No bands were detected (data not shown because
no bands were detected on the X-ray film). The membrane was also stripped and
incubated with anti-HMGB1 but no bands were detected under these conditions.
The same experiment was also repeated using the indirect method (Figure 2.4). As
suggested in the protocol provided by Pierce (Thermo Scientific website:
http://www.piercenet.com/browse.cfm?fldID=E5049863-D59D-B7A5-D057-
E23F7B22AD44), this method is preferred to the direct method when the protein of
interest is present at low concentration. For this experiment, proteins were
immunoprecipitated with anti-HMGB1 and then subjected to Western blot using
anti-HMGB1 to visualize not only the glutathionylated HMGB1 but the total
HMGB1, in order to understand the validity and how to optimize the IP conditions.
No bands were detected with this method (data not shown because no bands were
detected on the X-ray film).
125
3.4 Discussion
The aim of this part of work was to investigate whether HMGB1 is released from
activated macrophages in the glutathionylated form. Our hypothesis was that
glutathionylation could regulate the release and the inflammatory activity of
HMGB1. It is well known that HMGB1 is actively secreted by macrophages and
monocytes or released by necrosis. In addition, HMGB1 have a role in inflammation
acting as danger signal. The redox state of the three cysteines contained in HMGB1
(Cys23, Cys45 and Cys106) regulate the proinflammatory activity as demonstrated
by Yang et al. In particular, the Cys106 in the reduced form can interact with TLR4
and induce cytokines production (262). Therefore, reversible glutathionylation of
Cys106 could avoid irreversible oxidation of SH group and the reduction of
proinflammatory activity.
Preliminary experiments were performed to set-up the methods for the research. First
of all, we demonstrated that HMGB1 can undergo glutathionylation, at least in a cell-
free system. The glutathionylation of HMGB1, obtained by incubation of hrHMGB1
with BioGSSG was tested by Western blot using Streptavidin-POD and was used as
a positive control to demonstrate the validity of the method to detect the
glutathionylated HMGB1.
We then confirmed that the choice of BioGEE was appropriate to detect
glutathionylated proteins released into the medium of Raw 264.7 cells activated by
LPS that is our experimental model. In fact, results of Western blot of LPS
supernatants using Streptavidin-POD revealed a number of proteins labelled with
BioGEE while no bands were detected in the sample without preincubated with
BioGEE. Another similar experiment also confirmed the labelling of proteins with
BioGEE and the decrease of the label in reducing conditions, indicating that labelling
of these proteins was in fact due to glutathionylation.
A further preliminary experiment showed that HMGB1 was indeed secreted from
Raw 264.7 cells with BioGEE and then with LPS, when HMGB1 was measured by
Western blot using anti-HMGB1 antibodies. This confirms that our experimental
model is appropriate for studying HMGB1 release as described by Wang et al. (238).
126
The absence of the band in the sample affinity-purified with Streptavidin beads
suggests that the HMGB1 released was not glutathionylated, although we must be
aware of the limit of sensitivity of the method. In fact, the negative finding might be
due to the low concentration of proteins present in the conditioned medium, which
would make more difficult to detect the glutathionylated form of the released
HMGB1.
The fact that we were unsuccessful to detect any bands when HMGB1 was
immunoprecipitated could also be due to various reasons. Considering all the steps of
the immunoprecipitation, a possible reason of no bands detection could be connected
with the concentration of the target protein expressed in the sample. In this case, the
amount of glutathionylated HMGB1 released by activated Raw 264.7 could be too
low and then difficult to detect. Another possible reason could be an insufficient
concentration of the anti-HMGB1 used for the immunoprecipitation, or its poor
ability to bind the antigen in solution. Finally, the last step of the
immunoprecipitation includes the elution of the protein from the beads. Therefore,
inappropriate elution of the protein from the beads can explain the difficult to find
the protein in the sample analysed by Western blot.
Taken together, these results confirm the results by Wang et al. (238) showing that
HMGB1 is released by Raw 264.7 macrophages activated by LPS in our
experimental condition. However, the presence of glutathionylated HMGB1 in LPS
supernatants could not be demonstrated using any of these techniques.
3.5 Following chapter
In this chapter we have addressed the specific hypothesis of a glutathionylation of
one specific released protein, HMGB1. However, the set-up experiments have shown
that, indeed, LPS-stimulated Raw 264.7 release several potentially glutathionylated
proteins, as indicated by the Western blots shown in Figure 3.1 and Figure 3.2. We
therefore decided to undertake the identification of these proteins by redox
proteomics.
127
Chapter 4. Proteomic analysis of conditioned
medium from Raw 264.7 cells
128
4.1. Introduction to the identification of unknown
released proteins
Proteomics is a general term to indicate the study of “proteome”, the totality of
proteins present in a cell. A proteomic analysis can provide information about the
structure, localization, and function of a protein and also on its posttranslational
modifications. Currently, the techniques most used in proteomics consist in
proteolytic digestion, MS and bioinformatics analysis of the protein sample, either
with or without previous separation of the proteins (or peptides) by affinity
purification, 1 or 2-dimensional gel electrophoresis, or 1 or 2-dimensional
chromatography. A review of the commonly used proteomic techniques and their
application in haematology has been written by Cristea et al. (263).
The study of proteins secreted or released by cells can give important biological
information and could have biomedical application, for instance a specific protein
identified in body fluids could be used as potential biomarkers. A proteomic analysis
of rat fluids has led to a protein map of rat serum, urine, cerebrospinal fluid and
bronchoalveolar lavage fluid; in particular, Giannazza et al. found a different protein
pattern between serum from rats in acute inflammatory vs. normal conditions, which
makes it possible to distinguish the two conditions (264). In human serum 490
proteins have been identified by Adkins et al. (265). There are many studies
supporting the utility of proteomic techniques in the identification of released
proteins in cancer, for instance as showed by Vaughn et al. more than 200 proteins
were identified into growth media of follicular lymphoma-derived cells by MS (266),
while Martin et al. identified released proteins by neoplastic prostate epithelium
(267). MS was also used by Bin et al. to identify haptoglobin-derived α subunit in
sera of ovarian cancer patients (268). A more complete list of potential ovarian, liver,
lung and pancreatic cancer biomarkers identified by MS is summarized by Tessitore
et al. (269). The MS method is not only limited to identify proteins implicated in
cancer, but also useful for the diagnosis and monitoring of other diseases, such as
Alzheimer’s disease (270), heart disease (271), kidney disease (272) and metabolic
diseases (273).
129
Moreover, the identification of proteins by MS can contribute to characterize the
pathogenic mechanisms underlying diseases, which is a fundamental step in the
identification of novel targets and in the development of novel drugs and therapeutic
approaches. Lafon-Cazan et al. identified different proteins secreted, via the classical
or alternative pathways, in the conditioned medium of cultured astrocytes. These
proteins could have a significant role for the neuronal survival and function (274).
Catterall et al. developed a 2D proteomics approach to identify proteins secreted by
primary chondrocytes after stimulation with IL-1 and oncostatin M, cytokines that
have been shown to be up-regulated in RA (275). A combination of one-dimensional
electrophoresis and MS was used to study adipogenesis. In particular, results
identified different proteins secreted by 3T3-L1 preadipocytes or adipocytes (276). A
more comprehensive list of “adipokines” has been identified in the conditioned
medium from primary human adipocytes by Lehr et al. (277). A proteomic approach
led to establish a protein map of exosomes secreted by dendritic cells (278). Finally,
proteomics has been a useful tool to identify secretome composition of HIV-infected
human macrophages (279) and in a more recent work, to identify cytoskeleton
proteins involved in the cytopathicity of the virus (280).
The most recent advances in proteomics allowed us to identify proteins in low levels
or that undergoing posttranslational modification, which identification relies on the
detection of the peptide/s that contains the site of modification. In recent years
several modifications of the existing proteomics techniques have been developed in
order to identify proteins with specific redox modifications of their cysteines,
including glutathionylation. The term “redox proteomics” was first proposed in 2002
by Fratelli et al., who identified 38 proteins undergoing glutathionylation in oxidative
stressed T-Lymphocytes (177). Despite the fact that several studies identified
glutathionylated proteins by redox proteomics inside cells, little is known about the
proteins released in this form and the literature lacks a systematic proteomic study of
this topic.
When aiming at the identification of potential new cytokines in the secretome,
however, one has to face the problem that many of these proteins are secreted in
small amount and present at relatively low concentration in the cell supernatant. It is
important to note that most cytokines were discovered by following a protein activity
(such as T cell proliferation for interleukin-1, cytotoxic activity for TNF, or antiviral
130
activity for interferons), while others (for instance IL-21, IL-33 and IL-37) were
discovered in silico using bioinformatics approaches or analysing microarray data.
To my knowledge the only protein that was identified by analysing on gel
electrophoresis the differential expression of proteins in control versus LPS-
stimulated macrophages cultures was HMGB1. In fact, Kevin Tracey’s group
discovered this by looking at the expression of late-induced proteins in the
supernatant of Raw 264 mouse macrophage cells (238).
On the other hand, we use here a technique not previously used to analyse the
secretome. In fact, the proteomics techniques used previously were based on
identifying differential expression of proteins 2D gels by analysing the intensity of
stained gels. In some cases, the more recent technology of difference gel
electrophoresis (DIGE) has been applied. This technology is based on the staining of
two different samples with two different fluorescent dyes (that is, with different
excitation or emission wavelength). Two samples (for instance, one treated and one
control) are then mixed and run together on a 2D gel. This reduced variation between
two 2D-gels, and has been applied to the identification of cancer biomarkers
(reviewed in (281)) but also for in vitro studies (282). A further refinement is
represented by the technology using isotope-coded affinity tag (ICAT), where the
proteins in two samples are labelled with the same reagent having two different
isotope labels (heavy and light). The ICAT technology bypasses the need for 2D
electrophoresis and the samples can be subject to tryptic digestion and subsequent
HPLC-MS. Although MS is normally not a quantitative technique, using this
approach and looking at the ration between the same peptides tagged with the two
different isotopes provides a quantitative measure of the ratio in the expression of the
protein of interest in the two samples. This technique has also been widely used for
the identification of secreted proteins in cancer (283). As far as I know, none of these
techniques have been applied to the identification of novel cytokines or inflammatory
mediators in LPS-stimulated macrophages. Of note, a modification of the ICAT
technology has been developed to identify the redox state of protein thiols in specific
proteins. This technology is based on the use of a thiol reagent, such as
iodoacetamide, tagged with different isotopes (284). However, although in theory
these techniques could be applied to the study of the redox state of cysteines in
secreted proteins, it was not appropriate to test our hypothesis, that glutathionylated
proteins could be released by LPS-stimulated macrophages.
131
Carbonylated proteins have been extensively studied in inflammation or in diseases
associated with inflammation or neurodegeneration, and the use of redox proteomics
to identify them is reviewed by Butterfield et al (285). To my knowledge, the only
studies applying proteomics to identify proteins whose carbonylation is induced by
LPS in vivo have investigated intracellular proteins and not the secreted ones (286,
287).
Therefore the approach used in our experiments was an original one, in that no redox
proteomics studies had been performed on the LPS-induced macrophage secretome,
and particularly there were no studies attempting to identify released glutathionylated
proteins. We were hoping that the use of redox proteomics would lead to the
identification of previously non recognized proteins whose release was induced by
LPS although, clearly, the experimental approach was such that would only identify
proteins containing a free cysteine susceptible to glutathionylation.
4.2. Aim of Chapter 4
The purpose of the experiments described in this Chapter was to identify those
proteins released in the glutathionylated form in response to inflammatory stimuli
such as LPS by macrophages. Considering the scientific literature on the contribution
of MS (e.g. as described above for the diagnosis of diseases and the study of protein
function) and having a MS facility available for our research, a proteomic method
(combination of one-dimensional gel electrophoresis and MS) was used for this
project. MS could provide initial experimental data to identify released protein from
LPS-stimulated macrophages to be confirmed by other techniques as Western blot or
ELISA.
132
4.3. MS identification of proteins released by LPS-
stimulated Raw 264.7 cells
In order to study released proteins, experiments were performed using the BioGEE
method to label glutathionylated proteins as described in Methods (2.2.5). Results
were obtained using a combination of affinity purification, one-dimensional gel
electrophoresis and protein identification by MS. All the experiments of this chapter
(including cell treatments, pull-down with Streptavidin beads, SDS-PAGE and
staining for the MS experiments), were carried-out by myself, except for the MS and
MS data analyses, which were performed by Dr. Lucas Bowler.
Three independent experiments are described in this chapter and the experimental
procedure is shown in Figure 4.1. In particular, the experiments were performed
following 4 main steps.
Step 1 (BioGEE method): Raw 264.7 cells were preincubated with BioGEE,
treated with LPS and then the conditioned medium was concentrated with
Vivaspin centrifugal concentrator as described in Methods (2.2.5.1).
Step 2 (pull-down): the concentrated conditioned medium was incubated with
Streptavidin agarose beads and then captured biotinylated proteins were
eluted using the elution buffer as described in Methods (2.2.5.2).
Step 3: the proteins were separated onto 12% SDS-PAGE and the gel was
stained with Coomassie or Silver as described in Methods (2.2.5.3 and
2.2.5.4).
Step 4: proteins were identified by MS as described in Methods (2.2.5.5).
Considering that some are proteins are detectable only at higher concentrations (e.g.
25 ng) and furthermore, that the conditioned medium contains proteins in low
concentrations, it was necessary to find the most appropriate staining. For this
reason, the Comassie staining reagent used was the a GelCode Blue Stain reagent
solution, as it that has a sensitivity of 8 ng and is preferred to other traditional
homemade Coomassie dyes that only allow the detection of proteins in the
microgram range. However, the Coomassie staining solution was used for the first
part of experiment; mainly for practical reasons as Coomassie staining is performed
in only one step. The Silver staining was then an alternative option when only a few
133
of the bands were detected by Coomassie. In particular, as summarised in Figure 4.1,
proteins from Experiment 1 were separated onto 12% SDS-PAGE and then the gel
was stained using the Coomassie procedure. For Experiment 2, the Silver staining
was used, to optimize the visualization of the proteins. One more experiment
(Experiment 3) was performed as the previous experiments but MS analysis was
performed directly on-beads using NH4HCO3 without SDS-PAGE and staining.
134
Figure 4.1: Summary of the experimental procedure for the identification of the
released proteins. (1) Raw 264.7 cells were labelled with BioGEE and incubated with
LPS (BioGEE method). (2-3) Biotinylated proteins bound to Streptavidin beads were
resolved by SDS-PAGE and stained with Coomassie or Silver. One experiment was
also performed without protein separation on SDS-PAGE. (4) Proteins were
identified using MS.
135
Experiment 1
According to the protocol, a Coomassie gel staining (Figure 4.2) was performed to
visualize proteins from two samples. The two samples represent the first and the
second elution step of the same original sample (conditioned medium from
Raw 264.7 cells treated with BioGEE and LPS, and incubated with streptavidin
beads). Therefore, samples named sample 1 (first elution) and sample 2 (second
elution) were different from each other only for the elution step used. In particular
the first elution was performed for 30 min RT with SB 1X (+ 10 mM DTT) while for
the second elution the same volume of SB 1X (+ 10 mM DTT) was added to the
previous beads for 10 min at 100 °C.
From the Coomassie stained gel, only 3 major bands were visualized and then
excised for MS analysis. Lane 1 represents the Molecular Weight. Lane 2 represents
the first elution of the sample while lane 3 the second elution. One band at ~ 76 kDa
was excised from lane 2, one band at ~ 76 kDa and one more band at ~ 12 kDa from
lane 3.
Only a few bands were visible from Experiment 1, then in Experiment 2 the gel was
stained with Silver stain for best results. In particular, the experiment was performed
using Silver staining kit because of its greater sensitivity (sensitivity of 0.3 ng) than
the Coomassie stain. Furthermore, the kit used is compatible with MS because it does
not contain glutaraldehyde or formaldehyde in the sensitizer that can interfere with
trypsin digestion.
Experiment 2
This experiment was performed using the same experimental scheme as
Experiment 1. One more sample without preincubation with BioGEE was added as
control for the detection of non-specific binding. In Experiment 2 (Figure 4.3), more
bands were detected compared with the Coomassie stain experiment (Figure 4.2). In
addition more bands were observed in lanes 3 and 5 (sample with BioGEE)
compared to lanes 2 and 4 (sample without BioGEE). Even if more bands were
detected with Silver staining, the majority of the bands were non-specific since they
were also present in the sample without BioGEE. In order to eliminate false
positives, only one band (~ 24 kDa) was excised from the Silver gel (the band was
present only in the sample incubated with BioGEE).
136
HSP70 (70 kDa), profilin (15 kDa) and Trx1 (15 kDa) were found from the 3 bands
excised in Experiment 1 (Coomassie stain), while Prx1 was the protein identified in
Experiment 2 (Silver Silver).
137
Experiment 1
Figure 4.2: Biotinylated glutathionylated proteins in Raw 264.7 cells. Coomassie
stained of conditioned medium by LPS-stimulated macrophages. Raw 264.7 were
loaded with BioGEE and subsequently incubated with LPS for 24 hours. After
purification of biotin-labeled proteins on streptavidin agarose beads, proteins were
separated by a 12% SDS gel under reducing condition. Three major bands were
visible in Coomassie stained gel. The bands of interest were excised and HSP70,
profilin and Trx1 were identified by MS.
Lane 1 2 3
138
Experiment 2
Figure 4.3: Silver stained gel of conditioned medium from Raw 264.7 cells.
Confluent Raw 264.7 cells were cultured in serum free culture medium in the
absence (control) or presence of BioGEE and incubated with LPS. Supernatants were
prepared as described under Methods. Purified proteins were resolved by SDS-PAGE
and detected by Silver staining. Proteins were identified by MS. Molecular weight
markers (kDa) are indicated. Different bands were visible in Silver staining but only
one band at ~ 24 kDa was excised and Prx1 was identified by MS.
Lane 1 2 3 4 5
139
Experiment 3
In order to identify more proteins, the identification of the BioGEE labelled proteins
purified by streptavidin beads was also attempted by directly injecting into LC-MS
without electrophoretic separation (shotgun proteomics). The conditioned medium
was incubated with streptavidin beads and then the beads were washed firstly with
PBS, and then with NH4CO3. The beads were then subjected to tryptic digestion and
the SN was analysed by LC-MS as described in Methods (2.2.5.5). Indeed, in
Experiment 3, more proteins were identified as shown in Table 4.1. Furthermore,
several common proteins listed in the sample with BioGEE and in the sample
without preincubation with BioGEE showed the possible presence of released but not
glutathionylated proteins.
A key issue in this chapter was the need to remove possible false positives (non
biotinylated proteins that might bind non-specificically to streptavidin beads) from
our list. In order to solve this problem, only the proteins detected in the sample with
BioGEE were selected as shown in bold in Table 4.1. In particular 30 proteins were
identified from the sample without BioGEE and 40 proteins from the sample with
BioGEE. In particular, the HSP70 identified in Experiment 1 was found also in
Experiment 3, and it was only in the sample with BioGEE. Prx1 identified from the
band in Experiment 2 and profilin identified from Experiment 1 were considered non
specific for this experiment because they were also present in the sample without
BioGEE.
Table 4.2 summarizes the main function and the peptide coverage of the three
proteins selected as the candidate proteins to continue the project (HSP70, Prx2 and
vimentin). In addition, the peptide coverage of LDH is also indicated. The peptide
coverage was very low, only 5% for HSP70, 14% for Prx2, 5% for vimentin and
12% for LDH A. Among the proteins listed above, Prx2 was selected as the protein
of interest for further studies. The thesis presents the final results derived from the
research database as described in Methods (2.2.5.5) of the three experiments.
Furthermore, MS/MS spectra of HSP70, Prx2, vimentin and LDH A identified in
Experiment 3 are shown in Figure 4.4, Figure 4.5, Figure 4.6 and Figure 4.7
respectively.
140
Experiment 3
Sample without BioGEE (30 proteins) Sample with BioGEE (40 proteins) 6-phosphogluconate dehydrogenase 78 kDa glucose-regulated protein
78 kDa glucose-regulated protein Actin, cytoplasmic 1
Actin, cytoplasmic 1 Alpha-enolase
Actin-related protein 2/3 complex subunit 3 Bifunctional aminoacyl-tRNA synthetase
Adenosylhomocysteinase Cathepsin B
Alpha-enolase Elongation factor 1-gamma
D-3-phosphoglycerate dehydrogenase Elongation factor 2
Elongation factor 1-alpha1 Envelope polyprotein
Eukaryotic translation initiation factor 2
subunit 1 Exportin-1
GTP-binding nuclear protein Ran Fatty acid synthase
H-2 class I histocompatibility antigen, D-D alpha
chain Glucose-6-phosphate 1-dehydrogenase X
Heat shock cognate 71 kDa protein H-2 class I histocompatibility antigen, L-D alpha
chain
HSP90-alpha Heat shock 70 kDa protein 4
Hsc70-interacting protein Heat shock cognate 71 kDa protein
Isoform 1 of IL-1 receptor antagonist protein HSP90-alpha
Isoform C of Lamin-A/C Hypoxanthine-guanine
phosphoribosyltransferase
Isoform M2 of Pyruvate kinase isozymes M1/M2 Isoform 1 of Filamin-A
Monocyte differentiation antigen CD14 Isoform 1 of IL-1 receptor antagonist protein
Nucleolin Isoform 2 of Tropomyosin alpha-3 chain
Peptidyl-prolyl cis-trans isomerase B Isoform C of Lamin-A/C
Prx1 Isoform M2 of Pyruvate kinase isozymes M1/M2
Phosphoglycerate kinase 1 LDH A chain
Plastin-2 Multifunctional protein ADE2
Profilin-1 Nucleolin
Protein disulphide-isomerase A3 Peptidyl-prolyl cis-trans isomerase B
similar to Protein disulphide isomerase
associated 6
Prx1
T-complex protein 1 subunit beta Prx2
Triosephosphate isomerase Phosphoglycerate kinase 1
Tubulin alpha-1B chain Plastin-2
Tubulin beta-5 chain Profilin-1
Protein disulphide-isomerase A3
similar to Protein disulphide isomerase associated 6
Sulfated glycoprotein 1
T-complex protein 1 subunit epsilon
Transgelin-2
Transketolase
Triosephosphate isomerase
Tubulin alpha-1B chain
Tubulin beta-5 chain
Vimentin
Table 4.1: Proteins identified by MS in the experiment without separation with SDS-
PAGE (Experiment 3). Proteins identified only in the sample with BioGEE are
indicated in bold.
141
Protein name
Mass (kDa)
pI
Function MS
Peptides
Coverage
HSP70
72
4.8
Molecular chaperone
(288)
5%
(3 peptides)
Prx2
22
5.0
Antioxidant enzyme and
molecular chaperone (211)
14%
(2 peptides)
Vimentin
53
4.9
Intermediate filaments (289)
5% (2 peptides)
LDH A
36
7.6
Conversion of pyruvate to
lactate, regenerating NAD+
from reduced NADH
(290)
12%
(5 peptides)
Table 4.2: MS results of more interesting proteins identified in three independent
experiments. Some information (mass, function and peptide coverage) related to
HSP70, Prx2 and vimentin that were selected as candidate glutathionylated released
proteins from Raw 264.7 cells stimulated with LPS. LDH A was also identified from
MS.
142
MS/MS Spectra (Experiment 3)
Peptide 1)
Peptide 2)
Peptide 3)
Figure 4.4: Tandem mass spectra (MS/MS spectra) of HSP70 peptides obtained
during analysis of tryptic digests of samples by LC-MS. Each spectra represents the
ionic products of peptide (1-2-3) fragmentation produced by collision induced
dissociation (CID). Each labelled spectral peak indicates the 'b' (red) and 'y' (blue)
ion series identified and mapped to defined tryptic peptides contained within the
ipi.MOUSE.v3.72 database used for searching in this study. 'b' ions represent
charged peptide fragments that extend from the amino-terminus of the matched
peptide, the 'y' ions the fragments extending from the carboxy- terminus of the
peptide. Some neutral losses (loss of water or ammonia) may also be indicated
(green). Black peaks indicate unmatched ions; these will predominantly comprise
non-peptidic charged molecular species ('noise'). The sequence of the identified
peptide is determined by the mass differences observed between these peaks.
143
Peptide 1)
Peptide 2)
Figure 4.5: MS/MS spectra of 2 Prx2 peptides obtained during analysis of tryptic
digests of samples by LC-MS.
144
Peptide 1)
Peptide 2)
Figure 4.6: MS/MS spectra of 2 vimentin peptides obtained during analysis of
tryptic digests of samples by LC-MS.
145
Peptide 1)
Peptide 2)
Peptide 3)
146
Peptide 4)
Peptide 5)
Figure 4.7: MS/MS spectra of 5 LDH A peptides obtained during analysis of tryptic
digests of samples by LC-MS.
147
4.4. Discussion
We have successfully identified some proteins undergoing glutathionylation in cell
culture supernatants. In general, there have been many studies that led to the
identification of glutathionylated proteins using redox proteomics. All of these
studies have focused on intracellular proteins and, as far I know, ours is the first such
study on secreted proteins. The fact that most studies focused on cytosolic proteins is
not surprising considering that these proteins are known to have their cysteines
mostly in the reduced state that is not engaged in structural disulfide bonds, due to
the reducing environment in the cytosol. On the other hand, extracellular and
secreted proteins would have more structural disulfides and less free thiols because
the extracellular environment lacks reducing agents such as GSH (291) (292).
However, this distinction is not so strict and proteomics experiments have shown that
several cytosolic proteins can form transient disulphides, such as by glutathionylation
(293). Likewise, at least some serum/plasma protein, transthyretin and some serpins,
has been identified as glutathionylated (294) (295).
Proteins identified from the three experiments have two main common
characteristics: the presence of cysteines that can undergo glutathionylation and the
fact that they do not have a signal sequence and therefore must be released through
alternative pathway rather than classical secretory mechanisms. HSP70, profilin and
Trx1 identified by MS from Experiment 1 are proteins whose cysteine residues had
already been reported to undergo glutathionylation. In particular, under oxidative
stress glutathionylation of cysteine residues contained in Hsp70 (molecular
chaperone) can induce higher chaperon activity (69). Profilin is an actin-binding
protein (296), also identified as glutathionylated in oxidatively stressed human T
lymphocytes (187). Finally, Casagrande et al. identified glutathionylated Trx in
PBMC treated with diamide (187).
Furthermore, Trx1 and HSP70 lacking a signal peptide are secreted with a non-
classical secretory pathway as demonstrated by Tanudji et al. for Trx in transfected
CHO cells (237) and by Lancaster et al. (65) for Hsp70 released from PBMC using
an exocytotic pathway (exosomes-dependent). Prx1 identified in Experiment 2, also
shows cysteine residues and it is not secreted by ER/Golgi pathway. In Experiment 3,
148
we identified different proteins (e.g. Prx2 and vimentin) but only proteins identified
exclusively in the sample prepared with BioGEE were considered as glutathionylated
released protein by LPS-treated Raw 264.7 cells.
After several considerations, only 3 proteins (HSP70, Prx2 and vimentin) were
considered as possible candidates to continue future experiments. These proteins
were selected based on their relative interest from those identified by MS and Prx2
was chosen to start the project. Prx2, a member of the Prxs family could be secreted
by a non-classical secretory pathway (297) and its chaperone activity or the
peroxidase activity depends on the protein structure changes linked with
glutathionylation (211).
The presence of LDH (a cytosolic enzyme) among the proteins identified in the
supernatant might indicate that some cell necrosis takes place, and that the release of
at least some of the proteins identified by MS can be correlated with cell death. More
experiments, using an LDH will therefore be performed to confirm the presence of
LDH and then a possible cytotoxicity.
A post-translational modification such as glutathionylation can explain the low
peptide coverage of HSP70 (5%), Prx2 (14%) and vimentin (5%) indentified by MS.
Peptide identification by MS is heavily dependent on accurate mass determination as
deviation from expected mass will result in failure to find matches within sequence
databases. Search parameters used for database searching therefore usually allow for
a number of post-translational modifications although these are typically limited to
the most common and those most likely to be introduced by the sample preparation
methods used. In this study we restricted such optional post-translational
modifications to carboxyamidomethylation (on Cys residues), deamidation (on Asn)
and oxidation (on Met). It is not normal to include many more variable modifications
as this will considerably increase search times. Additionally, as naturally occurring
post-translational modifications are normally only present on small subsets of given
protein species, accurate identification can normally be made by matching of
unmodified peptide spectra only.
Nevertheless low sequence coverage can result from the presence of high levels of
(undefined) post-translation modification as suggested. However it is more likely that
149
the relatively low coverage is a result of sequence-dependent variability in the ease
of ionisation of the tryptic fragments generated (some peptides ionise more
efficiently than others, and abundant and readily ionisable species can prevent
ionisation and/or 'mask' the presence of others. Missed cleavage sites are also a
possible explanation as peptides outside a particular mass range (generally <5 amino
acids or 35> amino acids) are difficult to detect by MS (298).
4.5. Following chapter
I am well aware that identifying a protein by MS following labeling with BioGEE,
affinity purification and LPS stimulation is not a conclusive demonstration that the
protein is released in response to LPS, or even that it is glutathionylated, as the
finding could be due to non-specific binding to streptavidin or co-elution with other
proteins. Whether the release of the protein is actually increased by LPS needs to be
confirmed by other techniques (e.g. by Western blot with an anti-Prx2 antibody or by
ELISA) and its glutathionylation needs to be validated by different techniques (e.g.
immunoprecipitation followed by Western blot with an anti-GSH antibody), and
these are the experiments that will be part of the next Chapter.
150
Chapter 5. Prx2 release in LPS-stimulated mouse
and human macrophages
151
5.1. Introduction: extracellular Prx2
The presence of Prx1 and Prx2 among the proteins potentially released as
glutathionylated was of particular interest to us. In fact, in addition to be potentially
released in response to LPS, Prxs are important in redox regulation of inflammation.
In fact, Prxs are thioredoxin (Trx) peroxidases as described in chapter 2. They would
not be the only redox enzymes implicated in the metabolism of reactive oxygen
species or protein thiols that are released in inflammatory conditions. Earlier studies
have shown that the superoxide-generating enzyme, xanthine oxidase (XO), is
induced in animal models of inflammation, such as that induced by LPS injection,
and its concentration is increased in circulation (299). It is thought that XO may be
important in the pathogenesis of ischemia/reperfusion injury (162) and its inhibitors
are being tested in clinical trials in cardiovascular diseases (300-302).
Trx, the Prx substrate, is released by LPS-stimulated macrophages (303), and its
concentration is increased in the circulation in many inflammatory/infective diseases
(304). Thioredoxin may act as a reducing agent and thus help T cell proliferation
(305). Trx reductase, the enzyme that keeps Trx in the reduced state and is therefore
essential for its reducing activity, can also be released (306) is also increased in the
circulation of patients with rheumatoid arthritis (307).
As for Prxs, their secretion or release has already been the subject of some studies.
As mentioned in chapter 1, Prx4 has a signal peptide and is secreted through the
classical mechanism (308, 309). Prx4 is secreted rapidly, within 10 min according to
pulse-chase experiments (309). Extracellular Prx4 has heparin-binding properties and
binds to human umbilical vein endothelial cells; interestingly this binding is
diminished when Prx4 is oxidized by hydrogen peroxide or diamide (309). In terms
of biological activity, extracellular Prx4 protects cells from hydrogen peroxide
toxicity (309). Particularly in line with our findings that will be presented in Chapter
7, extracellular Prx4 activates NFkB in the human myeloid U937 cell line, and this is
followed by increased expression of inducible nitric oxide synthase (iNOS) and
Intercellular Adhesion Molecule 1 (ICAM1) (310). In the context of inflammation,
Prx4 has been detected in the plasma and synovial fluid from patents with
rheumatoid arthritis (311)
152
Prxs lacking the signal peptide are not secreted by the ER/Golgi pathway, but can be
secreted through alternative pathways. There are several study focused on the
extracellular Prxs. For instance, Chang et al. demonstrated that the secretion of Prx1
from A549 lung cancer cells was insensitive to brefeldin A and then using a non-
classical secretory pathway (312). Prxs are also secreted/released from other cells
such as astrocytes. In particular, proteomic analysis of extracellular medium showed
higher levels of Prx1 and Prx2 compared with the total cells extract. In addition, the
secretion was insensitive to brefeldin A (274).
Szabo-Taylor et al. investigated the intracellular and extracellular levels of Prxs in
patients with RA. In particular, lysate from Peripheral blood lymphocytes (RA
patients and healthy subject) was analysed by Western blot in non-reducing
conditions showing higher levels of Prx2 in RA lymphocytes while Prx3 levels were
similar in healthy and RA patients. Furthermore, Prx2 was detected in the redox
active form (only as dimer) as demonstrated by the absence of monomer
(overoxidated form). Prx2 was also identified on the surface of T cells in RA and in
particular on Th17 cells suggesting a possible role of Prx2 in the activation T cells in
RA. This study also demonstrated that Prx2 can be detected in plasma and synovial
fluid from RA patients. A trend for an increase in Prx2 levels was observed in
plasma of RA patients compared to healthy subject (313).
Prx2 was identified by 2D gel electrophoresis and MS in plasma from patients with
Severe Acute Respiratory Syndrome (SARS) while was not detected in normal
controls. In addition, Western blot of Prx2 in non-reducing conditions showed the
presence of a band at 44 kDa, corresponding to the dimeric form of Prx2 (314). High
levels of Prx2 in plasma of patients with Hepatitis B virus infection were also
observed, suggesting that Prx2 detection could be useful for early diagnosis of
infection diseases (315).
A recent article highlights the extracellular inflammatory role of Prxs in the ischemic
brain. Prxs released by necrotic brain cells are inactive as antioxidant due the
oxidation of the cysteines implicated in the peroxidase activity. Shichita et al.
showed that incubation of marrow-derived dendritic cells with brain lysate from
ischemic mouse brain induces proinflammatory cytokines such as IL-23 and then IL-
17. Then, the brain lysate was analysed by MS and different proteins such as Prxs
153
and HMGB1 were identified. Furthermore, the reduction of inflammation in
ischemic mice deficient of TLR2 and TLR4 suggested that Prxs act as DAMP
through the binding with TLR2 and TLR4. Treatments with Prx-specific antibodies
protects from stroke reducing the size of the infarct (316).
5.2. Aim of Chapter 5
The previous chapter described the identification of unknown released
glutathionylated proteins from LPS-stimulated macrophages. This chapter will focus
on Prx2, as the protein of interest for further studies. This chapter presents the results
obtained by Western blot to confirm the release of Prx2 in Raw 264.7 cells and in
mice (in vivo experiments). More experiments in human cells (human macrophages
and PBMC) to measure Prx2 levels were also performed.
154
5.3. Detection of released Prx2 from Raw 264.7 cells
According to the initial identification by MS analysis, the following step of this study
was to confirm the release of Prx2 by Western blot. In MS experiments, Raw 264.7
cells were incubated for 24 hours with LPS but a time course setup experiment,
followed by Prx2 Western blot was also performed for a better understanding of Prx2
release after LPS treatment. Briefly, cells were plated in 24-well plates at 0.3 x 106
with complete RPMI 1640 medium containing 10% FBS as described in Methods
(Table 2.2). After ON, the medium was removed and cells washed with PBS and
Opti-MEM I medium. Treatment with 100 ng/ml LPS was carried out in a minimum
of 200 µl of Opti-MEM I medium.
For the time course experiment (Figure 5.1) cells were incubated with LPS or only
medium (control) and analysed at different time points (0, 2, 4, 8 hours, ON).
Supernatants were centrifuged and pooled for each time point. Proteins were
subjected to 12% SDS-PAGE gel under reducing conditions followed by incubation
with anti-Prx2 as described in Methods (2.2.6.1). Results of the time course
experiment confirmed first of all the presence of Prx2 into medium of treated
Raw 264.7 cells as previously identified by MS. Furthermore, results showed that
released Prx2 levels were significantly increased in LPS-treated cells compared to
untreated cells. However, basal levels of Prx2 were also detected in untreated cells.
Comparing all time incubation, a higher band of Prx2 was detected with LPS
incubated ON, and released Prx2 was chosen as protein to continue the research.
More experiments were performed to investigate released Prx2 in Raw 264.7 cells
treated with LPS for 24 hours as previous MS experiments but without previous
incubation with BioGEE.
Another experiment treating Raw 264.7 cells with LPS for 4 hours (short incubation
time) and LPS 24 hours (long incubation time) was performed. Results showed a
difference between LPS 24 hours and untreated cells confirming once again higher
Prx2 signal with LPS long incubation time treatment. Furthermore, release of Prx2
with LPS 24 hours was higher compared to LPS 4 hours (Figure 5.2). On the basis of
our preliminary experiments, all subsequent experiments were conducted incubating
cells with LPS for 24 hours.
155
Figure 5.1: Time course of Prx2 release in Raw 264.7 cells. Raw 264.7 cells were
incubated with LPS at different time as described in Methods and supernatants were
separated on 12% SDS-PAGE under reducing conditions (10% 2-ME). Experiment
performed in triplicate and samples analysed as a pool for each time point. Upper
panel: Western blot of Prx2. Lower panel: Densitometric analysis.
156
Figure 5.2: Western blot showing the effect of LPS 24 and LPS 4 hours on Prx2
release by Raw 264.7 cells. Equals amounts of supernatants of Raw 264.7 cells,
treated as described in Methods were analysed by Western blot using an anti-Prx2
antibody. Values represent the mean of an experiment performed with duplicate
samples (n=2 per condition). Dark grey bar indicates mean values for control (values:
189-172; mean: 180.5), black bar indicates mean values for LPS 24 hours treatment
(values: 283-347; mean: 315) and light grey bar indicates mean values for LPS 4
hours treatment (values: 147-128; mean: 137.5).
157
Detection of released Prx2 by Western blot from Raw 264.6 cells in the experiments
described above (Figure 5.1 and Figure 5.2) confirmed the previous protein
identification by MS. However, more experiments were performed following the
final experimental scheme shown in Figure 5.3. Briefly, Raw 264.7 cells were plated
in 24-well plates at 0.3 x 106 in complete RPMI 1640 medium and treated in Opti-
MEM I medium with and without LPS for 24 hours, then collected, centrifuged and
analysed by Western blot.
The experimental scheme described above was also adopted for the detection of
intracellular Prx2. Cells were lysed directly in SB 1.5X with 10% 2-ME. Released
and intracellular Prx2 was separated by 12% SDS-PAGE in reducing conditions. In
particular, equal volumes of SN (20 µl/lane) or total cell lysate (5-10 µl/lane) were
loaded.
Results of five independent Prx2 Western blot are shown below (Figure 5.4, Figure
5.5, Figure 5.6, Figure 5.7 and Figure 5.8). Samples were analysed in single and
statistic (t-test) was applied only for experiments performed in triplicate and in
quadruplicate as described in Methods (2.2.13). In all 5 experiments, Prx2 bands
were detected by Western blot at ~ 22 kDa, quantified by densitometry and
expressed in arbitrary units (AU). In the first series of experiments (Experiments 1-2)
only released Prx2 from Raw 264.7 cells treated with LPS was examined, as
described above. Briefly, Experiment 1 was performed using two replicates per
biological condition (2 control and 2 LPS). Densitometric analysis in Figure 5.4
shows higher Prx2 levels (2.0-fold increase) in samples incubated with LPS
compared to the control. Experiment 2 was performed using three replicates per
biological condition (3 control and 3 LPS) and only weak bands of Prx2 were
detected in LPS-treated cells. However, an increasing trend in LPS-treated cells
when compared to the control was observed, as shown in Figure 5.5.
158
Figure 5.3: Experimental scheme used for LPS stimulation of Raw 264.7 cells.
Twenty-four hours after treatment, SN or cell lysate were analysed by Prx2 Western
blot.
159
Experiment 1
Figure 5.4: Western blot showing the effect of LPS on Prx2 release by Raw 264.7
cells (Experiment 1). Equals amounts of supernatants of Raw 264.7 cells, treated as
described in Methods were analysed by Western blot using an anti-Prx2 antibody.
Values represent the mean of an experiment performed with duplicate samples (n=2
per condition). Grey bar indicates mean values for control (values: 145-197;
mean: 171), and black bar indicates mean values for LPS treatment (values: 302-382;
mean: 342).
160
Experiment 2
Figure 5.5: Western blot showing the effect of LPS on Prx2 release by Raw 264.7
cells (Experiment 2). Upper panel: Western blot of Prx2 and lower panel:
densitometric analysis. Grey bar indicates mean values ± SD for control (n=3 per
condition), and black bar indicates mean values ± SD for LPS treatment. Control
(min-max range: 6-14; mean: 9). LPS (min-max range: 22-56; mean: 38). Error bars
represent the standard deviation from triplicate samples. *p<0.05 versus control.
161
This data were also confirmed in the second series of experiments (Experiments 3, 4
and 5), where intracellular Prx2 was examined as showed in Figure 5.6, Figure 5.7,
and Figure 5.8). In particular, a significant increase (3-fold compared to the control)
of released Prx2 levels is showed in Figure 5.6 and Figure 5.8, while similar
intracellular Prx2 levels were detected in samples with and without LPS treatment in
both experiments. Similarly, a significant increase of released Prx2 levels (6-fold
compared to the control) is showed in Figure 5.7 (Experiment 4) while higher
intracellular Prx2 levels (~2.4-fold compared to LPS) were observed in control cells.
The average of Prx2 increase in these five experiments is 3.6.
To summarize, in Experiments 3 and 5 the intracellular Prx2 levels were similar in all
samples (control and LPS) while experiment 4 shows lower intracellular Prx2 levels
in LPS treatment compared to the control. The variability in the data between the
three experiments may be due to the amount of total protein loaded for the Western
blot. In conclusion, these experiments did not allow us to demonstrate whether the
release of Prx2 from the cells is linked with a new protein synthesis. Overall, these
results clearly confirm that Prx2 is released in LPS-stimulated Raw 264.7 cells by
Western blot in five independent experiments.
162
Experiment 3
Figure 5.6: Effect of LPS on released and intracellular Prx2 levels in Raw 264.7
cells (Experiment 3). Released and intracellular detection of Prx2 by Western blot
(upper panel) and a corresponding densitometric analysis (lower panel). Grey bars
indicate mean values ± SD for control (n=3 per condition), and black bars indicate
mean values ± SD for LPS treatment. Supernatant: control (min-max range: 56-60;
mean: 65). LPS (min-max range: 138-226; mean: 190). Cells: control (min-max
range: 536-606; mean: 562). LPS (min-max range: 425-523; mean: 481). Error bars
represent the standard deviation from triplicate samples as described in the upper
panel. **p<0.01 versus control in supernatant.
163
Experiment 4
Figure 5.7: Effect of LPS on released and intracellular Prx2 levels in Raw 264.7
cells (Experiment 4). Western blot (upper panel) of an experiment in quadruplicate
(n=4 per condition) and a corresponding densitometric analysis (lower panel). Grey
bars indicate mean values ± SD for control, and black bars indicate mean values ±
SD for LPS treatment. Supernatant: control (min-max range: 12-19; mean: 15). LPS
(min-max range: 86-110; mean: 95). Cells: control (min-max range: 420-643;
mean: 547). LPS (min-max range: 207-259; mean: 229). Error bars represent the
standard deviation from quadruplicate samples as described in the upper panel.
***p<0.001 versus its control.
164
Experiment 5
Figure 5.8: Effect of LPS on released and intracellular Prx2 levels in Raw 264.7
cells (Experiment 5). Prx2 detection by Western blot (upper panel) and densitometric
analysis (lower panel) of an experiment performed with duplicate samples (n=2 per
condition). Grey bars indicate mean values for control, and black bars indicate mean
values for LPS treatment. Supernatant: control (values: 17-43; mean: 30). LPS
(values: 110-111; mean: 110). Cells: control (values: 761-884; mean: 822). LPS
(values: 602-657; mean: 630).
165
Incubation with LPS in Raw 264.7 cells as previous described were also performed
to study whether Prx2 was released as monomer, dimer or higher molecular weight
structure. For this purpose, released Prx2 was analysed by Western blot in non-
reducing conditions in supernatant and cell lysates. Experiments in Figure 5.9 show
the presence of a 38 kDa band (dimer) for both supernatant and cell lysates samples.
In contrast, Western blot of supernatant showed only the band at 38 kDa, suggesting
that released Prx2 existed mainly as a dimer, reduced to monomeric form by 2-ME.
166
Figure 5.9: Prx2 detection by Western blot in non-reducing conditions. Left panel:
Prx2 levels in supernatant. Right panel: Prx2 levels in cell lysates.
167
5.4. Passive release of Prx2
Many publications suggest that different proteins can be released passively from cell
death. For instance, Ireland et al. (317) showed that HSP70 was released by necrotic
cells. The release of HSP70 from necrotic cells was also detected by experiments in
Basu’s laboratory (64). Furthermore, many findings demonstrated that HMGB1 is
secreted actively by stimulated macrophages and monocytes (238) or released
passively from necrotic cells (240).
Since the previous data demonstrated Prx2 release in murine macrophages treated
with LPS, in order to better understand whether cell death was also implicated for the
release of Prx2, cell viability by CTB and LDH assay were performed in Raw 264.7
cells treated with 100 ng/ml LPS for 24 hours.
5.4.1. CTB assay
Cell viability was measured in Raw 264.7 cells treated with and without 100 ng/ml
LPS for 24 hours. Experiments were performed in 96-well plates following the
manufacturer’s instructions (2.2.2). Results shown in Figure 5.10 are the mean of
three independent experiments with triplicate samples for each condition and
suggested that treatment with LPS can decrease cell viability from 100% (control) to
72%.
5.4.2. LDH assay
According to the LDH identification by MS in the Experiment 3 of Chapter 4 (Table
4.1) the LDH release was also measured by the CytoTox96 kit assay in supernatant
of Raw 264.7 cells as described in Methods (2.2.3). The experiment was performed
using three replicates per biological condition (3 controls and 3 LPS). The result of
one experiment in triplicate showed in Figure 5.11 suggests that LPS had an effect on
LDH release. A significant 3-fold increase of LDH release was observed in
supernatant of untreated cells (control) when compared with LPS-treated cells (LPS).
168
Figure 5.10: Effect of LPS on Raw 264.7 cell viability. Cell viability was measured
as described in Methods with CTB and expressed at % control value. The cell
viability was 72% in cells treated with LPS when compared with the control. Bars
represent the mean ± SD of three independent experiments, each run in triplicate.
Values obtained from control cell (white bar) were taken as 100% viability. Control
(min-max range: 79-111; mean: 100). LPS (black bar) (min-max range: 65-89; mean:
72). Error bars represent the standard deviation from triplicate experiments.
***p<0.001 versus control.
169
Figure 5.11: LDH release by Raw 264.7 cells. (n=3 per condition) White bar
indicates mean values ± SD for control, and grey bar indicates mean values ± SD for
LPS treatment. The total LDH released (black bar) was considered 100%. Total LDH
(min-max range: 83-131; mean: 100). Control (min-max range: 1.9-4.3; mean: 2.7).
LPS (min-max range: 8-11.3; mean: 9.8). Error bars represent the standard deviation
from triplicate samples. **p<0.01 versus control.
170
5.5. Detection of released Prx2 from PBMC and human
macrophages
Following the results obtained by Western blot analysis of released proteins in
Raw 264.7 cells which had been stimulated for 24 hours with bacterial LPS, it was
next examined whether Prx2 was also released by human cells (PBMC and human
macrophages) treated with LPS. Prx2 concentration in the supernatant was measured
by the human Prx2 ELISA kit as described in Methods (2.2.12.3).
5.5.1. PBMC
The experiment was performed with five healthy volunteer donors. Briefly PBMC
were prepared as described in Methods and cells were plated in 96-well plates at
0.5 x 106
in 100 µl of complete RPMI (5% FBS). After ON incubation, a 100 µl of
2 X working solution of LPS (final concentration: 100 ng/ml) or only medium was
added to PBMC for 24 hours.
The results from each donor are expressed as independent experiments and showed
in one graph (Figure 5.12). Experimental results suggested that LPS increases Prx2
production not only in mouse macrophages as shown in previous experiments by
Western blot but also in PBMC analysed by ELISA. In particular, a 4-fold increase
in Prx2 levels was observed in Donor A, 1.5-fold in Donor B, 2.8-fold in Donor C,
2.1-fold in Donor D and 4.7-fold in Donor E. The average of Prx2 increase of these
five experiments is 3, a similar average increase was obtained by Western blot
analysis of LPS-treated or untreated Raw 264.7 cells.
Since basal levels of secreted Prx2 were also detected in untreated cells, it was
investigated whether cells were activated. To this purpose, in parallel with Prx2, the
TNF-α production was also measured. TNF-α levels were undetectable (<15 pg/ml)
in supernatants of untreated PBMC, whereas we could measure high levels of
secreted TNF-α in PBMC treated with LPS for 24 hours, as expected (Table 5.1).
171
In conclusion, even if different basal levels of Prx2 were detected in control samples,
an increasing common tendency of Prx2 release was observed in PBMC treated with
LPS for 24 hours. Furthermore, the low levels of TNF-α from untreated PBMC
compared to LPS-treated PBMC demonstrated that cells were not activated and that
the effect of Prx2 release in treated cells was only due to LPS treatment.
172
Figure 5.12: Effect of LPS on Prx2 release in PBMC. Results from five independent
donors performed in duplicate (PBMC from donor A), in quadruplicate (PBMC from
donor B) or in triplicate (PBMC from donor C, D and E) are shown above. Prx2
levels were measured by ELISA. Grey bars indicate mean values ± SD for control,
and black bars indicate mean values ± SD for LPS treatment. Donor A: control
(values: 1.6-1.6; mean: 1.6). LPS (min-max range: 6.1-7; mean: 6.5). Donor B:
control (min-max range: 6.5-7.5; mean: 7). LPS (min-max range: 9.5-12.2;
mean: 10.9). Donor C: control (min-max range: 0.5-0.7; mean: 0.7). LPS (min-max
range: 1.4-2.5; mean: 2) Donor D: control (min-max range: 1.2-1.8; mean: 1.6). LPS
(min-max range: 2.6-5; mean: 3.4). Donor E: control (min-max range: 3.2-4.1;
mean: 3.7). LPS (min-max range: 17-18; mean: 17.6). Error bars represent the
standard deviation. *p<0.05, **p<0.01, ***p<0.001 versus control.
173
5.5.2. Human macrophages
In an identical way to PBMC samples, Prx2 levels were also quantified by ELISA in
human macrophages. The experiment was performed with four healthy volunteer
donors and primary macrophages were differentiated from PBMC as described in
Methods (2.2.1). Briefly, cells were plated in 96-well plates at 0.2 x 106
in 100 µl of
complete RPMI (5% FBS) and incubated with LPS following the same experimental
scheme as described above for PBMC.
The results from each donor are expressed as an independent experiment and shown
in one graph (Figure 5.13). Experimental results suggested that LPS increases Prx2
production in human macrophages. In particular, 2.7-fold increase in Prx2 levels was
observed in human macrophages from Donor F treated with LPS compared to
control cells, only 1-fold in Donor G, 1.8-fold in Donor H and 1.4-fold in Donor I.
The average of Prx2 increase of these four experiments is 1.7-fold.
Furthermore, as in the previous PBMC experiments, no TNF-α (<15 pg/ml) was
detected by ELISA in supernatants from untreated macrophages in the four
experiments. All samples were pooled and tested in single for TNF-α production
(Table 5.1). All samples were pooled and tested in single for the TNF-α production.
This result support the initial idea that LPS induces Prx2 release, confirming then
that the effect of Prx2 release is only due to LPS treatment and was not associated
with previous cells activation.
In conclusion, these experiments showed that higher Prx2 levels were released by
human macrophages after LPS stimulation compared to control. However, Prx2
release observed in human macrophages was lower compared to Prx2 levels
evaluated from LPS-treated Raw 264.7 cells and PBMC.
174
Figure 5.13: Effect of LPS on Prx2 release in human macrophages. Results from
four donors were performed in quadruplicate (macrophages from donor F and G) or
in triplicate (macrophages from Donor H and I) as shown above. Prx2 levels were
measured by ELISA. Grey bars indicate mean values ± SD for control, and black
bars indicate mean values ± SD for LPS treatment. Donor F control (min-max range:
0.4-1.15; mean: 0.8). LPS (min-max range: 2-2.5; mean: 2.2). Donor G control (min-
max range: 4.2-5.2; mean: 4.8). LPS (min-max range: 4.3-6.2; mean: 5.3). Donor H
control (min-max range: 4.2-5.2; mean: 4.7). LPS (min-max range: 7.3-9.3;
mean: 8.5). Donor I: control (min-max range: 2.3-3; mean: 2.6). LPS (min-max
range: 3-4.5; mean: 3.8). **p<0.01, ***p<0.001 versus its Control.
175
TNF (pg/ml)
Control LPS
PBMC
Donor A <15 850
Donor B <15 430
Donor C <15 7500
Donor D <15 4240
Donor E <15 2490
Human macrophages
Donor F <15 2700
Donor G <15 7500
Donor H <15 5670
Donor I <15 5840
Table 5.1: TNF-α levels in supernatants from untreated (control) and LPS-stimulated
PBMC and human macrophages. Supernatants (duplicates from Donor A,
quadruplicates from Donor B, triplicates from Donor C-D-E, quadruplicates from
Donor F-G and triplicates from Donor H-I) were pooled together and assayed in
single by ELISA.
176
5.6. Detection of Prx2 levels in serum from LPS-treated
mice
The release of Prx2 in vitro (murine and human cells) was confirmed in the previous
experiments. To investigate whether Prx2 is also released in vivo, serum from mice
was analysed by Western blot as described in Methods (2.2.6.2).
Three experiments of mice treated with vehicle (saline) or LPS (400 µg/mouse) were
performed. In Experiment 1, mice (2 each group) were sacrificed at 90 min or
24 hours post treatment. Western blot results shown in Figure 5.14 suggested that
Prx2 levels in LPS-treated mice were 1.8-fold higher compared to vehicle-treated
mice. Basal levels of Prx2 were also observed in mice treated only with saline.
Furthermore, no differences were observed between the group treated with LPS for
90 min and the group treated for 24 hours.
One more experiment (Experiment 2, Figure 5.15) was also repeated at 90 min but
this result did not confirm an increase of LPS group compared to vehicle. For this
reason the treatment with LPS for 24 hours was preferred to the treatment with LPS
for 90 min.
In Experiments 3 and 4 (Figure 5.16 and Figure 5.17) mice were injected with
vehicle or LPS for 24 hours and serum was analysed by Western blot, as in the
previous experiments. In Experiment 3, Prx2 levels in LPS-treated mice were
increased of 4.5-fold compared to vehicle-mice while in Experiment 4 a non-
significant increase of 1.4-fold was observed. As seen by the results from all the
experiments, Prx2 levels in LPS-treated mice were higher than in the vehicle;
however this difference was not statistically significant. A result more accurate can
be obtained increasing the number of mice for experiment.
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Experiment 1
Figure 5.14: Western blot analysis of serum from LPS-treated mice (90 min or
24 hours) and vehicle-treated mice (Experiment 1). Upper panel: Western blot of
Prx2; Lower panel: Densitometric analysis. Serum was prepared as described under
Methods and Prx2 was visualized by Western blot. Each lane of the Western blot
represents serum from one single mouse. The bands corresponding to Prx2 were
quantified by densitometric analysis and values are expressed as arbitrary units (AU).
Values represent the mean of an experiment performed with duplicate samples (n=2
per condition). White bar indicates mean values for vehicle group, grey bar indicates
means values for LPS 90 minutes group, and black bar indicates mean values for
LPS 24 hours group. Vehicle (values: 102-210; mean: 156). LPS 90 minutes (values:
272-312; mean: 292). LPS 24 hours (values: 253-339; mean: 296).
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Experiment 2
Figure 5.15: Western blot analysis of serum from LPS-treated mice (90 min) and
vehicle-treated mice (Experiment 2). Upper panel: Western blot of Prx2; Lower
panel: Densitometric analysis. Serum was prepared as described under Methods and
Prx2 was visualized by Western blot. Each lane of the Western blot represents serum
from one single mouse. The bands corresponding to Prx2 were quantified by
densitometric analysis and values are expressed as arbitrary units (AU). Values
represent the mean of an experiment performed with triplicate samples (n=3 per
condition). White bar indicates mean values for vehicle group, grey bar indicates
means values for LPS 90 minutes group. Vehicle (min-max range: 264-360; mean:
300). LPS 90 min (min-max range: 188-264; mean: 246). Error bars represents the
standard deviation from triplicate samples. There was no statistically significant
(n.s).
179
Experiment 3
Figure 5.16: Serum Prx2 levels after vehicle (saline) or LPS (24 hours) injection in
mice (Experiment 3). Upper panel: Western blot of Prx2; Lower panel:
Densitometric analysis. Serum (n=4 for each group) was prepared as described under
Methods. One lowest value (the first value) of the vehicle group and one lowest
value (the third value) of the LPS group were removed from the average and from t-
test calculation. White bar indicates means values for vehicle group, and black bar
indicates means values for LPS 24 hours group. Vehicle (min-max range: 16-91;
mean: 59). LPS 24 hours (min-max range: 206-320; mean: 269). Error bars
represents the standard deviation from triplicate samples. **p<0.01 versus control.
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Experiment 4
Figure 5.17: Serum Prx2 levels after LPS (24 hours) injection in mice
(Experiment 4). Upper panel: Western blot of Prx2; Lower panel: Densitometric
analysis. White bar indicates mean values ± SD for vehicle group, and black bar
indicates mean values ± SD for LPS 24 hours group. Vehicle (min-max range: 212-
509; mean: 357). LPS 24 hours (min-max range: 337-687; mean: 488). There were
no statistically significant (n.s) differences between the vehicle group and LPS
group.
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5.7. Discussion
The presence of Prx2 in supernatants from macrophages activated by LPS was an
important finding. To support the identification of Prx2 obtained by MS, more
experiments were performed and Prx2 release was analysed by Western Blot. Results
confirmed the increased levels of Prx2 in supernatant from Raw 264.7 cells treated
with LPS for 24 hours. Furthermore, Western blot in non-reducing conditions
showed the dimer structure of extracellular Prx2, while both monomer and dimer
forms of Prx2 were present in the cell lysate.
Based on the information that release from dying cells can be an additional way of
release of danger signals (such as HMGB1, ATP or uric acid), we hypothesized a
possible passive release of Prx2 in a similar way to HMGB1. In fact, HMGB1 that is
a nuclear factor can be secreted from activated macrophages and monocytes or
passively released by necrotic but not apoptotic cells. Thus, on the basis of the cell
viability results in LPS-treatment that showed a cell death of 28% when compared
with control cells and considering the higher level of Prx2 in cells treated with LPS,
we concluded that we cannot exclude that cell death may contribute to the release of
Prx2. However, we believed that this release occurs mainly by non-classical
secretory pathway such as the exosome-dependent pathway. Exosome are small
vesicles derived from the fusion of endosomes/lysosomes with the plasma
membrane. Exosome-dependent pathway has been shown to be a new secretory
pathway for HSP70 secreted by endothelial cells (318) and PBMC (65). By a
proteomic analysis, Thery et al. identified different exosomal proteins (e.g. Trx
peroxidase II and Alix) derived from dendritic cells (278).
The study by Shichita et al. showed that Prxs are released following cellular necrosis.
Our finding indicate that Prx2 release can also be induced by LPS and suggesting
that cellular necrosis may not be the only explanation for this. For instance, in
Chapter 6, we showed that BSO, despite increasing cell death, does not augment, but
rather inhibit, Prx2 release. It should also be stressed that we are using exactly the
same experimental model used in the laboratory of Kevin Tracey to study LPS-
induced release of HMGB1 (238). As in the case of Prx2, HMGB1 is also released
not only actively, following LPS stimulation, but also passively as a result of cellular
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necrosis (240). We think that the many studies reporting the presence of Prx2 in the
secretome (discussed above in this chapter) strengthen the view that Prx2 release is
not just secondary to cell toxicity.
Of note this is different from what has been reported by Shichita et al. (316). In fact,
they reported that in cerebral ischemia there is an extracellular release of most Prxs,
but this was associated with an increased mRNA expression for most Prxs (although
no statistical analysis was provided for these data), which seems at variance with our
results. This can reflect a difference in the stimulus used (LPS in our study; ischemic
injury in Shichita et al (316)) as well as in the in vivo versus in vitro models.
Furthermore, Shichita et al detected mostly extracellular Prx6 and Prx1/2 expression
was evaluated by immunostaining in control brains only, thus not conclusively
demonstrating the release of Prx2 in the extracellular space.
Additional experiments confirmed the presence of Prx2 levels in conditioned
medium from human cells (human macrophages and PBMC) measured by ELISA.
Furthermore, a tendency to release Prx2 was observed in serum from mice injected
with LPS (in vivo experiments) by Western blot analyses. Since that Prx2 is the third
most abundant protein of red blood cells, the presence of Prx2 in mice serum in the
experiments of this thesis could be explained by hemolysis of red blood cells. The
lysis of these cells could cause release of Prx2 into the serum. However, in our
experiments, samples visibly hemolyzed were not analysed. In addition, the release
of Prx2 independently of hemolysis was demonstrated by Antunes et al. in red blood
cells and suggested in vivo release of Prx2 under physiological conditions (319).
5.8. Following chapter
In this chapter, we confirmed the release of Prx2 in Raw 264.7 cells previously
identified by MS. In addition, new experiments demonstrated the release of Prx2 in
human cells and the possibility if its release in vivo as shown by analysis of serum
from mice injected with LPS. The decrease in cell viability and the detection of LDH
into the culture medium of Raw 264.7 treated with LPS can suggest cell death as a
possible mechanism of Prx2 release. However, our hypothesis is that the release of
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Prx2 is not just due to cell death but that a non-classical release pathway is also
implicated. The following chapter reports work performed to confirm
glutathionylation of released Prx2 in LPS-activated macrophages and whether is
implicated in its release.
184
Chapter 6. Glutathionylated Prx2 released by
Raw 264.7 cells
185
6.1. Introduction
Most studies of protein glutathionylation focused on cytoplasmic proteins for several
reasons, including their high concentrations, ease of migration in gel electrophoresis.
But the main reason was probably that it is normally thought that, because
glutathione is present intracellularly in high concentrations, and free cysteines are
more abundant in cytoplasmic proteins where structural disulfides are relatively rare,
there will be a higher chance of formation of mixed disulfides (291, 292)(reviewed
in: (320)). Another point is that, while the main small molecular weight thiol in the
cytosol is GSH/GSSG, in the extracellular environment it is cysteine (137) (321). For
this reason, it is often thought that, extracellularly, if proteins are present as mixed
disulfides they will be cysteinylated, rather than glutathionylated (322). While the
main cysteinylated protein found extracellularly is albumin (323), it is particularly
important in the context of my work that a typical cytokine, migration-inhibitory
factor (MIF), also known as glycation inhibitory factor (GIF), is secreted by T cell in
the cysteinylated form (324). In that case, cysteinylation was particularly important
as the non-cysteinylated cytokine produced as a recombinant protein in Escherichia
coli is not biologically active (324). Cysteinylation of MIF must occur during or after
the process of secretion/release because intracellular MIF/GIF is not cysteinylated
and it is also inactive (324). On the other hand, the protein transthyretin, a thyroid
hormone carrier that has a signal peptide and presumably is secreted via a classical
mechanism (325), which is present in the circulation in significant amount, is
significantly glutathionylated in plasma and other biological fluids (294, 326, 327).
Several studies have shown that Prxs can be glutathionylated. In particular, Prx1 and
Prx5 are glutathionylated in human T lymphocytes treated with diamide as showed
by Fratelli et al. (177). Recent studies (211) also reported glutathionylation of Prx1
and Prx2 after treatment with H2O2 in HeLa cells and highlight the link between
glutathionylation and the regulation of the Prxs functions (328). Park et al. suggested
that glutathionylation of Prx1 induces structural change from decamer to dimer with
loss of chaperone activity and the highest peroxidase activity (211).
It should be noted, however, that all these studies were performed on intracellular
Prxs. Another enzyme in the same class of the protein thiol-disulfide oxidoreductase
186
that was reported to undergo glutathionylation is Trx (187). Also in this case,
however, glutathionylation was identified in the intracellular form of Trx in human T
cells secreted. Because Trx also lacks a signal peptide and is released through a yet
to be defined non-classical mechanism, and its release is also stimulated by LPS
(303), it will be interesting to establish whether the released form of Trx can also be
present in the glutathionylated form. Glutaredoxin can also form a glutathionylated
intermediate as part of the catalytic reaction (329).
A second aspect of the glutathionylation of Prxs is that, like any other form of
protein glutathionylation it is reversible. As discusses in section 1.5.2, Grxs are the
most studied de-glutathionylating enzymes. Inhibition of Grx1 by means of siRNA
causes hyperglutathionylation of proteins has been a widely used tool to study the
role of glutathionylation (for instance, of actin: see (330)). However, there are
evidences that Grx2 can act as a glutathionylating enzyme, at least in some
experimental models (322). Specifically, in the case of Prx1, sulfiredoxin seems to
be more important than Grx in its de-glutathionylation. In addition, protein
glutathionylation might be diminished in cells where GSH has been depleted. This
has been observed in endothelial cells where treatment with the GSH synthesis
inhibitor, BSO, inhibited glutathionylation of the p65 subunit of nuclear factor kappa
B (331).
This chapter reports the results of experiments designed to test two main hypotheses.
The first hypothesis was that the Prx2 would be glutathionylated when released from
LPS-stimulated macrophages. The second one was more aimed at studying the
importance of glutathionylation by investigating whether GSH levels can regulate the
release of Prx2.
6.2. Aim of Chapter 6
This chapter aims first at confirming that Prx2 released by LPS-stimulated Raw
264.7 macrophages is glutathionylated. This will be done using a widely-used
approach consisting of immunoprecipitation of Prx2 followed by Western blot using
187
anti-GSH antibodies. A second aim is to assess the relevance of glutathionylation in
the release of Prx2. For this purpose, I will use an inhibitor of GSH synthesis,
buthionine sulfoximine (BSO) and evaluate its effect on Prx2 release.
188
6.3. Immunoprecipitation of Prx2
As shown in Chapter 4, Prx2 was identified in Raw 264.7 cells preincubated with
BioGEE and treated with LPS. Proteins were glutathionylated with BioGEE and LPS
and then biotinylated glutathionylated proteins were separated from the others using
streptavidin beads. Then, biotinylated glutathionylated proteins were identified by
MS. In order to confirm the release of Prx2 in glutathionylated form, supernatant
from LPS-stimulated Raw 264.7 cells was immunoprecipitated as described in the
Methods section (2.2.7.2).
Immunoprecipitated proteins were subjected to 12% SDS-PAGE gel under non-
reducing conditions followed by incubation with anti-GSH to identify
glutathionylated immunoprecipitated Prx2 (Prx2-SSG) and then incubated with an
anti-mouse secondary antibody. As seen by the result shown in Figure 6.1, in non-
reducing conditions a band corresponding to the dimer of Prx2 was detected. The
signal for Prx2-SSG detected by GSH Western blot indicated that Prx2 was
glutathionylated.
189
Figure 6.1: Released glutathionylated Prx2 by Raw 264.7 cells treated with LPS.
The band corresponding to Prx2-SSG was detected in sample immunoprecipitated
with anti-Prx2 and then a Western blot under non-reducing conditions using an anti-
GSH antibody was performed.
190
6.4. Depletion of intracellular GSH
It is well known that post-translational modifications such as phosphorylation,
glycosylation, acetylation, methylation and glutathionylation can regulate protein
function, including their secretion/release. For instance, acetylation of HMGB1 can
affect its extracellular release (332). Following previous experiments to confirm the
release of glutathionylated Prx2 from Raw 264.7 cells treated with LPS, another aim
of this thesis was to understand whether glutathionylation of Prx2 could have an
effect, either negative or positive, on its release. To this purpose, first of all it was
necessary to set up the experimental conditions in which the concentration of BSO (a
GSH synthesis inhibitor) was able to decrease the GSH levels, but that was not toxic
in combination with LPS. Then Prx2 release was measured by Western blot in
conditions of GSH depletion after LPS treatment (following the same experimental
scheme as in the previous experiments).
The experimental scheme followed for the cell viability assay, GSH assay and
Western blot is shown in Figure 6.2. Briefly, Raw 264.7 cells were plated in
complete RPMI 1640 medium, after ON incubation, the medium was changed with
Opti-MEM I and cells were incubated with BSO (0, 125 µM, 250 µM) for 24 hours
and then stimulated with and without 100 ng/ml LPS for an additional 24 hours. At
the end of this period the cell viability, GSH assay or Western blot were performed
following their respective protocol.
191
Figure 6.2: Experimental scheme of BSO and LPS treatments of Raw 264.7 cells.
192
6.4.1. Cell viability
Raw 264.7 cells were plated in 96-well and cell viability was measured by CTB
assay as described in Methods (2.2.2). Results shown in Figure 6.3 are the mean of
three independent experiments with triplicate samples for each condition. In
conclusion, the results showed that treatments of Raw 264.7 cells with BSO (+/-LPS)
induced 20% cells death compared to control. Similar cell viability was also
observed in the previous set of experiments to confirm Prx2 release. In addition, no
differences were observed between 125 and 250 µM BSO. For this reason all the
experiments were performed with both concentrations.
193
Figure 6.3: Effect of BSO and LPS on Raw 264.7 cell viability. Cell viability was
measured as described in Methods with CTB at 125 µM and 250 µM BSO with and
without 100 ng/ml LPS. Bars represent the mean value ± SD of three independent
experiments, each run in triplicate. The control cell was considered 100% viable and
the cell death was ~ 20%. Control (min-max range: 87-108; mean: 100) LPS (min-
max range: 60-95; mean: 84). BSO 125 µM (min-max range: 77-96; mean: 85). BSO
250 µM (min-max range: 80-95; mean: 87). BSO 125 µM + LPS (min-max range:
70-91; mean: 82). BSO 250 µM + LPS (min-max range: 68-98; mean: 84).
Significance was established by the Tukey-Kramer multiple comparison test.
*p<0.05 for BSO 125 + LPS versus control. Non-significant change in cell viability
was observed by the Tukey Kramer’s test in all other groups
194
6.4.2. GSH assay
In order to ensure the decrease of intracellular GSH depletion by BSO, the GSH
levels were measured following the enzymatic protocol as described in Methods
(2.2.8). Raw 264.7 cells were plated in 6-well plates and preincubated with 125 µM
and 250 µM BSO for 24 hours. After BSO treatment, cells were exposed to LPS for
24 hours following the experimental scheme described above. Figure 6.4 shows the
results of three independent experiments performed in duplicate for each point.
These results were obtained measuring total glutathione (GSHtot) levels, which
include reduced plus oxidized form. Results were expressed as nmol/mg of protein.
GSHtot levels of 2.7 nmol/mg of protein were measured in untreated cells (control).
Treatment of Raw 264.7 cells with both concentrations of BSO (125 µM and
250 µM) resulted in a dramatic decrease of total GSH levels. Lower levels of total
GSH were also observed in cells treated with BSO and LPS.
195
Figure 6.4: Effect of BSO on GSHtot (GSH + GSSG) levels in Raw 264.7 cells. Cells
were seeded at a density of 1 x 106 in 6-well plate, incubated with BSO (125 µM or
250 µM) for 24 hours and then stimulated with LPS for a further 24 hours. Proteins
were precipitated with 6% TCA and supernatants were used in the GSH assay. GSH
levels are expressed as nmol/mg protein. Results are the mean ± SD of three
independent experiments performed in duplicate (n=6 for each group). Control (min-
max range: 1.8-3.6; mean: 2.7). LPS (min-max range: 3.4-7.9; mean: 5). BSO 125
(min-max range: 0.1-0.2; mean: 0.1). BSO 125 + LPS (min-max range: 0.1-0.4;
mean: 0.2). BSO 250 (min-max range: 0.1-0.2; mean: 0.1). BSO 250 + LPS (min-
max range: 0.1-0.5; mean: 0.2). *p<0.05; ***p<0.001 versus control.
196
6.4.3. Detection of glutathionylated Prx2 by Western blot
In order to study the effect of GSH depletion on Prx2 release in conditioned medium
from Raw 264.7 cells, pilot experiments were performed and Prx2 levels were
measured by Western blot. As shown in Figure 6.5 and Figure 6.6, BSO treatments
(250 µM and 125 µM) dramatically decreased the intracellular total GSH with a 20%
cell death (Figure 6.3).
Experiments performed with BSO 250 µM (Figure 6.5) or 125 µM (Figure 6.6) gave
the same results. In particular, the results of both experiments showed higher levels
of released Prx2 in cells treated with LPS compared to control, confirming once
again Prx2 release in LPS-stimulated macrophages. Furthermore, higher levels of
released Prx2 were found in supernatant from cells treated with LPS alone compared
to cells preincubated with BSO before LPS stimulation. In particular, a reduction of
1.5-fold was observed in the sample treated with 125 µM BSO + LPS compared with
LPS alone. A similar result (1.5-fold) was also observed in the sample treated with
250 µM BSO + LPS compared with LPS alone.
197
Figure 6.5: Effect of GSH depletion by BSO 250 µM on Raw 264.7 cells. Released
Prx2 levels decreased in response to BSO treatments as shown in the Western blot
(upper panel) and a corresponding densitometric analysis (lower panel). Bars
represent the mean value for sample treatment as described in the upper panel.
Control, BSO, LPS and BSO + LPS (n=2 per condition). Control (values: 409-618;
mean: 513). BSO 250 µM (values: 426, 430; mean: 428). LPS (values: 750-1158;
mean: 954). BSO250 µM + LPS (values: 610-688; mean: 649).
198
Figure 6.6: Effect of GSH depletion by BSO 125 µM on Raw 264.7 cells. Released
Prx2 levels decreased in response to BSO treatments as shown in the Western blot
(upper panel) and a corresponding densitometric analysis (lower panel). Bars
represent mean values for sample treatment as described in the upper panel. Control
(n=1), BSO (n=2), LPS (n=3) and BSO + LPS (n=3). Control value: 526.
BSO 125 µM (values: 283-472; mean: 378). LPS (min-max range: 589-859;
mean: 751). BSO125 µM + LPS (min-max range: 409-614; mean: 480).
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6.5. Discussion
The glutathionylation of released Prx2 was demonstrated by immunoprecipitation
with an antibody against Prx2 followed by Western using an anti-GSH antibody. The
results obtained from the immunoprecipitation showed a band which correspond to
the dimer form, implying that Prx2 was glutathionylated.
Of note, the glutathionylation form of Prx2 released migrates as a homodimer. In
fact, in order to detect protein glutathionylation, we run the electrophoresis and all
subsequent steps under non-reducing conditions. The Figure 6.7, from Shroeder et al.
(208) summarizes the various possible redox state of Prx2. As shown in the Figure,
Prx2 can exist under a decameric complex made of 5 dimers, where the dimers could
be either in the reduced form and or under different oxidation forms. The fact that we
could detect the dimer under non-reducing conditions (Figure 5.9) but not under
reducing conditions (for instance in Figure 5.4) indicates that the released dimer is
either the disulphides-bonded form (with two disulphide bridges) or the potentially
disulphides-bonded form (with only one disulphide bridges). A dimer can also exist
in the absence of disulphide bridges just by hydrophobic interactions and hydrogen
bonds (333), but it is unlikely that a non-covalently bound dimer would be stable
under the conditions used for our electrophoretic separation that, even if not
involving reducing agents, include the use of SDS and boiling the sample in sample
buffer. In this respect, it should also be noted that when we detect Prx2 as a
monomer, a monomer under denaturing conditions indicates a reduced form, not
necessarily a monomeric state in vivo, where Prx2 probably exist as a decameric
complex.
The fact that LPS induces Prx2 release in the form of a dimer might be explained in
two ways. The first possibility is that LPS induces a transient production of ROS and
this causes a shift towards the disulphide-linked forms of the dimer. This could be
consistent with the fact that LPS augments ROS production by macrophages (334)
(335). However, we cannot exclude that Prx2 is released as a monomer and forms a
dimer once it is in the culture medium that lacks reducing equivalents.
200
Figure 6.7: The different oxidation states of Prx2 and the effect of H2O2 treatment.
Figure from Shroeder et al. (208).
201
The effect of glutathionylation on Prx2 release was further investigated in this study
to understand whether GSH levels can induce a different release of Prx2. Results
showed that BSO (125 or 250 µM) preincubation was able to decrease GSH levels in
Raw 264.7 cells. Furthermore, preliminary results suggested that low levels of GSH
can affect Prx2 release in LPS-stimulated Raw 264.7 cells. In particular, we found
lower levels of Prx2 in cells treated with BSO + LPS compared to LPS treatment
alone. However, the experiment with BSO 125 µM was performed in triplicate and
statistical analysis between LPS and BSO 125 µM + LPS treatment showed that the
difference between the two groups was not statistically significant. This result may
be due to the dual effect of BSO that can reduce GSH levels increasing also oxidative
stress levels. In conclusion, this data suggests that the combination between reduced
intracellular GSH by BSO and higher levels of oxidative stress can produce opposite
effect on the Prx2 release. Further experiments will be needed to confirm the
correlation between GSH and Prx2 release.
6.6. Following chapter
The results obtained in the previous chapters confirmed the release of Prx2 from
LPS-stimulated macrophages, PBMC and human macrophages. The aim of the
following chapter will investigate the role of extracellular Prx2.
202
Chapter 7. Role of extracellular Prx2
203
7.1 Introduction
Several studies have reported the presence of extracellular Prxs in body fluids and
from mammalian cells. For instance, Gharib et al. identified Prx1, Prx5 and Prx6 in
human and mouse bronchoalveolar fluid but obtained contrasting results. In
particular, patients with acute lung injury have higher levels of Prx1 compared to
healthy controls, while in mice lower levels of Prx1 were observed during lung
injury compared with normal controls (336). This could mean that the significance
and mechanism of release of Prx1 is very different in mice and men, but one should
also keep in mind that the mouse models of acute lung injury used in the study by
Gharib et al. may not reproduce exactly the clinical setting. A further limitation of
that study was that 3 mice were studied at one time point. In a different study, Guo et
al. also identified Prxs (Prx1, Prx2, Prx4, Prx5 and Prx6) in bronchoalveolar fluid
from one single healthy mouse (337). In addition, Shichita et al. (316) as discussed in
Chapter 5, showed the release of Prxs with productions of proinflammatory
cytokines in the ischemic brain. Chang et al. demonstrated that transforming growth
factor-beta1 could induce what they defined as “non-classical secretion” of Prx1
from A549 cells (312). Riddell et al. showed secretion of Prx1 from immature bone
marrow-derived dendritic cells (338) and from prostate cancer (339) and provided
evidence that Prx1 stimulates cytokine secretion through a TLR4-dependent
mechanism (338). Taken together, these studies suggest an inflammatory role of
extracellular Prxs, where the possible molecular mechanism is involves the binding
of Prxs to TLRs and consequent cytokines production.
In general there seems to be a marked difference in the intracellular role of Prxs and
their extracellular one. Intracellularly, Prxs are thought to act primarily as antioxidant
enzymes, eliminating hydrogen peroxide. There are several papers showing that
ROS, including hydrogen peroxide, are implicated as mediators of inflammation.
Pioneering work by the group of Baeuerle has shown that ROS activate NF-kB, the
transcription factor that regulates the expression of several inflammatory mediators
and cytokines (140, 340). Along this line, they could also show that thiol
antioxidants such as N-acetyl-cysteine inhibit NF-kB activation (340). Interestingly,
oxidized GSH, GSSG, which can be a thiol oxidant through thiol-disulfide exchange
204
reactions, seems to promote NF-kB activation (341), thus supporting the hypothesis
that an oxidized state of cellular thiols leads to promotion of inflammation.
The role of intracellular Prxs in inflammation seems to agree with this picture.
Suppression of Prx4 results in an augmented NF-kB activation and TNF production
in response to bacterial products (342) and similar results have been obtained
studying the role of Prx5 in LPS-induced IL-6 production (343) and with Prx1 for
LPS-induced activation of NF-kB (344). Intracellular, endogenous Prx2 itself acts as
a negative regulator of inflammation. In fact, macrophages from Prx2 knock out
mice show an upregulated production of inflammatory cytokines, including TNF
(345). This is associated with an increased production of hydrogen peroxide in
response to LPS in Prx2-deficient mice, and ultimately results in an increased
susceptibility to the lethal effect of LPS (345). Interestingly, when recombinant Prx6
is added exogenously but transduced to have it located intracellularly, it behaves as
an inhibitor of NF-kB activation, which is also in agreement with an anti-
inflammatory role of intracellular Prxs (346). Although the published evidence for a
proinflammatory role, in terms of induction of inflammatory cytokines via TLR4
have been obtained with Prx1 (338), it was sensible to investigate whether the same
happened with Prx2.
Prx2 is an intracellular antioxidant enzyme that is different in structure, mechanism
of action and localization compared to other members of its protein family (205)
(204). For instance, Prx2 does not contain a signal peptide for the classical secretory
pathway (Prx4 is the only one of its family with a signal peptide). Prx1 and Prx2 are
not identical proteins, for Prx2 has three cysteines while Prx1 has an additional one
(205). However, according to the previous information on the role of secreted Prx1
and considering that both Prx1 and Prx2 are typical 2-Cys Prxs, it was hypothesized
that there could be a similar extracellular inflammatory role for Prx2.
205
7.2 Aim of Chapter 7
As seen in the previous chapters, Prx2 was identified in the culture supernatant of
LPS-stimulated Raw 264.7 cells and then its release was confirmed not only in
murine macrophages but also in human cells (PBMC and macrophages). This chapter
focuses on the role of the released form of Prx2 in inflammation by investigating its
potential inflammatory role. This is an important aspect of the project and led to a
new set of experiments looking at the effect of addition of recombinant Prx2 to
macrophages call culture and measuring their TNF production. I performed several
experiments were performed using murine macrophages (Raw 264.7 cells) treated
with hrPrx2 (prepared by Dr. Eva Maria Hanschmann in Prof. C.H Lillig’s
laboratory). Cytokine secretion (TNF-α) was measured by ELISA.
.
206
7.3 Peroxidase activity of hrPrx2
The enzyme activity of hrPrx2 was determined by Dr. Eva Maria Hanschmann
measuring consumed NADPH as described in Methods (2.2.9). As shown in Figure
7.1, NADPH concentration diminished in a time-dependent manner in the reaction
using Trx1 while in the control sample (without Prx2), the consumption of NADPH
was not observed. This data demonstrated that the hrPrx2 used for all the
experiments of this thesis was an enzymatically active preparation.
207
Figure 7.1: Peroxidase activity of hrPrx2. Figure by Dr. Eva Maria Hanschmann.
208
7.4 Removal of LPS from hrPrx2 and Western blot
analysis of LPS-free hrPrx2
Human and mouse Prx2 are 93 % similar in their amino acid sequences as shown
comparing the two different species using NCBI Basic Local Alignment Search Tool
(BLAST). On the basis of this homology, we decided to treat Raw 264.7 cells with
human Prx2.
Before performing the experiments investigating the effects of hrPrx2 on cytokine
production, we considered that endotoxin (lipopolysaccharide, LPS) contamination
can lead to an artefact of TNF-α production. This has always been a problem with
recombinant proteins produced in E. coli (a Gram-negative bacterium) and might
then represent a limiting factor for the use of hrPrx2 in the experiments. In order to
circumvent this problem, the recombinant protein was subjected to the following
three steps and then used for all experiments as summarized in Figure 7.2.
Step 1 hrPrx2 was passed through a Detoxi-Gel column (as described in
Methods, 2.2.10) containing PMB to remove a possible LPS contamination.
Step 2 the presence and integrity of the protein contained in the first eluate
fraction by Western blot was checked.
Step 3 (in parallel with step 2), a LAL test was performed to confirm the
absence of LPS contamination from the recombinant protein after Detoxi-Gel
purification. The LAL test is described in Methods (2.2.11).
Three eluate fractions were collected from the Detoxi-Gel column. After loading the
sample (200 µl hrPrx2 at the protein concentration of 9.7 mg/ml) onto column, eluate
fractions were obtained by adding 2 ml PBS for three times and three fractions were
obtained. The protein concentration of hrPrx2 (fraction 1-2-3) was determined using
the colorimetric assay, DC protein assay kit. A standard curve of BSA with 2-fold
serial dilutions from 1.5 mg/ml was used as reference as indicated in Figure 7.3 A.
The protein concentration in the first eluted fraction, that had highest concentration
of hrPrx2 (0.573 mg/ml), was determined compared to the second fraction
(0.129 mg/ml) and to the third fraction (undetectable) (Figure 7.3 B). Only the first
eluted fraction containing the higher concentration of purified proteins was analysed
at two concentrations (230 ng and 115 ng hrPrx2) by Western blot. The
209
concentration 10µg/ml hrPrx2 was used in all experiments to study the TNF-α
induction. Briefly, hrPrx2 was subjected to 12 % SDS-PAGE under reducing
conditions (10% 2-ME) followed by Western blot with anti-Prx2 antibody as
described in Methods section (2.2.6.1). A 22 kDa band was detected after the Detoxi-
Gel column purification (Figure 7.4), confirming the presence and integrity of the
protein. Furthermore, the densitometric analysis showed an intense band with the
high-amount 230 ng hrPrx2, while the signal was decreased by approximately half in
the sample containing 125 ng hrPrx2.
210
Figure 7.2: Scheme for the removal and checking of LPS contamination from hrPrx2
(1) hrPrx2 purification from LPS using PMB column (Detoxi-Gel). (2) The purified
hrPrx2 preparation was analysed by SDS-PAGE followed by Prx2 antibody and anti-
rabbit. (3) The possible LPS contamination of hrPrx2 was also determined using the
LAL test. (4). Cells were incubated with hrPrx2, previously passed through the
Detoxi-Gel column and then TNF-α concentration was measured in supernatant by
ELISA kit.
211
Figure 7.3: Protein determination of hrPrx2 after Detoxi-Gel filtration. (A) Standard
curve obtained using BSA as standard (2-fold serial dilutions from 1.5 mg/ml). (B)
Protein concentration of the three eluted fractions calculated from the standard curve.
The first eluate fraction contains the higher hrPrx2 concentration (0.573 mg/ml).
212
Figure 7.4: Western blot analysis of hrPrx2 after Detoxi-Gel filtration. Upper panel:
Western blot of hrPrx2; Lower panel: Densitometric analysis. The first eluted
fraction (the higher concentration of hrPrx2) obtained after column was diluted at
230 ng and 115 ng and analysed by Western blot. This data shows that the protein
was still present after the passage through the column, and the resulting hrPrx2 was
undamaged.
213
Once the presence and integrity of the protein was demonstrated by Western blot, the
absence of LPS in fraction 1 was verified with the LAL test to be sure that the
Detoxi-Gel column was able to remove the possible LPS contamination. To this
purpose, the Pyrogent Plus kit with a sensitivity of 0.06 EU/ml was used following
the manufacturer’s instructions. Protein and Control Standard Endotoxin (CSE) were
prepared in pyrogen free water as described in Methods (2.2.11). Following the FDA
(Food and Drugs Administration) reference for the standard endotoxin, 1 ng of
endotoxin is equivalent to 5 EU (Limulus lysate assay).
CSE and hrPrx2 were tested at different dilutions; 0.5 EU/ml, 0.125 EU/ml, 0.06
EU/ml and 0.03 EU/ml CSE and hrPrx2 (40 µg/ml, 20 µg/ml, 10 µg/ml and
2.5 µg/ml). RPMI 1640 medium and water were also tested to exclude a possible
contamination of these reagents used for the preparation of solution and cells culture.
All the solutions (CSE, hrPrx2, water and medium) were incubated for 1 hour at
37°C, the time necessary for the clot formation, as described in the Method section.
Results of the LAL test are shown in the Table 7.1. In particular, the clot was not
observed for 2.5 µg/ml and 10 µg/ml hrPrx2, confirming that less than 0.06 EU/ml
was contained in the recombinant proteins and that hrPrx2 was not contaminated by
LPS at least at these low concentrations. In contrast, a positive clot was observed for
20 µg/ml and 40 µg/ml hrPrx2, demonstrating the presence of LPS contamination at
these concentrations. In conclusion, the first elution fraction contained the higher
concentration of hrPrx2 collected from the Detoxi-Gel column (0.573 mg/ml hrPrx2)
and was negative to LAL test. Therefore, this preparation was used at a final
concentration of 10 µg/ml for all the cell treatments described in this chapter.
214
LAL Test Sample
Stable Clot
Endotoxin standard
0.5 EU/ml Yes 0.125 EU/ml Yes
0.06 EU/ml No
0.03 EU/ml No
Samples hrPrx2 (40 µg/ml) Yes
hrPrx2 (20 µg/ml) Yes
hrPrx2 (10 µg/ml) No hrPrx2 (2.5 µg/ml) No
Medium (RPMI 1640) No
Ultrapure water No
Table 7.1: LAL test for endotoxin contamination. Different dilutions of hrPrx2 and
endotoxin standard were prepared in ultrapure water and endotoxin contamination
was analysed by LAL test. Medium and ultrapure water were also tested. After
1 hour of incubation at 37°C the formation (Yes) or absence (No) of a stable clot was
observed in the sample as indicated above.
215
7.5 TNF-α production in Raw 264.7 cells treated with
hrPrx2
The experimental design to investigate the possible inflammatory activity of Prx2 in
mouse macrophages measuring TNF-α is shown in Figure 7.5. Raw 264.7 cells were
seeded in 96-well plates (as described in Methods) and on the following day the cells
were added with 10 µg/ml hrPrx2 (0.5 µM) for 4 hours. Supernatant was collected,
centrifuged and then TNF-α production was measured by ELISA.
All experiments of this chapter including the removal of LPS, protein check by
Western blot and cell treatments for the experiments to measure the TNF-α
production were carried out by the author, except for the ELISA assay, which was
performed by Dr. Manuela Mengozzi (Brighton and Sussex Medical School, UK).
In this chapter, two experiments are shown to study the effect of hrPrx2 on TNF-α
production. Furthermore, two additional controls (Prx2 preincubated with PMB and
boiled hrPrx2) were included to unequivocally rule out the possibility that the TNF-α
production was due to contamination by LPS. LPS is heat-resistant whereas the
proteins are inactivated by boiling.
Raw 264.7 cells were incubated with 10 µg/ml hrPrx2 for 4 hours or hrPrx2
previously boiled for 30 min (Figure 7.6). Results from this experiment showed a
significant increase (2.6-fold) of TNF-α level in hrPrx2-treated cells compared with
untreated (Control) cells. The effect on TNF-α production obtained with 10 µg/ml
hrPrx2 was similar to the effect obtained by stimulating cells with 10 pg/ml LPS.
Furthermore, boiled hrPrx2 was much less effective than hrPrx2. In particular, the
heating reduced the TNF-α production from a 2.6-fold increase observed after
treatment with hrPrx2 to a 1.3-fold increase with boiled hrPrx2.
These conclusions were confirmed in the experiments where we measured TNF
production by cells treated with LPS + hrPrx2 compared to those treated with LPS +
boiled hrPrx2 or LPS alone. In particular, in the sample LPS + boiled hrPrx2, the
boiled hrPrx2 was inactivated by the heating and the value of TNF-α production was
216
similar to the LPS-treatment alone. In contrast, in the sample LPS + hrPrx2 the effect
observed was the sum of these two treatments.
As a further check for LPS contamination, a second experiment was performed
following the same experimental design as the previous one but including a treatment
with the LPS inhibitor PMB. Raw 264.7 cells were treated with 10 µg/ml of a hrPrx2
preparation that was preincubated for 20 min at RT with and without PMB. Cells
were also treated with a higher concentration of LPS (50 pg/ml) preincubated with
and without PMB to confirm the inhibitory effect of PMB on LPS.
The treatment with hrPrx2 increased TNF-α levels and this effect was not reduced
when the Prx2 preparation had been preincubated with PMB (Figure 7.7). This
demonstrates, using a different approach from heat denaturation that the effect was
only due to the protein activity and not to the LPS contaminant. In contrast,
preincubation with PMB reduced markedly the TNF-inducing activity of 50 pg/ml
LPS. This indicates that, if the effect of Prx2 was due to a LPS contamination, we
would have been in the condition to observe an inhibition by PMB.
In conclusion, the results of the two experiments described above suggest that hrPrx2
has an inflammatory role in mouse macrophages, as detected by an increase in TNF-
α production. Furthermore, we also confirmed that Detoxi-Gel column successfully
removed LPS from the recombinant Prx2 preparation, as shown in the experiment
with boiled hrPrx2 and PMB. Therefore, the induced TNF-α production in mouse
macrophages was only due to the effect of the recombinant protein.
217
Figure 7.5: Scheme of the experiment for TNF-α production in Raw 264.7 cells
evaluated by ELISA. Cells were seeded at a density of 3 x 104 in 96-well plates as
described in Methods and incubated with hrPrx2 for 4 hours. Cells were also treated
with boiled hrPrx2 or hrPrx2 preincubated with PMB as an additional control for
protein endotoxin contamination.
218
Figure 7.6: Effect of hrPrx2 on TNF-α production by Raw 264.7 cells using boiled
hrPrx2 as control for Prx2 contamination by endotoxin. Cells were seeded in 96-well
plates, incubated with hrPrx2 or boiled hrPrx2 for 4 hours. The same incubation was
performed with LPS in the media. Data shown are mean values ± SD from an
experiment performed in quadruplicate. Control (min-max range: 212-280; mean:
244). hrPrx2 (min-max range: 559-678; mean: 636). Boiled hrPrx2 (min-max range:
284-407; mean: 328). LPS (min-max range: 546-674; mean: 608). LPS + hrPrx2
(min-max range: 909-1178; mean: 1054). LPS + boiled hrPrx2 (min-max range: 559-
870; mean: 687). *p< 0.05 versus control; ***p<0.001 versus control; # # # p<0.001
versus LPS alone.
219
Figure 7.7: Effect of hrPrx2 on TNF-α production by Raw 264.7 cells using PMB to
investigate endotoxin contamination. As a further check for endotoxin
contamination, cells were preincubated both with and without PMB (inhibitor of
LPS). TNF-α concentration was measured by ELISA. Data shown are mean value ±
SD from an experiment performed in quadruplicate. Control (min-max range: 235-
295; mean: 240). PMB (min-max range: 196-272; mean: 240). hrPrx2 (min-max
range: 444-497; mean: 481). hrPrx2 + PMB (min-max range: 473-647; mean: 552).
LPS (min-max range: 556-879; mean: 682). LPS + PMB (min-max range: 306-384;
mean: 333). **p<0.01 versus control; *** p< 0.001 versus control; # # p<0.01 versus
LPS alone.
220
7.6 TNF-α production in human macrophages
treated with hrPrx2
The effect of hrPrx2 on TNF-α production was also studied in human macrophages
by ELISA. The experiment was performed with four healthy volunteer donors and
primary macrophages were differentiated from PBMC as described in Methods
(2.2.1). Cells were plated in 96-well plates at 0.2 x 106
in 100 µl of complete RPMI
1640 (5% FBS) and incubated with and without 10 µg/ml hrPrx2 following the same
experimental scheme as described above for Raw 264.7 cells.
The results from Donors 1, 2, 3 and 4 (control and hrPrx2-treatment) are shown in
one graph in Figure 7.8. Experimental results showed in human macrophages a
significant difference of TNF-α production between the control (<15 pg/ml) cells and
hrPrx2 treatment for 4 hours. In conclusion, hrPrx2 was able to induce TNF-α
production not only in murine cells (Raw 264.7) but also in human macrophages as
showed below.
221
Figure 7.8: Effect of hrPrx2 on TNF-α production by human macrophages. Human
macrophages from 4 donors were plated in 96-well plates, incubated with or without
hrPrx2 for 4 hours. Data represent mean values ± SD of sample treatment as
described above (n=3 per condition). TNF-α concentration for the Control cells is
< 15 pg/ml. Donor 1: hrPrx2 (min-max range: 290-405; mean: 358). Donor 2: hrPrx2
(min-max range: 360-480; mean: 403). Donor 3: hrPrx2 (min-max range: 761-1041;
mean: 863). Donor 4: hrPrx2 (min-max range: 341-423; mean: 389). ***p< 0.001
versus its control.
222
7.7 Discussion
Prxs not only exist as intracellular proteins but can also be released in the
extracellular environment and can have a different role depending on their
localization. For instance, Prxs can have intracellular antioxidant or chaperone
activity while a new biological role of released Prxs is currently studied. Other
studies reported a possible inflammatory role of extracellular Prxs (316) (312) (338)
(339) (313) and our hypothesis is that Prx2 can be involved in inflammation when it
is present in the extracellular compartment in a similar way as demonstrated for
Prx1. Therefore, to support the hypothesis that Prx2 can induce the production of
proinflammatory cytokines, macrophages (human macrophages and Raw 264.7 cells)
were treated with hrPrx2 and then we measured TNF-α production in supernatant by
ELISA. The results obtained from our experiments showed higher levels of TNF-α
production from cells treated with hrPrx2 compared to untreated cells.
Furthermore, to remove any possible doubt that the cytokine production was
associated with LPS contamination of hrPrx2, two internal controls (boiled hrPrx2
and PMB) were used. The results showed that the low level of TNF-α production
after cells treatment with boiled hrPrx2 was due to inactivation of hrPrx2 by boiling,
whereas LPS that is heat-resistant, was not present in hrPrx2 preparation, and then
was not able to induce cytokine production. To support our results, preincubation of
hrPrx2 with PMB was used to inactivate any possible effect of LPS on cytokine
production. TNF-α levels from cells treated with hrPrx2 preincubated with PMB
were similar to hrPrx2 without preincubation with PMB demonstrating once again
that the effect was only due to the hrPrx2. Clearly, our results indicate that Prx2 has
an inflammatory role inducing secretion of TNF-α in murine and human
macrophages.
We devoted particular attention to rule out the possibility that LPS contamination
might give false results as this happened with various studies in the past. Having
used different and complementary approaches, including the removal of LPS using
PMB-based Detoxi-Gel, the use of boiling (to inactivate the protein but not the
possible LPS contaminant), and the use of PMB (to inactivate the contaminating LPS
223
only, if any was present), I am now confident that the proinflammatory activity of
Prx2 that I have observed is real.
224
Chapter 8. Conclusions and discussion
225
8.1 Summary of thesis results
This chapter is divided into four main parts and contains a summary of results (Table
8.1). In addition, potential future experiments with helpful suggestions on how to
strengthen our data and add further knowledge to the previous results are included.
The first part of the results shows experiments designed to evaluate a possible redox
regulation of HMGB1, a protein well-known to be involved in inflammation. Based
on the negative results obtained in the first part of the experiments, the research on
HMGB1 was not pursued further. The second part of the results was described in
Chapter 4. A new set of experiments focused on the identification by MS of
unknown released proteins that could be involved in inflammation and immunity
were performed. After identification and selection of Prx2 as the protein of interest,
the project was developed to use appropriate methods to confirm Prx2 release in
LPS-activated human and mouse macrophages and extended to in vivo studies
measuring Prx2 levels in serum from mice after LPS injection (Chapter 5). Since
Prx2 is an important enzyme involved in the cells redox regulation, the
glutathionylation of this protein could be important in the modulation of one or more
of its activities. To better understand this hypothesis, the aim of the third part of the
results was to confirm Prx2 glutathionylation as reported in Chapter 6. Until now, the
fact that Prx2 is released by macrophages was unknown, and the role of Prx2 was
studied mainly as an antioxidant enzyme in the context of the intracellular
environment. The presence of released Prx2 from macrophages treated with LPS
suggested a new role of extracellular Prx2 different from its antioxidant and
chaperone activity. For this purpose, the fourth part of the results was focused on the
inflammatory role of Prx2. Results of this set of experiments are reported in the
Chapter 7.
226
AIMS RESULTS
To study a possible redox regulation of
released HMGB1
Negative results
(Chapter 3)
To identify the proteins released from LPS
stimulated macrophages
Prx2 was identified and selected from the
list.
(Chapter 4)
To confirm Prx2 release in human and mouse macrophages stimulated by LPS
Prx2 is released by human and mouse macrophages
(Chapter 5)
To confirm the glutathionylation of released
Prx2
Prx2 released from LPS-stimulated Raw
264.7 cells is glutathionylated (Chapter 6)
To investigate the possible inflammatory role
of Prx2
Prx2 has an inflammatory role inducing
TNF-α production (Chapter 7)
Table 8.1: Aims and results from each chapter of this thesis.
227
The present study used various approaches to test the hypothesis that
glutathionylated proteins may be released during inflammation. The study was
challenging because to date there were no studies on the inflammatory secretome
specifically directed at identifying glutathionylated proteins. Indeed, formation of
mixed disulfides with glutathione or cysteine has been reported in some secreted or
released proteins. However, this was only described after the proteins were identified
as secreted or released. In particular, transthyretin, a secreted protein present in
plasma was identified as the target of glutathionylation by its microeterogeneity
(327) (326) (294). In contrast to Prx2, transthyretin has a N-terminal signal peptide
according to the annotation in the Uniprot database
(http://www.uniprot.org/uniprot/P02766). Another protein undergoing thiol oxidation
is serum albumin, a well-known protein secreted in the classical way and having a
signal peptide. Albumin has also been detected in the cysteinylated form (323). More
relevant to this thesis is the observation that one of the first cytokines to be
described, macrophage migration-inhibitory factor (MIF) also known as
glycosylation-inhibitory factor (GIF) is secreted in the cysteinylated form (324). MIF
does not have a signal peptide but one report shows that its secretion requires the
Golgi-associated vesicular transport factor p115 (347)
228
8.2 Glutathionylation and release of HMGB1
In the present study, we started by investigating the specific hypothesis that HMGB1
might be released as glutathionylated. However, although we could demonstrate that
a recombinant preparation of HMGB1 can be glutathionylated by GSSG in a cell-
free system, we could not find any evidence that this occurs and the protein released
by macrophages. In fact, the literature reports indirect evidence, based on its
susceptibility to the reducing activity of Grx, that HMGB1 can undergo
glutathionylation (as reported in Chapter 1). It was suggested that this might be
important for the nuclear-cytoplasmic transport of HMGB1, which is a prerequisite
for the release of this protein that lacks a signal peptide and therefore does not use
the classical secretory pathway. However, we could not detect glutathionylated
HMGB1 in the macrophage supernatants. Furthermore, HMGB1 was not present
among the proteins identified in the redox proteomics experiments described in
Chapter 3. While we should not expect that the identification of proteins by tryptic
digestion and MS would be equally effective toward all proteins, because this
depends on the susceptibility to trypsin, the generation of peptides that fly well in
MS and other variables. However, it must be said that HMGB1 is one of the most
abundant proteins released by LPS-stimulated Raw 264.7 cells (238) or by necrotic
cells (240). It is still possible, however, that, if HMGB1 is glutathionylated in the
nucleus or cytoplasm, it is then deglutathionylated by Grx during the process of its
secretion/release.
229
8.3 Released Prx2 by LPS-stimulated macrophages
A recent review of Ishii et al. (297) summarizes the importance of Prxs in cancer,
inflammation and immunity. Studies on cancer showed the presence of Prxs in
tumour tissue and serum. For instance, Karihtala et al. found higher levels of Prxs in
tissue from patients with breast cancer (348). In addition, higher levels of Prx were
also found in cell lung cancer (349). These studies are important because Prxs are
antioxidant enzymes and could confer an advantage to cancer cells in terms of
resistance to oxidative stress and radiation toxicity. Prxs could have an important role
as cancer biomarkers and thus be helpful for early detection. Furthermore, there is
also considerable interest in the inflammatory role of Prxs. For instance, Prx1
released form activated cells can induce production of cytokines, suggesting a role in
inflammation and immunity (297). In addition, anti-Prx2 antibodies were found in
serum from patient with systemic vasculitis suggesting that Prx2 can be implicated as
an antigen in inflammation of autoimmune origin (350).
Like HMGB1, Prx2, might be released following LPS stimulation but also by cell
necrosis. In fact, we could detect some toxicity when Raw 264.7 macrophages were
exposed to LPS, although this issue had never been addressed in the studies with
HMGB1 (238). Along this line, it is interesting to note that a marker of cellular
necrosis, LDH, was detected by MS in the supernatants. The release of LDH might
be due to some cellular toxicity. LDH might then have been identified in our redox
proteomics experiments as some studies indicated that it may be susceptible to
glutathionylation (351) (352), S-nitrosylation (353), and other form of disulphide
formation (354).
In this context, it is particularly important to discuss the release of Prx2 in relation to
that of other redox enzymes, Trx in particular. Trx is one of the earliest examples of
a protein with a double action. It was identified by Holmgren as the cofactor for
ribonucleotide reductase (183). About the same period, two different groups were
characterizing the cytokines implicated in the growth of leukemia cells and identified
a factor that, released by T-cell leukemia, augmented the expression of the IL-2
receptor. The factor was named “adult T-cell leukemia-derived factor” (ADF) and
suggested to be a third isoform of IL-1 (355). Yodoi and colleagues identified ADF
230
as the same protein as Trx (356). Subsequently, other studies extended the list of the
cytokine-like activities of Trx. In particular, it was shown to be identical to
eosinophil cytotoxicity-enhancing factor (ECEF) (357) and induces eosinophil
migration (358). Trx is also chemotactic for monocytes, neutrophils and T cells in
vitro and in vivo (359) and, particularly relevant to our findings, it is a costimulus to
TNF production by monocytes (360).
Because Trx and Prx are both secreted, this raises the interesting question of the role
of their enzymatic activity in their proinflammatory activities. At first one may think
that Trx might not be enzymatically active extracellularly as it lacks the complete
“Trx system” which includes TrxR, NADPH and a NADPH-regenerating system. On
the other hand, Angelini et al. have shown that Trx released by dendritic cells can act
as a reducing agent for T lymphocytes (305). To further complicate this issue, Trx, in
the absence of TrxR and NADPH and in an oxidizing environment such as bacterial
periplasm, can act as an oxidant (361). Despite all these studies, the mechanisms of
the cytokine/chemokine-like actions of extracellular Trx remain elusive, although
one target for the redox activity of extracellular Trx has been identified as the TNF
receptor superfamily member 8 (TNFRSF8/CD30) (362). However, studies using
redox-inactive mutants of Trx lacking one active-site Cys have shown that the
eosinophil migration activity of Trx depends on the enzyme activity (358) and most
of the chemotactic action on leukocytes (359) suggesting that, indeed, the enzymatic
activity might be important for the cytokine-like activities of redoxins.
Another analogy with the release of Trx is that both Trx and Prx2 lack a signal
peptide. In the case of Trx, some of it is released as a truncated form (Trx80) (363)
although our Western blot experiment do not suggest the existence of a shorter form
of Prx2 in the released from LPS-stimulated macrophages.
231
8.4 Glutathionylated Prx2
Once I confirmed that Prx2 is released by macrophages stimulated with LPS, in the
second part of the project, I performed additional experiments to further enhance
knowledge of its role in the context of inflammation. In particular, the next step of
the project focused on experiments to confirm firstly the main hypothesis that Prx2
was released as glutathionylated in conditioned medium and secondly to better
understand whether intracellular level of GSH affects Prx2 release.
In our study, we have not addressed the issue whether glutathionylation of Prx2 is
implicated in its release and further studies using mutants lacking specific cysteines
are being planned to investigate this hypothesis. In fact, glutathionylation might
change the physico-chemical properties of Prx2. For instance, addition of a molecule
of GSH, due to its Glutamic acid, moiety, would likely cause an acidic shift in the
isoelectric point of any protein. Furthermore, in contrast to the previous method that
decreased GSH levels by using BSO, in the new set of experiment, we plan to
increase the levels of protein glutathionylation by blocking deglutathionylation with
small interference RNA (siRNA) against Grx.
232
8.5 Inflammatory role of released Prx2
In the last part of this thesis, we studied the potential extracellular role of Prx2 in
inflammation, based on previous knowledge that extracellular enzymes can display
cytokine-like activities, as in the case of Trx (359) (363). The possibility that Prx2 is
released by necrotic cells and act as an inflammatory mediator enlist this protein
among the DAMPS that, like HMGB1, can be released following LPS stimulation or
necrosis. One study suggested that Prxs (338), like HMGB1, induce inflammatory
cytokines via an interaction with TLR4.
Because, as discussed above, Prxs are released along with their substrate, Trx, we
cannot exclude that the enzymatic activity of Prx2 is involved in its cytokine-
stimulating action. Future studies will be required to investigate whether hrPrx2 act
as an oxidizing or a reducing agent towards macrophages and whether its actions are
mediated by an interaction with Trx, whether the released one or the membrane-
associated form that has been described in the past (364). In this respect, however, it
is important to note that the hrPrx2 preparation that I have obtained from my
collaborators in Germany is enzymatically active (as described in Chapter 7 by the
experiment on peroxidase activity measured by Dr. Eva Maria Hanschmann).
To better understand the inflammatory role of Prx2, future studies could examine the
secretion of other proinflammatory cytokines (e.g. IL-1 and IL-6) from macrophages
treated with hrPrx2. Experiments could be extended from in vitro to animal models,
for instance proinflammatory cytokines could be measured in serum from hrPrx2-
treated mice or Prx2 could be injected locally to evaluate possible inflammatory
infiltration, as it was done with Trx (359). Furthermore, another important part of
future research could investigate whether Prx2 levels in patients with inflammatory
disease correlate with the severity of the disease. Other important experiments could
be performed to study Prx2 release from macrophages stimulated with different TLR
ligands. For instance, Poly I:C (TLR3L), R848 (TLR7/8L), and PAM3 Cys (TLR2L)
(365).
233
8.6 Prx2 in the context of danger signals and
oxidative stress
My findings add one new piece of knowledge to the mechanism by which
inflammation can be triggered be endogenous molecules. The main finding in this
context has been the fact that Prx2, released as a glutathionylated protein, acts as a
proinflammatory danger signal. The present thesis’ work has focused on its release
induced by LPS, but clearly cell necrosis may also induce release of significant
amounts of Prx2. In fact, Prxs are abundant proteins and Prx2, in cell lines, can
amount to 0.33% of the total protein content (366). This puts Prx2 in the same class
as HMGB1 whose role as an inflammatory mediator was originally discovered
following the finding that it is released in response to LPS (238) but that was
subsequently reported to account for the inflammatory response induced by cell
necrosis (240). Like HMGB1, Prx2 mRNA is not induced by LPS. This seems not to
be the case for all Prxs and for all conditions. The study by Shichita et al. (316)
reports that while Prxs present in cell lysates have inflammatory activity in that they
act as IL-23 inducers, their expression is increased in the model of cerebral ischemia
used in that study. The response to LPS seems to be different, probably because it is
not associated with tissue injury, unlike cerebral ischemia. In fact, a previous study
analysed by Western blot the intracellular expression of various Prxs in LPS-treated
Raw 264.7 cells and Prx5 was the only one whose intracellular levels where
increased by LPS, while Prx2 was unaffected (343). In fact, we noted, at least in
some experiments, that the extracellular release of Prx2 induced by LPS is associated
with a decrease in the intracellular form. Thus, releasing Prx2 may not only result in
the release of an inflammatory danger signal but could also lead to a decrease of the
intracellular breakdown of hydrogen peroxide. This may in turn contribute to ROS-
mediated LPS signalling according to the floodgate hypothesis by which Prxs
regulate signalling by hydrogen peroxide (367, 368). It seems reasonable to
hypothesize that a loss of intracellular Prx may contribute to the increased hydrogen
peroxide levels observed in macrophages treated with LPS.
Another important feature of the release of Prx2 is that it is associated with cysteine
oxidation to form a glutathionylated dimer. It should be noted that these two
modifications might not necessarily interest the same percentage of the secreted
234
Prx2. The experimental approach used to verify whether the secreted protein is
glutathionylated (immunoprecipitation followed by Western blot) is not quantitative
and does not provide information as to how much of the secreted Prx2 is
glutathionylated. This will require other experiments using different approaches (for
instance the ratio between glutathionylated vs. non-glutathionylated peptides by
Mass Spectrometry). On the other hand, the almost exclusive existence of
extracellular Prx2 as a dimer is demonstrated by the Western blot experiments
carried out using gel electrophoresis under non-reducing conditions. It is therefore
safe to conclude that Prx2 secretion is associated with cysteine oxidation to form a
dimer and, at least to a certain extent, a glutathionylated dimer.
This is somewhat similar to what observed with HMGB1. Urbonaviciute at al. (369)
reported that the HMGB1 released upon cellular necrosis is oxidized on at least one
of its cysteine residues. Although the technique used does not distinguish between
the possible forms of oxidation (whether glutathionylation or another for of mixed
disulfide, or by an intramolecular disulfide), and dimeric forms of HMGB1 have
never been reported, the techniques used in that study might be applied in the future
to the study of released Prx2. Another paper reported that HMGB1 could undergo
glutathionylation (57). HMGB1 is probably released though a more complex
mechanism than Prx2. As a nuclear protein, HMGB1 needs to be first mobilized
from the nucleus to the cytoplasm and then released. While no connection has been
made between HMGB1 glutathionylation and the release of HMGB1, a triple
cysteine mutation shows impaired nuclear retention leaving the possibility open for a
role of cysteine oxidation in its release. It should also be noted that, although we
have not been able to demonstrate whether HMGB1 is secreted under the
glutathionylated form, this couldn’t be absolutely excluded.
In the future, it will be important to investigate whether cysteine oxidation,
glutathionylation and/or disulfide-linked dimerization) are important in its LPS-
induced secretion. For instance, members of the transforming-growth factor beta
superfamily form disulfide-linked dimers before secretion (370). However, these
proteins are secreted by the classical pathway to that their secretion does not help in
formulating hypotheses on the mechanism of secretion of Prx2. On the other hand, in
any case, the release due to cell necrosis would not have any special requirement as,
235
being Prx2 cytosolic, it will be released with the cell lysis irrespectively of the redox
state.
A second aspect is whether cysteine oxidation is important in the proinflammatory
activity of Prx2 in terms of cytokine production. For instance, it was previously
reported that the chemotactic activity of Trx depends on its catalytic cysteines and
mutation of those cysteines lead to a non-chemotactic form of Trx (359). I did not
have available Prx2 mutants to test this possibility. However, the proinflammatory
activity of the related Prx1 depends, at least in part on the presence of Cys83 as the
serine mutant of Cys83 was a much less effective cytokine inducer, while mutation
of Cys52 did not have an effect (338). This may not be generalized because the study
by Shichita showed that mutation of various cysteines did not impair the IL-23-
inducing activity of Prx5. Furthermore, to follow on the analogy with HMGB1, the
proinflammatory action of HMGB1 seems to depend on its redox properties because
mutation of specific cysteines reduces its ability to activate NF-kB and induce TNF
production (262). To address the question of the role of the oxidation state of Prx2 in
its proinflammatory activity the existing literature suggests the approach of using of
cysteine mutants as well as treatment of the recombinant protein with reducing
agents or with alkylating agents to block its enzyme activity.
Finally, it will be important to investigate whether oxidants, by promoting Prx2
dimerization and/or glutathionylation, induce the release of Prx2. In fact, formation
of the disulfide-linked dimer is promoted by oxidants (371). If increased
dimerization resulted in an increased release, that might be particularly important as
a mechanism that contributes to the inflammatory response observed in diseases,
such as ischemia/reperfusion injury, associated with oxidative stress but in the
absence of other inducers of the inflammatory cascade such as infection or
autoimmunity.
236
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