Oxidative tissue injury in experimental disease

Oxidative tissue injury in experimental disease models
of multiple sclerosis
Doctoral thesis at the Medical University of Vienna
for obtaining the academic degree
Doctor of Philosophy
Submitted by
Mag. Cornelia Schuh
Supervisor:
O. Univ. Prof. Dr. Hans Lassmann
Center for Brain Research, Spitalgasse 4, 1090 Vienna
Vienna, 02/2014
ii
Declaration
Hereby, I declare that this submitted thesis is my own work for obtaining the academic degree
Doctor of Philosophy from the Medical University of Vienna and has not been previously
submitted by me at any other University.
The experimental part of the thesis was performed at the Department of Neuroimmunology at
the Centre for Brain Research of the Medical University of Vienna. My work comprised
immunohistochemical analysis and quantification of MS lesions and different animal models of
MS including material collected in our lab during the last decades. I conducted the analysis of
microglia activation patterns, oxidative damage and iron accumulation. I performed T cell
cultures and the induction of passive transfer EAE with the support of Monika Bradl. The tissues
derived from other animal models originate from collaborative studies. The induction of MOGEAE in DA rats was performed in collaboration with Maria Storch. Tissues from C57BL/6 mouse
MOG-EAE were derived from the group of Lesley Probert. The coronavirus-injections were
conducted together with Helmut Wege. The mouse MOG-EAE material originates from the group
of Lesley Probert. The CD8+ T cell transfer was conducted in the lab of Roland Liblau. The LPSinjections were done by the group of Ken Smith. The Theiler´s virus injections were studied in
cooperation with Claudia Lucchinetti. The cuprizone experiments were performed in the lab of
Anne-Marie Van Dam. For all the mentioned models, original histopathological analysis was
performed in close collaboration with Hans Lassmann. The original papers describing the
respective animal models are cited correspondingly in the thesis. The antibody recognising
oxidised phospholipids was provided by Christoph Binder. Barbara Scheiber-Mojdehkar shared
the ferrocene protocol with us. Jan Bauer supported my work on frozen material. I performed the
experiments on myelinating spinal cord cultures at the University of Glasgow in the group of
Chris Linington. My stay in Glasgow was enabled by a MSIF Du Pré grant.
Hans Lassmann provided his expertise in neuropathology on human and animal material and
supervised this thesis study.
iii
Table of contents
DECLARATION ........................................................................................................................................................ ii
TABLE OF CONTENTS ..............................................................................................................................................iii
LIST OF FIGURES .................................................................................................................................................... vii
LIST OF TABLES ..................................................................................................................................................... viii
ABSTRACT .............................................................................................................................................................. ix
ZUSAMMENFASSUNG ............................................................................................................................................ xi
PUBLICATIONS ARISING FROM THIS THESIS ......................................................................................................... xiii
ABBREVIATIONS ................................................................................................................................................... xiv
ACKNOWLEDGEMENTS ......................................................................................................................................... xv
1
INTRODUCTION.............................................................................................................................................. 1
1.1
HALLMARKS OF MS ........................................................................................................................................... 1
1.2
CLINICAL COURSE ............................................................................................................................................... 1
1.3
MECHANISMS OF TISSUE INJURY IN MS .................................................................................................................. 3
1.3.1
Oxidative stress and mitochondrial injury ................................................................................................ 4
1.3.2
Footprints of oxidative damage in MS ..................................................................................................... 7
1.3.3
Iron in MS ............................................................................................................................................... 10
1.4
OXIDATIVE BURST, OXIDATIVE DAMAGE AND IRON ACCUMULATION IRON IN MS........................................................... 14
1.5
EXPERIMENTAL MODELS ................................................................................................................................... 16
1.5.1
Animal models used in this thesis ........................................................................................................... 17
1.5.1.1
Passive transfer of MBP-specific T cells in Lewis rats ..................................................................................... 17
1.5.1.2
Active immunisation with MOG/CFA in DA rats ............................................................................................. 18
1.5.1.3
Active immunisation with MOG35-55 in C57/BL6 mice .................................................................................... 18
1.5.1.4
Passive transfer of hemagglutinin (HA)-specific CD8 T cells in HA-transgenic BALB/c mice ......................... 19
1.5.1.5
Intraspinal LPS-injection in SD rats ................................................................................................................. 19
1.5.1.6
Cuprizone diet-induced demyelination in C57BL/6 mice ............................................................................... 19
1.5.1.7
Mouse hepatitis virus (MHV, JHM strain)-induced demyelinating disease in Lewis rats................................ 20
1.5.1.8
Theiler´s murine encephalomyelitis virus (TMEV, DA strain)-mediated demyelinating disease in SJL mice ..22
+
1.5.2
Oxidative injury in animal models .......................................................................................................... 22
1.5.3
Outlook to MS therapy ........................................................................................................................... 23
iv
1.6
2
3
4
AIMS OF THIS THESIS ........................................................................................................................................ 24
RESULTS ....................................................................................................................................................... 25
2.1
IRON ACCUMULATION IN THE RAT CNS DURING AGEING .......................................................................................... 25
2.2
ACUTE CD4 T CELL-MEDIATED EAE IN AGED LEWIS RATS ....................................................................................... 28
2.3
CHRONIC RELAPSING MOG-INDUCED EAE IN DA RATS ........................................................................................... 37
2.4
CHRONIC RELAPSING MOG-INDUCED EAE IN C57BL/6 MICE .................................................................................. 39
2.5
INFLAMMATORY DEMYELINATION INDUCED BY CYTOTOXIC T CELLS............................................................................. 40
2.6
INNATE IMMUNITY-DRIVEN INFLAMMATORY DEMYELINATING LESIONS........................................................................ 42
2.7
TOXIC CUPRIZONE-INDUCED DEMYELINATION ........................................................................................................ 43
2.8
MHV-JHM CORONAVIRUS-INDUCED ENCEPHALOMYELITIS ...................................................................................... 44
2.9
THEILER'S MURINE ENCEPHALOMYELITIS VIRUS (TMEV)-INDUCED DISEASE................................................................ 49
2.10
IN VITRO OXIDATION ......................................................................................................................................... 57
2.11
IRON LOADING OF GLIAL CELL CULTURES ............................................................................................................... 59
+
DISCUSSION ................................................................................................................................................. 71
3.1
GENERAL DISCUSSION ....................................................................................................................................... 71
3.2
CONCLUSION AND FUTURE PROSPECTS ................................................................................................................. 79
MATERIALS .................................................................................................................................................. 80
4.1
MATERIALS FOR HISTOLOGY ............................................................................................................................... 80
4.1.1
Buffers .................................................................................................................................................... 80
4.1.1.1
Boric acid, 50 mM ........................................................................................................................................... 80
4.1.1.2
Citrate buffer 10x stock pH 6.0, 10 mM ......................................................................................................... 80
4.1.1.3
Dako buffer ..................................................................................................................................................... 80
4.1.1.4
EDTA buffer 20x stock pH 8.5 ......................................................................................................................... 80
4.1.1.5
PBS buffer 4x stock ......................................................................................................................................... 80
4.1.1.6
Sodium acetate buffer pH 4.9, 0.05 M ........................................................................................................... 80
4.1.1.7
Sodium potassium phosphate buffer pH 7.4, 5 mM ....................................................................................... 81
4.1.1.8
Sörensen buffer pH 7.4, 0.2 M........................................................................................................................ 81
4.1.1.9
TBS 20x stock, 1 M .......................................................................................................................................... 81
4.1.1.10
TBS/CaCl2 (2 mM) ........................................................................................................................................... 81
4.1.1.11
Tris/HCl buffer pH 8.5, 0.1 M .......................................................................................................................... 81
4.1.2
Stock solutions and developing agents .................................................................................................. 81
4.1.2.1
Amino ethylcarbazole reagent (AEC) developing agent ................................................................................. 81
4.1.2.2
CSA stock solution .......................................................................................................................................... 82
4.1.2.3
DAB developing agent .................................................................................................................................... 82
4.1.2.4
DTPA 100x stock, 5 mM .................................................................................................................................. 82
4.1.2.5
Eosin solution ................................................................................................................................................. 82
v
4.1.2.6
Fast blue developing agent ............................................................................................................................. 82
4.1.2.7
Geltol mounting medium ............................................................................................................................... 83
4.1.2.8
HCl/ethanol .................................................................................................................................................... 83
4.1.2.9
Iron detection reagent (Ferrocene assay) ...................................................................................................... 83
4.1.2.10
Luxol fast blue (LFB) and periodic acid Schiff (PAS) staining solutions ........................................................... 83
4.1.2.11
Methanol/H2O2 ............................................................................................................................................... 83
4.1.2.12
Proteinase K solution ...................................................................................................................................... 83
4.1.2.13
Scott’s solution ............................................................................................................................................... 84
4.2
MEDIA FOR CELL CULTURES................................................................................................................................ 84
4.2.1
4.2.1.1
Restimulation medium for T cells ................................................................................................................... 84
4.2.1.2
TCGF medium for T cells ................................................................................................................................. 84
4.2.1.3
Freezing medium for T cells ............................................................................................................................ 84
4.2.2
5
Media for T cells ..................................................................................................................................... 84
Media for glial cells ................................................................................................................................ 85
4.2.2.1
Mixed glia medium for microglia isolation ..................................................................................................... 85
4.2.2.2
Mixed glia medium for oligodendrocyte isolation .......................................................................................... 85
4.2.2.3
Digestion medium .......................................................................................................................................... 85
4.2.2.4
Sato medium .................................................................................................................................................. 85
4.2.2.5
Neurosphere medium (NSM) ......................................................................................................................... 86
4.2.2.6
Hormone mix 10x ........................................................................................................................................... 86
4.2.2.7
Astrocyte medium .......................................................................................................................................... 86
4.2.2.8
SD solution (soybean trypsin inhibitor) .......................................................................................................... 86
4.2.2.9
Plating medium .............................................................................................................................................. 87
4.2.2.10
Differentiation medium (DM) ......................................................................................................................... 87
METHODS .................................................................................................................................................... 87
5.1
MS PATIENTS.................................................................................................................................................. 87
5.2
EXPERIMENTAL MODELS .................................................................................................................................... 89
5.2.1
+
CD4 T cell-mediated EAE in Lewis rats .................................................................................................. 89
5.2.1.1
Immunisation with MBP/CFA and T cell line preparation............................................................................... 89
5.2.1.2
Induction of MBP-specific T cell EAE in Lewis rats .......................................................................................... 90
5.2.2
Chronic relapsing MOG EAE in DA rats ................................................................................................... 90
5.2.3
Chronic relapsing MOG EAE in C57BL/6 mice ......................................................................................... 91
5.2.4
Acute CD8 T cell-mediated EAE ............................................................................................................. 91
5.2.5
LPS injection-induced inflammation ....................................................................................................... 91
5.2.6
Curpizone-induced demyelination .......................................................................................................... 92
5.2.7
Mouse hepatitis virus strain JHM (MHV-JHM) coronavirus-mediated demyelinating disease............... 92
5.2.8
Theiler's Murine Encephalomyelitis Virus (TMEV)-mediated demyelinating disease ............................. 93
5.2.9
Control animals ...................................................................................................................................... 93
+
vi
5.3
HISTOLOGY..................................................................................................................................................... 93
5.3.1
5.3.1.1
General staining protocol ............................................................................................................................... 94
5.3.1.2
E06 staining for oxidised phospholipids ......................................................................................................... 96
5.3.1.3
Double stainings ............................................................................................................................................. 96
5.3.1.4
Quantification of immunohistochemistry ...................................................................................................... 97
5.3.1.5
Pre-absorption of p22phox antibody ............................................................................................................. 98
5.3.2
Histochemistry ........................................................................................................................................ 98
5.3.2.1
Haematoxylin and eosin staining (H&E) ......................................................................................................... 98
5.3.2.2
Luxol fast blue myelin stain ............................................................................................................................ 98
5.3.2.3
Histochemistry for non-heme tissue iron ....................................................................................................... 99
5.3.3
In vitro oxidation of native fresh frozen tissue sections ......................................................................... 99
5.3.4
Ferrocene assay for iron quantification ................................................................................................ 100
5.4
GLIAL CELL CULTURE TECHNIQUES .................................................................................................................... 101
5.4.1
Mixed glia cultures ............................................................................................................................... 101
5.4.2
Microglia cultures ................................................................................................................................. 101
5.4.3
Oligodendrocyte cultures ..................................................................................................................... 101
5.4.4
Myelinating spinal cord cultures .......................................................................................................... 102
5.4.4.1
Neurosphere-derived astrocytes .................................................................................................................. 103
5.4.4.2
Dissociated spinal cord cultures ................................................................................................................... 103
5.4.5
Iron loading of cells .............................................................................................................................. 104
5.4.6
Immunofluorescence of cell cultures .................................................................................................... 104
5.5
6
Immunohistochemistry ........................................................................................................................... 94
STATISTICAL ANALYSIS..................................................................................................................................... 105
REFERENCES ................................................................................................................................................106
CURRICULUM VITAE ............................................................................................................................................132
vii
List of figures
FIGURE 1: PATHOLOGICAL HALLMARKS OF MS RELATED WITH THE CHANGE FROM RRMS (PINK) TO SPMS (GREEN) ............................. 3
FIGURE 2: SUMMARY OF REACTIVE OXYGEN AND NITROGEN SPECIES DETOXIFICATION ....................................................................... 5
FIGURE 3: THE RESPIRATORY CHAIN OF MITOCHONDRIA .............................................................................................................. 7
FIGURE 4: THE PRODUCTION OF ROS AND SUBSEQUENT LIPID PEROXIDATION ................................................................................. 8
FIGURE 5: SUMMARY OF HOW ROS CAN CONTRIBUTE TO MS PATHOGENESIS............................................................................... 10
FIGURE 6: PROPOSED MECHANISMS OF IRON TRANSPORT IN THE BRAIN ....................................................................................... 13
FIGURE 7: THE EXPRESSION OF P22PHOX AND INOS AND THE ACCUMULATION OF OXIDISED PHOSPHOLIPIDS (E06) AND IRON IN MS
LESIONS .................................................................................................................................................................. 15
FIGURE 8: SCHEMATIC SUMMARY OF MECHANISMS OF TISSUE INJURY IN ACTIVE MS LESIONS. .......................................................... 24
FIGURE 9: IRON ACCUMULATING HOTSPOTS IN THE AGED LEWIS RAT. .......................................................................................... 26
FIGURE 10: IRON HISTOCHEMISTRY AND BIOCHEMICAL IRON QUANTIFICATION IN THE RAT SPINAL CORD. ............................................ 26
FIGURE 11: HISTOCHEMICAL IRON QUANTIFICATION IN THE RAT BASAL GANGLIA............................................................................ 27
FIGURE 12: CORRELATION OF TWO DIFFERENT METHODS OF IRON QUANTIFICATION. ...................................................................... 28
+
FIGURE 13: WEIGHT LOSS AND EAE DISEASE STAGES IN ACUTE CD4 T CELL TRANSFER EAE IN YOUNG AND AGED LEWIS RATS. ............. 29
FIGURE 14: IMMUNOHISTOCHEMISTRY OF MARKERS FOR INFLAMMATION AND NEURODEGENERATION IN THE SPINAL CORDS OF YOUNG AND
+
OLD LEWIS RATS SUFFERING FROM ACUTE CD4
MBP-SPECIFIC T CELL TRANSFER EAE. ......................................................... 30
FIGURE 15: QUANTIFICATION OF STAININGS FOR INFLAMMATORY AND NEURODEGENERATIVE MARKERS IN THE LUMBAR SPINAL CORDS OF
+
YOUNG AND OLD LEWIS RATS SUFFERING FROM ACUTE CD4
MBP-SPECIFIC T CELL TRANSFER EAE. ........................................ 31
+
FIGURE 16: OXIDATIVE BURST AND OXIDATIVE INJURY IN ACUTE CD4 T CELL TRANSFER EAE IN LEWIS RATS. ..................................... 32
+
FIGURE 17: THE EXPRESSION OF P22PHOX AND INOS IN ACUTE CD4 T CELL-MEDIATED EAE IN YOUNG AND OLD ANIMALS. ................ 33
+
FIGURE 18: IRON DEPOSITION IN CD4 MBP-SPECIFIC T CELL TRANSFER EAE IN YOUNG AND OLD LEWIS RATS. .................................. 34
+
FIGURE 19: INFLAMMATION AND NEURODEGENERATION IN IRON-ACCUMULATING HOTSPOTS IN CD4 T CELL TRANSFER EAE. .............. 35
FIGURE 20: OPTICAL DENSITOMETRY OF TBB STAININGS OF IRON ACCUMULATING HOTSPOTS IN OLD EAE AND CONTROL ANIMALS......... 36
FIGURE 21: OXIDATIVE BURST, OXIDATIVE INJURY AND IRON ACCUMULATION IN CHRONIC RELAPSING EAE OF MOG-IMMUNISED DA RATS.
............................................................................................................................................................................. 38
FIGURE 22: OXIDATIVE BURST, OXIDATIVE INJURY AND IRON ACCUMULATION IN CHRONIC RELAPSING EAE OF MOG-IMMUNISED C57BL/6
MICE. ..................................................................................................................................................................... 40
+
FIGURE 23: OXIDATIVE BURST, OXIDATIVE INJURY AND IRON ACCUMULATION IN CD8 T CELL-MEDIATED EAE IN MICE. ........................ 41
FIGURE 24: OXIDATIVE BURST, OXIDATIVE INJURY AND IRON ACCUMULATION INDUCED BY LPS INJECTION INTO THE SPINAL CORD DORSAL
COLUMN OF SD RATS. ............................................................................................................................................... 43
FIGURE 25: OXIDATIVE BURST, OXIDATIVE INJURY AND IRON ACCUMULATION IN CUPRIZONE DIET-INDUCED ACTIVE DEMYELINATION IN MICE.
............................................................................................................................................................................. 44
viii
FIGURE 26: OXIDATIVE BURST, OXIDATIVE INJURY AND IRON ACCUMULATION IN MHV-JHM CORONAVIRUS-INDUCED ENCEPHALOMYELITIS
IN LEWIS RATS.......................................................................................................................................................... 47
FIGURE 27: OXIDATIVE BURST, OXIDATIVE INJURY AND IRON ACCUMULATION IN TMEV-INFECTED MICE 3 DPI. ................................... 50
FIGURE 28: OXIDATIVE BURST, OXIDATIVE INJURY AND IRON ACCUMULATION IN TMEV-INFECTED MICE 45 DPI. ................................. 51
FIGURE 29: QUANTIFICATION OF MICROGLIAL NODULES IN TMEV-INDUCED INFLAMMATORY DEMYELINATING DISEASE. ....................... 52
FIGURE 30: OXIDATIVE BURST, OXIDATIVE INJURY AND IRON ACCUMULATION IN TMEV-INFECTED MICE 90 DPI. ................................. 53
FIGURE 31: OPTICAL DENSITOMETRY OF E06 IMMUNOREACTIVITY IN MEDULLARY LESIONS OF TMEV-INFECTED MICE AT DIFFERENT DISEASE
STAGES. .................................................................................................................................................................. 54
FIGURE 32: OXIDATIVE BURST, OXIDATIVE INJURY AND IRON ACCUMULATION IN TMEV-INFECTED MICE 180 DPI. ............................... 55
FIGURE 33: OXIDATIVE BURST, OXIDATIVE INJURY AND IRON ACCUMULATION IN TMEV-INFECTED MICE 220 DPI. ............................... 56
FIGURE 34: E06 STAINING AFTER IN VITRO OXIDATION. ............................................................................................................ 57
FIGURE 35: 8OHDG STAINING AFTER IN VITRO OXIDATION........................................................................................................ 58
FIGURE 36: IRON LOADING OF PURIFIED OLIGODENDROCYTE CULTURES........................................................................................ 59
FIGURE 37: CHARACTERISTICS OF MYELINATING SPINAL CORD CULTURES ...................................................................................... 60
FIGURE 38: DOSAGE-EFFECT OF IRON CHLORIDE ON THE INTEGRITY OF MYELINATING SPINAL CORD CULTURES. .................................... 61
FIGURE 39: IRON ACCUMULATION IN MYELINATING SPINAL CORD CULTURES AFTER IRON CHLORIDE TREATMENT. ................................. 62
FIGURE 40: IRON ACCUMULATION IN MICROGLIA IN MYELINATING SPINAL CORD CULTURES AFTER IRON CHLORIDE TREATMENT. .............. 63
FIGURE 41: IRON ACCUMULATION IN MICROGLIA CELLS OF MYELINATING SPINAL CORD CULTURES AFTER SHORT-TERM FERRITIN TREATMENT.
............................................................................................................................................................................. 64
FIGURE 42: OLIGODENDROCYTE AND MYELIN INTEGRITY AND FERRITIN ACCUMULATION AFTER SHORT-TERM FERRITIN TREATMENT. ........ 65
FIGURE 43: IRON ACCUMULATION IN MICROGLIA OF MYELINATING SPINAL CORD CULTURES AFTER LONG-TERM FERRITIN TREATMENT. ..... 66
FIGURE 44: OLIGODENDROCYTE AND MYELIN INTEGRITY AND FERRITIN ACCUMULATION AFTER LONG-TERM FERRITIN TREATMENT........... 67
FIGURE 45: LOSS OF MICROGLIA UPON IRON CHLORIDE TREATMENT. ........................................................................................... 68
FIGURE 46: MICROGLIA NUMBERS IN DIFFERENT EXPERIMENTAL SETUPS OF FERRITIN TREATMENT. ................................................... 69
FIGURE 47: IRON ACCUMULATION IN IRON CHLORIDE- AND FERRITIN-TREATED MICROGLIA CULTURES. ............................................... 70
List of tables
TABLE 1: QUANTIFICATION OF MARKERS FOR INFLAMMATION AND OXIDATIVE INJURY IN DIFFERENT EXPERIMENTAL MODELS FOR CNS
INFLAMMATION AND MS ........................................................................................................................................... 48
TABLE 2: CLINICAL DATA OF MS CASES AND CONTROLS OF THE STUDY COHORT. ............................................................................ 88
TABLE 3: PRIMARY ANTIBODIES AND ANTIGEN RETRIEVAL PROCEDURES FOR IMMUNOHISTOCHEMISTRY .............................................. 95
TABLE 4: SECONDARY ANTIBODIES FOR IMMUNOHISTOCHEMISTRY.............................................................................................. 95
TABLE 5: PRIMARY ANTIBODIES FOR IMMUNOFLUORESCENCE STAININGS OF CELL CULTURES........................................................... 105
ix
Abstract
Multiple sclerosis (MS) is a chronic disease of the central nervous system (CNS)
characterised by inflammation and demyelination. Major advances have been made in
unravelling the mechanisms of inflammatory and neurodegenerative processes underlying the
disease. A number of different animal models of MS contributed not only to a better
understanding of MS pathology but also explained fundamental immunological concepts.
Additionally, experimental models enabled the development of immune-modulatory therapies for
MS patients.
In recent years, growing evidence for a major role of oxidative injury in demyelination and
neurodegeneration in MS has been emerging. Hence, we aimed to characterise the nature and
the extent of oxidative damage in different experimental models in comparison to MS. For this
purpose, we emphasised the expression of enzymes involved in reactive species production (the
essential NADPH oxidase subunit p22phox and inducible nitric oxide synthase; iNOS), the
accumulation of oxidised phospholipids and iron, which is a potential amplifier of oxidative
damage. The different animal models were triggered by diverse inflammatory mechanisms, each
representing distinct aspects of MS pathology.
The only experimental model accumulating oxidised phospholipids to a comparable
extent as MS lesions was the coronavirus-triggered demyelinating encephalomyelitis. In MS as
well as in the coronavirus-model, oxidative injury was associated with a profound activation of
microglia and macrophages. This was characterised by the pronounced expression of p22phox
but only scattered expression of iNOS. The second virus model in our study, Theiler´s murine
encephalomyelitis virus-induced chronic demyelination, did not reach the level of microglia and
macrophage activation, regarding p22phox expression, observed in the coronavirus-model.
Further, a staining for oxidised phospholipids was detectable but only to a lesser extent
compared with the coronavirus-induced disease. In both virus models, iNOS expression was
minor. In contrast, animals suffering from passive T cell transfer experimental autoimmune
encephalomyelitis (EAE) exhibited profound p22phox and iNOS expression at the peak of the
disease. Similarly, we found p22phox-expressing macrophages in animals suffering from chronic
T cell and macrophage-induced disease, but few iNOS+ cells. In contrast, chronic lesions of
x
animals affected by antibody-mediated demyelination contained phagocytosing macrophages
lacking p22phox and iNOS expression. Further, acute lesions induced by CD8+ T cell transfer or
by mechanisms of the innate immune system showed p22phox expression in activated microglia
and macrophages, but iNOS reactivity was marginal. Additionally to oxidative damage and
activation patterns in microglia and macrophages concerning iNOS and p22phox, we assessed
the impact of iron in experimental demyelination. Age-dependent iron accumulation and
subsequent lesion-associated cellular iron release, as observed in the human brain, was only
poorly reflected in the central nervous system of rodents. Therefore, we used a complex cell
culture-based approach to study a possible amplification of neurodegeneration by iron
accumulation in oligodendrocytes. Unfortunately, we were only able to induce iron accumulation
in microglia but not in any other cell population.
The presented study reports a diverging extent of oxidative injury and underlying
mechanisms in different models for inflammatory demyelination. We conclude that rodent
models seem to lack amplification processes that have been suggested to play a role in MS
pathogenesis. Therefore, established experimental models appear to reflect the aspect of
oxidative injury in the human disease only to a minor extent.
xi
Zusammenfassung
Multiple Sklerose (MS) ist eine chronische durch Demyelinisierung gekennzeichnete
entzündliche Erkrankung des zentralen Nervensystems (ZNS). Große Fortschritte konnten in der
Aufklärung der Entzündung und Neurodegeneration zugrunde liegenden Mechanismen erzielt
werden. Eine Vielzahl von verschiedenen Tiermodellen trug zu einem besseren Verständnis der
MS-Pathologie und fundamentalen immunologischen Konzepten bei. Außerdem ermöglichten
experimentelle Modelle die Entwicklung von immunmodulatorischen Therapien für MS
Patienten.
Die Forschung der letzten Jahre erbrachte wichtige Hinweise dafür, dass oxidativer
Schaden eine wichtige Rolle im Hinblick auf die Neurodegeneration in der MS spielt. Daher war
unser Ziel, die Beschaffenheit und das Ausmaß von oxidativem Schaden in verschiedenen
Tiermodellen zu ermitteln und direkt mit MS Läsionen zu vergleichen. Hierbei konzentrierten wir
uns auf die Expression von freien Radikale-produzierenden Enzymen (die essentielle
Untereinheit der NADPH Oxidase p22phox und die induzierbare Stickstoffmonoxid Synthetase,
iNOS), die Anreicherung von oxidierten Phospholipiden und Eisen, das oxidativen Schaden
verstärken kann. Die Erkrankungen in den verschiedenen Tiermodellen wurden durch
unterschiedliche inflammatorische Mechanismen induziert, die jeweils andere Aspekte der MS
Pathologie repräsentieren.
Das einzige Modell, in dem wir annähernd so viel oxidativen Schaden wie in MS
Läsionen detektierten, war die durch Coronavirus-Infektion hervorgerufene demyelinisierende
Encephalomyelitis. In der MS als auch im Coronavirus-Modell war der oxidative Schaden mit
einer massiven Aktivierung der Mikroglia und Makrophagen verbunden. Diese zeichnete sich
durch ausgeprägte Expression von p22phox und geringes Vorkommen von iNOS aus. Das
zweite in unsere Studie eingeschlossene Virusmodell war eine chronische Demyelinisierung, die
durch Theiler´s Virus (TMEV) verursacht wurde. In diesem Fall erreichte jedoch die Expression
von p22phox in aktivierten Mikroglia und Makrophagen nicht das Ausmaß des CoronavirusModells. Außerdem konnten wir im TMEV-Modell oxidative Schaden zwar detektieren, aber zu
einem geringeren Grad als im Coronavirus-Modell. Die iNOS Expression erschien in beiden
Modellen ähnlich gering. Läsionen, die durch den passiven Transfer von T-Zellen
xii
hervorgerufenen wurden, zeigten massive p22phox und iNOS Expression in der akuten Phase
der Entzündung. Ähnlich dazu fanden wir p22phox exprimierende Makrophagen in Tieren mit
chronischen Läsionen, die durch T-Zellen und Makrophagen hervorgerufen wurden, aber im
Gegensatz dazu wenige iNOS positive Zellen. Chronische Läsionen von Antikörpern mediierter
Demyelinisierung enthielten phagozytisch aktive Makrophagen, die aber keine Expression von
p22phox oder iNOS zeigten. In akuten Läsionen, die durch den Transfer von CD8+ T-Zellen oder
Mechanismen des angeborenen Immunsystems hervorgerufen wurden, detektieren wir die
Expression von p22phox in Mikroglia und Makrophagen. Im Gegensatz dazu war iNOS nur
geringfügig zu finden. Zusätzlich zum oxidativem Schaden und den verschiedenen
Expressionsmustern von p22phox und iNOS, untersuchten wir auch den Einfluss von Eisen in
den
jeweiligen
experimentellen
Modellen.
Die
alters-abhängige
Eisenanreicherung
in
Oligodendrozyten und dessen Freisetzung, die man in MS Läsionen findet, werden nur wenig in
Tiermodellen widergespiegelt. Wir versuchten die Rolle von Eisen in der Neurodegeneration
mittels Zellkultur zu studieren, konnten eine Eisen Anreicherung aber nur in Mikroglia und nicht
in Oligodendrozyten hervorrufen.
Unsere Studie zeigt, dass in den verschiedenen Tiermodellen der MS auch ein
unterschiedliches Muster von oxidativem Schaden zu finden ist, dessen Ausmaß aber in den
wenigsten Fällen jenes der MS erreicht. Dies liegt möglicherweise daran, dass den Tiermodellen
mögliche Amplifizierungsmechanismen fehlen, die in der MS Pathogenese eine wichtige Rolle
spielen.
xiii
Publications arising from this thesis
Revised manuscript resubmitted to Acta Neuropathologica
Oxidative tissue injury in multiple sclerosis is only partly reflected in experimental
disease models
Cornelia Schuh1, Isabella Wimmer1, Simon Hametner1, Lukas Haider1, Anne-Marie Van Dam2,
Roland S Liblau3, Ken J Smith4,Lesley Probert5, Christoph J Binder6, Jan Bauer1, Monika Bradl1,
Don Mahad7, Hans Lassmann1
1: Department of Neuroimmunology, Centre for Brain Research, Medical University of Vienna,
Vienna, Austria
2: Department of Anatomy & Neurosciences, VU University Medical Center, Amsterdam, The
Netherlands
3: Inserm, U1043 - CNRS, U5282 - Université de Toulouse, Centre de Physiopathologie de
Toulouse Purpan (CPTP), Toulouse, F-31300, France
4: Department of Neuroinflammation, University College London Institute of Neurology, United
Kingdom
5: Laboratory of Molecular Genetics, Hellenic Pasteur Institute, Athens, Greece
6: Department of Laboratory Medicine, Medical University of Vienna, Vienna, Austria
7: Centre for Neuroregeneration, University of Edinburgh, Edinburgh, United Kingdom
xiv
Abbreviations
BBB
blood-brain barrier
CFA
complete Freud´s adjuvants
CNS
central nervous system
DA
dark agouti
Dpi
days post injection
EAE
experimental autoimmune encephalomyelitis
iNOS
inducible nitric oxide synthase
LPS
lipopolysaccharide
MBP
myelin basic protein
MHC
major histocompatibility complex
MHV
mouse hepatitis virus
MOG
myelin oligodendrocyte glycoprotein
MS
multiple sclerosis
MPO
myeloperoxidase
NAWM
normal-appearing white matter
NO
nitric oxide
NWM
normal white matter
PLP
proteolipid protein
PPMS
primary progressive multiple sclerosis
ROS
reactive oxygen species
RRMS
relapsing remitting multiple sclerosis
RT
room temperature
SD
Sprague Dawley
SPMS
secondary progressive multiple sclerosis
TBB
DAB-enhanced turnbull blue
TMEV
Theiler´s murine encephalomyelitis virus
xv
Acknowledgements
I very much enjoyed being a PhD student at the Centre for Brain Research thanks to
many wonderful people who made this time of my life a good one. I am not only grateful for
seeing myself as a part of the Centre for Brain Research, but also of the PhD program Cell
Communication in Health and Disease which is embedded in the Medical University of Vienna.
In the first place, I want to express my gratitude to Prof. Hans Lassmann who is a patient
teacher. His door is always open for us and he is never weary of advising us on the most
important aspects of our work or supporting us wherever he can. Prof. Lassmann is very kindly
leading his team. Additionally to Prof. Lassmann, Prof. Monika Bradl and Prof. Jan Bauer also
guided me through these years of learning and developing.
Our success is based on the supporting pillars of the lab, the busy hands of Marianne
Leisser, Ulli Köck and Angela Kury. Without their expertise and help, I would have been lost in
histology. Further, without the organisation skills of Gerti Makkos and Regina Hirnschall I would
have descended into total chaos.
My fellow students deserve a huge hug for thanking them for being my friends, spending
coffee and beer breaks with me but also for valuable discussions about projects, prospects,
spelling mistakes, actually everything that is important in the life of a student. My thanks go to
Isabella, Simon and Maja for being my PhD student buddies and to all my peers from the
Neuroimmunology lab, the CCHD and the Centre for Brain Research: Lukas, Maria, Isi, Josef,
Josephine, Michi, Hend, Itziar, Phillip, Riem, Justyna, Marco, Carol-Ann, Babsi, Caro, Flo,
Gabriel, Markus, Christoph, Fabian, Simona, Bleranda, Niko, Joana, Taro, Tobias, Simon,
Marko, Maila, Till, Mesi, Eva and Rhaki. More thanks go to Buena Vista Social Club and Early
Grey.
My stay at the University of Glasgow would not have been possible without Prof. Chris
Linington. He and his wife Allison adopted me and gave me shelter and support. Further, Maren,
Christina, Katja and Daniela made my stay in Scotland unforgettable.
xvi
I owe my deepest gratitude to my family for their encouragement throughout my life. My
parents and my brother always believe in me. My parents always told me that I can do
everything I wanted and made my whole education possible.
I thank my partner Matthias for his absolute confidence in me, which is very encouraging.
I would not have the opportunity to write these acknowledgements without him. He always
believes in my ideas, irrespective of how crazy they might be.
I want to thank all the people I met during this journey, including people I did not mention
in particular. During a Phd student’s life, also short encounters or moments can be a big help
and support.
Introduction 1
1 Introduction
1.1 Hallmarks of MS
Multiple sclerosis (MS) is the most prevalent non-traumatic neurological disorder among
young adults in Europe and North America with more than 2 million people worldwide suffering
from the disease. MS is occurring with a frequency of 2:1 regarding female versus male patients
(Hirtz et al, 2007; Noseworthy et al, 2000). In 2008, the world health organisation together with
the multiple sclerosis international federation estimated the worldwide prevalence of MS as 30
per 100000 human beings. They reported that the risk to develop MS is highest in Europe (80
per 100000), followed by the Eastern Mediterranean (15.9 per 100000), America (8.3 per
100000), the Western Pacific (5 per 100000), South-East Asia (2.8 per 100000) and Africa (0.3
per
100000)
(Atlas
multiple
sclerosis
resources
http://www.who.int/mental_health/neurology/Atlas_MS_WEB.pdf,
in
the
14.01.2014).
world
2008,
To
current
opinion, an autoimmune inflammatory process is driving tissue injury in MS (Lassmann et al,
2007). Although an autoimmune pathogenesis of MS is widely accepted and reasonable, the
causing autoimmune mechanism has not yet been nailed down. Therefore, the trigger of MS
remains unknown (Lassmann, 2008; Lipton et al, 2007; Noseworthy et al, 2000).
1.2 Clinical course
In most patients, MS starts with a relapsing/remitting (RRMS) disease course from which
they suffer a number of years. When they reach a certain threshold of irreversible neurological
damage, they enter a secondary progressive disease (SPMS) (Confavreux et al, 2003;
Confavreux et al, 2000; Leray et al, 2010). Generally, RRMS changes to SPMS between an age
of 35 to 50 years. A small portion of patients starts with a primary progressive disease course
(PPMS), frequently in the time span of life when RRMS patients enter SPMS (Confavreux &
Vukusic, 2006; Lassmann et al, 2007).
Clinical data propose that inflammation and the formation of new white matter lesions are
driving the disease in RRMS. The development of new lesions is rare in SPMS but this
progressive stage is rather characterised by diffuse grey and white matter atrophy (Miller et al,
2002). These experiences suggest that inflammation is driving the early disease while a
Introduction 2
neurodegenerative process, developing at least partly independently of inflammation, is
underlying the progressive stage of MS (Zamvil & Steinman, 2003).
MS was first described as an inflammatory process with concomitant focal plaques of
primary demyelination in the white matter of the brain and spinal cord (Charcot et al, 1880). At all
disease stages, MS is associated with inflammation comprising primarily perivascular and
parenchymal lymphocytes and activated macrophages or microglia (Frischer et al, 2009; Prineas
& Wright, 1978). Moreover, CD8+ T cells outnumber other T cell populations, B cells or plasma
cells (Babbe et al, 2000; Frischer et al, 2009).
Active lesions, as typical for RRMS, are characterised by a disturbance of the blood-brain
barrier (BBB) (Hochmeister et al, 2006; Kirk et al, 2003), which can be visualised with magnetic
resonance imaging (MRI) by gadolinium enhancement (Gaitan et al, 2011; Grossman et al,
1988; Miller et al, 1988). In contrast, BBB damage is not detectable anymore by gadoliniumenhanced MRI in PPMS and SPMS despite inflammation, active demyelination and
neurodegeneration. This indicates an inflammatory process that is compartmentalised behind a
relatively closed BBB (Lassmann et al, 2012). Active lesions are associated with the local
expression of pro-inflammatory cytokines, chemokines and their respective receptors (Cannella
& Raine, 1995; Huang et al, 2000). Demyelination is associated with acute axonal injury and
axonal loss (Ferguson et al, 1997; Trapp et al, 1998) which is to some extent counterbalanced
by remyelination (Kornek et al, 2000).
In progressive MS patients (SPMS and PPMS), new and active demyelinating lesions are
scarce. However, pre-existing established plaques can expand slowly at their rims (Prineas et al,
2001) which entails pronounced cortical demyelination and is related with major diffuse injury of
the normal appearing white and grey matter (Lassmann et al, 2012). The lesion expansion is
accompanied by a moderate infiltration of inflammatory cells, primarily T cells, and high-grade
microglial activation. Myelin degradation product-containing cells are rare, which proposes a
very low level of demyelination. Moreover, diffuse inflammation and activated microglia
accompany axonal injury and secondary demyelination in the normal appearing white matter
(NAWM) of progressive MS patients (Allen & McKeown, 1979; Allen et al, 2001; Kutzelnigg et al,
2005; Lassmann et al, 2007). Additionally to severe white matter injury, the cortex is also
affected (Bo et al, 2003; Peterson et al, 2001). Moreover, MS is characterised by astrocytic scar
formation (Lassmann et al, 2007).
Introduction 3
The most common lesion types in all MS stages, particularly prevailing in patients with
long disease duration, are inactive lesions. They are characterised by a lack of myelin, axonal
loss and astrocytic scar tissue. In the lesions, T and B cells are sparse and the number of
microglia in the lesion is lower than in the NAWM (Lassmann et al, 2012). Figure 1 summarises
the pathological hallmarks of different MS courses in relation to disease duration and age.
Figure 1: Pathological hallmarks of MS related with the change from RRMS (pink) to SPMS (green)
During disease duration, inflammation, BBB disturbance and the formation of new lesions decrease,
whereas already emerged lesions expand and cortical demyelination and diffuse tissue injury occur.
Reprinted by permission from Macmillan Publishers Ltd: [Nature Reviews Neurology] (Lassmann et al,
2012), copyright (2012)
1.3 Mechanisms of tissue injury in MS
Along with different players of the innate and the adaptive immune system, CD8+ T cells
(Na et al, 2008; Saxena et al, 2008) and autoantibodies specific for neuronal or glial antigens
(Linington et al, 1988; Mathey et al, 2007) drive demyelination, oligodendrocyte death, axonal
and neuronal injury. Moreover, cytokines and reactive oxygen or nitric oxide species released by
activated macrophages and microglia damage the tissue, especially the vulnerable myelin
sheaths (Lassmann et al, 2012). The different patterns of tissue injury in RRMS lesions correlate
with the heterogeneous components of the immune system causing demyelination. The most
common patterns of tissue injury are antibody- and complement-associated pathology (pattern
II) and hypoxia-like tissue injury (pattern III). In pattern III lesions, demyelination is induced by
oligodendrocyte apoptosis reminiscent of white matter stroke lesions. Another pattern (pattern I)
Introduction 4
represents patients bearing lesions associated with T cell infiltration and microglia activation only
(Lucchinetti et al, 2000). The majority of MS lesions fail to remyelinate, especially in PPMS and
SPMS, even though oligodendrocyte precursors and axons are present (Franklin & FfrenchConstant, 2008).
1.3.1
Oxidative stress and mitochondrial injury
A major role for oxidative injury in demyelination and neurodegeneration in MS has been
suggested (Lassmann et al, 2012). Oxidative damage can be induced by reactive oxygen
species (ROS). ROS have unpaired electrons and are thus very reactive. Furthermore, they can
give rise to more new reactive species. The most abundant forms of ROS in cells are superoxide
(O2-), the related hydroxyl radical (OH·) and hydrogen peroxide (H2O2) (van Horssen et al, 2011).
Superoxide is produced by NAD(P)H oxidases and can be transformed into hydrogen peroxide
by superoxide dismutases (SODs) (Deby & Goutier, 1990) or give rise to other forms of ROS
(Fridovich, 1978). Hydrogen peroxide can, together with divalent transition metal ions, catalyse
the formation of reactive hydroxyl radicals via the Fenton reaction. Superoxide and nitric oxide
(NO), which is produced by nitric oxide synthase (NOS, e.g. inducible nitric oxide synthase,
iNOS), give rise to peroxynitrite (ONOO-) (Beckman, 1991) (Fig. 2). In our bodies, immune cells
produce ROS during the so-called oxidative burst in order to fight pathogens and facilitate
phagocytosis (Forman & Torres, 2001; Hampton et al, 1998).
Introduction 5
Figure 2: Summary of reactive oxygen and nitrogen species detoxification
−
Superoxide (O2· ) is converted by superoxide dismutases (SODs) into hydrogen peroxide (H2O2). In
presence of nitric oxide (NO), superoxide can form peroxynitrite (ONOO·). Hydrogen peroxide can be
detoxified by endogenous antioxidant enzymes (catalase, glutathione peroxidase, peroxiredoxins) but in
combination with transition metals also catalyse the formation of highly toxic hydroxyl radicals (OH·);
adapted from [Biochimica et Biophysica Acta] (van Horssen et al, 2011) copyright (2011) and [European
Journal of Clinical Nutrition] (van Meeteren et al, 2005) copyright (2005).
In line with the emerging role for oxidative injury in MS pathogenesis (Lassmann et al,
2012), tissue injury in MS is characterised by activation of microglia and macrophages (Prineas
et al, 2001) which can produce oxidizing radicals (Colton & Gilbert, 1993). Inflammation-linked
oxidative burst in activated microglia and macrophages emerges to drive oxidative injury
(Fischer et al, 2012; Fischer et al, 2013). Moreover, different subunits of the NADPH oxidase
were shown to be expressed in active MS lesions, as for example the essential subunit p22phox
(Fischer et al, 2012). Oxidative burst by activated macrophages and microglia promotes
demyelination and axonal injury. Additionally to the lesion areas, the NAWM of progressive MS
patients contains activated microglia, which also form microglia nodules (Lassmann et al, 2012).
Introduction 6
The expression of enzymes crucial for different pathways of reactive species production
and their products are highly increased in active MS lesions. Nitrotyrosine is a footprint of
peroxynitrite-mediated cell damage and nitric oxide (NO) production. The strong oxidant
peroxynitrite is generated by the reaction of NO with superoxide at sites of inflammation (Cross
et al, 1998). Myeloperoxidase and inducible nitric oxide synthase (iNOS) were detected in acute
MS lesions primarily in microglia (Liu et al, 2001; Marik et al, 2007; Stadelmann et al, 2005).
Moreover, the expression and activity of myeloperoxidase (MPO), which catalyses the formation
of the cytotoxic oxidant hypochlorous acid, is increased in MS tissue (Gray et al, 2008).
As the patterns of demyelination and axonal injury in acute MS lesions are reminiscent of
stroke lesions, energy deficiency and hypoxia-like tissue injury could play an important role in
MS pathology (Aboul-Enein et al, 2003; Trapp & Stys, 2009). Mitochondria are vulnerable to
injury induced by reactive oxygen and nitrogen species (Fig. 3) (Bolanos et al, 1997; Higgins et
al, 2010). Accordingly, mitochondrial injury could potentially expand oxidative injury (Campbell et
al, 2011; Mahad et al, 2008). In MS lesions, mitochondrial injury was detected by defective
complex I (NADH dehydrogenase) and compensatory increased complex IV activity (Fig. 3) (Lu
et al, 2000). Gene expression studies of motor cortex tissue of MS patients also indicate
mitochondrial damage (Dutta et al, 2006). Furthermore, oxidative damage was also shown to
affect mtDNA (DNA derived from cytoplasmic compartment of cells, mitochondrial DNA) in
chronic active plaques (Lu et al, 2000). Immunohistochemical studies on respiratory chain
proteins discovered major mitochondrial injury in active MS lesions, primarily affecting complex
IV (Fig. 3; cytochrome c oxidase, COX) (Mahad et al, 2008). In contrast, increased mitochondrial
quantities and enzyme activity were found in inactive lesions (Mahad et al, 2009; Witte et al,
2009). This increase in terms of mitochondria number seems to compensate for the augmented
need for energy in demyelinated axons (Lassmann et al, 2012). Defective mitochondria would
account for pathological hallmarks in MS, as reduced energy production can be detrimental for
demyelinated axons with impaired activity of ion channels (Bechtold et al, 2006; Black et al,
2006; Friese et al, 2007). Small diameter axons are more likely damaged than thicker axons as
they contain a lower number of mitochondria in relation to their surface (Evangelou et al, 2001;
Stys, 2005; Trapp & Stys, 2009). In oligodendrocytes, mitochondrial injury leads to apoptosis via
the activation of poly-ADP-ribose polymerase (PARP) which was shown to be the mechanism for
oligodendrocyte death in the cuprizone-induced model for demyelination (Veto et al, 2010).
Defective mitochondria and inflammation-induced oxidative stress generate free radicals
(Bedard & Krause, 2007; Love, 1999; Murphy, 2009). As mentioned before, this can be amplified
Introduction 7
by the presence of extracellular divalent metal ions which can catalyse the formation of oxygen
radicals via the Fenton reaction (Jomova & Valko, 2011).
Figure 3: The respiratory chain of mitochondria
The mitochondrial respiratory chain comprises four complexes (I-IV) and complex V, the latter being
responsible for ATP synthesis. NO can irreversibly damage complexes I and IV through peroxynitrite
-
(ONOO ). Additionally, NO competitively inhibits complex IV; adapted from [Brain] (Mahad et al, 2008)
copyright (2008).
1.3.2
Footprints of oxidative damage in MS
As mentioned before, high levels of ROS are generated during the pathogenesis of MS.
The expression of ROS-producing enzymes is increased in active MS lesions (Lassmann et al,
2012). Antioxidant protective mechanisms are overwhelmed which leads to oxidative stress.
Consequently, ROS harm polyunsaturated fatty acids in membrane lipids, proteins and
DNA/RNA. The CNS is particularly prone to lipid peroxidation as it employs a lot of oxygen and
comprises high levels of polyunsaturated fatty acids (van Horssen et al, 2011). Molecules
indicative for oxidative stress and decreased amounts of antioxidant enzymes and substances
were observed in the blood and cerebrospinal fluid of MS patients during acute disease bouts,
for example increased levels of malonedialdehyde, oxidised gluthatione, lipid hydroperoxides
(Ferretti et al, 2005; Karg et al, 1999) and glutathione reductase but decreased activity of
Introduction 8
glutathione peroxidase (Calabrese et al, 1994). Moreover, markers of oxidative damage were
found in MS lesions, for example 4-hydroxy-2-nonenal arising through lipid peroxidation (Fig. 4)
was shown to accumulate in macrophages and reactive astrocytes in active MS lesions (van
Horssen et al, 2008). Another marker for oxidised phospholipids (E06) was detected in
oligodendrocytes, myelin, neurons and axonal spheroids in active MS lesions (Fischer et al,
2013;
Haider
et
al,
2011).
Additionally,
malonedialdehyde
(MDA2)
was
found
in
oligodendrocytes, myelin and astrocytes in active MS lesions. The presence of 8-hydroxy-2deoxyguanosine (8OHdG, Fig. 4) refers to oxidative damage to nucleotides and was detected in
nuclei of oligodendrocytes, astrocytes and phagocytosing macrophages (Haider et al, 2011).
Figure 4: The production of ROS and subsequent lipid peroxidation
The production of ROS through NADPH oxidases and mitochondria, the detoxification through SODs and
the further reaction with metals via the Fenton reaction is depicted. The generated OH radical can be
involved in lipid peroxidation and DNA damage and give rise to adducts of oxidative damage like 8OHdG,
malondialdehyde and 4-hydroxynonenal; adapted from [Toxicology] (Jomova & Valko, 2011) copyright
(2011)
Introduction 9
Mitochondria are permanently producing ROS as a result of electron leakage in their
electron transfer cascade which is normally counterbalanced by local antioxidant enzymes
(Boveris & Chance, 1973; van Horssen et al, 2011). ROS induce the transcription of various cellprotective enzymes. Genes encoding these proteins have the common promoter element ARE
(antioxidant response element). ARE-controlled gene transcription is orchestrated by the
transcription factor Nrf2 (nuclear factor E2 related factor 2), which translocates to the nucleus
under conditions of oxidative stress (Itoh et al, 2004; Itoh et al, 2003; Motohashi & Yamamoto,
2004). Nrf2 drives gene expression of proteins involved in detoxification and antioxidant
defense, like SODs (McCord & Edeas, 2005), glutathione peroxidases (Thimmulappa et al,
2002), peroxiredoxins (Kim et al, 2007) and catalase (Dringen et al, 2005). MS lesions show
expression of Nrf2-induced enzymes, including SOD1, SOD2 and catalase in macrophages and
astrocytes (van Horssen et al, 2008).
In summary, there is accumulating evidence speaking for an important role of oxidative
stress in different steps in the formation and persistence of MS lesions (Fig. 5).
Counterbalancing the redox state in the MS brain is an attractive therapeutic target. Apparently,
endogenous Nrf2-regulated protective mechanisms are not sufficient enough to protect the brain
from ROS-induced cell destruction. One way to increase the activation of the Nrf2 system could
be further induction of Nrf2 through reagents that promote the transcription of endogenous
antioxidant enzymes, such as tert-butylhydroquinone, dimethylfumarate, sulforaphane or
hydroxycoumarin (Nguyen et al, 2003; van Horssen et al, 2011; Wierinckx et al, 2005).
Strikingly, BG00012, an oral form of dimethylfumarate, showed anti-inflammatory and
neuroprotective effects in clinical trials (van Horssen et al, 2011).
Introduction 10
Figure 5: Summary of how ROS can contribute to MS pathogenesis.
(1) Monocytes produce ROS, which facilitates infiltration of more monocytes by tight junction destruction
and cytoskeletal modifications. (2) Activated microglia and macrophages produce ROS contributing to
demyelination and oligodendrocyte death. Furthermore, ROS promote intracellular myelin degradation in
those activated cells. (3) Microglia- and macrophage-derived ROS can mediate axonal degeneration and
(4) contribute to mitochondrial dysfunction in affected axons. Subsequently, increased ROS and
decreased ATP levels are produced in demyelinated axons. (5) Increased need for energy in chronically
demyelinated axons leads to accumulation of dysfunctional mitochondria which in turn produce more ROS
and further promote axonal injury. [Biochimica et Biophysica Acta] (van Horssen et al, 2011) copyright
(2011)
1.3.3
Iron in MS
Additionally to mitochondrial injury, iron accumulating in the brain during aging and its
release from intracellular departments in active MS lesions (Hametner et al, 2013) could
potentially further add up to oxidative injury. Inflammation is related with a disturbed metal iron
homeostasis, which can exacerbate oxidative stress. This is thought to play an important role in
neurodegenerative diseases such as MS but also in Parkinson´s disease, Alzheimer´s disease
and Huntington´s disease (Crichton et al, 2011).
Iron accumulates in the normal brain during ageing, reaching a steady level between the
age of 40 to 50 years (Hallgren & Sourander, 1958). Interestingly, this is also the time span of
Introduction 11
life when patients typically develop PPMS or SPMS (Confavreux et al, 2000). The highest
amount of iron in the human as well as the rodent brain is found in certain areas, such as the
substantia nigra, the basal ganglia and the cerebellar nuclei. The predominant cell type that
stains for iron is the oligodendroglia (Benkovic & Connor, 1993; Meguro et al, 2007; Todorich et
al, 2008). They store iron mostly as non-heme iron in the classical iron storage molecule ferritin
(Connor & Menzies, 1995; Dwork et al, 1988; Hallgren & Sourander, 1958). Iron accumulation
during ageing does not seem to have apparent adverse effects but a surplus of iron can be as
detrimental as iron shortage (Crichton et al, 2011). Iron loading enhances the vulnerability of
oligodendrocytes to cytokine toxicity (Zhang et al, 2005), whereas a lack of iron, for example
during development, causes hypomyelination (Beard, 2008; Ortiz et al, 2004; Wu et al, 2008).
Iron is essential for many metabolic steps in the CNS including myelin synthesis,
neurotransmitter production, nitric oxide metabolism and oxygen transport. Further, iron is
indispensable for oxygen consumption and ATP production during oxidative phosphorylation,
because it is part of the catalytic centre of the iron-sulphur clusters of NADH dehydrogenase
(complex I), succinate dehydrogenase (complex II), cytochrome bc1 (complex III), cytochrome c
oxidase (complex IV) (Crichton et al, 2011; Kim et al, 2012; Todorich et al, 2009).
Iron is conveyed through our circulation by transferrin which binds two atoms of ferric iron
3+
(Fe ). A major way of cellular iron uptake is receptor-mediated endocytosis of transferrin-iron
and subsequent recycling of the transferrin receptors. Transferrin binds its receptor which leads
to internalisation into the endosome. The acidification of the pH during the endosome maturation
to the lysosome leads to the release of iron from transferrin. Subsequently, iron is transported
into the cytoplasm through the divalent metal transporter 1 (DMT1) (Fleming et al, 1998; Harding
& Stahl, 1983; Raub & Newton, 1991; Sipe & Murphy, 1991). DMT1, also known as NRAMP2,
can also be found in the plasma membrane of cells and used for iron (Fe2+) import (Fleming et
al, 1998; Fleming et al, 1997; Gunshin et al, 1997). In contrast to the several known iron import
mechanisms, there is just one established route for iron export. This includes the only known
iron exporter ferroportin and the ferroportin-coupled ferroxidases hephaestin and ceruloplasmin.
Ferroportin exports Fe2+ which is subsequently oxidised by hephaestin or ceruloplasmin to Fe3+.
Without the ferroxidases, iron export is not possible and it accumulates in the cytoplasm (De
Domenico et al, 2008; Donovan et al, 2000; Jeong & David, 2003; Vulpe et al, 1999).
Excess cellular iron is retained in the intracellular iron storage protein ferritin. Ferritin is a
heteropolymer of 24 subunits of the two different ferritin chains ferritin heavy (H) and light (L)
(Theil, 2004). The ferroxidase activity of ferritin H and the ferritin L ability to nucleate iron to the
Introduction 12
mineral iron core collaborate to store iron. The central core of ferritin can store up to 4500 iron
atoms (Crichton et al, 2011; De Domenico et al, 2008).
Iron transport into the brain is tightly regulated by the BBB. It mostly involves transferriniron and transferrin receptors on the blood capillary endothelial cells but the precise mechanisms
of further iron shuttling remain unclear. Astrocytes at the BBB most probably play an important
role as iron importers into the brain. After crossing the BBB, iron could be released from
transferrin near the astrocytic end-feet by hydrogen ions, ATP and citrate (Morgan, 1977;
Morgan, 1979). Cells in the CNS could then take up citrate-, ATP- or transferrin-bound iron
according to their possibilities and needs (Crichton et al, 2011).
Iron is crucial for oligodendrocyte function. Thus, many studies focussed on the
mechanisms of iron uptake. In vivo and in vitro studies implicated that transferrin-iron is
indispensable for oligodendrocyte development and myelination (Adamo et al, 2006; Badaracco
et al, 2008; Escobar Cabrera et al, 1997; Espinosa-Jeffrey et al, 2002; Espinosa de los Monteros
& Foucaud, 1987; Paez et al, 2006a; Paez et al, 2006b; Saleh et al, 2003). Although transferrin
is a fundamental ingredient of oligodendrocyte cell culture medium (Bottenstein, 1986), several
publications argue for an iron-independent trophic effect of transferrin on oligodendrocyte
survival (Adamo et al, 2006; Badaracco et al, 2008; Escobar Cabrera et al, 1997; Paez et al,
2006a; Paez et al, 2006b). The expression of transferrin receptors was detected on
oligodendrocyte precursors in vitro, but they are down-regulated during maturation (Escobar
Cabrera et al, 1997; Paez et al, 2004). Furthermore, oligodendrocytes cultures depleted from
transferrin were shown to take up iron (Takeda et al, 1998). In more recent studies, ferritin H
was reported to bind to oligodendrocyte progenitors (Hulet et al, 1999a; Hulet et al, 2000; Hulet
et al, 2002; Hulet et al, 1999b) and to be imported by receptor-mediated endocytosis (Hulet et al,
2000). Tim-2 (T cell immunoglobulin mucin domain 2 protein) appeared to be the receptor for
ferritin H on oligodendrocytes (Chen et al, 2005; Todorich et al, 2008) and ferritin H to be a
major iron source for oligodendrocytes (Todorich et al, 2011). Microglia were suggested to act as
iron distributors (Todorich et al, 2009) because they accumulate iron early postnatally and loose
it again while oligodendrocytes accumulate it (Cheepsunthorn et al, 1998; Connor et al, 1995).
Upon injury, oligodendrocytes are replaced by proliferating and ultimately maturing NG2+
oligodendrocyte progenitors (Keirstead & Blakemore, 1997; Schonberg et al, 2012). This can be
affected by activated macrophages and microglia releasing iron and ferritin (Schonberg &
McTigue, 2009; Schonberg et al, 2007; Zhang et al, 2006). Indeed, NG2+ oligodendrocyte
progenitors take up ferritin derived from activated macrophages in vivo (Schonberg et al, 2012).
Introduction 13
While ferritin appears to be a major source for iron for oligodendrocytes in vivo, they can also be
loaded with iron chloride in combination with ascorbate in vitro (Schulz et al, 2011).
In summary, while there are advances in knowledge about iron transporters and the
translational regulation of iron-related proteins and systemic iron homeostasis, many precise
mechanisms involving iron management between neurons and the different glial cells remain
unexplained (Crichton et al, 2011). A summary of proposed iron shuttling mechanisms in the
CNS is depicted in Figure 6.
Figure 6: Proposed mechanisms of iron transport in the brain
Transferrin-bound iron binds to transferrin receptors on the endothelial cells of the BBB. This leads to
receptor-mediated endocytosis. Subsequently, iron can be transported into the cytosol by DMT1 and then
stored in ferritin or used for physiological functions (for example respiratory chain function in
mitochondria). The exact mechanism how iron is crossing the BBB is not clear yet. After overcoming the
BBB, iron can be oxidised by ceruloplasmin in astrocyte membranes and again bind to transferrin. This
can be used as iron source by neurons which express transferrin receptors. Oligodendrocytes appear to
import iron bound to ferritin with the receptor Tim2. Microglia presumably take up iron through
phagocytosis. [Biochimica et Biophysica Acta] (Leitner & Connor, 2012) copyright (2012)
Introduction 14
1.4 Oxidative burst, oxidative damage and iron accumulation iron in
MS
As described before, macrophage and microglia activation in MS is characterised by the
expression of p22phox and iNOS (Fischer et al, 2012; Fischer et al, 2013; Marik et al, 2007). We
differentiate between microglia and macrophages by their morphological phenotype as there is
no microglia-specific marker to date. In the human control brain, p22phox is expressed by
scattered microglia (Fig. 7a, Table 1, page 48). In the NAWM of MS patients, p22phox is
detected in microglial nodules (Fig. 7b) and in areas of initial lesions (Barnett & Prineas, 2004;
Fischer et al, 2012) (Fig. 7c). At later lesion stages, p22phox is only present in cells with
macrophage morphology (Fig. 7d). iNOS is expressed by individual cells in the NAWM and in
initial lesions (Fig. 7e, f) and by a considerable number of microglia and macrophages in active
MS lesions (Marik et al, 2007) (Fig. 7g, Table 1, page 48).
Oxidative burst in active MS lesions has been related with the occurrence of oxidised
phospholipids (Fischer et al, 2012; Fischer et al, 2013; Haider et al, 2011). In human controls,
only low levels of oxidised lipids are detected (Fig. 7h) whereas intense immunoreactivity is
found in initial and active MS lesions (Fig. 7i-l). Oxidised phospholipids accumulate in
oligodendrocytes and myelin (Fig. 7i, j), in astrocytes (Fig. 7k) and in damaged axons and
neurons (Fig. 7l) (Haider et al, 2011). The release of iron from oligodendrocytes and myelin in
active MS lesions could potentially amplify oxidative injury (Hametner et al, 2013) (Fig. 7m-q). As
described above, oligodendrocytes store iron during ageing (Fig. 7n, o). Staining for iron is found
in the extracellular space in active MS lesions (Hametner et al, 2013). Subsequently, iron is
imported by microglia and macrophages (Fig. 7p, q).
Introduction 15
Figure 7: The expression of p22phox and iNOS and the accumulation of oxidised phospholipids
(E06) and iron in MS lesions
Introduction 16
p22phox expression in the normal white matter (NWM) of a human control brain is found in scattered
microglia (a). The NAWM shows activation of microglia by forming microglial nodules which are p22phox
+
(b). Macrophages and microglia in the initial (c) and active lesion (d) of an acute MS case exhibit profound
p22phox expression. Individual microglia stain for iNOS in the NAWM (e) and initial lesions (f) in acute
MS. In active lesions, primarily macrophages express iNOS (g). Oxidised phospholipids (E06) do not
accumulate in the NWM of human controls (h). In contrast, high intensity of E06 is detected in initial
lesions of acute MS (i). Double stainings for E06 (blue) and cell-specific markers (brown) confirm the
accumulation of oxidised phospholipids in oligodendrocytes (TPPPp25; j) and cytoplasmic granules of
astrocytes (GFAP; k). Also dystrophic neurons show E06 staining (l). Iron staining of an MS brain shows
iron accumulation in the subcortical white matter (m; arrow heads) and at the rim of active lesions (m;
arrows) from a SPMS patient. Established lesions exhibit less intense iron staining (m, asterisks).
Oligodendrocytes in young human controls accumulate a low amount of iron (n; age: 30), whereas the
aged human control (o; age: 84) and the MS patient (m; age: 57) reveal intense iron staining in
oligodendrocytes and myelin. Microglia at the lesion rim of a RRMS patient accumulate iron (p). The
centre of an active lesion shows decreased iron staining, mainly in (perivascular) macrophages (q). scale
bar: 50 µm except for m = 1 cm
1.5 Experimental Models
Experimental autoimmune encephalomyelitis (EAE) is the most commonly used model
for inflammatory demyelination of the CNS (Gold et al, 2006). These animal models driven by
autoimmunity against CNS antigens explained fundamental immunological concepts (Steinman,
2003), including T cell immune surveillance in the CNS, immune-mediated tissue injury,
demyelination,
neurodegeneration
and
molecular
mechanisms
participating
in
brain
inflammation. Based on this knowledge, established anti-inflammatory therapies for MS have
been developed. EAE can be induced in many species including mice, rats, guinea pigs, rabbits,
rhesus monkeys and marmosets, by active immunisation with myelin and non-myelin antigens
(for example myelin basic protein (MBP), proteolipid protein (PLP), myelin oligodendrocyte
glycoprotein (MOG) or S-100β) (Hohlfeld & Wekerle, 2004; Kipp et al, 2009). The transfer of
organ-specific activated autoimmune CD4+ T cells can induce EAE in healthy animals (Ben-Nun
et al, 1981). Furthermore, autoantibodies against the surface-exposed myelin antigen MOG
potentiate T cell-induced demyelination (Linington et al, 1988). Additionally to CD4+ T cells,
myelin-specific CD8+ T cells were shown to have encephalitogenic potential (Huseby et al,
2001). Therefore, these animal models support the notion that MS is an autoimmune disease
(Kipp et al, 2009).
Introduction 17
MS is a complex disease with heterogeneous pathological mechanisms in different
patients and different disease stages with variable clinical outcomes and immunological
phenotypes (Gold et al, 2006). One obvious difference between EAE and MS is that MS
develops spontaneously whereas EAE has to be induced. Further, EAE experiments are
performed in inbred animal strains under standardised conditions. Therefore, EAE can never
reflect the genetic heterogeneity in the MS population (Gold et al, 2006). None of the existing
experimental animal models covers the complexity of MS. Therefore, a variety of different
models is used to unravel different aspects of MS pathology (Kipp et al, 2009).
1.5.1
Animal models used in this thesis
A number of well-characterised animal models exist that account for understanding
pathological mechanisms in MS and are necessary to test new therapies before applying them in
clinical trials. In order to study oxidative injury, oxidative damage and iron accumulation, we
chose different models of MS, each reflecting different aspects of tissue injury.
1.5.1.1 Passive transfer of MBP-specific T cells in Lewis rats
This model originates from a major achievement in the 1980s (Ben-Nun et al, 1981). In
this work, the autoimmune nature of EAE was confirmed by inducing EAE with in vitro
propagated MBP-specific T cells in naïve syngeneic recipient animals. In the Lewis rat, clinical
disease starts 3-4 days and reaches a peak 5-6 days after T cell transfer associated with axonal
injury (Aboul-Enein et al, 2006). The disease is primarily mediated by CD4+ TH1 T cells and is
associated with pronounced microglial activation and recruitment of blood-derived monocytes
primarily in the spinal cord (Hickey et al, 1992). Finally the disease ends with a complete
remission of the animals about 12 days after T cell transfer. Although this model established that
MBP-specific T cells can induce autoimmune CNS disease, it is important to note its limitations.
The EAE course is monophasic and demyelination is sparse (Aboul-Enein et al, 2006). Hence, a
T cell response alone is insufficient to induce extensive demyelination and chronic disease,
which are both characteristic for MS. Despite these limitations, passive transfer EAE is a very
reproducible model and has contributed to unravel basic mechanisms involved in T cellmediated CNS inflammation. This was essential for the development of anti-inflammatory
therapies (Gold et al, 2006).
Introduction 18
1.5.1.2 Active immunisation with MOG/CFA in DA rats
Myelin oligodendrocyte glycoprotein (MOG) is a special myelin auto-antigen because
MOG-immunisation induces the production of demyelinating antibodies additionally to an
encephalitogenic T cell response in susceptible species (Linington et al, 1988). These anti-MOG
antibodies exacerbate disease severity and cause fulminant demyelination in mice, rats and
primates (Genain et al, 1995; Linington et al, 1988; Schluesener et al, 1987). Dark agouti (DA)
rats actively immunised with MOG1-125 exhibit a chronic relapsing-remitting disease reflecting
many pathological aspects of MS including axonal injury. First disease bouts are characterised
by perivenous and subpial inflammation and mediated by CD4+ T cells, macrophages and
granulocytes being also associated with demyelination. Lesion formation is observed in the
spinal cord but also in the forebrain and optic nerves (Storch et al, 1998). Chronic disease is
associated with large demyelinating lesions. Anti-MOG antibodies induce dismantling of myelin
through complement activation and antibody-mediated cytotoxicity (Piddlesden et al, 1991;
Piddlesden et al, 1993). This combination of mechanisms of tissue injury is reminiscent of the
pathology found in early MS patients with Pattern II lesions characterised by the precipitation of
immunoglobulins and complement (Gold et al, 2006).
1.5.1.3
Active immunisation with MOG35-55 in C57/BL6 mice
Additionally to DA rats, a chronic-progressive MOG35-55 EAE can also be induced in
C57/BL6 mice representing the most commonly used animal model in MS research. To
induce MOG-EAE in mice, pertussis toxin injections are necessary to boost disease. In contrast
to DA rats, primary tissue damage in C57/BL6 mice is not associated with an autoantibody
response. This is due to their inefficient complement cascade and MHC haplotype-associated
inability to mount an antibody response to rodent MOG (Bourquin et al, 2003; Gold et al, 2006).
MOG35-55-induced EAE in C57/BL6 mice is characterised by demyelination caused by T cells
and macrophages which is reminiscent of pattern I MS lesions (Calida et al, 2001; Gold et al,
2006). In our model, clinical disease arises 10 to 15 days after sensitization. This acute episode
appears to be similar to the disease peak of acute T cell transfer EAE. The first disease bout is
followed by a short remission and a relapse with a progressive increase of clinical disease
related to demyelination over the following weeks (Taoufik et al, 2011).
Introduction 19
1.5.1.4 Passive transfer of hemagglutinin (HA)-specific CD8+ T cells in HA-transgenic BALB/c
mice
CD8+ T cells were shown to be a major population of inflammatory cells in MS lesions
(Babbe et al, 2000; Neumann et al, 2002). Mouse models showed that CD8+ T cells can be
encephalitogenic in vivo (Cabarrocas et al, 2003; Ford & Evavold, 2005; Huseby et al, 2001).
This is only possible when the tight control of CD8+ T cells is circumvented. For our study, we
used a model where EAE was triggered by passive transfer of influenza virus peptide
hemagglutinin (HA)-reactive CD8+ T cells into transgenic BALB/c mice expressing HA
specifically in oligodendrocytes (Saxena et al, 2008). At variable time points after T cell-injection,
demyelinating lesions with partial preservation of axons occur in brain and spinal cord. The
lesions contain CD8+ T cells and activated microglia and macrophages. It was shown that CD8+
T cells attack oligodendrocytes with granzyme B, which is associated with oligodendrocyte
apoptosis and demyelination (Saxena et al, 2008).
1.5.1.5 Intraspinal LPS-injection in SD rats
A model to induce inflammation and primary demyelination in the CNS is intraspinal
lipopolysaccharide (LPS)-injection in Sprague Dawley (SD) rats (Felts et al, 2005). This causes
focal inflammation within the dorsal column 12-24 hours after the injection. Early infiltrates
mainly comprise granulocytes and a small number of macrophages and T cells. About one week
after LPS injection, this early inflammatory phase is succeeded by focal primary demyelination
with relative axonal preservation. This was accompanied by profound activation of microglia and
macrophage infiltration (Felts et al, 2005). In this model, demyelination is presumably induced
indirectly by LPS-activated microglia. LPS can activate microglia via mechanisms of the innate
immune system (Toll-like receptor 4, TLR4), which causes the production of cytokines as well as
reactive oxygen and nitrogen species. Lesions caused by LPS-injection are to some extent
reminiscent of pattern III MS lesions, which show oligodendrocyte apoptosis due to hypoxia-like
tissue injury (Felts et al, 2005; Gold et al, 2006; Marik et al, 2007; Sharma et al, 2010).
1.5.1.6 Cuprizone diet-induced demyelination in C57BL/6 mice
In contrast to above-mentioned animal models, the cuprizone model represents a toxininduced disease and is a common model to study mechanisms related to de- and re-myelination
in the CNS (Carlton, 1966; Kipp et al, 2009). Cuprizone (bis-cyclohexanone oxaldihydrazone) is
Introduction 20
a copper chelator causing demyelination by primary oligodendrocyte apoptosis rather than by a
destruction of myelin sheets (Kipp et al, 2009). Continuous cuprizone-supplemented diet (0.20.5%) causes progressive demyelination in C57BL/6 mice starting around day 10 (Morell et al,
1998; Van Strien et al, 2011). Large parts of the corpus callosum and the periventricular white
matter are demyelinated around 35 days in C57BL/6 mice after cuprizone diet onset. Additionally
to the corpus callosum also other parts of the brain can be affected by demyelination as for
example the deep cerebellar nuclei (Groebe et al, 2009), the hippocampus (Hoffmann et al,
2008; Norkute et al, 2009) and the putamen (Kipp et al, 2009). The demyelination pattern in SJL
mice has been reported to differ from that in C57BL/6 mice (Taylor et al, 2009). The cuprizone
model is an example for a reversible demyelination and remyelination system as, after changing
the mice to normal chow, spontaneous remyelination occurs. Moreover, remyelination can be
restricted by a prolonged cuprizone diet (Kipp et al, 2009).
The mechanism of cuprizone-induced oligodendrocyte apoptosis is not completely
explained but it definitely involves a pathological change of mitochondrial morphology.
Therefore, a dysfunction of the mitochondrial respiratory chain seems to be evident (Arnold &
Beyer, 2009). Additionally, the activity of carbonic anhydrase II (CA II) decreases in cuprizonefed animals already before the onset of demyelination (Cammer et al, 1995; Komoly et al, 1987).
CA II is involved in buffering acute pH shifts in the brain. Therefore, a perturbation of the
intracerebral pH (acidosis) could potentially contribute to oligodendrocyte pathology (Kida et al,
2006). Further, it is still under debate if the copper-chelating property of cuprizone is involved in
its neurotoxic effect or if an inhibition of oligodendrocyte differentiation (Cammer, 1999) or
microglia or macrophages-derived pro-inflammatory cytokines are accountable (Kipp et al, 2009;
Pasquini et al, 2007). The primary oligodendrocyte death occurring in the cuprizone model is
accompanied by microglia activation. This is reminiscent of MS lesion formation. Especially type
III MS lesions are believed to be in part mimicked by the cuprizone model (Barnett & Prineas,
2004; Kipp et al, 2009; Lucchinetti et al, 2000).
1.5.1.7 Mouse hepatitis virus (MHV, JHM strain)-induced demyelinating disease in Lewis rats
Virus infections are suspected to be one of the possible risks or causative factors for the
development of MS as viral and also bacterial infections, apart from genetic predisposition, are
considered to increase the immunogenicity of autoantigens. A common idea is that a virus could
induce molecular mimicry, which leads to a shift from a microbial epitope- to a self-directed
immune reaction. In contrast, tissue damage upon viral infections can also be a bystander or
Introduction 21
nonspecific effect caused by a local inflammatory milieu (Kamradt et al, 2005; Lipton et al, 2007;
Morahan & Morel, 2002; Wucherpfennig, 2001). Moreover, other virus infections of the CNS
such as subacute sclerosing panencephalitis (SSPE, caused by measles virus) (Payne et al,
1969; Tellez-Negal & Harter, 1966) and progressive multifocal leukoencephalopathy (PML,
caused by JC polyomavirus) (Padgett et al, 1971; Zurhein & Chou, 1965) are characterised by
demyelination.
Rodent CNS pathogens causing chronic demyelination include two well-described RNA
virus models: mouse hepatitis virus (MHV), a member of the enveloped Coronaviridae
(Bergmann et al, 2006; Lavi et al, 1984) and Theiler´s murine encephalomyelitis virus (TMEV), a
member of the non-enveloped Picornaviridae (Brahic et al, 2005; Lipton, 1975). Although both
viruses induce an acute CD8+ T cell response in their host, they manage to circumvent immune
surveillance and cause chronic CNS infection with concomitant myelin loss (Bergmann et al,
2006).
Typically, MHV infects the gastrointestinal tract of mice as a natural pathogen (Bergmann
et al, 2006). Direct intra-cerebral injection of the neurovirulent coronavirus mouse hepatitis virus
strain JHM causes a chronic inflammatory demyelinating disease in Lewis rats (Barac-Latas et
al, 1997; Korner et al, 1991; Wege et al, 1998; Zimprich et al, 1991). Virus persistence is
accompanied by chronic CNS inflammation and prevailing primary demyelination (Stohlman &
Weiner, 1981). Viral replication is supported by macrophages, microglia, astrocytes and
oligodendrocytes (Bergmann et al, 2006). Virus infection leads to different disease courses.
Animals can develop an acute inflammatory disease primarily affecting the grey matter with viral
antigens being expressed in neurons, astrocytes and oligodendrocytes. Other animals suffer
from a late-onset disease which starts around 30 days after virus injection. These animals
exhibit either a chronic panencephalitis of the grey and white matter or a chronic demyelinating
encephalomyelitis of the brain stem and the spinal cord white matter. At this late disease stage,
virus antigen is mainly expressed in glial cells (Zimprich et al, 1991). MHV infection of the CNS
induces a diversity of immune responses causing tissue injury including neutrophils,
macrophages, NK cells (Bergmann et al, 1999; Zhou et al, 2003), CD8+ and CD4+ T cells
(Dandekar et al, 2004; Haring et al, 2001; Pewe & Perlman, 2002; Stohlman et al, 2008), innate
immunity (TLRs) and anti-viral antibodies (Lin et al, 1999; Tschen et al, 2002; Zimprich et al,
1991).
Introduction 22
1.5.1.8 Theiler´s murine encephalomyelitis virus (TMEV, DA strain)-mediated demyelinating
disease in SJL mice
Theiler´s
murine
encephalomyelitis
virus
(TMEV)-induced
demyelination
is
an
established model to study MS-related symptoms (Lipton, 1975; Rodriguez et al, 1986). TMEV
causes neurological and enteric diseases in susceptible mouse strains such as SJL (Lipton &
Jelachich, 1997). Intracerebral infection with the TMEV DA strain induces a biphasic disease in
susceptible mouse strains, such as SJL (Daniels et al, 1952; Lipton, 1975; Lipton & Dal Canto,
1979). An early phase of acute encephalitis and concomitant primary demyelination is followed
by a chronic demyelinating disease (Dal Canto & Lipton, 1975; Drescher et al, 1997). TMEVresistant mouse strains, such as C57BL/6, only exhibit early acute disease and clear the virus
quickly without developing chronic demyelinating disease (Lipton, 1975; Lorch et al, 1981). In
contrast, virus clearance fails in TMEV-susceptible strains and the virus is lingering in
macrophages, microglia, astrocytes and oligodendrocytes (Clatch et al, 1990; Jelachich et al,
1995; Qi & Dal Canto, 1996; Stroop et al, 1981; Tsunoda & Fujinami, 1996). Virus infection
induces a complex immune response including CD8+ T cells (Ruby & Ramshaw, 1991;
Zinkernagel & Althage, 1977), TH1 helper T cells (Lipton et al, 2005), natural killer cells (Paya et
al, 1989), macrophages (Lipton et al, 2005) and anti-viral antibody production (Rodriguez et al,
1988a; Roos et al, 1987)
1.5.2
Oxidative injury in animal models
Like in MS, reactive oxygen and nitric oxide species contribute to the pathogenesis of
tissue injury in EAE (Nikic et al, 2011; Ruuls et al, 1995). As oxidative damage appears to be a
key mechanism of tissue damage in MS, antioxidant therapies are apparent ways to reduce
disease progression. Animal experiments discovered that administration of antioxidants like
lipoic acid (Chaudhary et al, 2006; Schreibelt et al, 2006), the peroxynitrite scavenger uric acid
(Kean et al, 2000), flavonoids (Hendriks et al, 2004; Muthian & Bright, 2004), free radical
scavengers (Moriya et al, 2008), the antioxidant N-acetylcysteine amide (Gilgun-Sherki et al,
2005), green tea epigallocatechin-3-gallate (Aktas et al, 2004) and propolis caffeic acid
phenethyl ester (Ilhan et al, 2004) limited progression and clinical signs of EAE. In contrast, to
date antioxidant therapies in MS patients were of limited success presumably because most
antioxidant substances cannot cross the BBB and have a short therapeutic window. Moreover,
high dosages of antioxidants are needed to have a protective effect even in EAE (van Horssen
et al, 2011).
Introduction 23
Referring to endogenous antioxidative defence mechanisms, animals suffering from EAE
show decreased catalase activity (Guy et al, 1989a), whereas catalase administration decreased
clinical disease and loss of BBB integrity (Guy et al, 1989a; Guy et al, 1989b; Ruuls et al, 1995).
Furthermore, intraperitoneal injections with peroxidase increased BBB integrity in EAE (Guy et
al, 1989b). SOD2 expression was found in astrocytes and microglia in EAE animals (Qi et al,
1997). Hence, endogenous antioxidant enzymes are also potential therapeutic targets. There
have been several studies showing protective effects of Nrf2 in vitro and in vivo (Shih et al,
2005a; Shih et al, 2005b; Yang et al, 2009; Zhao et al, 2006).
1.5.3
Outlook to MS therapy
To date, MS can be at least party treated in early disease stages by anti-inflammatory
and immune-modulatory treatments that reduce the severity and frequency of demyelinating
bouts. In contrast, the success of therapies is limited in patients once they are suffering from
progressive disease. Moreover, the development of neuroprotective therapies has proven to be
difficult. Many new therapeutic approaches showed promising result in T cell-mediated
experimental animal models but had no or exacerbating affects in MS patients (Friese & Fugger,
2005; Gold et al, 2006; Hohlfeld & Wekerle, 2004). The divergence in therapeutic success
between EAE and MS can be at least partly ascribed to the different nature of inflammation
(Friese & Fugger, 2005). CD4+ auto-reactive T cells drive inflammation in EAE, whereas mainly
CD8+ T cells are found in MS lesions (Booss et al, 1983; Hayashi et al, 1988). Furthermore, in
MS mostly MHC I- restricted CD8+ T cells undergo clonal expansion (Babbe et al, 2000) and
MHC I molecules are expressed in MS lesions in inflammatory cells, neurons and glia
(Hoftberger et al, 2004). After all, EAE is a valuable model but clearly not a perfect analogue of
MS and does not reproduce the heterogeneity of the disease (Hohlfeld & Wekerle, 2004).
Further, the target antigen(s) in MS are not known, in contrast to, for example, neuromyelitis
optica with aquaporin 4 as the target antigen in the majority of patients (Jarius et al, 2008;
Lennon et al, 2004). To adress the problem of therapy limitations for progressive MS patients,
one has to discover the mechanisms driving the disease and to develop animal models that
mimic this chronic disease more closely (Lassmann et al, 2012; Noseworthy et al, 2000).
Introduction 24
1.6 Aims of this thesis
As described before, inflammation with concomitant oxidative burst and mitochondrial
injury are proposed to me major mechanisms of tissue injury in MS. Oxidative damage can
potentially be amplified by the presence or iron (Fig. 8).
The aims of this thesis were to study the expression of p22phox and iNOS, as two
sources for reactive oxygen and nitrogen species in different rodent models of inflammatory
demyelination. Further, analysis of iron accumulation in inflammatory lesions of these models
was of interest. Finally, we wanted to compare the extent of oxidative injury in rodent models
with that in MS tissue relying on identical tools.
Additionally, we intended to study iron accumulation in rodents during aging and the
effect of iron on neurodegeneration in aged animals. Additionally, we were interested in in vitro
iron loading experiments of a complex cell culture system, as the established in vivo animal
models showed only limited iron accumulation.
Figure 8: Schematic summary of mechanisms of tissue injury in active MS lesions.
Oxidative damage mediated by oxidative burst of activated macrophages and microglia (expression of
p22phox and iNOS), radical-mediated mitochondrial injury (which leads to energy failure) and extracellular
iron can sum up to the profound oxidative damage in MS lesions.
Results 25
2 Results
2.1 Iron accumulation in the rat CNS during ageing
Iron accumulates in the human brain in oligodendrocytes and myelin (Connor & Menzies,
1995) (Hallgren & Sourander, 1958) (Hametner et al, 2013). The increase of iron is associated
with tissue damage, as iron can catalyse the Fenton reaction to potentiate the generation of
oxygen radicals (Crichton et al, 2002). Hence, the liberation of iron from oligodendrocytes and
myelin in active MS lesions is regarded as a potential amplification factor for oxidative injury
(Hametner et al, 2013). Iron accumulation in the rodent brain was described before (Benkovic &
Connor, 1993). In order to study the role of iron in the amplification of neurodegeneration in CNS
inflammation, we first identified iron-accumulating regions in the CNS of aged Lewis rats. After
pinpointing iron-storing hotspots, we analysed the level of neurodegeneration in these regions in
rats suffering from MBP-specific T cell transfer EAE.
In order to study iron accumulation in the rat CNS during ageing, we analysed the iron
content of tissues from Lewis rats from the age of 2 to 14 months with two different methods: the
histochemical staining DAB-enhanced turnbull blue (TBB) and the biochemical quantification by
the ferrocene assay. Only in rats above an age of 12 months, we could find iron deposition in
areas which are known to accumulate iron deposition in the human and rat brain (Benkovic &
Connor, 1993; Hallgren & Sourander, 1958). We detected TBB+ oligodendrocytes in the basal
ganglia and the cerebellar nuclei and defined the regions depicted in Figure 9 as the so-called
iron accumulation hotspots.
Results 26
Figure 9: Iron accumulating hotspots in the aged Lewis rat.
Iron accumulates in oligodendrocytes and myelin in the basal ganglia and the cerebellar nuclei. Shown are
brain slices of a 16 months old Lewis rat stained for iron with DAB-enhanced turnbull blue (TBB). CPU =
corpus striatum; POA = lateral preoptic area; SN = substantia nigra; IP = interpeduncular nucleus; ND =
nucleus dendatus, scale bar = 1 mm.
Besides the brain, we tested also the spinal cord for iron accumulation. TBB stainings of
rat spinal cords did not reveal any iron accumulation in particular cells (Fig. 10a, b). Likewise,
the ferrocene assays did not reveal any increase of iron content (Fig. 10c).
Figure 10: Iron histochemistry and biochemical iron quantification in the rat spinal cord.
TBB stainings revealed that iron did not accumulate in particular cells of the spinal cords of Lewis rat
during aging. Depicted are spinal cord sections of a 25 days (a) and of a 12 months old Lewis rat (b).
scale bar = 0.5 mm. Iron quantification by ferrocene assay revealed no significant increase of the iron
concentration in spinal cords of aged rats (c, 2-4 months: n = 16; 12-14 months: n = 17).
Results 27
Furthermore, we performed TBB stainings of the basal ganglia of Lewis rats (Fig. 11b).
Optical densitometry of equally sized pictures revealed a significant increase of iron staining
intensity (Fig. 11a, p ≤ 0.001) in representative areas (Fig. 11c, d).
Figure 11: Histochemical iron quantification in the rat basal ganglia.
Quantification of TBB stainings by optical densitometry revealed an increase in iron staining in the basal
ganglia of aged rats. Equally sized pictures were analysed (a; 2-4 months: n = 20; 12-14 months: n = 13; p
≤ 0.001 indicated by ***). TBB stainings of the basal ganglia of a 14 months (b, c) and 2 months (d) old
Lewis rat are depicted. scale bars b = 0.25 cm; c, d = 50 µm
For methodological reasons, we correlated the outcome of our two different methods for
iron detection. We compared the DAB-enhanced Turnbull blue staining (TBB) as an established
histological detection of iron (Meguro et al, 2007) and the ferrocene assay for the biochemical
quantification of the iron contents of fresh tissue (Fish, 1988). For that purpose, we analysed the
optical density of equally sized pictures taken from the dentate nucleus of the cerebellum, which
revealed iron accumulation with increasing age. As depicted in Figure 12, we correlated the
Results 28
results of TBB staining quantification with those of the ferrocene assay of whole cerebella and
found a linear correlation (r = 0.485, p < 0.001, n = 52).
Figure 12: Correlation of two different methods of iron quantification.
Histochemical and biochemical iron quantification of rat cerebella correlate. Optical densitometry of DABenhanced Turnbull blue staining (x-axis) was compared with iron quantification by ferrocene assay (y-axis)
of rat cerebella from animals aged 2 to 20 months (n = 52, r = 0.485, p < 0.001).
2.2 Acute CD4+ T cell-mediated EAE in aged Lewis rats
After pinpointing the iron hotspots, we induced passive transfer EAE in 2 and 14 months
old female Lewis rats and compared their disease course and histopathology. For that purpose,
we first prepared MBP-specific CD4+ T cells as described in the Material and Methods part. We
injected activated MBP-specific T cells and monitored the disease course by weighing the
animals and by staging the EAE according to criteria described in the Material and Methods part
(Aboul-Enein et al, 2006). The transfer of CD4+ MBP-specific T cells elicited a monophasic
disease in both, young and old animals, starting 3 days after T cell transfer. As depicted in
Figure 13, both age groups showed a similar loss of weight (Fig. 13a) and disease course (Fig.
13b) after the transfer of the same quantity of activated MBP-specific CD4+ T cells.
Results 29
+
Figure 13: Weight loss and EAE disease stages in acute CD4 T cell transfer EAE in young and
aged Lewis rats.
Young (2 months indicated by the blue line) and aged (14 months indicated by the red line) Lewis rats
exhibited a similar disease course according to loss of weight (a) and EAE disease staging (b), mean ±
SD).
Animals were sacrificed at the peak of the disease (day 6 after T cell transfer) or in the
recovery phase (day 13 after T cell transfer) in order to study inflammation, neurodegeneration
and iron accumulation. At the peak of the disease, CNS inflammation comprised of pronounced
T cell infiltration which was associated with microglial and macrophage activation and transient
axonal dysfunction (Aboul-Enein et al, 2006). We quantified inflammation by counting the
number of CD3+ T cells in the lumbar spinal cord slides of young (Fig. 14a, b) and old animals
(Fig. 14c) at the acute and recovery phase of EAE. At the acute disease phase, we found a
significantly higher number of T cells in the spinal cords of young rats compared with old ones
(Fig. 15a, p = 0.005). The percentage of area covered by ED1+ macrophages and microglia was
determined by densitometric analysis of pictures taken of whole lumbar spinal cord sections from
young (Fig. 14d, e) and old (Fig. 14f) animals. We did not find a difference in macrophage and
microglia activation between old and young animals (Fig. 15b), neither at acute nor recovery
phase. Furthermore, we counted APP+ axonal spheroids in the lumbar spinal cord sections as a
measure for neurodegeneration in young (Fig. 14g, h) and old (Fig. 14i) animals. In this case, old
animals showed a significantly higher number of axonal spheroids at the acute phase of EAE
than young animals (Fig. 15c, p = 0.039) which indicated enhanced axonal injury.
Results 30
Figure 14: Immunohistochemistry of markers for inflammation and neurodegeneration in the
+
spinal cords of young and old Lewis rats suffering from acute CD4 MBP-specific T cell transfer
EAE.
+
At the peak of EAE , CD3 T cells infiltrate the lumbar spinal cords of young (a, b, 2 months) and old (c, 14
+
+
months) Lewis rats. ED1 macrophages in young (d, e) and old (f) animals and APP axonal spheroids in
young (g, h) and old (i) animals at the EAE peak are depicted. scale bar = 0.5 mm except for b, e, h = 50
µm
Results 31
Figure 15: Quantification of stainings for inflammatory and neurodegenerative markers in the
+
lumbar spinal cords of young and old Lewis rats suffering from acute CD4 MBP-specific T cell
transfer EAE.
+
Young (2 months) animals exhibited a higher number of infiltrating CD3 T cells at the acute phase of EAE
compared with old (14 months) animals (a; young acute: n = 5, young late: n = 6, old acute: n = 8, old late:
n = 6; p = 0.005 indicated by **). The percentage of area of lumbar spinal cords covered by ED1
+
macrophages and microglia was similar in young and old animals in the respective EAE stage. The
+
number of APP axonal spheroids was higher in old animals at the acute stage of EAE than in young
animals (c; young acute: n = 5, young late: n = 6, old acute: n = 6, old late: n = 9; p = 0.039 indicated by *).
As described in the introduction, oxidative damage plays a central role in the
pathogenesis of MS. Therefore, we wanted to study the type and extent of oxidative injury in
EAE. At the acute phase of EAE, demyelination in CD4+ transfer EAE was rare (Fig. 16a)
(Aboul-Enein et al, 2006). Cells with microglial morphology did not express p22phox or iNOS
(Fig. 16b-e). In contrast, cells with macrophage morphology in the lesions intensely stained for
p22phox and iNOS (Fig. 16d, e). We did not find any evidence for the formation of microglia
nodules in the NAWM. In the recovery phase of EAE, cells with macrophage morphology
expressed the phagocytosis-associated molecule ED1 but neither p22phox nor iNOS (data not
Results 32
shown). A specific staining for oxidised phospholipids was neither detectable in young nor in
aged EAE animals (Fig. 16f). In one aged control animal, we found degenerating neurons
specifically staining for oxidised phospholipids (Fig. 16f, insert) confirming that the epitope of
E06 in rodents was detectable using the same staining protocol as for MS tissue.
+
Figure 16: Oxidative burst and oxidative injury in acute CD4 T cell transfer EAE in Lewis rats.
+
Demyelination was rare in acute CD4
mediated EAE (a). Inflammation was associated with the
expression of Iba-1 (pan-microglia/macrophage marker) (b, c) but p22phox (d) and iNOS (e) expression
were limited to cells with macrophage morphology. Inflammatory lesions were not associated with
immunoreactivity for oxidised phospholipids (E06). We detected single E06-positive degenerating neurons
in an aged control animal. scale bars = 50 µm except a, b = 0.5 mm
Analysing macrophage activation in lumbar spinal cord lesions of CD4+ transfer EAE, we
observed comparable expression of p22phox and iNOS at the acute phase of EAE in young and
old animals (Fig. 17).
Results 33
+
Figure 17: The expression of p22phox and iNOS in acute CD4 T cell-mediated EAE in young and
old animals.
Quantification of p22phox and iNOS expression by optical densitometry of spinal cord slides in young (2
months) and old (14 months) acute EAE animals did not reveal any significant differences (young n = 6,
old n = 8).
We analysed iron deposition in lumbar spinal cords of young and old Lewis rats at the
acute and the recovery stage of EAE. In inflammatory lesions, very few macrophages showed
iron accumulation. Quantification revealed the highest number of iron-accumulating cell clusters
in old animals at the late stage of disease (Fig. 18a). The TBB+ cells were mostly perivascular
and had a macrophage-like morphology (Fig. 18b)
Results 34
+
Figure 18: Iron deposition in CD4 MBP-specific T cell transfer EAE in young and old Lewis rats.
Quantification of TBB stainings of spinal cord sections revealed that old animals at the recovery stage of
EAE show the highest number of iron deposits compared with all the other groups (a; young acute n = 6,
young late n = 8, old acute n = 5, old late n = 6; p ≤ 0.001). A representative picture of iron deposition in
the spinal cord at the late stage of EAE from an old animal is depicted (b). scale bar = 50 µm
In order to study the effect of iron on neurodegeneration, we additionally quantified
inflammation and neurodegeneration in the iron accumulating hotspots in passive CD4+ T cell
transfer EAE of animals with 2 and 14 months of age. Generally, inflammation was less
pronounced in the brain than in the spinal cord. We analysed T cell and macrophage/microglia
activation and neurodegeneration in the acute (day 5 after T cell transfer) and the recovery
phase (day 13 after T cell transfer, data not shown). Analysis of the single iron accumulating
areas in the brain of EAE animals did not reveal any pronounced differences in infiltrating cells
and the number of APP+ axonal spheroids was very low (Fig. 19).
Results 35
+
Figure 19: Inflammation and neurodegeneration in iron-accumulating hotspots in CD4 T cell
transfer EAE.
2
+
The number of T cells per mm (a), the area fraction covered with ED1 macrophages/microglia (b) and
+
2
the number APP axonal spheroids per mm (c) were analysed for each hotspot of iron accumulation. We
studied 6 young (2 months) acute and 8 old (14 months) acute EAE animals. Depending on the section´s
plain the n numbers of the single hotspots differed (young: CPU n = 5, IP = 4, ND = 1, POA = 3, SN = 5;
old: CPU = 8, IP = 7, ND = 5, POA = 7, SN = 8). Quantification of T cells, macrophages and axonal
spheroids in iron accumulating hotspots did not reveal major differences between young and old animals.
As iron was shown to accumulate in neurodegenerative diseases as Alzheimer´s and
Parkinson´s disease and MS (Adams, 1988; Craelius et al, 1982; LeVine, 1997; Loeffler et al,
1995), we densitometrically quantified iron in the hotspots in old EAE and control animals. We
did not find any significant differences in the optical density of TBB stainings between EAE and
control animals (Fig. 20).
Results 36
Figure 20: Optical densitometry of TBB stainings of iron accumulating hotspots in old EAE and
control animals.
Quantification of TBB staining of iron hotspots of aged control animals, old acute EAE animals and old
animals in the recovery phase did not reveal any differences. We studied 4 aged controls, 8 acute EAE
and 6 recovery phase EAE animals. Depending on the section´s plain the n numbers of the single
hotspots differed (control: CPU n = 4, IP n = 3, ND n = 3, POA n = 4, SN n = 4; acute EAE: CPU n = 8, IP
n = 6, ND n = 5, POA n = 7, SN n = 8; recovery phase: CPU n = 6, n = IP 5, ND n = 5, POA n = 6, SN n =
5).
In summary, we compared MBP-specific CD4+ T cell transfer EAE in 2 and 14 months
old Lewis rats and found comparable EAE courses. We detected a higher incidence of T cell
infiltration in young animals, comparable macrophage infiltration but a higher number of APP+
axonal spheroids in lumbar spinal cords of aged animals. Moreover, we found an increased
number of iron-positive macrophages in old animals in the recovery phase of EAE. In the brain,
we generally found low grade inflammation and neurodegeneration with no difference in the
optical density of iron. Our findings about the expression of p22phox and iNOS and the
occurrence of oxidised phospholipids (E06) and iron accumulation are summarised in Table 1
(page 48).
In order to expand our study to a larger spectrum of different models for CNS
inflammation, we analysed archival material that had been used in former studies done in
collaboration with our laboratory. We investigated oxidative injury, oxidative burst and iron
accumulation in different rodent models that are based on a variety of disease mechanisms and
phenotypes.
Results 37
2.3 Chronic relapsing MOG-induced EAE in DA rats
Active sensitisation of DA rats with MOG1-125 is to date the model reflecting MS pathology
most closely (Storch et al, 1998). MOG immunisation first resulted in extensive primary
inflammatory demyelination (Fig. 21a-c, g) which was followed by a chronic relapsing disease
(Fig. 21d-f, h) (Storch et al, 1998). In initial stages of active lesions, mostly granulocytes but also
macrophages expressed p22phox (Fig. 21b) and few macrophages stained positively for iNOS
(Fig. 21g). In chronic active lesions, despite massive on-going demyelination (Fig. 21d) and the
presence of activated macrophages/microglia expressing ED1 (Fig. 21f), p22phox or iNOS
reactivity was largely absent (Fig. 21e, h). We did not observe microglia nodules in the NAWM.
Furthermore, animals suffering from MOG-induced chronic relapsing EAE did not accumulate
oxidised phospholipids in active or inactive lesions (Fig. 21i) with one exception. In one animal
out of 20, we found small clusters of E06+ neurons in the spinal cord anterior horn showing also
central chromatolysis (Fig. 21i, insert). Control animals did not show any TBB staining in brain
and spinal cord tissue but in most EAE animals we detected iron accumulation in perivascular
macrophages in the lesions (Fig. 21j). Our findings about the expression of p22phox and iNOS
and the occurrence of oxidised phospholipids (E06) and iron accumulation in MOG-induced EAE
in DA rats are summarised in Table 1 (page 48).
Results 38
Figure 21: Oxidative burst, oxidative injury and iron accumulation in chronic relapsing EAE of
MOG-immunised DA rats.
Initial lesions showed a characteristic perivenous pattern of demyelination (a) with clear p22phox (b) and
iNOS (g) reactivity. Mainly granulocytes expressed p22phox (b, insert). The chronic phase of the disease
was characterised by extensive active demyelination in the spinal cord grey and white matter (d) with the
presence of macrophages containing myelin degradation products (d, insert). These lesions comprised of
+
high numbers of ED1 macrophages (f) but their majority did neither express p22phox (e) nor iNOS (h).
Only few macrophages in the lesions expressed p22phox (e, insert). The vast majority of lesions did not
accumulate oxidised phospholipids (i). In one out of 20 animals, only individual neurons with
+
morphological evidence for retrograde degeneration were E06 (i, insert). Single macrophages and
microglia in the lesions accumulated iron (j). scale bars = 0.5 mm except for g-h = 50 µm
Results 39
2.4 Chronic relapsing MOG-induced EAE in C57BL/6 mice
Active MOG35-55-immunization of C57BL/6 mice represents the most frequently used
animal model for MS and is associated with profound demyelination and extensive axonal injury
and loss in the chronic disease stage. Inflammation is dominated by perivenous T cell and
macrophage infiltration, as well as microglial activation (Mendel et al, 1995; Taoufik et al, 2011).
In our model, acute disease started 10 to 15 days after MOG-sensitization with pathology
resembling the disease peak of acute MBP-specific T cell transfer EAE. After a short remission,
the animals suffered from a relapsing disease with a progressive deterioration. Confluent
demyelinating plaques appeared in the brain and the spinal cord around day 20 and propagated
until day 34. Demyelination was associated with axonal injury and loss. In the present study, we
evaluated animals suffering from active demyelination and neurodegeneration at day 21, 27 and
35 after MOG-immunization (Taoufik et al, 2011). Tissue damage was characterised by
infiltration of CD3+ T cells (Fig. 22a) and severe microglia activation (Fig. 22b). The expression
of p22phox was confined to cells with a macrophage-like morphology (Fig. 22c), whereas
activated Iba-1+ microglia around the lesion were devoid of p22phox staining. Moreover, we
could not detect any microglial nodules in the NAWM (Fig. 22b, c). We identified single cells with
a macrophage-like morphology expressing iNOS (Fig. 22d). A minor reactivity for oxidised
phospholipids (E06) was observed in few dystrophic axons (Fig. 22e). We did not find any iron
accumulation in oligodendrocytes or myelin, but only in scattered iron-containing perivascular or
meningeal macrophages in individual lesions (Fig. 22f). Our findings about p22phox and iNOS
expression and the accumulation of oxidised phospholipids (E06) and iron in MOG-induced EAE
in C57BL/6 mice are summarised in Table 1 (page 48).
Results 40
Figure 22: Oxidative burst, oxidative injury and iron accumulation in chronic relapsing EAE of
MOG-immunised C57BL/6 mice.
+
+
Active lesions were associated with the infiltration of CD3 T cells (a) and Iba-1 cells with a macrophageand microglia-like morphology (b). Numerous macrophages in the lesions expressed p22phox (c) while
positive iNOS staining was restricted to few perivascular cells (d). E06 reactivity (oxidised phospholipids)
was only detected in single axonal spheroids (e, insert). Scattered lesion-associated meningeal
macrophages accumulated iron (f).
2.5 Inflammatory demyelination induced by cytotoxic T cells
Inflammation in MS lesions is dominated by CD8+ cytotoxic T cells (Babbe et al, 2000).
Hence, we studied oxidative injury in a CD8+ T cell-driven EAE model (Saxena et al, 2008)
characterised by demyelination (Fig. 23a) and CD3+ T cell infiltration (Fig. 23c). Microglial
activation was evident (Fig. 23b, d) but p22phox was not expressed on microglia in the NAWM.
In and around the lesions, a considerable proportion of cells with a microglia as well as a
macrophage morphology expressed p22phox (Fig. 23e). We detected iNOS immunoreactivity in
restricted perivascular macrophages (Fig. 23f). Microglial nodules were absent from the NAWM
(Fig. 23b). Similarly to MOG-induced EAE, we did not detect any hint for oxidised phospholipids
in CD8+ T cell-mediated CNS inflammation (Fig. 23g). Iron accumulation was absent from the
NAWM of the brain and the spinal cord as well as the majority of active lesions (Fig. 23h). In
single lesions, perivascular macrophages accumulated iron in 4 out of 7 animals studied (Fig.
23i). Our results about p22phox and iNOS expression and the occurrence of oxidised
phospholipids (E06) and iron accumulation in CD8+ T cell-induced EAE in DA rats are
highlighted in Table 1 (page 48).
Results 41
+
Figure 23: Oxidative burst, oxidative injury and iron accumulation in CD8 T cell-mediated EAE in
mice.
Demyelination (a) and perivascular T cell infiltration (c) were associated with profound microglial activation
(b, d). Perivascular macrophages and to a lower degree also surrounding microglia (both based on
+
morphological hallmarks) expressed p22phox (e) while only restricted perivascular cells were iNOS (f).
There was no immunoreactivity for oxidised phospholipids (E06) detectable (g). Iron accumulation
occurred only in single lesions in perivascular macrophages (i). scale bars = 50 µm except for a, b = 0.5
mm
Results 42
2.6 Innate immunity-driven inflammatory demyelinating lesions
A major role for innate immune mechanisms in the pathogenesis of MS has been
suggested (Barnett & Prineas, 2004; Marik et al, 2007). Consequently, we analysed
inflammatory demyelinating lesions induced by the focal injection of bacterial LPS into the spinal
cord white matter of SD rats (Felts et al, 2005). At early stages of inflammation, some
macrophages and microglia were shown to express iNOS (Marik et al, 2007). In these initial
lesions, p22phox was expressed mainly by granulocytes (Fig. 24g), which were succeeded by
p22phox+ macrophages during lesion progression (not shown). Demyelination and axonal injury
were observed 7 to 9 days after LPS injection and were most pronounced after 12 to 15 days
(Felts et al, 2005). At this disease stage, active demyelinating lesions (Fig. 24a) were
characterised by massive microglial activation (Fig. 24b, c, e) and low numbers of CD3+
infiltrating T cells (Fig. 24d). Mostly macrophages and few cells with microglial morphology
expressed p22phox in the lesions (Fig. 24f) whereas in the lesion centres p22phox expression in
macrophages was minor (Fig. 24h). Microglial nodules were absent from the NAWM. We
detected rare iNOS expression in cells with microglia and macrophages morphology (Fig. 24i). In
lesions at day 12 after LPS injection, myelin was very weakly positive for E06 (Fig. 24j). Iron
stainings showed to be largely negative besides individual perivascular iron-positive
macrophages in the lesions (Fig. 24k). The results of our stainings for p22phox, iNOS, E06 and
iron accumulation in LPS-injection-driven CNS inflammation are summarised in Table 1 (page
48).
Results 43
Figure 24: Oxidative burst, oxidative injury and iron accumulation induced by LPS injection into
the spinal cord dorsal column of SD rats.
At the peak of active demyelination (a) (12 days after LPS injection), lesions showed massive macrophage
+
infiltration (b, c, e) but only individual CD3 T cells (d). In these lesions, macrophages and single cells with
a microglial morphology at the lesion edges expressed p22phox (f). They were preceded by p22phox
+
granulocytes in the initial disease stages (g, 1-3 days after LPS injection). In the lesion centres, p22phox
expression on macrophages was minor (h). In contrast to pronounced p22phox expression in active
lesions, very little iNOS reactivity was detected (i). Oxidised phospholipids appeared to a minor extent at
the lesions (j). Iron positive macrophages were sparse (k). scale bars = 50 µm except for a, b = 0.5 mm
2.7 Toxic cuprizone-induced demyelination
Cuprizone-supplemented diet causing demyelination in mice is regarded as an adequate model
of demyelination in MS (Kipp et al, 2009). Actively demyelinating lesions (Fig. 25a) in the corpus
callosum 35 days after disease induction exhibited oligodendrocyte apoptosis, microglial
Results 44
activation and astrocyte gliosis (Van Strien et al, 2011). The lesions contained Mac-3-expressing
macrophages, which indicates active phagocytosis, and cells with microglial morphology (Fig.
25b). In contrast, p22phox reactivity was completely absent (Fig. 25c) and iNOS was only
expressed by individual cells with microglial morphology (Fig. 25d). Neither stainings for oxidised
phospholipids (Fig. 25e) nor for iron (Fig. 25f) revealed any positive signal. Our findings about
the expression of p22phox and iNOS and the deposition of oxidised phospholipids (E06) and
cuprizone diet-induced demyelination are summarised in Table 1 (page 48).
Figure 25: Oxidative burst, oxidative injury and iron accumulation in cuprizone diet-induced active
demyelination in mice.
Loss of myelin in the corpus callosum (a) was associated with the presence of phagocytosing
macrophages and microglia (b). In contrast, macrophages did not express p22phox (c) and iNOS only to a
minor extent (d). We detected no deposition of oxidised phospholipids (e, E06) or iron (f). scale bars = 50
µm
2.8 MHV-JHM coronavirus-induced encephalomyelitis
Virus infections have been suggested to be one of the possible risk or causative factors
for the development of MS (Lipton et al, 2007). We analysed CNS lesions of Lewis rats infected
with MHV-JHM mouse hepatitis virus which leads to a complex immune response comprising
CD8+ T cells (Dandekar et al, 2004; Pewe & Perlman, 2002) and specific anti-viral antibodies
(Lin et al, 1999; Tschen et al, 2002; Zimprich et al, 1991). Intracerebral injection of MHV-JHM
coronavirus (Barac-Latas et al, 1997; Korner et al, 1991; Wege et al, 1998; Zimprich et al, 1991)
induced an initial panencephalitis characterised by profound CNS inflammation and tissue
Results 45
damage and virus spread in neurons and glial cells (Korner et al, 1991; Zimprich et al, 1991).
This initial disease was succeeded by a subacute or a chronic disease. In the chronic stage,
virus infection was mainly found in glial cells associated with demyelination and axonal damage
(Fig. 26a-f) (Korner et al, 1991; Wege et al, 1998; Zimprich et al, 1991). Inflammation was mainly
mediated by CD8+ T cells (Fig. 26d, i) and activated macrophages and microglia (Fig. 26i, j, k).
Microglial activation had been shown to be profound throughout the CNS in all disease stages
(Korner et al, 1991; Zimprich et al, 1991). In this virus-induced model, we observed microglial
nodules in the NAWM that expressed Iba-1 and p22phox (Fig. 26j-m). The density of microglia
increased towards the lesion edge (Fig. 26k, left side), was at a maximum at the area of initial
demyelination (Fig.26k, centre) and decreased towards the lesion centre (Fig. 26k, right side). In
the lesion centre, many cells exhibited macrophage morphology. Immunohistochemical stainings
for p22phox revealed a staining pattern very reminiscent of the Iba-1 staining in the NAWM (Fig.
26l) and the lesion (Fig. 26m). The expression of p22phox was most prominent in active lesions
and the surrounding NAWM, but was minor on macrophages in the inactive lesion centre (Fig.
26l, m). Stainings for iNOS were largely negative and, if positive, only found in individual cells
with macrophage morphology (Fig. 26n). Strikingly, we detected intense E06 staining within
active lesions (Fig.26o, p, q) and to a lesser extent in the NAWM. Additionally, we detected
positive stainings for oxidised DNA (8OHdG) in the coronavirus-induced lesions (Fig. 26r). Iron
deposition in the lesions was restricted to perivascular macrophages. Our observations
concerning the expression of p22phox and iNOS and the accumulation of oxidised lipids and
iron in MHV-JHM coronavirus-induced CNS disease are highlighted in Table 1 (page 48).
Results 46
Results 47
Figure 26: Oxidative burst, oxidative injury and iron accumulation in MHV-JHM coronavirusinduced encephalomyelitis in Lewis rats.
Virus infection induced extensive demyelination (a) with relative axonal preservation (b, f). Virus antigen
was expressed in the periplaque white and grey matter (c) in neurons (g) and in glial cells in the white
matter (h). Active lesions were characterised by T cell-mediated inflammation (d), the majority being CD8
+
+
(blue; i) and ED1 macrophages (brown, i). Active demyelination was associated with severe microglial
activation (k). In the NAWM, the activated microglia clustered to form microglia nodules (j) expressing also
p22phox (l). Microglia numbers increased towards the lesion edge (k; left side), reached a peak at areas
of initial demyelination (k, centre) and decreased in the lesion centres, where most cells were of
macrophage morphology (k; right side). Staining for p22phox in serial-cut sections exhibited a staining
pattern very reminiscent of the result of the Iba-1 stainings (l, m). iNOS expression was rare or absent (n).
Oxidised phospholipids (E06) accumulated in the lesions in myelin (o), in cells with apoptotic nuclei, in
macrophage granules (p) and in axonal spheroids (q). Additionally, evidence for oxidised DNA (8OHdG)
was discovered (r). Most lesions were negative for iron staining (s), although we found iron deposition in
individual lesions in perivascular macrophages. scale bars = 50 µm except for a-d = 0.5 mm
Results 48
Table 1: Quantification of markers for inflammation and oxidative injury in different experimental models for CNS inflammation and MS
model
acute pt EAE young
acute pt EAE aged
chronic MOG EAE mouse
chronic MOG EAE rat
CD8 EAE
LPS injection
cuprizone diet
coronavirus MHV-JHM
encephalomyelitis
acute MS
young control animals
old control animals
human controls
mechanism
CD4+ T cells
CD4+ T cells
CD4+ T cells
CD4+ T cells +
demyelinating antibodies
CD8+ T cells
innate immunity
toxic demyelination
virus, CD8+ T cells,
innate immunity
unknown
n.p.
n.p.
n.p.
Iba-1
10.90 (5.23)
8.59 (4.87)
11.12 (17.66)
p22phox
10.46 (3.66)
9.10 (3.83)
3.28 (8.33)
active lesion
iNOS
E06
2.25 (2.10)
122 (44)
2.99 (1.62)
130 (37)
0.02 (0.56)
185 (46)
9.33 (16.18)
0.30 (3.90)
0.01 (0.06)
124 (172)
0.01 (0.73)
7.58 (7.42)
6.90 (17.30)
11.97 (7.03)
1.17 (2.53)
0.80 (3.70)
0.04 (0.04)
0.00 (0.31)
0.02 (0.86)
0.00 (0.00)
233 (97)
200 (35)
130 (46)
0.05 (0.32)
0.01 (1.00)
0.01 (0.01)
15.81 (13.16)
3.79 (4.73)
0.00 (0.09)
644 (138)
0.01 (0.70)
12.60 (18.86)
8.69 (12.12)
Iba-1
p22phox
0.47 (2.58)
553 (313)
NWM/NGM
iNOS
E06
2.55 (1.69)
3.28 (2.98)
4.41 (5.58)
0.00 (0.01)
0.00 (0.01)
a
2.47 (1.31)
0.00 (0.00)
0.00 (0.00)
a
0.04 (0.02)
125 (48)
110 (39)
a
336 (132)
iron
0.00 (0.01)
0.00 (0.00)
0.18 (0.92)
*
iron
BG
SC
0.03 (0.04)
2.94 (2.08)
*
0.01 (0.02)
0.01 (0.02)
*
Quantification of Iba-1, p22phox, iNOS, oxidised phospholipids (E06) and iron staining in active lesions of different rodent models for CNS
inflammation in comparison with acute MS cases and human controls. Depicted are values (median (range) derived from optical densitometry (area
fraction for Iba-1, p22phox, iNOS and iron and integrated density for E06) of equally sized pictures taken under standardised conditions of the
respective animal model or MS case. Bold numbers indicate a significant increase compared to control animals, or in case of MS compared to human
controls, using Mann-Whitney U post hoc tests and Bonferroni-Holm correction. In case of iron staining, lesions were compared with the respective
control tissue.
a
indicates a significant increase compared to animal controls. Iron staining in basal ganglia (BG) was significantly increased in old
compared with young controls. * Iron accumulation in MS and human controls was not quantified in the present study, as it is dependent on location
and age as described elsewhere (Hametner et al, 2013). NWM = normal white matter; NGM = normal grey matter; EAE = experimental autoimmune
encephalomyelitis; pt = passive transfer; BG = basal ganglia; SC = spinal cord; LPS = lipopolysaccharide; n.p. = not present; (young control animals n
= 6; old control animals n = 6; acute EAE young n = 6; acute EAE aged n = 8; chronic MOG EAE mouse n = 15; chronic MOG EAE rat n = 14; CD8
EAE n = 6; LPS injection n = 20; cuprizone diet n = 5; coronavirus encephalomyelitis n = 11; acute MS n = 7; human controls n = 6)
Results 49
2.9 Theiler's Murine Encephalomyelitis Virus (TMEV)-induced disease
In order to extend our results on animals with a chronic virus-triggered disease, we
studied a second established virus-induced model for CNS inflammation, the Theiler´s murine
encephalomyelitis virus (TMEV) infection (Lipton, 1975; Rodriguez et al, 1986). Intracerebral
infection with TMEV resulted in a biphasic disease in susceptible mouse strains (Daniels et al,
1952; Lipton, 1975; Lipton & Dal Canto, 1979). An early phase of acute encephalitis with primary
demyelination was succeeded by chronic demyelinating disease (Dal Canto & Lipton, 1975;
Drescher et al, 1997). We analysed CNS lesions of SJL mice infected with TMEV from day 3 to
220 after virus injection. Virus infection caused a complex immune response involving cytotoxic
T cells (Ruby & Ramshaw, 1991; Zinkernagel & Althage, 1977), TH1 helper T cells (Lipton et al,
2005), natural killer cells (Paya et al, 1989), macrophages (Lipton et al, 2005) and antibody
production (Rodriguez et al, 1988a; Roos et al, 1987). Intracerebral injection of TMEV induced
encephalitis (Fig. 27a-g) including profound CNS inflammation and tissue damage and virus
localisation in neurons (Fig. 27d) (Dal Canto & Lipton, 1982). In this initial disease stage at 3
days post injection (dpi), the injection site was devoid of Iba1+ cells (Fig. 27a) and iNOS
expression (Fig. 27c) but in the NAWM microglia showed signs of activation (Fig. 27a, insert).
The injection site was related with profound p22phox expression primarily in granulocytes (Fig.
27b). Initial tissue damage was associated with E06 immunoreactivity in macrophages (Fig. 27e,
f). The iron deposition occurred due to leakage of blood upon injection trauma (Fig. 27g).
Results 50
Figure 27: Oxidative burst, oxidative injury and iron accumulation in TMEV-infected mice 3 dpi.
+
The injection site was devoid of Iba1 cells (a). In the NAWM, microglia showed an activated morphology
(a, insert). Mainly granulocytes expressed p22phox (b) whereas we found no iNOS reactivity (c). In this
early disease stage, the virus infected neurons (d). Tissue damage at the injection site was associated
with E06 immunoreactivity in macrophages (e, f). Iron deposition occurred at the injection site due to
bleeding (g). scale bars = 50 µm.
With progression of the disease (45 dpi), inflammation and tissue damage spread to the
spinal cord (Fig. 28a-h) where mainly glial cells were virus-infected (Fig. 28d) (Aubert et al,
1987; Dal Canto & Lipton, 1982; Lipton, 1975; Lipton et al, 2005; Rodriguez et al, 1983).
Inflammation was mediated by activated macrophages and microglia (Fig. 28b, c, e) (Mack et al,
2003). Similarly to coronavirus-induced disease, TMEV infection caused the formation of
microglial nodules in the NAWM. They expressed Iba-1 (Fig. 28b, c) but, in contrast to
coronavirus-infected animals, not p22phox. The number of microglial nodules (Fig. 28c)
increased with disease duration and reached a peak at 120 dpi (Fig. 29). The expression of
p22phox was minor although macrophages and microglia showed signs of phagocytosis (Fig
28e, Mac-3 expression). Lesions were largely iNOS negative. Sparse iNOS reactivity was only
found in individual cells with macrophage morphology (Fig. 28g). Iron deposition in the lesions in
spinal cord (Fig. 28h) and brain (Fig. 28i) was restricted to perivascular macrophages.
Results 51
Figure 28: Oxidative burst, oxidative injury and iron accumulation in TMEV-infected mice 45 dpi.
Demyelination and inflammation spread to the spinal cord (a-h). Demyelination was associated with
profound microglia activation (b) and the formation of microglial nodules (c). Virus infection was mainly
found in cells with glial morphology (d). Activation of macrophages and microglia, indicated by the
expression of Mac-3, was not associated with the expression of p22phox (f) or iNOS (g). Iron accumulated
in individual perivascular macrophages in spinal cord (h) and brain (i) lesions. scale bars 0 50 µm except
for a, b = 0.5 mm
Results 52
Figure 29: Quantification of microglial nodules in TMEV-induced inflammatory demyelinating
disease.
The number of microglial nodules increased with disease duration. At 35 dpi, we counted significantly
more microglial nodules (p = 0.044 indicated by *) than 3 dpi. The number of microglial nodules reached a
peak 120 days dpi (p = 0.006 compared to 3 dpi, indicated by **; p = 0.03 compared with unsusceptible
animals 45 dpi, indicated by *), (n = 5 except for 45 dpi n = 4).
During chronic disease (90 dpi), the animals exhibited profound lesions in the medulla
(Fig. 30) characterised by demyelination (Fig. 30a) and microglial activation (Fig. 30b). Despite
morphological hints for microglial activation, we did not detect p22phox (Fig. 30c) or iNOS (Fig.
30d) expression. In contrast, the lesions were associated with E06 immunoreactivity (Fig. 30e, g,
h). Quantification of E06 optical density of medullary lesions revealed an increase of oxidised
phospholipids with disease duration with first significant results at 90 dpi (p ≤ 0.001) compared
with TMEV-injected but non-susceptible animals at 45 dpi (Fig. 31). Spinal cord lesions only
partly accumulated oxidised phospholipids (Fig. 30i, j)
Results 53
Figure 30: Oxidative burst, oxidative injury and iron accumulation in TMEV-infected mice 90 dpi.
Animals exhibited profound demyelination (a) and inflammation (b), especially in lesions in the medulla.
Inflammation did not co-localise with the expression of p22phox (c) or iNOS (d). Oxidised phospholipids
accumulated in macrophages in medullary lesions (e, g, h). Spinal cord lesions were only partly
associated with E06 immunoreactivity (i, j). Iron accumulated infrequently in individual perivascular
macrophages (f). scale bars = 50 µm.
Results 54
Figure 31: Optical densitometry of E06 immunoreactivity in medullary lesions of TMEV-infected
mice at different disease stages.
Densitometric quantification of equally sized regions of interest taken under standardised conditions (for
example Fig. 30e) revealed an increase of oxidised phospholipids with disease duration (p ≤ 0.001 for 90
dpi, p = 0.004 for 120 dpi, p = 0.005 for 220 dpi when compared with 45 dpi non-susceptible control;
indicated by *** or ** respectively).
Later chronic disease stages were characterised by pronounced lesion formation in the
spinal cord (180 dpi, Fig. 32), the cerebellum and the medulla (220 dpi, Fig. 33). Demyelinating
lesions in the spinal cord (Fig. 32a) were associated with virus-infected cells (Fig. 32b), activated
macrophages and microglia (Fig. 32c, Mac-3) and T cell infiltration (Fig. 32d). Cells with
macrophage morphology stained weakly for p22phox (Fig. 32e). iNOS was only expressed in
individual macrophages (Fig. 32f). Only single perivascular macrophages accumulated iron (Fig.
32g).
Results 55
Figure 32: Oxidative burst, oxidative injury and iron accumulation in TMEV-infected mice 180 dpi.
Spinal cord lesions displayed demyelination (a) and virus infection in glial cells (b). The lesions were
associated with activated macrophages (c) and T cell infiltration (d). Inflammation was related with weak
+
p22phox expression in cells with macrophage morphology and with individual iNOS macrophages (f). The
majority of lesions was devoid of iron deposition (g). scale bars = 50 µm except for a-c = 0.5 mm.
The profound lesions in the cerebellum and the medulla at late disease stages (day 220
dpi, Fig. 33) were associated with T cell infiltration (Fig. 33a) and macrophage and microglia
activation (Fig. 33b, c). The lesions were largely negative for p22phox (Fig. 33d) and iNOS (Fig.
33e) expression. Oxidised phospholipids were detected in macrophages (Fig. 33g). Only single
cells stained positively for virus antigen (Fig. 33h). Iron accumulation was sparse in the lesions
(Fig. 33f) but was detected in oligodendrocytes and myelin in the basal ganglia (Fig. 33i).
Results 56
Figure 33: Oxidative burst, oxidative injury and iron accumulation in TMEV-infected mice 220 dpi.
Cerebellar and medullary lesions were associated with T cell infiltration (a) and macrophage/microglia
activation (b, c). Inflammation was mainly not related with p22phox or iNOS reactivity (d, e). Lesionassociated macrophages accumulated oxidised phospholipids (g). Only individual cells were virus-infected
(h). The majority of lesions was devoid of iron deposition (f) but iron accumulated in oligodendrocytes and
myelin of the basal ganglia (i). scale bars = 50 µm except for a-c = 0.5 mm.
Results 57
2.10 In vitro oxidation
Although antibodies recognising oxidation-specific epitopes had been used before in
rodent tissue (Chou et al, 2009; Horkko et al, 1996; Palinski et al, 1996), we tested if the
epitopes could be produced and detected in our experimental settings. For that purpose, we
oxidised native slides of fresh frozen rat brain in vitro and stained for oxidised phospholipids
(E06) or for oxidised DNA (8OHdG). We found that the epitope for E06 could be produced by
iron-induced lipid peroxidation (Ribeiro et al, 2007; Takatsu et al, 2009). Densitometric
quantification of equally sized regions of interest taken under standardised conditions (Fig. 34a)
revealed an increase of oxidised phospholipids upon iron treatment (Fig. 34b). This could not be
enhanced by addition of H2O2.
Figure 34: E06 staining after in vitro oxidation.
The E06 epitope of oxidised phospholipids could be generated by in vitro oxidation of fresh-frozen native
brain sections via free-radical generating systems containing iron (c, e). Densitometric quantification of
E06 staining revealed an increase of E06 staining intensity in iron-treated compared with control sections
(f; n = 6, p ≤ 0.001, indicated by ***)
Depicted are representative unmodified pictures taken under standardised conditions prior to
densitometric quantification. a = untreated (TBS control), b = ascorbate, c = FeSO4/ascorbate, d = H2O2, e
= FeSO4/ascorbate + H2O2; scale bar = 50 µm
Results 58
After using a similar iron-induced peroxidation system (Lodovici et al, 2001), we also
detected oxidised DNA (8OHdG) by immunohistochemistry. DNA oxidation was induced by
incubating native brain slides with the most effective treatment being iron chloride (Fig. 35g) and
iron sulphate (Fig. 35e, f) compared to controls (Fig. 35a-d). In summary, both epitopes shown
to occur in MS tissue could be produced by iron-induced in vitro oxidation and detected with
immunohistochemistry on rat brain tissue.
Figure 35: 8OHdG staining after in vitro oxidation.
The epitope of oxidised DNA could be generated by in vitro oxidation of native brain cryo-sections via
free-radical generating systems containing iron. Iron chloride treatment (g) was most effective compared
to iron sulphate treatments (e, f) and controls (a-d). a = untreated (TBS control), b = ascorbate, c
=glutathione (GSH), d = H2O2, e = FeSO4/ascorbate, f = FeSO4/ascrobate + H2O2, g = FeCl/GSH; scale
bar = 50 µm
Results 59
2.11 Iron loading of glial cell cultures
Iron accumulation in the studied rodent models for CNS inflammation appeared to be
minimal compared to the human brain. As they were not appropriate to study the effect of iron on
neurodegeneration, we chose an in vitro approach. Iron loading of glial cells is established to
study iron management in the brain (Rathore et al, 2012; Schulz et al, 2011). In preliminary
experiments, we treated purified oligodendrocyte cultures with iron chloride together with
ascorbate (in a relation of 1:44, as published (Rathore et al, 2012; Schulz et al, 2011)) and found
iron accumulation as visualised by TBB staining (Fig. 36).
Figure 36: Iron loading of purified oligodendrocyte cultures.
Treatment of oligodendrocyte cultures with iron chloride (10 µM FeCl3) for 6 h resulted in iron
accumulation, as depicted by double staining of TBB with galactocerebroside (GalC, marker for
oligodendroglial lineage) (b) compared to untreated cells (a). scale bar = 50 µm
As it was published that cytokine toxicity in oligodendrocytes could be enhanced by iron
(Zhang et al, 2005), our aim was to study the effect of iron accumulation on inflammatory
demyelination and secondary neurodegeneration. For this purpose, we chose the in vitro model
of myelinating spinal cord cultures (Elliott et al, 2012; Sorensen et al, 2008). These complex
long-term cultures comprised a confluent feeder layer of neurosphere-derived astrocytes (Fig.
37a, GFAP+ cells), on top of which neurons spread (Fig. 37b, SMI31+ cell processes) that were
partly surrounded by mature myelin sheaths (Fig. 37b, MOG+ myelin). On top of this, the cultures
also contained microglia (Fig. 37c, Iba-1+ cells). Our aim was to primarily load the
oligodendrocytes in these cultures with iron in order to study the impact of iron on demyelination
and possibly secondary neurodegeneration.
Results 60
Figure 37: Characteristics of myelinating spinal cord cultures
Mature myelinating spinal cord cultures (28 days after cell seeding) contained a confluent layer of
+
+
neurosphere-derived astrocytes (a, GFAP ) on top of which neurons spread (b, SMI31 ) that were partly
+
myelinated (b, MOG mature myelin sheaths). A considerable number of microglia covered the cultures (c,
+
Iba-1 ). scale bar = 50 µm
For iron loading of myelinating spinal cord cultures, we considered two experimental
strategies. Treatment with iron chloride (Rathore et al, 2012; Schulz et al, 2011) was shown to
cause iron accumulation in purified glial cultures. We confirmed this in our preliminary results for
oligodendrocytes (Fig. 36). Our second approach was treating the myelinating spinal cord
cultures with ferritin, which was shown to be a major source for iron for oligodendrocytes in vitro
and in vivo (Schonberg et al, 2012; Todorich et al, 2011).
To target our first strategy, we tested the effect of different iron chloride dosages on the
integrity of myelinating spinal cord cultures by staining for mature oligodendrocytes and myelin
(PLP) and neurons (SMI31) (Fig. 38). Cultures treated with 250 µM iron chloride (Fig. 38b)
appeared comparable to control treated cells (Fig. 38a). In contrast, cultures treated with higher
dosages (500 or 750 µM) showed signs of cell damage, as for example disturbed transport of
neuronal processes (Fig. 38c, d; SMI31+ swellings).
Results 61
Figure 38: Dosage-effect of iron chloride on the integrity of myelinating spinal cord cultures.
The integrity of myelinating spinal cord cultures was primarily assessed by staining for mature
oligodendrocytes and myelin (PLP) and neuronal processes (SMI31). Myelinating spinal cord cultures
treated with ascorbate only (control, a) and 250 µM FeCl3 appeared similar, whereas the cultures integrity
+
was disturbed by higher dosages (c, 500 µM, d, 750 µm) as reflected by swellings of SMI31 neuronal
processes. All treatments were performed overnight. scale bar = 50 µm
We continued further experiments by treating the myelinating spinal cord cultures
overnight with 250 µM iron chloride. In order to assess iron accumulation, we performed TBB
stainings which revealed TBB+ structures after iron chloride treatment (Fig. 39a) compared to
ascorbate (Fig. 39b) and untreated (Fig. 39c) controls.
Results 62
Figure 39: Iron accumulation in myelinating spinal cord cultures after iron chloride treatment.
Iron accumulation in myelinating spinal cord cultures was visualised by TBB staining. Overnight treatment
+
with 250 µM FeCl3 (a) resulted in an increase of TBB structures compared to ascorbate (b) and untreated
(c) control. scale bar = 50 µm
In order to identify the iron accumulating cell type, we performed double labellings with
oligodendrocyte and microglia markers and found that the only staining co-localising with TBB
was Iba-1 (Fig. 40a, TBB was converted from the brown TBB depicted in Fig.40b to the false
colour blue in order to enable overlay with fluorescent stainings). In control-treated cultures, we
did not detect any iron accumulation (Fig. 40c, d, the same colour conversion as in Fig.a and b is
depicted).
Results 63
Figure 40: Iron accumulation in microglia in myelinating spinal cord cultures after iron chloride
treatment.
Overnight treatment of myelinating spinal cord cultures with 250 µM FeCl3 caused iron accumulation in
microglia (a, b). In order to enable an overlay of light microscopy and fluorescence, TBB was converted
from brown (b, TBB) to the false colour blue (a). There was no iron deposition in the control-treated
cultures (c, d). scale bar = 50 µm
In conclusion, treating myelinating spinal cord cultures with iron chloride resulted in iron
accumulation in microglia cells but not in oligodendrocytes.
Ferritin was shown to be a major source of iron for oligodendrocytes (Todorich et al,
2011). Moreover, it was transferred from macrophages to oligodendrocyte precursors in vivo
(Schonberg et al, 2012). In myelinating spinal cord cultures, iron accumulation due to ferritin
loading appeared after 48 h of treatment (Fig. 41). TBB+ cells appeared to be microglia (Fig.
41a, in c TBB was converted to the false colour green in order to enable an overlay with Iba-1
staining in d). As representative result for short time treatment 48 h ferritin treatment is depicted,
but 72 h incubation led to a similar result. Control cultures did not show any sign of iron
accumulation (Fig. 41b).
Results 64
Figure 41: Iron accumulation in microglia cells of myelinating spinal cord cultures after short-term
ferritin treatment.
Ferritin treatment (5 µg/ml, 48 h) of myelinating spinal cord cultures induced iron accumulation in microglia
+
(Iba-1 ). TBB staining (a) was converted to the false colour green (c) to enable an overlay with Iba-1 (d).
Control cells did not show iron staining (b). scale bar = 50 µm
Furthermore, we analysed oligodendrocyte and myelin integrity of ferritin-loaded cultures.
As depicted in Fig. 42, PLP+ oligodendrocytes and myelin appeared similar in control (a) and
ferritin-treated cultures (b). Ferritin treatment increased ferritin levels (ferritin light chain, FTL) in
cells with microglial morphology (Fig. 42b) compared to controls (Fig. 42a). FTL staining colocalised with ED1 (Fig. 42d), confirming that microglia accumulated ferritin. We could not detect
FTL+ cells in control-treated cultures (Fig. 42a, c).
Results 65
Figure 42: Oligodendrocyte and myelin integrity and ferritin accumulation after short-term ferritin
treatment.
Ferritin (b, 5 µg/ml, 48 h) did not have a toxic effect on oligodendrocytes or myelin when compared with
control cultures (a). Ferritin treatment induced ferritin accumulation (ferritin light chain, FTL) in cells with
microglial phenotype (b), which was confirmed by double staining of FTL with ED1 (d). Control cultures did
+
not contain FTL cells (a, c). scale bar = 50 µm
In conclusion, one short-term ferritin treatment of myelinating spinal cord cultures caused
ferritin accumulation in microglia. Another possibility to load myelinating spinal cord cultures with
ferritin was treatment over a longer time span. As depicted in Fig. 43, repeated long-term
exposure with ferritin from day 16 to day 23 after cell seeding led again to iron accumulation in
microglia (Fig. 43a, in c TBB was converted to the false colour green in order to enable an
overlay with Iba-1 staining in d). We could not detect any iron accumulation in control-treated
cultures (Fig. 43b).
Results 66
Figure 43: Iron accumulation in microglia of myelinating spinal cord cultures after long-term
ferritin treatment.
Repeated ferritin treatment (1 µg/ml ferritin with every feeding from day 16 until day 23 after cell seeding)
+
of myelinating spinal cord cultures induced iron accumulation in microglia (Iba-1 , d). TBB staining (a) was
converted to the false colour green (c) to enable an overlay with Iba-1 (d). We could not detect iron
staining in control cultures (b). scale bar = 50 µm
Additionally, we analysed oligodendrocyte and myelin integrity of long-term ferritinexposed cultures. As depicted in Fig. 44, PLP+ oligodendrocytes and myelin appeared
comparable in control (a) and ferritin-treated cultures (b). Long-term exposure to ferritin induced
ferritin accumulation in cells with microglial morphology (Fig. 44b) compared to controls (Fig.
44a). Ferritin (FTL) staining co-localised with ED1 (Fig. 44d), confirming that microglia
accumulated ferritin. We could not detect FTL+ cells in control cultures (Fig. 44a, c).
Results 67
Figure 44: Oligodendrocyte and myelin integrity and ferritin accumulation after long-term ferritin
treatment.
Repeated exposure to ferritin (b, 1 µg/ml ferritin with every feeding from day 16 until day 23 after cell
+
seeding) did not have a toxic effect on oligodendrocytes or myelin (PLP ) when compared with control
cultures (a). Long-term ferritin treatment induced ferritin accumulation (ferritin light chain, FTL) in cells with
microglial phenotype (b), which was confirmed by double staining of FTL with ED1 (d). Control cultures did
+
not contain FTL cells (a, c). scale bar = 50 µm
In summary, both short and long-term ferritin exposure induced iron and ferritin
accumulation in microglia but not in oligodendrocytes which was similar to our observation in
cultures incubated with iron chloride. As the phenomenon of iron-accumulating dystrophic
microglia was reported in Huntington’s disease (Simmons et al, 2007) and Alzheimer’s disease
(Lopes et al, 2008; Streit & Xue, 2013), we assessed the number of microglia in iron chloride
and ferritin-treated cultures. We found a significant decrease of microglia numbers in cultures
treated overnight with 250 µM iron chloride compared to control cultures (Fig. 45).
Results 68
Figure 45: Loss of microglia upon iron chloride treatment.
Treatment of myelinating spinal cord cultures with iron chloride (b, overnight, 250 µM FeCl3) significantly
+
decreased the number of microglia compared to ascorbate-treated control cultures (a). Iba1 microglia
cells were counted from 10 pictures taken per coverslip with 3 coverslips analysed per experiment. The
number of microglia in iron chloride and ascorbate control treated cultures was normalised to the number
of microglia in untreated control cultures (p ≤ 0.001, n = 4). scale bar = 50 µm
Additionally, we assessed the number of microglia in different short and long-term
treatments of myelinating spinal cord cultures with different concentrations of ferritin. In all
experimental setups of ferritin treatment, we could not detect a decrease in microglia numbers
(Fig. 46).
Results 69
Figure 46: Microglia numbers in different experimental setups of ferritin treatment.
Short and long-term exposure of myelinating spinal cord cultures with different concentrations of ferritin
did not lead to a decreased number of microglia. In some conditions an increase of microglia was
observed (b, as one example of cells treated with 0.5 µg/ml ferritin for 72 h) compared to control cultures
+
(a). Iba1 microglia in different experimental approaches of ferritin treatment were counted from 10
pictures taken per coverslip with 3 coverslips analysed per experiment. The number of microglia was
normalised to control cultures (c). The results of single experiments are depicted. scale bar = 50 µm
Taken together, different in vitro iron loading approaches of myelinating spinal cord
cultures with iron chloride or iron bound to ferritin induced iron accumulation in microglia but not
in any other cell type. Iron loading with iron chloride decreased the number of microglia whereas
ferritin did not have a toxic effect.
As myelinating spinal cord cultures are complex and contain many other cells types, we
tested if iron loading with iron chloride and ferritin could be reproduced in purified microglia
cultures. Treatment of microglia cultures with iron chloride and ferritin induced iron accumulation,
detected by TBB, compared to control cells (Fig. 47).
Results 70
Figure 47: Iron accumulation in iron chloride- and ferritin-treated microglia cultures.
TBB stainings revealed that treatment of purified microglia cultures overnight with iron chloride (b, 10 µM
FeCl3) and ferritin (c, 5 µg/ml) induced iron accumulation compared with untreated cells (a). scale bar = 50
µm
In summary, our iron chloride and ferritin loading experiments with myelinating spinal
cord cultures induced iron accumulation in microglia but not in oligodendrocytes although
purified oligodendrocytes basically can accumulate iron (Fig. 36) and were shown to take up
ferritin (Schonberg et al, 2012). Iron chloride treatment of myelinating spinal cord cultures led to
a decrease of microglia numbers whereas ferritin had no toxic effect. Furthermore, we could
reproduce iron chloride and ferritin uptake in purified microglia cultures (Fig. 47).
Discussion 71
3 Discussion
3.1 General discussion
Our investigations indicate major differences between MS and a selection of different
animal models of inflammatory demyelination regarding microglia activation patterns and the
extent of oxidative injury and iron accumulation. We studied the expression of enzymes essential
for oxygen and nitric oxide radical production, p22phox and iNOS, respectively. The expression
pattern of p22phox and iNOS is variable between the different investigated models and
apparently dependent on the nature of the disease-inducing stimulus and the primary
inflammatory response. The expression of enzymes involved in radical-generating cascades has
been reported in inflammatory infiltrates of different models of CD4+ T cell-mediated
inflammatory demyelination (Cross et al, 1996; Koprowski et al, 1993; Misko et al, 1995; Ruuls
et al, 1995; Van Dam et al, 1995; van der Goes et al, 1998). More precisely, iNOS mRNA
expression and activity were increased in the spinal cords of mice after passive transfer of MBPspecific T cells in mice (Cross et al, 1996). iNOS mRNA and protein expression were also
detected in rats after active immunisation with MBP (Koprowski et al, 1993; Van Dam et al,
1995). Further, microglia and macrophages isolated from rats after active immunization with
guinea pig spinal cord homogenate were shown to produce ROS in vitro (Ruuls et al, 1995). The
expression of enzymes involved in reactive species production was primarily reported in
macrophages (Okuda et al, 1995; Van Dam et al, 1995) and was generally associated with
increased disease severity (Cross et al, 1996; Koprowski et al, 1993; Misko et al, 1995).
Apart from antimicrobial and toxic effects, NO can be protective because it inhibits T cell
proliferation (Fu & Blankenhorn, 1992). Further, iNOS serves in negative feedback regulation in
TH1-mediated autoimmune reactions (Bogdan, 2001). The administration of different NOS
inhibitors during EAE has led to conflicting results regarding disease severity and progression.
Some investigators reported that pharmacologic inhibition of iNOS exacerbated disease severity
and/or prevalence in rat EAE (Ruuls et al, 1996; Zielasek et al, 1995). Likewise, Gold et al.
reported an aggravation of the disease in actively immunized animals upon iNOS inhibition. In
contrast, blocking of iNOS had protective effects in passive T cell transfer EAE. These
experiments were performed in parallel in Lewis rats and the target antigen in both cases was
MBP (Gold et al, 1997). Interestingly, also Cross et al. reported that iNOS inhibition reduced
Discussion 72
disease pathology in adoptive transfer EAE in mice (Cross et al, 1994). Hence, it was concluded
that iNOS can play two different roles in EAE depending on the way of disease induction. In
case of passive T cell transfer EAE, NO production by encephalitogenic T cells is suggested to
be mostly tissue-damaging. In contrast, NO seems to have rather disease-dampening effects
after active immunisation (Bogdan, 1998; Gold et al, 1997). Strikingly, this does not hold true for
other experiments, as Brenner et al. reported a protective effect of iNOS inhibition in passive and
active MPB EAE in mice (Brenner et al, 1997). In contrast, NOS2 (the gene encoding for iNOS)
knockout mice exhibited a higher disease incidence and a more fulminant disease course than
wildtype controls after active immunisation with MOG35-55 or MBP (20-mer peptide) in different
mouse strains (Fenyk-Melody et al, 1998; Sahrbacher et al, 1998).
In summary, species- (rat versus mouse) and mouse strain-dependent factors seem to
account for the different roles or effects of iNOS in EAE. Furthermore, the specificity of applied
iNOS inhibitors and their bioavailability in the target tissue may be an issue (Bogdan, 1998;
Fenyk-Melody et al, 1998). The previously mentioned finding about iNOS inhibition-mediated
disease amelioration in T cell transfer EAEs is in line with our observation of profound iNOS
expression in the acute phase of our MBP-specific T cell transfer EAE. Further, the species- and
disease mechanism-dependent role of iNOS is also reflected in our data. As previously reported,
we detected iNOS expression mainly in cells with macrophages morphology (Okuda et al, 1995;
Van Dam et al, 1995) although iNOS was detected also in astrocytes (Tran et al, 1997). In
combination with ROS, NO-related reactive species can account for axonal injury (Aboul-Enein
et al, 2006; Choi et al, 2012; Moreno et al, 2011) through the production of peroxynitrite (Cross
et al, 1997).
In our models, we found different microglia and macrophage activation patterns
according to the expression of the essential NADPH oxidase subunit p22phox and iNOS. While
we detected profound expression of iNOS and p22phox in the acute MBP T cell transfer EAE,
CNS inflammation induced by T cells and macrophages (MOG35-55-immunised C57BL/6 mice),
by CD8+ T cells (passive transfer of HA-specific T cells into HA-transgenic mice) or by innate
immunity (LPS injection into the spinal cord white matter) appeared differently. At time points of
acute demyelination, mainly macrophages stained intensely for p22phox but iNOS was only
detected in scattered cells. In case of the LPS-injection model, iNOS expression in very early
stages of inflammation (day 3 after injection) in microglia and macrophages has been published
before. In line with iNOS inhibition experiments mentioned above, iNOS seems to play a role
rather in the induction phase of the disease after LPS-injection than in the very fulminant phase.
Discussion 73
In the inflammatory demyelinating disease triggered by active immunisation of DA rats
with MOG1-125 resulting in myelin-reactive antibody production, p22phox expression was
primarily detected in early demyelination stages in granulocytes. In this early disease stage,
iNOS was expressed by individual macrophages. In later chronic demyelinating disease stages,
p22phox expression was limited to single macrophages and iNOS+ cells were very rare. This is
in contrast to the profound expression of the phagocytosis-related marker ED1 by macrophages
in the chronic demyelinating lesions. In animals suffering from cuprizone-induced demyelination,
p22phox and iNOS expression were both very minor despite a marked staining for the
phagocytosis-related marker Mac-3 within macrophages in the demyelinating areas. Throughout
our study, we distinguished microglia from macrophages by their morphological appearance as
there is currently no microglia-specific marker available. Thus, our study does not allow
conclusions, whether macrophages within the lesions were derived from the microglial cell pool
or from recruited monocytes.
The minor expression of p22phox and iNOS was an unexpected finding since proinflammatory cytokines, which are presumably extensively produced in the lesions, were shown
to induce the expression of NADPH oxidase and iNOS in microglia in vitro (Cheret et al, 2008;
Lijia et al, 2012; Misko et al, 1995). Like other phagocytes, microglia express the phagocyte
NADPH oxidase (Sankarapandi et al, 1998), an enzyme comprising four regulatory cytosolic
subunits (p47phox, p67phox, p40phox, rac proteins) and a transmembrane heterodimer of
p22phox and NOX2, which generates superoxide (Cheret et al, 2008). LPS, for example,
induces the production of superoxide which was shown to enhance the expression of iNOS and
interleukin-1beta secretion (Cheret et al, 2008).
In the MHV-JHM coronavirus-induced demyelinating disease, iNOS expression was
described in macrophages in acute lesions (5-7 days after infection) and astrocytes in
demyelinated lesions. In contrast to our model, this was found by in situ hybridization after
intranasal inoculation with the virus (Grzybicki et al, 1997). We studied iNOS protein expression
by immunohistochemistry in intrathecally infected animals and found sparse iNOS+ microglia or
macrophages. We did not detect astrocytic iNOS staining in our experimental setup. NO was
shown to suppress viral replication in vitro (Lane et al, 1997) and iNOS mRNA was expressed
after MHV-JHM infection in vivo (Wege et al, 1998). Nevertheless, iNOS does not seem to have
a function in controlling viral replication in the CNS as inhibition of iNOS either had no or a
delaying effect on the progression of MHV-induced demyelination (Lane et al, 1999; Wu et al,
2000). In contrast to iNOS, microglia and macrophages in the demyelinating lesions stained
Discussion 74
intensely for p22phox. Additionally, we found p22phox-expressing microglial nodules in the
NAWM. The profound expression of NADPH oxidase could be one reason for the considerable
accumulation of oxidised phospholipids (E06) only in this model.
The second virus-induced model included in our study was TMEV-mediated
demyelinating disease. In contrast to MHV-JHM, TMEV-infection triggered p22phox expression
only shortly after virus-injection mostly in granulocytes. Moreover, TMEV did not induce the
expression of p22phox in microglial nodules, which were therefore only identifiable by Iba-1
expression. The number of microglial nodules increased with disease progression but they never
showed p22phox expression. Despite considerable staining for the phagocytosis-related
molecule Mac-3, the expression of p22phox as well as iNOS was rare in cells with macrophage
morphology in TMEV-induced lesions. In our experimental setup, we could reproduce the
reported iNOS expression in macrophages but not in astrocytes (Oleszak et al, 1997).
Our findings support the common opinion that microglia activation in the context of an
integrated immune response in vivo is more multifaceted than an activation of purified microglia
in vitro (Boche et al, 2013; Colton & Wilcock, 2010; Raivich et al, 1999). This view is reflected by
the many different possible ways of inducing NADPH oxidases. They were shown to be
activated by Toll-like receptor (TLR2) signaling in a model for spinal nerve injury (Lim et al,
2013) and by a number of cytokines with proposed anti-inflammatory functions, as for example
interleukin 4 and 13 (Nam et al, 2012; Park et al, 2008; Park et al, 2009). Further, thrombin was
reported to activate both NADPH oxidases and iNOS (Choi et al, 2005). Other factors involved in
NADPH oxidase activation are cholesterol load (Rackova, 2013) and divalent cations such as
zinc (Higashi et al, 2011; Kauppinen et al, 2008). Further, a relationship between sodium influx
and ROS production has been shown (Hossain et al, 2013). All these listed factors could act
together in the complex inflammatory process in the CNS.
Oxidative injury and mitochondrial damage are thought play a major role in tissue
destruction in MS. Furthermore, treatments addressing oxidative injury or mitochondria
protection have been shown to have a beneficial effect in EAE. Orally administered resveratrol, a
naturally occurring polyphenol, decreased neuronal damage in PLP-immunised SJL mice
without modulating the immune response (Shindler et al, 2010). A similar effect of resveratrol
was reported in MOG-immunised C57BL/6 mice (Fonseca-Kelly et al, 2012). Resveratrol mainly
has antioxidant and anti-inflammatory properties but was also shown to act pro-oxidant in the
presence of transition metal ions (de la Lastra & Villegas, 2007). This may be an issue
Discussion 75
concerning MS therapies as iron was shown to accumulate in the human brain and might
augment oxidative damage (Hallgren & Sourander, 1958; Hametner et al, 2013).
Mitochondrial CoQ10 (MitoQ), a derivative of coenzyme Q10 having anti-oxidative potential
in vitro (Manczak et al, 2010) and in vivo (Smith & Murphy, 2010) reduced disease course and
inflammation in MOG-immunised C57BL/6 mice (Mao et al, 2013). Moreover, an adenovirusbased gene-transfer of SOD2 in mouse EAE (DBA mice immunised with spinal cord
homogenate) had neuroprotective effects (Qi et al, 2007). Uric acid , a scavenger of
peroxynitrite, reduced clinical signs in EAE (MBP-immunised PLSJL mice) in contrast to ascorbic
acid which had no effect (Hooper et al, 1998; Spitsin et al, 2002). Interestingly, MS patients were
reported to have lower levels of uric acid in their serum than controls (Hooper et al, 1998).
Idebenone, a synthetic analog of coenzyme Q10, was shown to have protective effects in
Friedreich´s ataxia and cerebral ischemia (Di Prospero et al, 2007; Rego et al, 1999). In
contrast, it had no impact on EAE (in MOG35-55 immunised C57BL/6 mice) (Fiebiger et al, 2013).
Nevertheless, oxidative injury plays a role in animal models, as antibodies specific for
oxidised phospholipids were detected in MS patients as well as in MOG35-55 immunised C57BL/6
mice (Qin et al, 2007). We did not detect an accumulation of oxidised phospholipids in the
majority of the selected models but this does not generally exclude a role of oxidative injury.
However, our study reveals a major quantitative difference in the extent of oxidative damage
between MS lesions and inflammatory demyelination in animals. This could be explained by a
number of differences between rodents and humans. The observation that NADPH oxidases and
iNOS are already expressed to a certain extent in the normal human brain, but not in rodents,
could be one possibility to explain a higher susceptibility of humans to oxidative damage than
rodents. Another point is that laboratory animals are housed under standardised low pathogen
conditions avoiding infections and therefore peripheral immune stimulation (Perry & Teeling,
2013).
Another important point is that the basic level of microglia activation increases during
ageing (Lopes et al, 2008; Streit et al, 2004). Moreover, other age-related factors and the
intensifying lesion burden possibly add up to oxidative injury in MS patients. One potential
contributing factor could be iron accumulation in oligodendrocytes and myelin during ageing
(Hallgren & Sourander, 1958). Iron is proposed to boost oxidative injury occurring in MS lesions
when it is liberated from degenerating oligodendrocytes or myelin upon demyelination
(Hametner et al, 2013). In our study, iron accumulation is limited to aged rodents and only
Discussion 76
detectable in certain areas of the brain such as the brain stem nuclei. In rodent CNS
inflammation, we detected iron deposition only in perivascular macrophages, most likely caused
by leakage of blood into inflamed areas (Forge et al, 1998). Aged Lewis rats did not differ from
young rats regarding EAE course and severity and showed similar amounts of infiltrating
macrophages. Aged animals exhibited a lower number of infiltrating T cells but a higher number
of APP+ axonal spheroid, which is a measure for neurodegeneration. Although this could be
caused by aged-related factors, it cannot be accounted to iron accumulation. Iron does not
accumulate during ageing in the spinal cord which is mostly affected by inflammation. Due to the
low CNS iron burden of rodents, established animal models are not appropriate to study the
effect of iron on the amplification of oxidative injury. New models have been designed to
increase the iron load in the rodent CNS, for example ceruloplasmin knockout mice.
Ceruloplasmin is a ferroxidase essential for ferroportin-mediated iron export, linked with iron
detoxification and management. Ceruloplasmin knockout mice show iron accumulation in
astrocytes in the cerebellum at an advanced age which is accompanied by neurodegeneration
(Jeong & David, 2006). These animals do not accumulate iron in the non-injured spinal cord.
However, upon spinal cord injury, knockout animals showed augmented iron accumulation and
radical-mediated injury and reduced functional recovery due to enhanced neurodegeneration
(Rathore et al, 2008). Humans bearing a mutation in the gene encoding ceruloplasmin
(aceruloplasminemia) accumulate iron in astrocytes and suffer from neurodegeneration. In
patients suffering from aceruloplasminemia, oxidative stress is represented by the presence of
4-hydroxynonenal on neurons and astrocytes (Kaneko et al, 2002; Miyajima et al, 1987).
Another model associated with iron accumulation is the so-called sex-linked anemia (sla)
mouse. Sla mice have, due to a partial deletion, a dysfunctional hephaestin gene. In these
animals, spinal cord grey matter oligodendrocytes show increased iron content with ageing, as
hephaestin is another ferroxidase indispensable for iron export (Schulz et al, 2011).
Iron chelator treatment during EAE was reported to reduce disease severity in different
EAE models. Bowern et al. reported disease amelioration due to subcutaneous desferrioxamine
treatment before disease onset (in Lewis rats immunised with spinal cord homogenate) (Bowern
et al, 1984). Likewise, desferrioxamine injection at the EAE peak in MBP-immunised SJL mice
reduced clinical signs (Pedchenko & LeVine, 1998). Oral administration of deferiprone (Mitchell
et al, 2007) as well as intramuscular injections of apoferritin (LeVine et al, 2002) reduced
disease activity in PLP-immunised SJL mice. This disease-ameliorating effect was ascribed to
reduced oxidative stress and immune cell proliferation (Mitchell et al, 2007). The positive effect
of iron-scavenging substances in EAE should be translated to MS therapy with reservation,
Discussion 77
because the brain iron load and resulting oxidative stress is considerably higher in humans than
in rodents.
In order to nail down a possible amplification of neurodegeneration by increased iron
load, we chose an in vitro approach. It has been reported that purified oligodendrocytes can be
loaded with iron in vitro (Rathore et al, 2012; Schulz et al, 2011) and that increased iron levels in
oligodendrocytes renders them more vulnerable to cytokine toxicity (Zhang et al, 2005). This
was only shown for oligodendrocyte precursors and does not ultimately mean that iron has the
same
effect
on
mature
oligodendrocytes,
demyelination
or
even
on
secondary
neurodegeneration. This, however, cannot be studied in purified oligodendrocyte cultures.
Therefore, we chose the complex cell culture system of myelinating spinal cord cultures.
Although we successfully reproduced iron loading in purified oligodendrocytes using iron
chloride, the only cell population accumulating iron in the myelinating cultures were microglia.
Neurons, astrocytes and oligodendrocytes did not incorporate iron. Iron chloride treatment
reduced the number of microglia in these cultures. This is reminiscent of a phenomenon called
microglia dystrophy or senescence which was associated with iron toxicity and described in the
aged brain, Alzheimer´s and Huntington´s disease (Lopes et al, 2008; Simmons et al, 2007;
Streit et al, 2009). Moreover, microglial dystrophy in MS lesions and microglial loss in the center
of active lesions were reported in MS (Hametner et al, 2013). Apart from iron chloride, ferritin
was shown to be an iron source for oligodendrocytes in vitro and in vivo (Schonberg et al, 2012;
Todorich et al, 2011). Similarly to iron chloride loading experiments, ferritin treatments induced
iron accumulation only in microglia. In contrast to pure oligodendrocyte cultures (Todorich et al,
2011), oligodendrocytes in the myelinating cultures could not be forced to take up iron or
possibly to keep it. Moreover, oligodendrocytes were shown to take up ferritin from
macrophages which were preloaded with ferritin and injected into the spinal cord of rats
(Schonberg et al, 2012). Possibly, the ferritin-treated myelinating spinal cord cultures were
lacking an additional stimulus in order to trigger ferritin uptake in oligodendrocytes. Another
explanation for the lack of iron and ferritin accumulation in oligodendrocytes within this particular
culture system could be that they shuffle iron according to their needs. They could also lack
some factors that in vivo oligodendrocytes have. In order to analyse the iron management
properties of myelinating spinal cord cultures, the expression of transferrin and ferritin receptors
in the different cell populations has to be characterised.
Another factor important factor in MS pathology is the increasing lesion burden. A
smouldering activation of microglia may be intensified by retrograde degeneration from distant
Discussion 78
chronic lesions. This notion is supported by a study suggesting that cortical areas in the MS
brain are more prone to develop new lesions when they are connected with distant former
lesions (Kolasinski et al, 2012). However, in case of animal models, even the profound
demyelination, axonal loss and retrograde degeneration found in DA rat MOG EAE (Storch et al,
1998) did not induce microglia activation in the normal appearing grey and white matter.
The only animal model exhibiting an accumulation of oxidative damage to a similar extent
as MS patients was the chronic inflammatory demyelination triggered by intrathecal injection of
the MHV-JHM virus. One reason for that could be the profound expression of NADPH oxidases
and the resulting production of ROS in the lesions. In the TMEV-infected animals, we found a
higher reactivity for E06 (oxidised phospholipids) than in non-susceptible control animals but to a
lower degree compared with MHV-JHM virus-infected animals. The MHV-JHM-induced chronic
inflammatory demyelination and MS have some aspects in common. The virus causes a chronic
progressive inflammatory process leading to extensive lesions of primary demyelination with a
variable degree of axonal loss and diffuse damage in the NAWM (Barac-Latas et al, 1997;
Zimprich et al, 1991). Similar to MS lesions, CD8+ T cells dominate inflammatory infiltrates
(Dandekar et al, 2004; Pewe & Perlman, 2002), which is associated with a high activation level
of microglia and macrophages (Wu & Perlman, 1999). In the MHV-JHM- as well as the TMEVmediated disease, we found clusters of activated microglia, commonly called microglia nodules,
in the NAWM. Such microglia nodules are typical for MS pathology (Barnett & Prineas, 2004;
Prineas et al, 2001; van Horssen et al, 2012). In case of MHV-JHM, many mechanisms of tissue
injury have been elucidated involving CD8+and CD4+ T cells (Dandekar et al, 2004; Haring et al,
2001; Pewe et al, 2002; Pewe & Perlman, 2002; Stohlman et al, 2008) and anti-viral antibodies
(Lin et al, 1999; Tschen et al, 2002; Zimprich et al, 1991). Similarly, also the TMEV-induced
demyelination is characterised by CD8+ T cells (Ruby & Ramshaw, 1991; Zinkernagel & Althage,
1977), NK cells (Paya et al, 1989), antibody secretion (Rodriguez et al, 1988a; Roos et al, 1987),
macrophages and TH1 T cells (Lipton et al, 2005). Moreover, a continuous virus infection in the
CNS could provide a persistent stimulus for microglia activation via Toll-like receptors. This
should be true for both virus models, although lesions in MHV-JHM-infected rats exhibit a higher
degree of oxidative damage than TMEV-infected mice. This could be due to a higher infection
rate of MHV-JHM, a better persistence of the virus or possibly a difference due to species (rat in
MHV-JHM versus mouse in TMEV infection). Nevertheless, the chronicity and long-lasting
disease duration of virus-induced models for inflammatory CNS demyelination could provide
certain similarities in the pathology compared with MS.
Discussion 79
3.2 Conclusion and future prospects
In order to promote the development of new MS therapies it is essential to study the
constitution and extent of oxidative injury in rodent animal models of inflammatory demyelination
and to compare it directly to MS. For that purpose, we studied a broad sample of different acute
and chronic inflammatory diseases of the rodent CNS relying on identical tools as for the MS
tissue.
Our work could have an important impact on the design of new therapeutics for MS and
the preceding experimental studies. The remarkable differences concerning the mechanisms of
tissue injury between established animal models and MS could account for discrepancies in
outcome for neuroprotective therapeutics between animals and humans in trials. One possible
example for a new therapeutic intervention would be the boost of endogenous anti-oxidant
defence mechanisms, for example by Nrf2-mediated pathways. Nrf2-induced transcription of
oxidative stress defence mechanisms was suggested to take place in active MS lesions.
Apparently, this endogenous response is not able to cope with ROS-induced cellular damage in
MS (van Horssen et al, 2010). This strategy could prove to be efficient in established EAE
models but not necessarily in situations of profound oxidative injury when endogenous defence
mechanisms may be already exhausted. Thus, we propose the design of new experimental
animal models reflecting oxidative damage in MS more closely than classic EAE experiments.
Possibly, animals suffering from chronic virus-induced demyelination could provide new insights.
Additionally, new transgenic animals having a higher base level of microglia activation, for
example by overexpressing NADPH oxidases, could improve the situation. Different approaches
would be animals with impaired mitochondrial function or profound iron accumulation in the
CNS.
Apart from rodents, EAE can be induced in nonhuman primates, such as marmoset and
rhesus monkeys, by active immunisation with MBP, PLP or MOG. Dependent on the species
and the immunisation procedure, EAE in nonhuman primates reflects various clinical signs of
MS including acute, relapsing-remitting, primary progressive and secondary progressive
disease. In contrast to rodents, monkey colonies are outbred and exhibit a higher variation within
experimental groups. Further, they are phylogenetically closer to man than rodents. Marmoset
EAE turned out to be a valuable model for the preclinical validation of anti-inflammatory and
immune-modulatory MS therapies (Gold et al, 2006; t Hart et al, 2000). The characterisation of
the nature and the extent of oxidative damage in nonhuman primate EAE could have a major
impact on the development of new MS therapies.
Materials and Methods 80
4 Materials
4.1 Materials for histology
4.1.1
Buffers
4.1.1.1 Boric acid, 50 mM
0.1545 g boric acid (Merck) was dissolved in distilled water. The pH was adjusted to 8.0
with sodium hydroxide (NaOH). Distilled water was added to a final volume of 50 ml.
4.1.1.2 Citrate buffer 10x stock pH 6.0, 10 mM
For the stock solution, 21 g citric acid monohydrate (Fluka) were dissolved in distilled
water and the pH was adjusted to 6.0 with NaOH. Distilled water was added to a final
volume of 1 l. This stock solution (0.1 M) was diluted 1:10 in distilled water for the
working buffer (10 mM), which was used for antigen retrieval.
4.1.1.3 Dako buffer
For the working solution of Dako buffer 100 ml Dako cytomation wash buffer 10x (Dako,
S3006) were diluted with 900 ml distilled water.
4.1.1.4 EDTA buffer 20x stock pH 8.5
1.21 g Tris base (AppliChem, 10 mM) and 0.37 g EDTA (Merck, 1 mM) were dissolved
in 50 ml distilled water. The pH was adjusted to 8.5 (or 9.0) with HCl.
4.1.1.5 PBS buffer 4x stock
90 g NaCl (Merck) were dissolved in 2.5 l 0.2 M Sörensen buffer.
4.1.1.6 Sodium acetate buffer pH 4.9, 0.05 M
6.8 g sodium acetate (Merck) were dissolved in distilled water. The pH was adjusted to
4.9 with acetic acid. Distilled water was added to a final volume of 1 l.
Materials and Methods 81
4.1.1.7 Sodium potassium phosphate buffer pH 7.4, 5 mM
0.68 g potassium dihydrogen phosphate (Merck) were dissolved in distilled water. The
pH was adjusted to 7.4 with NaOH and distilled water was added to a final volume of 1 l.
For homogenization of tissue used for protein analysis (e.g. western blot), 1 µl of 1 M
phenylmethylsulfonylfluorid (PMSF) can be added per 1 ml sodium potassium
phosphate buffer.
4.1.1.8 Sörensen buffer pH 7.4, 0.2 M
13.8 g NaH2PO4 (Merck) and 71.2 g Na2HPO4 (Merck) were dissolved in 2.5 l distilled
water.
4.1.1.9 TBS 20x stock, 1 M
60.57 g Tris base (AppliChem) and 180 g NaCl (Merck) were dissolved in 500 ml
distilled water. The pH was adjusted to 7.5 with 400 ml 1M HCl (Fluka). Distilled water
was added to a final volume of 1 l.
4.1.1.10 TBS/CaCl2 (2 mM)
0.294 g CaCl2 were mixed with 50 ml TBS 20x stock and 950ml distilled water.
TBS/CaCl2 was stored at 37 °C for proteinase K digestion.
4.1.1.11 Tris/HCl buffer pH 8.5, 0.1 M
12.1 g Tris base (AppliChem) were dissolved in distilled water and the pH was adjusted
to 8.5 with HCl. Distilled water was added to a total volume of 1 l. Tris/HCl (0.1 M) buffer
was stored at 37 °C for fast blue development.
4.1.2
Stock solutions and developing agents
4.1.2.1 Amino ethylcarbazole reagent (AEC) developing agent
The AEC stock solution was prepared by dissolving 780 mg AEC (Sigma-Aldrich) in 45 ml
dimethylformamide (DMF, can be stored up to 2 months at 4 °C). After thoroughly rinsing the
slides with distilled water, 1.15 ml AEC stock were added to 50 ml sodium acetate buffer pH 4.9.
Finally, 25 µl H2O2 were added and the development was performed without prior filtration of the
solution.
Materials and Methods 82
4.1.2.2 CSA stock solution
CSA stock was prepared by dissolving 15 mg sulpho-NHS-LC-Biotin (Pierce, 21335) in 6 ml
borate buffer. 4.5 mg tyramine (Sigma, T-7255) were added and the solution was stirred
overnight at RT. The solution was filtered with a 0.45 µm filter and small aliquots were stored at 20°C.
4.1.2.3 DAB developing agent
For the 3,3’-diaminobenzidine (DAB) stock solution, 1 g DAB (Fluka, 32750) was dissolved in 40
ml PBS buffer and stored at -20 °C in 1 ml aliquots. The DAB working solution for one cuvette
comprised of 1 ml DAB stock solution for 50 ml PBS. After filtration, 16.5 µl H2O2 were added.
4.1.2.4 DTPA 100x stock, 5 mM
197 mg diethylenetriamine-pentaacetic acid (DTPA, Sigma-Aldrich, D1133) were dissolved in
distilled water. The pH was adjusted to 7.4 with NaOH and distilled water was added for a final
volume of 100 ml. For chelating extracellular iron, the 100x stock was diluted in medium and
cells were incubated for 5 min at RT.
4.1.2.5 Eosin solution
For the eosin 10x stock solution, 10 g Eosin Y (Merck) were dissolved in 100 ml distilled
water. For the working eosin solution, 2.5 ml eosin 10x stock solution were diluted with
250 ml distilled water and with 12 drops of glacial acetic acid (Fluka).
4.1.2.6 Fast blue developing agent
4% NaNO2 was prepared by dissolving 0.2 g NaNO2 (Merck) in 5 ml distilled water (can be
stored for 2 weeks at RT). For 100 ml fast blue working solution, 100 ml Tris/HCl were
maintained at 37 °C. 12.5 mg naphtol-AS-MX (Sigma) were dissolved in 615 µl
dimethylformamide (Fluka) and mixed with the Tris/HCl buffer. Next, 25 mg fast blue salt
(Sigma) were mixed with 615 µl 2N HCl (Merck) and 615 µl 4% NaNO2. This was added to the
Tris/HCl mixture and stirred until the solution was clear. The fast blue mixture was filtrated. At
last, 154 μl of levamisole (1 M in Tris/HCl, Sigma) were added. The sections were developed at
37°C.
Materials and Methods 83
4.1.2.7 Geltol mounting medium
Sections developed with fast blue or AEC were mounted with geltol. The geltol solution was
prepared by thorough mixing of 6 g glycerine (Life Technologies) with 2.4 g mowiol (Calbiochem)
in 6 ml distilled water. 12 ml 0.2 M Tris/HCl buffer were added and the solution was stirred for 10
minutes at 50 °C. The solution was centrifuged for 15 minutes at 5000 x g and stored at -20 °C.
(For mounting of fluorescent stainings, 2.1 g gallate (Sigma) were dissolved in 10 ml geltol.)
4.1.2.8 HCl/ethanol
0.5 ml concentrated HCl were added to 100 ml 70% ethanol.
4.1.2.9 Iron detection reagent (Ferrocene assay)
The iron detection reagent contained 7.04 g ascorbate (Sigma-Aldrich), 7.76g
ammonium acetate (Merck), 64 mg ferrocene (Fluka, 82950) and 64 mg neocuproine
(Sigma-Aldrich). First, ascorbate and ammonium acetate had to be dissolved in distilled
water. Subsequently, ferrocene and neucuproine were added ant distilled water was
added to a total volume of 20 ml.
4.1.2.10 Luxol fast blue (LFB) and periodic acid Schiff (PAS) staining solutions
The 0.1% LFB solution was prepared by dissolving 1 g Luxol Fast Blue (Chroma) in 1 l
96% ethanol overnight at 57 °C. The 0.1% lithium carbonate solution was made by
dissolving 1 g lithium carbonate (Merck) in 1 l distilled water. For the periodic acid, 4 g
periodic acid (Merck) were solved in 500 ml distilled water. For the sulfite solution, 5 ml
concentrated hydrochloric acid (HCl, Fluka) were mixed with 20 ml 10% potassium
disulfite (Fluka) and 500 ml distilled water.
4.1.2.11 Methanol/H2O2
1 ml H2O2 (30%) was added to 150 ml methanol.
4.1.2.12 Proteinase K solution
10 mg proteinase K (Sigma, P0390) were dissolved in 1 ml TBS and 100 µl aliquots were stored
at -20 °C. For the digestion of snap-frozen and acetone/PFA-fixed slides, 50 µl proteinase K
were mixed in 50 ml warm (37 °C) TBS/CaCl2.
Materials and Methods 84
4.1.2.13 Scott’s solution
2 g potassium hydrogen carbonate (Merck) and 20 g magnesium sulphate heptahydrate
(Merck) were dissolved in a final volume of 1000 ml distilled water.
4.2 Media for cell cultures
4.2.1
Media for T cells
4.2.1.1 Restimulation medium for T cells
RPMI-1640 (Lonza, BE12-167F)
1% non-essential amino acids (Gibco, 11140-035)
1% L-glutamine (Lonza, BE17-605E)
1% penicillin/streptomycin (Lonza, DE17-602E)
1% sodium-pyruvate (Gibco, 11360-039)
1% rat serum (own stock)
300 µl β-mercaptoethanol stock/100 ml medium (for the stock, Sigma-Aldrich 99% βmercaptoethanol was diluted in RPMI-1640 1:500)
4.2.1.2 TCGF medium for T cells
RPMI-1640 (Lonza)
10% FCS (PAA, S0115)
10% supernatant of IL-2 producing MLA-IL-2-cells (own stock)
1% non-essential amino acids (Gibco)
1% L-glutamine (Invitrogen)
1% penicillin/streptomycin (Lonza)
1% sodium-pyruvate (Gibco)
0.3% β-mercaptoethanol (as described above)
4.2.1.3 Freezing medium for T cells
45% RPMI-1640 (Lonza)
45% FCS (PAA)
10% DMSO (Sigma-Aldrich)
Materials and Methods 85
4.2.2
Media for glial cells
4.2.2.1 Mixed glia medium for microglia isolation
RPMI-1640 (Lonza, BE12-167F)
10% FCS (PAA, S0115)
1% L-glutamine (Lonza, BE17-605E)
1% penicillin/streptomycin (Lonza, DE17-602E)
4.2.2.2 Mixed glia medium for oligodendrocyte isolation
DMEM (Lonza, BE12-709F)
10% Horse Serum (HS, PAA, B02309-7031)
1% L-glutamine (Lonza)
1% penicillin/streptomycin (Lonza)
4.2.2.3 Digestion medium
2.5 mg/ml papain (Sigma-Aldrich, P4762)
240 µg/ml L-cystein (Sigma-Aldrich, C7352)
40 µg/ml DNAse I (Roche Diagnostics, 104159)
In HBSS (Lonza)
1 ml aliquots were stored at -20 °C.
4.2.2.4 Sato medium
16.1 µg/ml putrescine (Sigma-Aldrich, P5780)
60 ng/ml progesterone (Sigma, P0130)
100 µg/ml BSA (PAA)
0.5 ng/ml sodium selenite (Sigma-Aldrich, S5261)
400 ng/ml tri-iodothyroxine (Sigma-Aldrich, T6397)
400 ng/ml L-thyroxine (Sigma-Aldrich, T1775)
50 µg/ml transferrin (Sigma-Aldrich, T8158)
5 µg/ml insulin (Sigma-Aldrich, I1882)
0.5% horse serum (PAA)
1% L-glutamine (Invitrogen)
in DMEM (Lonza)
Aliquots of stock solutions were stored at -20 °C except for transferrin and insulin that were
dissolved immediately before medium preparation.
Materials and Methods 86
4.2.2.5 Neurosphere medium (NSM)
DMEM/F12 (Invitrogen, 21331-020)
Hormone Mix (own 10x stock)
0.6% glucose (Sigma-Aldrich)
0.1% NaHCO3 (Sigma-Aldrich)
5 mM HEPES (Sigma-Aldrich)
1% L-glutamine (Invitrogen)
1% penicillin/streptomycin (Invitrogen)
0.01 % BSA (Sigma-Aldrich)
4.2.2.6 Hormone mix 10x
250 ml DMEM/F12 supplemented with:
250 mg apo-human transferrin (Sigma-Aldrich, I-9278)
25 ml of the mixture of 6.25 ml human recombinant insulin (Sigma-Aldrich, I-9278) plus 19.75 ml
distilled water
25 ml putrescine (9.6 µg/ml in distilled water, Sigma-Aldrich, 7505)
25 µl selenium (0.5 mg/ml in distilled water, Sigma-Aldrich, S-9133)
25 µl progesterone (0.625 mg/ml in 95% ethanol, Sigma-Aldrich, P6149)
0.6% glucose (Sigma-Aldrich, G7021)
0.1% NaHCO3 (Sigma-Aldrich, S5761)
5 mM HEPES (Sigma-Aldrich, H4034)
The 10x stock hormone mix was stored in 25 ml aliquots at -80 °C.
4.2.2.7 Astrocyte medium
DMEM (1 g/l glucose, Invitrogen, 21885-025)
10% FCS (Sigma-Aldrich, F9665)
0.5% L-glutamine (Invitrogen)
1% penicillin/streptomycin (Invitrogen)
4.2.2.8 SD solution (soybean trypsin inhibitor)
25 ml Leibovitz L15 medium (Invitrogen, 11415-049) were supplemented with:
13 mg trypsin hinhibitor (Sigma-Aldrich, T9003)
1 mg DNAse I (Sigma-Aldrich, D4263)
75 mg BSA (Sigma-Aldrich)
The solution was sterile filtrated and 2 ml aliquots were stored at -20 °C.
Materials and Methods 87
4.2.2.9 Plating medium
DMEM (4.5 g/l glucose, Invitrogen, 61965-026)
25% Horse serum (Sigma-Aldrich, H1270)
25% HBSS (Invitrogen)
1% L-glutamine (Invitrogen)
4.2.2.10 Differentiation medium (DM)
DMEM (4.5 g/l glucose)
1 µg/ml Biotin (Sigma-Aldrich, B4501)
0.5% N1 supplement (Sigma-Aldrich, N6530)
50 nM hydrocortisone (Sigma-Aldrich, H0396)
10 µg/ml Insulin (Sigma-Aldrich, I1882)
1% penicillin/streptomycin (Invitrogen)
5 Methods
5.1 MS patients
The stainings were performed on archival formalin-fixed paraffin-embedded tissue of MS
patients collected at the Centre for Brain Research during the last three decades. The study was
approved by the Ethical Committee of the Medical University of Vienna (EK. No. 535/2004 and
213/06/2013). The MS study cohort contained autopsy tissues from 15 controls and 24 MS
cases, including 8 acute MS patients, 3 relapsing remitting MS (RRMS) patients, 6 secondary
progressive MS (SPMS) patients and 7 primary progressive MS (PPMS) patients. The
corresponding clinical data of MS patients and controls are depicted in Table 2.
Materials and Methods 88
Table 2: Clinical data of MS cases and controls of the study cohort.
case number
age (years)
sex
clinical course MS
MS 1
34
f
acute
disease duration
(months)
4
MS 2
35
m
acute
1.5
MS 3
45
m
acute
0.6
MS 4
45
m
acute
0.2
MS 5
51
f
acute
5
MS 6
52
m
acute
1.5
MS 7
78
m
acute
2
MS 8
69
f
acute
1
MS 9
44
f
RRMS
261
MS 10
40
f
RRMS
120
MS 11
57
f
RRMS
156
MS 12
46
f
SPMS
444
MS 13
56
m
SPMS
372
MS 14
41
m
SPMS
137
MS 15
61
f
SPMS
288
MS 16
62
f
SPMS
144
MS 17
34
m
SPMS
120
MS 18
77
f
PPMS
168
MS 19
55
f
PPMS
60
MS 20
67
m
PPMS
87
MS 21
53
m
PPMS
168
MS 22
34
f
PPMS
204
MS 23
54
f
PPMS
72
MS 24
71
f
PPMS
264
control 1
97
f
-
-
control 2
65
m
-
-
control 3
92
f
-
-
control 4
71
f
-
-
control 5
80
f
-
-
control 6
71
f
-
-
control 7
72
m
-
-
control 8
88
f
-
-
control 9
83
m
-
-
control 10
46
m
-
-
control 11
36
f
-
-
control 12
39
f
-
-
control 13
42
f
-
-
control 14
30
f
-
-
control 15
37
m
MS = multiple sclerosis; f = female; m = male; RRMS = relapsing-remitting MS; SPMS = secondary
progressive MS; PPMS = primary progressive MS
Materials and Methods 89
5.2 Experimental models
For the analysis of rodent models of MS-like inflammatory demyelination in the CNS, we
used rat tissue from experimental autoimmune encephalomyelitis (EAE) experiments performed
in our lab and rat and mouse tissue from former studies performed in collaboration with our
group, as stated in the corresponding sections below. Tissues containing inflammatory lesions
with active demyelination or neurodegeneration were selected from a larger collection of archival
material. They were defined by the presence of macrophages with early myelin or neuronal
degradation products. The following experimental models were analysed:
5.2.1
CD4+ T cell-mediated EAE in Lewis rats
Lewis rats were bred and housed under standardised conditions in the Decentral
Facilities of the Institute for Biomedical Research at the Medical University of Vienna. All animals
had access to food and water ad libitum. Our experiments were approved by the ethic
commission of the Medical University of Vienna and performed with the license of the Austrian
Ministry for Science and Research.
5.2.1.1 Immunisation with MBP/CFA and T cell line preparation
Lewis rats aged 8-10 weeks were immunised subcutaneously at the base of tail with 200
µl of a 1:1 mixture of myelin basic protein (MBP) (2 mg/ml in RPMI-1640; guinea pig myelin
basic protein, Sigma-Aldrich) and complete Freund´s adjuvants (CFA; incomplete Freund´s
adjuvants (IFA) supplemented with 4 mg/ml mycobacterium tuberculosis H37Ra (Difco
Laboratories)) under isoflurane (Abbott) anaesthesia.
MBP-specific T cell lines were established as described (Aboul-Enein et al, 2006; BenNun et al, 1981). Shortly summarized, animals were sacrificed with an overdose of CO2 9-11
days after immunisation. The draining lymph nodes were collected, dissociated through a mesh
(100 µm mesh size) and taken up in 5 ml restimulation medium containing 10 µg/ml MBP
antigen and seeded in 60 mm dishes. The cells were maintained at 37 °C and 5% CO2.
For propagation of MBP-specific T cells, cells from the primary culture were transferred to
100 mm dishes and 5-10 ml TCGF medium was added. This was done after 1-2 days of primary
culture when the cells had formed clusters and increased in size. In TCGF medium, the cells
underwent a proliferation phase. The end of this growth phase was noticed by the lack of cell
clusters and smaller size. At this point, the cells were restimulated.
Materials and Methods 90
For T cell restimulation, thymocytes were isolated from Lewis rats aged 8-10 weeks. The
thymus was excised and dissociated through a mesh (100 µm mesh size). Cells were collected
in RPMI-1640 and irradiated with 3000 rad in order to prevent thymocyte overgrowth of the T cell
culture. Finally, freshly isolated thymocytes were centrifuged, taken up in restimulation medium
and adjusted to a density of 1-1.5 x 107 cells/ml. Prior to restimulation, the cultured T cells were
centrifuged, resuspended in restimulation medium and adjusted to a cell number of 1-1.5 x 106
cells/ml. Finally, 1 ml of thymocyte suspension was added to 4 ml of T cell suspension in a 60
mm dish including 10 µg/ml MBP antigen. The MBP-specific T cells were again activated
noticeable by an increase in size and formation of clusters after approximately 48 hours. At this
stage, the cells could be used for T cell transfer into rats to induce EAE, for further T cell
propagation or frozen for later EAE induction.
For freezing, T cells were harvested by centrifugation and resuspended in freezing
medium. Cryopreservation was performed using a cell freezing container with isopropyl alcohol
at -80 °C. Subsequently, cells were stored in liquid nitrogen.
5.2.1.2 Induction of MBP-specific T cell EAE in Lewis rats
EAE was induced in female Lewis rats by passive transfer (intraperitoneal injection) of
4.5 x 106 activated MBP-specific T cells in RPMI-1640. The clinical scores and body weight were
monitored daily. Clinical signs were accessed according to following criteria: 0 = no clinical
signs, 0.5 = partial loss of tail tonus; 1 = complete loss of tail tonus; 2 = hind limb weakness; 3 =
complete hind limb paralysis (termination criterion); 4 = hind and fore limb paralysis; 5 =
moribund. Animals were sacrificed by an overdose of CO2 and perfused with 4% phosphatebuffered paraformaldehyde (PFA). Brain, spinal cord and organs were dissected and post-fixed
for 18 h in PFA. The PFA-fixed samples were routinely embedded in paraffin. Brain and lumbar
spinal cord tissue was analysed at the peak (6 days) and at the recovery phase (12 days) after T
cell transfer.
5.2.2
Chronic relapsing MOG EAE in DA rats
Archival material originating from collaboration studies with Prof. Maria Storch was
analysed. Chronic relapsing EAE was induced by active sensitisation with myelin
oligodendrocyte glycoprotein (MOG 1-125) in complete Freund’s adjuvant in dark agouti (DA)
rats as published (Storch et al, 1998). Chronic active lesions in the spinal cord from the first bout
or subsequent relapses occurring 12 to 55 days after sensitization were studied in the current
Materials and Methods 91
project. This model is characterised by a chronic stage with large confluent plaques of
demyelination, loss of oligodendrocytes and a variable extent of axonal injury in parallel with
very early perivenous lesions. Tissue injury is known to be triggered by encephalitogenic CD4+ T
cells and amplified by demyelinating anti-MOG antibodies (Lorentzen et al, 1995; Weissert et al,
1998).
5.2.3
Chronic relapsing MOG EAE in C57BL/6 mice
Chronic relapsing EAE in C57BL/6 mice was induced in the lab of Prof. Lesley Probert
and was used for collaborative studies (Taoufik et al, 2011). C57BL/6 mice were actively
sensitised with rat MOG35-55 peptide together with complete Freund’s adjuvant. Additionally, mice
were intraperitoneally injected with pertussis toxin on days 0 and 2 after MOG-immunization.
Typically, clinical disease started 10 days after immunization and continuously increased the
following 3 weeks. The early disease phases (around day 10 to 18 after immunization), was
characterised by perivenous inflammation by T cells related with microglia activation and
macrophage infiltration. Around 20 days after immunization, the animals developed profound
demyelinating lesions, which grew in size and number until day 34. Demyelination was
accompanied with axonal injury and loss. In the current study, we included animals from day 21,
27 and 34 after immunization.
5.2.4
Acute CD8+ T cell-mediated EAE
Acute
demyelinating
encephalomyelitis
was
induced
by
passive
transfer
of
+
hemagglutinin-reactive CD8 T cells into transgenic BALB/c mice expressing hemagglutinin
specifically in oligodendrocytes. This was performed by the group of Prof. Roland Liblau
(Saxena et al, 2008) and histological analysis of CNS tissue was performed in collaboration with
our group. In the current study, brain and spinal cord were analysed during the active phase of
inflammation and demyelination 9 to 28 days after T cell transfer. Inflammation was shown to be
induced by the infiltration of CD8+ cytotoxic T cells leading to profound microglia activation,
direct T cell-mediated killing of oligodendrocytes and demyelination (Saxena et al, 2008).
5.2.5
LPS injection-induced inflammation
LPS-injections into spinal cord of SD rats were performed by the group of Prof. Kenneth
Smith (Felts et al, 2005) and spinal cord specimens were used in former collaboration studies
Materials and Methods 92
(Sharma et al, 2010). Acute demyelinating myelitis was established by focal injection of
lipopolysaccharide (LPS) into the dorsal column of the spinal cord of Sprague Dawley (SD) rats
leading to focal inflammation in the dorsal column 12 to 24 hours after LPS injection. The
disease started with an inflammatory phase, which was followed by a demyelinating phase
where focal demyelinated lesions with relative axonal preservation appeared. This was
associated with massive microglia activation and macrophage recruitment. The peak of active
demyelination occurred 9-12 days after disease induction. In our study, tissue from the
inflammatory as well as the demyelinating phase of the disease was analysed. Control animals
were injected with saline instead of LPS and sampled in the same way as LPS-injected animals.
5.2.6
Curpizone-induced demyelination
Cuprizone-induced demyelination experiments were performed by the group of Prof.
Annemarie van Dam (Van Strien et al, 2011). Active demyelination was induced by a 0.2%
cuprizone (bis-cyclohexanone oxaldihydrazone)-supplemented diet in C57BL/6 mice. In our
study, archival material of animals 35 days after the initiation of the cuprizone diet was studied.
At this stage of the disease active demyelination of the corpus callosum was observed.
5.2.7
Mouse hepatitis virus strain JHM (MHV-JHM) coronavirus-mediated demyelinating
disease
Chronic inflammatory demyelinating disease was induced by intracerebral infection of
Lewis rats with MHV-JHM coronavirus. These experiments were done in collaboration with Prof.
Helmut Wege (Barac-Latas et al, 1997; Korner et al, 1991; Wege et al, 1998; Zimprich et al,
1991). The virus infection caused different disease courses. One group of animals was suffering
from an acute inflammatory disease mainly affecting the grey matter of the CNS with
pronounced virus antigen expression in neurons, astrocytes and oligodendrocytes. Other
animals showed a late-onset type of disease reflected either by a chronic panencephalitis
affecting grey and white matter, or a chronic inflammatory demyelinating encephalomyelitis
mainly involving the brain stem and the spinal cord white matter. At these late stages of the
disease, the virus antigen was primarily seen in glial cells (Zimprich et al, 1991). In the present
study, we analysed animals with active lesions from all disease stages between 30 and 60 days
after infection (Barac-Latas et al, 1997).
Materials and Methods 93
5.2.8
Theiler's Murine Encephalomyelitis Virus (TMEV)-mediated demyelinating disease
Chronic inflammatory demyelinating disease was induced by intracerebral inoculation of
mice with the Daniels (DA) strain of Theiler´s murine encephalomyelitis virus (TMEV). The
stainings were performed on experimental material from our collaborators Dr. Claudia
Lucchinetti and Prof. Moses Rodriguez (Drescher et al, 1998; Rodriguez et al, 1986; Rodriguez
et al, 1988a; Rodriguez et al, 1988b). TMEV injections were performed with SJL mice that are
susceptible for the virus infection (Drescher et al, 1997; Lipton, 1975; Rodriguez et al, 1983).
Animals were sacrificed at different time points after virus injection (days post injection = dpi, 3,
7, 14, 21, 35, 45, 90, 120, 180, 220). For control reason, injections were also performed in not
susceptible C57BL/6 mice (Drescher et al, 1997) which were analysed 45 dpi.
5.2.9
Control animals
In order to validate the stainings on diseased animals, brain, spinal cord and lymphatic
tissues of untreated young and aged Lewis rats, untreated and saline-injected SD rats and DA
rats and C57BL/6 mice without inflammatory lesions were analysed.
5.3 Histology
Experimental material was routinely fixed in 4% paraformaldehyde (PFA) and embedded
in paraffin. Neuropathological evaluation was based on haematoxylin/eosin and Luxol Fast blue
myelin stains and Bielschowsky silver impregnation. Inflammation was evaluated by
immunohistochemistry for CD3. Macrophage activation was analysed using antibodies against
Iba-1 (human, rat and mouse tissue), CD68 (human), ED1 (rat) and Mac-3 (mouse).
Demyelination was assessed by immunohistochemistry for myelin basic protein (MBP). For the
detection of oxidative injury, four different markers were used: E06 (recognising oxidised
phospholipids), p22phox (subunit of NADPH oxidase complexes), iNOS (Nos2; inducible nitric
oxide synthase) and 3,3’-diaminobenzidine (DAB)-enhanced Turnbull reaction (non-heme tissue
iron).
Materials and Methods 94
5.3.1
Immunohistochemistry
5.3.1.1 General staining protocol
Immunohistochemistry was performed on 5 µm sections as was previously described (Bauer et
al, 2007; King et al, 1997). Sections were deparaffinised for 2x 15 min in xylene, rinsed twice in
96% ethanol and endogenous peroxidase activity was blocked in Methanol/H2O2 for 30 min.
Then, the sections were rehydrated via 90%, 70% and 50% ethanol series and finally rinsed in
distilled water. Prior to staining, antigen retrieval was performed in a steamer for 60 min in
plastic cuvettes. Antigen retrieval procedures and primary antibodies are listed in Table 3. After
washing with TBS, non-specific antibody binding was blocked by incubating the sections with
10% fetal calf serum (FCS, Lonza) in Dako buffer (Dako) for 20 min. Primary antibodies were
applied in 10% FCS/Dako buffer at 4 °C overnight. After washing with TBS, secondary
antibodies (Table 4) were applied for 1 h at room temperature (RT) in 10% FCS/Dako buffer.
Finally, the sections were incubated with avidin peroxidase (1:100, Sigma Aldrich, A3151) in
10% FCS/Dako buffer for 1 h at RT. Antibody binding was routinely visualised using the
chromogen DAB (Sigma-Aldrich). The reaction was stopped by rinsing the sections in tab water.
The sections were then counterstained with haematoxylin (Merck) for 20 sec, rinsed in tap water
and differentiated with HCl/ethanol and Scott´s solution. Finally, the sections were rinsed in tap
water, dehydrated via 50%, 70% and 90% ethanol series and n-butyl acetate and mounted in
eukitt (Gröpl Elektronenmikroskopie) under glass coverslips. To validate the staining results,
appropriate primary antibody positive controls and blanks were used.
In case of CD3 antibody, the staining was amplified with biotinylated tyramine
enhancement (CSA) (Bauer et al, 2007; King et al, 1997). After the avidin peroxidase step, the
slides were incubated for 20 min with CSA (stock 1:1000) in PBS with H2O2 (1:1000). Slides
were rinsed in PBS followed by a second incubation with avidin peroxidase 1:100 for 30 min.
Finally, the staining was developed with DAB.
Materials and Methods 95
Table 3: Primary antibodies and antigen retrieval procedures for immunohistochemistry
Antibody
Origin
Target
Dilution
Antigen
retrieval
Source
8OHdG
goat (pAB)
8-Hydroxy-2′-deoxyguanosine
(oxidised DNA)
1:500
St (E)*
Abcam, ab10802
APP
mouse (mAb)
amyloid precursor protein
1:1000
St (C)
CD3
rabbit (mAb)
T cells
1:2000
St (E)
CD68
mouse (mAb)
phagocytic macrophages
1:100
St (E)
DA virus
rabbit (pAB)
virus antigen
1:200
0
E06
mouse (mAb)
oxidised phospholipids
1:200
0
ED1
mouse (mAb)
rat CD68
1:1000
St (E)
GFAP
rabbit (pAB)
1:3000
St (E)
Iba-1
rabbit (pAb)
glial fibrillary acidic protein
ionized calcium binding
adaptor molecule 1
1:3000
St (E)
rabbit (pAb)
inducible nitric oxide synthase
1:30000
St (E)
rabbit (pAb)
inducible nitric oxide synthase
1:375
St (C)
Mac-3
rat (pAb)
mouse CD68
1:200
St (C)
MBP
N 556
Ox8
rabbit (pAb)
mouse (mAb)
mouse (mAb)
myelin basic protein
virus nucleocapsid
anti-rat CD8
1:2500
1:50
1:400
0
0
0
p22phox
rabbit (pAb)
NADPH oxidase protein
1:100
St (C)
TPPP/p25
rat (pAB)
tubulin polymerization
promoting protein
1:3000
St (E)
iNOS antihuman
iNOS antirat
Chemicon;
MAB348
Neomarkers; RM9107
Dako; M0814
Stained by Mayo
Clinic
Avanti; 330001
Serotec;
MCA341R
Dako; Z0334
Wako Chemicals;
019-19741
Chemicon;
AB5384
Chemicon;
AB1631
Serotec;
MCA2293FB
Dako; A0623
Wege et al, 1984
Seralab; MAS041
Santa Cruz
Biotech; sc-20781
Hoftberger et al,
2010
0 = no antigen retrieval; St = steaming using the indicated buffer for 1 h, C = citrate buffer (pH 6.0), E =
EDTA buffer (pH 8.6); mAb = monoclonal antibody; pAb = polyclonal antibody
Antibodies were used for human, rat and mouse tissue with the following exceptions: CD68 for human,
ED1 and Ox8 for rat, Mac-3 for mouse tissue and iNOS anti rat for rat and mouse. * For 8OHdG staining
of snap-frozen tissue, the slides were pre-treated with proteinase K.
Table 4: Secondary antibodies for immunohistochemistry
Antibody
Dilution
Source
biotinylated anti-mouse from sheep
1:500
Jackson ImmunoResearch, 515-065-003
biotinylated anti-rabbit from donkey
1:2000
Jackson ImmunoResearch, 711-065-152
biotinylated anti-sheep/goat from donkey
1:200
Amersham/ Healthcare RPN 1025
biotinylated anti-rat from donkey
1:1500
Jackson ImmunoResearch, 712-065-156
alkaline phosphatase-conjugated antimouse from donkey
1:200
Jackson ImmunoResearch, 715-055-151
Materials and Methods 96
5.3.1.2 E06 staining for oxidised phospholipids
The sections were deparaffinised for 2x 15 min in xylene and rehydrated via 90%, 70%
and 50% ethanol series and finally rinsed in distilled water. The sections were rinsed in
phosphate buffer (PB)/Tween20 (PB/Tween, Sörensen buffer plus 0.05% Tween20 (Dako). Nonspecific antibody binding was blocked by incubating the sections for 20 min in Dako REALTM
antibody diluent (Dako, S2022). After washing with PB, further blocking was performed using the
avidin-biotin-blocking system from Dako (Dako, X0590). Avidin was blocked for 10 min by
incubation with avidin-block (1:100 in PB) followed by 10 min biotin-block (1:10 in PB). The
slides were rinsed with PB and the E06 primary antibody was applied in Dako-diluent at 4 °C
overnight. After washing with PB/Tween, endogenous peroxidase activity was blocked in 3%
H2O2 in PB for 10 min. After rinsing with PB/Tween, biotinylated secondary antibody was applied
for 30 min at RT in Dako-diluent. After washing in PB/Tween, the sections were incubated with
avidin-conjugated horse radish peroxidase complex (1:1000, Sigma-Aldrich, S2438) in PB for 30
min at RT. Antibody binding was routinely visualised using the chromogen DAB. For
counterstaining, dehydration and mounting, the general protocol for immunohistochemistry was
followed.
5.3.1.3 Double stainings
Double stainings were performed as previously described (Haider et al, 2011; Marik et al,
2007). For double labelling of E06 and TPPP/p25 for oligodendrocytes or GFAP for astrocytes,
E06
staining
was
basically
performed
as
described
in
the
general
protocol
for
immunohistochemistry but without antigen retrieval through steaming. E06 primary antibody was
applied, followed by alkaline phosphatase-conjugated anti-mouse secondary antibody (1:200,
Jackson ImmunoResearch) incubation and fast blue development (blue reaction product; see
materials part). For antigen retrieval of TPPP/p25 and GFAP, the sections were steamed in
EDTA buffer followed by TPPP/p25 or GFAP primary antibody application, species-specific
biotinylated anti-rat (1:1500, Jackson ImmunoResearch) or anti-rabbit antibody (1:2000, Jackson
ImmunoResearch), avidin-peroxidase incubation and development in amino ethylcarbazole
reagent (AEC, red reaction product).
In case of E06 double staining with APP, E06 primary antibody incubation was followed
by alkaline phosphatase-conjugated anti-mouse secondary antibody and fast blue development.
For antigen retrieval of APP, the sections were steamed in citrate buffer, followed by APP
Materials and Methods 97
primary
antibody
application,
biotinylated
secondary
antibody
(1:500,
Jackson
ImmunoResearch) and avidin-peroxidase incubation and AEC development.
Double labelling of Ox8 and ED1 was performed starting with ED1 primary antibody
application, biotinylated secondary antibody and avidin-peroxidase incubation and DAB
development, following the general immunohistochemistry protocol. Subsequently, the sections
were incubated with Ox8 primary antibody which was followed by alkaline phosphataseconjugated anti-mouse secondary antibody and fast blue development. In this case, no
intermediary steaming step was necessary, as DAB blocks all prior antibody sites.
5.3.1.4 Quantification of immunohistochemistry
Quantification of immunohistochemistry was performed by manual counting of CD3+ cells
or APP+ axonal spheroids. ED1+, Iba-1+, p22phox+, iNOS+ or iron-positive macrophages and
microglia were quantified by densitometric analysis of pictures taken under standardised
conditions (identical microscope settings controlled by white balance values, gain 1.2 and
exposure time of 6 ms) with a Reichert Polyvar 2 microscope using Nikon NIS-Elements D3.10.
Area fraction was determined using ImageJ. E06 and iron stainings were quantified by taking
pictures under the same standardised conditions. Optical density was determined by ImageJ
after converting the pictures to 8-bit, inverting and setting the scale to 1024 = 1. The stainings
were quantified from equally sized regions of interest in all cases.
The following macros were designed to optimise quantification of area fraction using
ImageJ. The macro “Maske” was designed to determine the total number of spinal cord pixels.
The macro “Analyze objects” was used to select the positively stained cells and determine their
pixel counts.
macro "Analyze objects [y]" {
a = getInfo("image.filename");
run("Colour Deconvolution", "vectors=[H DAB]");
b = a+"-(Colour_2)";
selectWindow(b);
//print(b);
run("8-bit");
run("Invert");
run("Set Measurements...", " mean redirect=None decimal=3");
run("Measure");
wert = getResult("Mean", (nResults-1));
//print(wert);
run("Subtract...", "value=wert");
run("Invert LUT");
setThreshold(60, 255);
run("Analyze Particles...", "size=0-Infinity circularity=0.00-1.00 show=Outlines display summarize");
Materials and Methods 98
}
macro "Maske [1]" {
run("8-bit");
run("Enhance Contrast", "saturated=0.35");
run("Invert");
run("Invert LUT");
setThreshold(60,255);
run("Convert to Mask");
run("Fill Holes");
run("Create Selection");
run("Set Measurements...", "area mean integrated redirect=None decimal=3");
run("Measure");
}
5.3.1.5 Pre-absorption of p22phox antibody
In order to reduce unspecific neuronal stainings caused by the primary antibody for
p22phox on rodent tissue, a pre-absorption of the antibody with Lewis rat cortex was performed.
300 mg cortex were homogenised per 1 ml TBS. The antibody was diluted in the homogenate
1:100 and incubated at 37 °C for 1 h in a rotating hybridisation oven. Then, the homogenate was
centrifuged for 10 min at 300 g. The supernatant was directly used for primary antibody
incubation.
5.3.2
Histochemistry
5.3.2.1 Haematoxylin and eosin staining (H&E)
Sections were deparaffinised 2x 15 min in xylene and rehydrated via 90%, 70% and 50%
ethanol series and finally rinsed in distilled water. The sections were incubated in haematoxylin
(Merck) for 5 min and rinsed in tap water. Then, the staining was differentiated in HCl/ethanol
and rinsed again in tap water. The sections were incubated in Scott´s solution for 5 min and after
another wash in tap water counterstained with eosin solution for 5 min. Finally, dehydration was
performed in ascending series of ethanol and n-butyl acetate. Sections were mounted in Eukit
(Gröpl Elektronenmikroskopie) with glass coverslips.
5.3.2.2 Luxol fast blue myelin stain
Sections were deparaffinised and rehydrated as described above. They were incubated
in 0.1% LFB solution overnight at 56 °C. After cooling to RT, the slides were washed in 96 %
Materials and Methods 99
ethanol and rinsed in tap water. This was followed by 5 min incubation in 0.1% lithium carbonate
for 5 min and differentiation in 70% ethanol (Differentiation is finished when only myelin sheaths
remain stained.). After rinsing the sections in tap water, they were placed in 0.8% periodic acid
for 10 min, rinsed in tap water and incubated in Schiff´s reagent (Merck) for 20 min. At last,
sections were treated with sulfit solution 2 times for 3 min. After rinsing the sections for 10 min in
running tap water, they were dehydrated and mounted as described above.
5.3.2.3 Histochemistry for non-heme tissue iron
Basically, the DAB-enhanced Turnbull blue staining (TBB) protocol was applied as
described (Meguro et al, 2007). Paraffin sections were deparaffinised and incubated in
ammonium sulphide (1:10 dilution in distilled water; Merck; ammonium sulphide should not be
stored longer than 6 months) for 90 min. Thereby, all Fe3+ is reduced to Fe2+. After thorough
rinsing in distilled water, the sections were treated with an aqueous solution of 10% potassium
ferricyanide (Merck) in 0.5% HCl, giving rise to the insoluble blue complex called Turnbull Blue.
Endogenous peroxidase activity was blocked in methanol with 0.01 M sodium azide and 0.3%
hydrogen peroxide for 60 min. Finally, the staining signal from Turnbull Blue was enhanced by a
20 min incubation in 0.025% DAB and 0.0005% H2O2 in 0.1 M phosphate buffer.
This protocol was established for PFA-fixed and TritonX-permeabilised cell cultures or
frozen sections. The treatments can be basically performed as described above with the
exception that endogenous peroxidases are blocked in PBS with sodium azide and 0.3% H2O2
instead of methanol. Iron-treated cells and control cells were washed with 5 µM DTPA in
medium for 5 min at RT in order to chelate extracellular iron. For cell culture double-stainings for
iron and cellular markers, the iron staining was performed until the peroxidase blocking.
Subsequently, the antibody staining was performed and finally the iron staining was enhanced
with DAB.
5.3.3
In vitro oxidation of native fresh frozen tissue sections
Although the antibodies recognising the E06 (oxidised phospholipids) and the 8OHdG (8Hydroxy-2′-deoxyguanosine, oxidised DNA) epitopes had been used before in rodent tissue, we
tested if the epitope could be produced and detected with similar sensitivity and specificity in
rodent tissue in our experimental settings. Wild-type Lewis rats were perfused with phosphate
buffer and brains were snap-frozen in pre-cooled isopentane. Sections (7 µm) were cut and
stored at -20 °C. For producing the E06 and 8OHdG epitopes, we used iron-induced oxidative
Materials and Methods 100
damage in vitro. Fe2+/ascorbate- and H2O2-induced lipid peroxidation was performed similarly as
described (Ribeiro et al, 2007; Takatsu et al, 2009). The sections were treated overnight at 37
°C with 20 µM FeSO4 (Merck), 1 mM ascorbate (Sigma Aldrich) and 500 µM tbH2O2 (Luperox
Sigma Aldrich) in TBS buffer. DNA oxidation was performed in a similar way using additionally
FeCl3/glutathione (GSH, 3 µM FeCl3 and 15 mM GSH) as a free-radical generating system as
described (Lodovici et al, 2001).
After in vitro oxidation, the slides were fixed in cold acetone/0.1% PFA at 4 °C for 20 min.
In case of E06, this was followed by delipidation in 50%, 70% and 90% increasing ethanol series
and xylene, each for 15 min. Subsequently, rehydration was performed by going through
decreasing ethanol series. In case of 8OHdG, the sections were pre-treated with proteinase K
for 15 min at 37 °C. Subsequently, the slides were blocked in 0.1 % BSA/TBS for 20 min and
primary antibodies for E06 and 8OHdG were applied overnight. Endogenous peroxidase activity
was blocked in 3% H2O2/TBS for 10 min and secondary antibodies were applied in 1%
BSA/TBS. Avidin peroxidase (for E06, 1:100, Sigma-Aldrich, A3151) or avidin alkaline
phosphatase (for 8HdG, 1:500, Sigma-Aldrich, AP020114754) were diluted in 0.1% BSA/TBS
(avidin peroxidase 1:100, avidin alkaline phosphatase 1:500) and development was performed
with DAB or fast blue, respectively.
5.3.4
Ferrocene assay for iron quantification
Rats aged from 2 to 20 months were sacrificed with an overdose of CO2 and perfused
with PBS. Cerebella were homogenised in sodium potassium phosphate buffer (pH 7.4, 5 mM).
The iron content of rat cerebella was determined using the colorimetric ferrocene assay (Fish,
1988). Iron was released from proteins and stable iron complexes by acidic permanganate
treatment at 60 °C for 2 hours. For 20 ml of this iron-releasing reagent, 10 ml 1.2 M HCl were
mixed with 10 ml 4.5% KMnO4 (Sigma-Aldrich). Subsequently, iron was reduced by ascorbic
acid to Fe2+ and quantitatively complexed by ferrocene. In order to catch interfering copper ions,
neocuproine was added to the reaction. This was accomplished by the iron detection reagent.
Absorption was measured at 540 nm using a Powerwave X-340 plate reader. Data were
normalised to protein concentration determined by total protein measurement using a Nanodrop
2000 spectrophotometer.
Materials and Methods 101
5.4 Glial Cell Culture Techniques
5.4.1
Mixed glia cultures
Mixed glia cultures were produced as described (Sharma et al, 2010). Mixed glia cultures
were obtained from brains of P1 Lewis rats (less than 48 h old). Postnatal pubs were
decapitated. The brains were removed from the scull and collected in RPMI medium. After
carefully removing the meninges using forceps (no. 5), the brains were transferred into a dish
containing mixed glia medium. The brains were dissociated by gentle trituration with a 10 ml
pipette and seeded in T75 cm2 cell culture flasks (one brain per flask). Prior to seeding, the
flasks were coated with 5 µg/ml poly-L-lysine (Sigma, P9155) in PBS (Lonza) for at least 3 hours
at 37 °C and 5% CO2. The flasks were washed three times with PBS before adding the cell
suspension. The cells were maintained in 10 ml mixed glia medium at 37 °C and 5% CO2. The
medium was replaced one day after seeding and then every other day. The cells constituted a
confluent layer after approximately one week.
5.4.2
Microglia cultures
After 5-7 days of mixed glia cell culture, a confluent layer mainly consisting of astrocytes
and some ependymal cell colonies covered the bottom of the flask with microglia and O2A
precursor cells growing on top of them. To detach the loosely adherent microglia from the
confluent cell layer, the culture flasks were tightly sealed and placed on a shaker for at least 1 h
at 37 °C and 170 rpm. Afterwards, the supernatant was collected and selective adherence was
performed on cell culture plastic dishes for 5-10 min at 37 °C and 5% CO2. Microglia attached to
the plastic bottom and contaminating astrocytes or O2A cells were carefully washed off with prewarmed mixed glia medium. Microglia were maintained in mixed glia medium until iron loading
experiments. For experiments including stainings, microglia were seeded on AclarTM plastic
coverslips (GE healthcare, PAA20012x).
The remaining mixed glia culture could be used for isolation of astrocytes or maintained
at 37 °C and 5% CO2 and used for at least one more shake-off.
5.4.3
Oligodendrocyte cultures
The protocol for oligodendrocyte cultures was established based on the combination of
different protocols and similar to (Chen et al, 2007). Mixed glia cultures for oligodendrocyte
Materials and Methods 102
isolation were produced from P1 Lewis rats. Postnatal pubs were decapitated. The brains were
removed from the scull and collected in HBSS (PAA, H15-007) on ice. The brains were
dissected by removing the cerebellum, separating the cortices, removal of the midbrain including
basal ganglia and peeling off the meninges using forceps (no. 5). The cortices were cut into
small pieces with a scalpel. The pieces from up to 10 cortices were collected in a 50 ml tube,
excess of HBSS was taken off and the tissue was gently triturated using a 1000 µl pipette tip in 1
ml of digestion medium. The tissue was digested for 30 min at 37 °C in a waterbath.
Subsequently, the tissue was homogenised to single cell suspension with a 1000 µl pipette tip
before stopping the digestion with the 10-fold volume of pre-warmed mixed glia medium for
oligodendrocyte cultures (DMEM/10% HS). The cells were centrifuged at 200 g for 10 min at 4
°C. The cell pellet was resuspended in DMEM/10% HS and seeded in PLL-coated flasks at a
density of 1.5 brains per T75 cm2 flask. The cells were maintained at 37 °C and 10% CO2 for at
least 10 days with the first medium change earliest after 3 days and then every other day.
For seeding of oligodendrocytes, glass cover glasses (Thermo Fisher, 12 mm,
004710183) were coated with PLL (100 µg/ml in PBS). After 10-12 days of mixed culture, the
flasks were put on an orbital shaker for 1 h at 37 °C at 180 rpm. The supernatant containing the
bulk of microglia was discarded and the monolayer comprising of astrocytes with O2A
precursors on top was washed with pre-warmed DMEM (without serum). Then, 10 ml of prewarmed DMEM (without serum) were added to each flask. The O2A precursors were shaken off
by hand and collected in plastic dishes with untreated surface (usually used for bacteria culture).
After 20 min adherence at 37 °C and 10% CO2, the dishes were swirled gently. The
supernatants were taken off, filtered through a 40 µm cell strainer and centrifuged at 200 g for
10 min in 15 ml tubes. The cells were thoroughly resuspended in Sato medium and 5-7 x 104
cells were seeded in 24-well plates on the PLL-coated cover glasses. For the first 2 days in
culture, the Sato medium was supplemented with 10 ng/ml PDGF (PeproTech, 100-13A) and
bFGF (R&D Systems, 233-FB) in order to support oligodendrocyte precursor proliferation.
Subsequently, the medium was replaced by Sato medium containing 0.5% HS but no PDGF or
bFGF anymore in order to enable oligodendrocyte maturation. The oligodendrocytes were
maintained at 37 °C and 10% CO2 with half of the medium exchanged every other day.
5.4.4
Myelinating spinal cord cultures
Myelinating spinal cord cultures were generated as described previously (Sorensen et al, 2008).
Materials and Methods 103
5.4.4.1 Neurosphere-derived astrocytes
Neurospheres were generated as described (Zhang et al, 1998) and differentiated into
astrocytes as outlined in (Sorensen et al, 2008). Neurospheres were produced from the corpus
striatum of P1 SD rats. The brains were removed and cut sagittal with a scalpel to separate the
two hemispheres. The corpus striatum was dissected by one cut at the frontal tip of the corpus
callosum and one at the lateral ventricle. The corpus striatum was then removed using curved
forceps and collected in L-15 medium. The tissue was dissociated by gentle trituration using a
glass Pasteur pipette and centrifuged at 140 g for 5 min. The resulting pellet from 2 brains was
resuspended in NSM and seeded in T75 cm2 flasks in 20 ml NSM supplemented with 5 ng/ml
EGF (Peprotech, 315-09) in order to promote sphere formation. The Neurospheres were
maintained at 37 °C and 7.5% CO2 and fed twice a week by adding 5 ml NSM and 5 ng/ml EGF
(for the total volume of medium). After approximately 1 week, the neurospheres reached a
certain size and started to attach to the bottom of the flask. Then, in order to differentiate the
neurospheres to astrocytes, the neurospheres were centrifuged at 140 g for 5 min. The cells
were resuspended in astrocyte medium (DMEM/10% FCS) by gentle trituration with a glass
Pasteur pipette and seeded on PLL-coated glass coverslips in 24-well plates. The coverslips
(13mm, VWR, 631-0150) were coated with 13.3 ng/ml PLL (Sigma-Aldrich, P1274) for at least 1
h at 37 °C, rinsed thoroughly in sterile distilled water, placed into 24-well plates and left to air dry
before use. Depending on the neurosphere culture quality, from one flask of neurospheres 4-6
24-well plates were plated for differentiation into astrocytes. The astrocytes were maintained at
37 °C and 7.5% CO2 with half of the medium exchanged twice a week until the cultures were
confluent (approximately 7 days).
5.4.4.2 Dissociated spinal cord cultures
Dissociated spinal cord cultures were generated from E15.5 SD rat embryos. Embryos
were taken out from the gravid uterus and decapitated. The skin covering the spinal cord was
gently removed and the spinal cord was carefully dissected. The meninges were thoroughly
removed and 3-5 spinal cords were collected per 1 ml HBSS (Invitrogen) on ice. After addition of
100 µl trypsin (2.5%, Sigma-Aldrich, T4549) and collagenase (1% in L15, Invitrogen, 17100-017)
to 1 ml HBSS, the tissue was gently dissociated with a Pasteur pipette and incubated for 15 min
at 37 °C and 7.5% CO2. The digestion was stopped by adding 2 ml SD solution and the tissue
was triturated into a single cell suspension using a Pasteur pipette. The cells were centrifuged at
Materials and Methods 104
200 g for 5 min and resuspended in plating medium. A live cell count was performed using a
haemocytometer and trypan blue and diluted in plating medium to 3x106 cells/ml.
The dissociated spinal cord cells were plated on top of the astrocyte monolayer of
neurosphere-derived astrocytes. Three coverslips of astrocytes were placed into a 35 mm petri
dish and 50 µl (150.000 cells) of spinal cord cells were placed onto each coverslip. The cells
were left to attach for approximately 2 h at 37 °C and 7.5% CO2. Subsequently, 600 µl DM+ and
450 µl plating medium was added to each dish. The cultures were maintained for up to 30 days
at 37 °C and 7.5% CO2 and fed every other day with DM by exchanging half of the medium.
After 12 days, insulin was omitted from the medium in order to promote oligodendrocyte
differentiation.
5.4.5
Iron loading of cells
Iron loading of cells was performed as published (Rathore et al, 2012; Schulz et al,
2011). Irrespective of the type of culture, the cells were washed with medium devoid of serum or
transferrin prior to iron loading. Subsequently, the cells were incubated for 1 h at 37 °C in 5 or
7.5% CO2 (depending on the culture) in medium without serum or transferrin. The cells were
treated with FeCl3 plus ascorbate, prepared in a ratio of 1:44 in distilled water. Ferritin (Sigma
F4503) treatments were performed without serum/transferrin starvation by directly adding the
respective dilution to the medium. The iron concentration of ferritin was according to the
manufacturer between 6.97 mg/ml und 14.9 mg/ml.
5.4.6
Immunofluorescence of cell cultures
Cells were fixed with 4% PFA for 20 min at RT, then washed in PBS and permeabilised
for 10 min in 0.5% TritonX100/PBS. After 30 min blocking in blocking buffer (1% BSA, 10%
Horse Serum in PBS), the primary antibodies were applied for 45 min at RT. For details about
primary antibody dilutions and species see Table 5. After washing in PBS, the secondary
antibodies were applied 1:400 for 15 min at RT. Finally, the coverslips were washed 3 times in
PBS and once in distilled water and mounted in mowiol plus Dapi.
For live stainings, primary antibodies were applied in cold DMEM for 30 min at 4 °C. After
a gentle wash in cold DMEM, the cells were fixed in 4% PFA and the staining protocol was
continued as described above.
Materials and Methods 105
Table 5: Primary antibodies for immunofluorescence stainings of cell cultures
Antibody
Species
Isotype
Dilution
Fixation
Source
GalC
mouse
IgG3
1:100
PFA + Triton
Millipore, MAB342
ED1
mouse
IgG1
1:100
PFA + Triton
Abcam, ab3160
FTL
rabbit
IgG
1:200
PFA + Triton
Proteintech, 10727-1-AP
GFAP
rabbit
IgG
1:500
PFA + Triton
Dako, Z0334
Iba-1
rabbit
IgG
1:1000
PFA + Triton
Wako, 01-1974
MOG/Z2
mouse
IgG2a
1:100
live
Linington lab
PLP/O10
mouse
IgM
1:100
PFA + Triton
Linington lab
SMI31
mouse
IgG1
1:1000
PFA + Triton
Abcam
5.5 Statistical analysis
Descriptive statistical analysis involved median value and range, box plots and scatter
plots. The linear dependence of samples was classified by Pearson correlation test. For
comparison of multiple groups of normally distributed data, one-way ANOVA followed by pairwise comparisons and Tukey adjustment was calculated. In case of multiple comparison of
not normally distributed data derived from densitometry, Kruskal-Wallis group testing
was followed by Mann-Whitney U post hoc tests and Bonferroni-Holm correction. Active
lesions of experimental models were compared with the corresponding age control,
although there was no significant difference detected between young and aged control
animals (except for iron staining in basal ganglia which was considered separately).
A p-value ≤ 0.05 was considered as statistically significant. Statistical analyses was performed
using PASW Statistics 18 (SPSS Inc.).
References 106
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References 131
Zinkernagel RM, Althage A (1977) Antiviral protection by virus-immune cytotoxic T cells: infected target
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644-651
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Disease. Science 148: 1477-1479
Appendix 132
Curriculum Vitae
Cornelia Schuh
PERSONAL DATA
rd
Date of Birth:
January 23 , 1985
Place of Birth:
Oberpullendorf, Burgenland, Austria
Citizenship:
Austria
Contact:
Oelweingasse 30/12
1150 Vienna
 cornelia.schuh@gmx.at
 +43 664 75073300
EDUCATION
06/2012 – 12/2012
Stay at the University of Glasgow, Institute of Infection, Immunity and
Inflammation; funded by a Du Pre´ grant of Medical and Scientific Research
Multiple Sclerosis International Federation
03/2010 – 02/2014
PhD thesis within the PhD program “Cell Communication in Health and Disease”
(CCHD) of the Medical University of Vienna
07/2008 – 07/2009
Diploma thesis in the department of Tumour Immunology, Children´s Cancer
Research Institute (CCRI), Vienna (“Immune Suppressive Mechanisms in Tolllike-Receptor 4 (TLR4) activated Dendritic Cells”)
10/2003 – 12/2009
Studies of Molecular Biology, University of Vienna, Austria
Main focus on Immunology and Cell Biology
09/1995 – 06/2003
Secondary School, BG/BRG Oberschützen
WORK EXPERIENCE
03/2010 – 02/2014
PhD study
Medical University of Vienna
Centre for Brain Research, Department of Neuroimmunology
Supervisor: Univ. Prof. Dr. Hans Lassmann
“Oxidative tissue injury in experimental disease models
of multiple sclerosis”
Appendix 133
07/2008 – 07/2009
Diploma Thesis
Children Cancer Research Institute (CCRI)
Department of Tumour Immunology, Supervisor: Dr. Thomas Felzmann
“Immune Suppressive Mechanisms in Toll-like-Receptor 4 (TLR4) activated
Dendritic Cells”
10/2007 – 11/2007
Internship Novartis Austria
Supervisor: Dr. Tamas Schweighoffer
“Characterisation of antibodies specific for degranulated mast cells”
“Expression of transglutaminase 2 and microtiter plate transglutaminase assay”
08/2007 – 09/2007
Internship Max F. Perutz Laboratories University Vienna
Department of Chromosome biology, Supervisor: Ao.Univ. Prof. Dr. Michael
Jantsch
“Finding activators and inhibitors of ADAR”
02/2007 – 03/2007
Internship Children Cancer Research Institute (CCRI)
Department of Tumour Immunology, Supervisor: Dr. Thomas Felzmann
“Immune stimulatory capacity of IDO silenced dendritic cells”
08/2006
Internship CCRI
Department of Molecular Biology, Supervisor: Univ.-Prof. Dr. Heinrich Kovar
“Jagged1 cloning of the Jagged1 promoter sequence into pGL4-Luc2”
PARTICIPATION IN CONFERENCES
10/2013
07/2013
09/2012
06/2012
09/2011
th
29 Congress of the European Commitees for Treatment and Research in
Multiple Sclerosis (ECTRIMS), Copenhagen, Denmark; “Iron accumulation and
oxidative damage in various models for inflammation/degeneration of the central
nervous system.” (poster presentation)
th
11 European Meeting on Glial Cells in Health and Diseases, Berlin, Germany;
“Development of an in vitro model to study the impact of iron in inflammatory
demyelinating diseases.” (poster presentation)
European Congress on Immunology, Glasgow, Scotland; “The role of iron in
inflammatory demyelinating diseases: development of experimental in vitro and in
vivo models.” (poster presentation)
th
8 PhD-Symposium of the Medical University of Vienna, Vienna, Austria; “The
role of iron in inflammatory demyelinating diseases: development of experimental
in vitro and in vivo models.” (poster presentation)
th
10 European Meeting on Glial Cells in Health and Diseases, Prague, Czech
Republic; “Iron accumulation in models for inflammation/degeneration of the
central nervous system: Implications in chronic multiple sclerosis?” (poster
presentation)
Appendix 134
07/2011
06/2011
12/2010
09/2010
06/2010
09/2009
11/2008
th
11 ESNI Course, European School of Neuroimmunology, Glasgow, Scotland;
“Iron accumulation in models for inflammation/degeneration of the central nervous
system: Implications in chronic multiple sclerosis?” (Abstract Book)
th
7 PhD-Symposium of the Medical University of Vienna, Vienna, Austria; “Iron
accumulation in models for inflammation/degeneration of the central nervous
system: Implications in chronic multiple sclerosis?” (poster presentation)
Annual meeting of Austrian Society for Allergology and Immunology (ÖGAI),
Vienna, Austria; “Iron accumulation in central nervous system inflammation and
degeneration.” (poster presentation)
FEBS & EFIS Workshop Inflammatory Diseases and Immune Response: Basic
Aspects, Novel Approaches and Experimental Models, Vienna, Austria; “Iron
accumulation in models for inflammation/degeneration of the central nervous
system.” (poster presentation)
th
6 PhD-Symposium of the Medical University of Vienna, Vienna, Austria; “Iron
accumulation and inflammation/degeneration in the central nervous system.”
(poster presentation)
International Joint Symposium of four Collaborative Research Centres from Berlin
and Hannover: Tolerance and Immune-Regulation, Berlin, Germany; “Toll-like
Receptor 4 and CD40 Signalling in Dendritic Cells Induce a PHD Finger Family
Transcription Factor with Strong Immune Suppressive Potential.” (poster
presentation)
Bridging Innovation and Translation in Paediatric Oncology, Vienna, Austria;
“Identification of Potential Immune Regulatory Master Switches Induced by Tolllike Receptor 4 and CD40 Signalling in Dendritic Cells.”
PUBLICATIONS
1. C Schuh, I Wimmer, S Hametner, L Haider, AM Van Dam, R Liblau, KJ Smith, CJ Binder, J
Bauer, M Bradl, D Mahad, H Lassmann. (2014). “Oxidative tissue injury in multiple sclerosis is
only partly reflected in experimental disease models”
2. A M Dohnal, A Halfmann, M Le Bras, S Vittori, C Schuh, R Luger, D Stoiber, A Kotlyarov, M
Gaestel, T Felzmann. (2014, in revision). “The MAPK-activated kinase MK2 attenuates dendritic
cell-triggered TH1 and TH17 immune responses”
3. M Lindner, K Thümmler, A Arthur, S Brunner, C Elliott, H Mohan, A Williams, J Edgar, C Schuh,
C Stadelmann, S Barnett, H Lassmann, S Mücklisch, M Mudaliar, N Schaeren-Wiemers, E Meinl,
C Linington (2014, in revision) “Fibroblast growth factor signalling in multiple sclerosis: inhibition of
myelination and induction of pro-inflammatory signalling environment by FGF”
AWARDS
th
PhD-Symposium of the Medical University of
th
PhD-Symposium of the Medical University of
06/2012
Vienna
Poster prize at the 8
06/2010
Vienna
Poster prize at the 6
FUNDING
07/2013
07/2012
th
Travel grant for 29 Congress of the European Commitees for Treatment and
Research in Multiple Sclerosis (ECTRIMS), Copenhagen, Denmark
Du Pré Fellowship of Medical and Scientific Research Multiple Sclerosis
International Federation (MSIF) for a 6 months stay at University of Glasgow;
Institute of Infection, Immunity and Inflammation
Appendix 135
07/2012
07/2011
01/2010
“Top Stipendium des Landes Niederösterreich” for 6 months stay at the University
of Glasgow, Institute of Infection, Immunity and Inflammation
th
Travel grant for the 11 ESNI Course, European School of Neuroimmunology,
Glasgow, Scotland
“Top Stipendium des Landes Niederösterreich” for successful graduation in
Molecular Biology at the University of Vienna
TEACHING EXPERIENCE
04/2013 - 05/2013
Basics of Neuroscience (University of Vienna)
300113; UE Basics of Neuroscience
Including conception of taught subjects, the corresponding lectures and handouts
and exam questions
03/2013
“Brain Awareness Week” in the Centre for Brain Research Vienna
Education of secondary school pupils
03/2012
“Brain Awareness Week” in the Centre for Brain Research Vienna
Education of secondary school pupils
ADDITIONAL QUALIFICATIONS
02/2014
th
Member of organizing committee of the 7 international workshop “Bridging the
Gap” hosted by the students of the CCHD PhD program
02/2012
th
Member of organizing committee of the 5 international workshop “Bridging the
Gap” hosted by students of the CCHD PhD program
02/2011
th
Member of organizing committee of the 4 international workshop “Bridging the
Gap” hosted by students of the CCHD PhD program
06/2002
Cambridge First Certificate of English, Grade A
FURTHER INTERESTS
Playing the saxophone in the brass marching band “MV Gschaidt”
Martial arts Ebmas Wing Tsun, travelling (especially to Scotland)
Hiking, cross-country skiing