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STUDIES OF CELL DEATH IN PARKINSON’S
DISEASE USING ORGANOTYPIC CELL
CULTURES
TUYET TB TRAN
B.Sc (Hons)
Discipline of Health Sciences, School of Medical Sciences
The University of Adelaide
September 2008
A thesis submitted to the University of Adelaide in fulfillment of the requirements for the
degree of Doctor of Philosophy
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DECLARATION
This work contains no material which has been accepted for the award of any other
degree or diploma in any university or other tertiary institution and that, to the best of my
knowledge and belief, the thesis contains no material previously published or written by
another person, except where due references has been made in the text.
I give consent to this copy of my thesis, when deposited in the University library, being
made available for photocopying and loan
Tuyet Tran
Date:
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PUBLICATIONS AND PRESENTATIONS
The following articles have been published or accepted for publication or presentation
during the period of my PhD candidature, and sections of these articles are included in
the present thesis.
Publications under review:
Tuyet T.B Tran, Peter C. Blumbergs and James A. Temlett (2008)
The effects of MPTP and rotenone on Dopaminergic neurons in Organotypic cell culture.
Journal of Neurotoxicology
Abstracts:
Tuyet T.B Tran, Peter C. Blumbergs and James A. Temlett (2007)
The Effects of MPTP and rotenone on Dopaminergic neuronal growth in Organotypic
Cell Culture. Abstracts of the 8th International Conference AD/PD 2007, Salzburg,
Austria. March 14 – 18, p327
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ACKNOWLEDGEMENTS
I wish to convey my gratitude and appreciation to the following people.
Professor James Temlett, my primary supervisor, thank you for providing me the
opportunity to undertake my PhD.
Professor Peter Blumbergs, my co-supervisor, for your time and helpful critique of my
thesis and the never-ending support.
Renee Turner and Emma Thornton, for your friendship and support. Through the good
and bad times you guys were there for me, I will cherish our friendship (especially all the
baking and Friday lunches) whether you are near or far. Wish you guys all the best for
the future and research careers.
James Donkin and Islam Hassan, for your friendship and sharing your PhD and masters
experience with me. I wish you guys the best for the future.
The Eye-boys: John Wood, Glyn Chidlow and Michael Schober, for your support and
helpful tips in thesis writing, culturing, and immunostaining. Also providing me an
opportunity to work and gain experience for my research career.
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Ghafar Sarvestani, for your confocal expertise and taking your time teaching me how to
use the expensive equipment.
The Neuroscience Lab- Jim Manavis, for your immunohistochemistry expertise, your
jokes and support, Kathryn, Yvonne, Sven, Cathy, Bernice, Sophie and Felicity, for you
assistance when needed and friendship, I’ll miss our international days in the tea room.
My family and friends for your support, encouragement and understanding through the
never-ending years of studying.
Finally, very special thanks to a particular person who put up with all my whinging and
complaining during the few remaining years of my PhD, Luke Francis, for your
unconditional love and support.
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ABBREVIATIONS
AA -ascorbic acid
AA(A/D)C -aromatic l-amino acid decarboxylase
Ab -antibody
ACh -acetylcholine
AChE -acetylcholinesterase
AD -Alzheimer’s disease
AIF -apoptosis-inducing factor
α-syn -alpha synuclein
ALDH -aldehyde dehydrogenase
AMPA -alpha-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid
AMPT -alpha-methyl-p-tyrosine
ANOVA -analysis of variance
AP -antero posterior
APTP -1-amino-4-phenyl-1,2,3,6-tetrahydropyridine
APP+ -1-amino-4-phenyl-pyridinium
ARs -aldose reductase
ART -artemin
ATP -adenosine triphosphate
Bax -�Bcl-2-associated protein X
BDNF -brain-derived neurotrophic factor
bFGF -basic fibroblast growth factor
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BG -basal ganglia
BJAB -B lymphoma cell line
BMPs -bone morphogenic factors
BSA -bovine serum albumin
BSS -balanced salt solution
Ca2+ -calcium
cAMP -3'-5'-cyclic adenosine monophosphate
CGC -cerebellar granule cells
CoQ(10) -coenzyme Q(10)
COMT -catechol-O-methyl transferase
CNS -central nervous system
Cu-SOD -copper superoxide dismutase
cGMP -Guanosine 3,5-cyclic monophosphate
DA -dopamine
DAergic -dopaminergic neurons
D1 -dopamine receptor, type 1
D2 -dopamine receptor, type 2
DAB -3,3’-diaminobenzidine-Tetrahydrochloride
DAPI -4,6-diamino-2-phenolindol dihydrochloride
DAT -dopamine transport
DBS -deep brain stimulation
DCF -2’,7’-dichlorofluorescin
DIV -days in vitro
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DMEM -modified basal medium, Eagle
DMSO -dimethyl sulfoxide
DLBD -dementia with LB disease
DOPAC -3,4-dihydroxyphenylacetic acid
DOPAL -dihydroxyphenylacetaldehyde
DOPET -3, 4-dihydroxyphenylethanol
DPX -DePex mounting medium
DTC -dithiocarbamate
DV -dorso ventral
E -embryonic
EDTA -ethylenediaminetetraacetic acid
ELISA -enzyme-linked immunosorbent assay
En-1 -engrailed
Enk -enkephalin
Epo -erythropoietin
ERKs -extracellular signal-regulated kinases
ETC -electron transport chain
FADD -Fas-associated death domain
FGF8 -fibroblast growth factor-8
FD -fluorodopa
FJC -fluoro jade C
Foxa2 -Forkhead box protein A2
FP -floor plate
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FPD - familial Parkinsons Disease
GABA -gamma-aminobutyric acid
GADPH - glyceraldehyde-3-phosphate dehydrogenase
GBSS -Geys balanced salt solution
Gbx2 -gastrulation brain homeobox 2
GDNF -glial-derived neurotrophic factor
GDF5 -growth/differentiation factor 5
GFAP -glial fibrillary acidic protein
GLU -glutamate
GM-CSF -granulocyte macrophage colony-stimulating factor
GP -globus pallidus
GPe -external segment of the globus pallidus
GPi -internal segment of the globus pallidus
GPx -glutathione peroxidase
GSSG -oxidised glutathione
GSH -glutathione
h/hr -hour
H&E -haemoatoxylin and eosin
HBSS -Hank’s balanced salt solution
HB-EGF -heparin-binding epidermal growth factor
Hesc -human embryonic stem cells
HL-60 -human promyelocytic leukemia cell line
HP100 - H2O2-resistant cell line
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HRP -horseradish peroxidase
HT1080 -human fibrosarcoma cell line
H2O2 -hydrogen peroxide
H2SO4 - sulphuric acid
5-HT -serotonin
6-OHDA -6-Hydroxydopamine
Fe 2+ -iron
IBMX -isobutylmethylxanthine
IMVS -Institute of Medical and Veterinary Science
IFN-� -interferon gamma
IL-1β -interleukin-1β
IL-6 interleukin-6
iNOs -inducible nitric oxide synthase
IPD -idiopathic Parkinsons’s disease
JNK -Jun N-terminal kinase
LB -Lewy body
L-dopa -levodopa
LDH -lactate dehydrogenase
Lmx1 -Lim-domain transcription factors
LPS -lipopolysaccharide
MAPK -Mitogen-activated protein (MAP) kinases
mDNs -midbrain/mesencephalic dopaminergic neurons
M -molar
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MAO-B -monoamine oxidase B
MHB -mid-hindbrain organizer
MHJ -midbrain–hindbrain junction
μm -micromolar
μl -microlitres
mg -milligrams
mins -minutes
MN9D -midbrain-derived dopaminergic neuronal cell line
Mn-SOD -magnesium superoxide dismutase
Mn–EBDC -manganese ethylene-bis-dithiocarbamate
MPDP -1-methyl-4-phenyl-2,3-dihydropyridinium
MPPC -1 -methyl-4-phenylpyridine
MPPP -1-methyl-phenyl-propion-oxypiperidine
MPP+ -N-methyl-1-4-phenylpyridinium ion
MPTP -1-methyl-4-phenl-1,2,3,6-tetrahydropyridine
Mw -molecular weight
NA -noradrenaline
NAC -N-acetylcysteine
NDI1 -single-subunit nicotinamide adenine dinucleotide (reduced)
dehydrogenase of Saccharomyces cerevisiae
NMDA -N-methyl-D-aspartic acid
NADPH -Nicotinamide adenine dinucleotide phosphate
NADH -Nicotinamide adenine dinucleotide
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NADH-DH -NADH-dehydrogenase
3-NT -3-Nitrotyrosine
n -number
nM -nanomolar
NGF -nerve growth factor
NM -neuromelanin
NO• -nitric oxide
NTF -neurotrophic factor
NRT -neurturin
NSAIDs -nonsteroidal anti-inflammatory drugs
NT-3 -neurotrophin-3
NT-4/5 -neurotrophin-4/5
NMDA -N-methyl-D-aspartate
NF- �B -nuclear factor �B
Nurr1 -Nuclear receptor related 1
O2 -oxygen
OH• -hydroxyl radical
•O2- -superoxide anion radical (superoxide)
ONOO– -peroxynitrite
OS -oxidative stress
Otx2 -homeobox protein
PARP -poly (ADP-ribose) polymerase
Pax- -paired-like homeodomain proteins
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PBS -phosphate buffered saline
PC12 cells -rat pheochromocytoma cell line
PD -Parkinson’s disease
P7 -postnatal day 7 (seven day old)
P75 -low affinity neurotrophin receptor
PBS -phosphate buffered saline
PET -positron emission tomography
Pitx3 -paired-like homeodomain transcription factor 3
PKC -Protein Kinase C
PPX -pramipexole
PSP -persephin
Ref -reference
ROI -reactive oxygen intermediates
ROP -ropinirole
ROS -reactive oxygen species
RRF -retrorubral field
SAPK -stress-activated protein kinase
SD -standard deviation
SEM -standard error of the mean
sFas - soluble Fas
Shh - Sonic hedgehog
SH-SY5Y -human dopaminergic cells
SK-N-SH -human neuroblastoma cell line
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SN -substantia nigra (A9)
SNpc -substantia nigra pars compacta
SNpr -substantia nigra pars reticulate
SOD -superoxide dismutase
SP -substance P
SPC -Pierce peroxidase conjugated streptavidin tertiary
STN -subthalamic nucleus
ST -striatum
RPM -revolutions per minute
TBARS -thiobabituric acid reactive substances
TBS -Tris buffered saline
TH -tyrosine hydroxylase
TH-ir -tyrosine hydroxylase immunoreactive
TGF-� -transforming growth factor beta
TMB -3,3�,5,5�-Tetramethylbenzidine
TNF -tumor necrosis factor
TNFα-R1 - tumor necrosis factor alpha receptor 1
TRADD -TNFRSF1A-associated via death domain
TUNEL -terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling
Trk - tyrosine receptor kinase
UV -ultraviolet
VM -ventral mesencephalon
VMAT -vesicular monoamine transporter
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VMB -ventral midbrain
VTA -ventral tegmental area (A10)
Wnt1 -Wingless-type MMTV integration site family
WT -wild type
Zn-SOD -zinc superoxide dismutase
Z-DEVD-fmk -Caspase-3 Inhibitor
oC -degrees Celsius
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TABLE OF CONTENTS
CHAPTER 1: …………………………………………………………………….……....1
OVERVIEW OF PARKINSON’S DIESEASE…………………………………….….....1
1A
1.0 INTRODUCTION………………………………………………………………...…2
1.1 Incidence and prevalence of PD) …………………………………………………...2
1.2 Clinical Characteristics of PD…………………………………………………….…3
1.3 Neurochemical and neuropathological Features of PD…………………………....4
1.3.1 Cell loss in PD………………………………………………………………6
1.3.2 Lewy Bodies in PD………………………………………………………….6
1.4 Etiology of PD……………………………………………………………………...…6
1.4.1 Environmental Factors…………………………………………...............….6
1.4.2 Genetic Factors………………………………………………………….......9
1.4.3 Ageing and PD development……………………………………………....12
1.5 Mechanisms of Parkinsonian cell death…………………………………..………12
1.5.1 Oxidative stress (OS) ……………………………………………………...12
1.5.2 Iron……………………………………………………………....................14
15.3 Neuromelanin (NM) ………………………………………………………..16
1.5.4 Glutathione (GSH) ………………………………………………………...17
1.5.5 Mitochondrial Dysfunction………………………………………………...19
1.5.6 Excitotoxicity………………………………………………………………20
1.5.7 Nitric Oxide………………………………………………………………..22
1.5.8 Inflammation……………………………………………………………….23
1.6 Cell death in PD…………………………………………………………………….25
1.6.1 Apoptosis…………………………………………………………………..25
1.6.2 Apoptotic markers in PD…………………………………………………..25
1.7 Current treatments and potential therapies for PD……………………………...27
1.7.1 Symptomatic Treatment……………………………………………………27
1.7.2 Cell transplantation …………………………………………………….….28
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1.7.3 Gene therapy …………………………………………………………..…..29
1.7.4 Surgical methods …………………………………………………………..30
1B
1.8 BASAL GANGLIA………………………………………………………………....33
1.8.1 Introduction…………………………………………………………………….....33
1.8.2 Striatum (ST) …………………………………………………………………..…34
1.8.2.1 Dopamine receptors (D1R & D2R) ……………………………………..35
1.8.3 Pallidal complex………………………………………………………………..…36
1.8.3.1 Entopeduncular nucleus (medial globus pallidus) ………………………….36
1.8.3.2 Globus pallidus (lateral globus pallidus) …………………………………...38
1.8.4 Substantia nigra (SN) ……………………………………………….…………...39
1.8.4.1 Substantia nigra pars compacta (SNpc) ………………………………….....39
1.8.4.2 Substantia nigra pars reticulate (SNpr) …………………………………......40
1.8.5 Subthalamic nucleus……………………………………………..….................…41
1.8.6 The basal ganglia circuitary……………………………………………………...42
1.8.6.1 The direct and indirect pathway model…………………………………..42
1.8.6.2 BG changes in PD…………………………………………..…………....44
1 C
THE NIGROSTRIATAL PATHWAY……………………………………...………...49
1.9 Introduction…………………………………………………………………....……49
1.9.1Ontogeny of Midbrain Dopaminergic neurons (mDA) ………………………...49
1.9.2 In vivo development of the nigro-striatal pathway……………………...……...50
1.9.3 In vivo development of patch/matrix innervation……………………………....51
1.9.4 In vitro modelling of nigro-striatal pathway……………………………….……52
1.10 FACTORS INFLUENCING THE DEVELOPMENT OF MESENCEPHALIC
DOPAMINE NEURONS (mDNs) ……………………………………….……………54
1.10.1 Early development………………………………………………..…………..…54
1.10.2 Specification of mitotic mesencephalic precursors to the mDN
fate………………………………………………………………..……………………...55
1.10.3 Postmitotic development of mDNs…………………………………………..…56
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1.10.4 Functional maturation and axonal pathfinding……………………………….59
1.11 Neurotrophins and the development of mDNs………………………………..…62
1.11.1 Glial-cell-line-derived neurotrophic factor…………………………….…63
1.11.2 GDNF family-ligands…………………………………………………….64
1.11.2.1 Neurturin………………………………………………..………………65
1.11.2.2 Neublastin/artemin and persephin……………………………………...65
1.11.3 Neurotrophins………………………………………………………….....66
1.11.4 Local factors………………………………………………………………67
1.11.4.1 Transforming growth factor-beta…………………………………….…68
1.11.5 Low molecular weight compounds……………………………………….68
1.11.6 Immunophilin ligands…………………………………………………….69
1.12 Changes in cytokine levels, decline of essential NTFs and links with
PD………………………………………………………………………………………..70
1.12.1 Human evidence…………………………………………………………..70
1.12.2 Evidence from experimental PD models ………………………...………71
CHAPTER 2: …………………………………………………………………………...73
IN VIVO PARKINSONIAN MODELS: MPTP & ROTENONE…….………………..73
2.0 INTRODUCTION……………………………………………………………….…74
2.1 MPTP………………………………………………………………..........................74
2.2 Biochemistry of MPTP…………………………………………………………..…75
2. ANIMAL MODELS OF PD USING MPTP……………………………..………....78
2.3.1 Factors influencing the neurotoxic action of MPTP………………..…...…78
2.3.2 Behavioural changes……………………………………………….…..…..78
2.3.3 Histopathological changes…………………………………………...….....80
2.3.4 Neurochemical changes…………………………………..……..…………81
2.3.5 Invertebrates……………………………………………………………......82
2.3.6 Genetic mouse models and MPTP………………………………..………..83
2.4 Mechanisms of MPTP neurotoxicity……………………………………................83
2.4.1 Mitochondrial impairment……………………………………………........83
2.4.2 Energy Failure………………………………………………..………….....85
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2.4.3 Calcium homeostasis……………………………………………………....85
2.4.4 Glutamate release………………………………………………..................86
2.4.5 Reactive oxygen (ROS) and nitrogen species (NOS) ………………..........86
2.4.6 Cytokines and inflammatory processes…………………………...……….88
2.4.7 MPTP and apoptosis…………………………………………………….…90
2.5 DA agonists and MPTP………………………………………………..……...…....92
2.6 ROTENONE…………………………………………………………..…………….95
2.6.1 Introduction………………………………………………….................…..95
2.6.2 Biochemistry of rotenone……………………………………..……………95
2.7 Rotenone-treated animal PD Models……………………………………...............96
2.7.1 Rodents……………………………………………………..……………...96
2.7.2 Invertebrates……………………………………………...……………...…98
2.8 Rotenone and PD Pathology…………………………………………………..…...99
2.9 Rotenone and L-DOPA………………………………………………………..….100
2.10 Mechanisms of Rotenone toxicity………………………………..……………...100
2.10.1 The Complex 1 Inhibitor………………………………………...............100
2.10.2 Rotenone and OS……………………………………………………......102
2.10.3 Rotenone and Inflammation……………………………………………..124
2.10.4 Rotenone and Apoptosis………………………………………………...103
2.11 Other Parkinsonian-inducing neurotoxins………………..……………104
2.11.1 6-hydroxydopamine (6-OHDA) ……………………………………...…104
2.11.2 Maneb…………………………………………………………………...106
2.11.3 Paraquat………………………………………………………………….108
2.12 Summary……………………………………………………………………….....110
CHAPTER 3: ………………………………………………………….........................112
IN VITRO MODELS OF PD………………………………………………………….112
3.0 Introduction…………………………………………...…………………………...113
3.1 Dissociated cultures…………………………………………………….................113
3.2 Reaggregate cultures……………………………………………………………. 115
3.3 Organotypic explant cultures………………………………………….………….116
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3.4 Organotypic slice cultures…………………………………………..…………….117
3.5 In Vitro Systems And Their Relevance To PD……….………………………….121
3.5.1 MPTP & In Vitro Studies………..……………..…………………………121
3.5.1.2 DAergic Neurons-Target Specific Toxicity………………………….…123
3.5.1.3 Complex 1 Inhibitor…………………………………….…...………….124
3.5.1.4 Oxidative Stress (OS) …………………………………….…................125
3.5.1.5 Inflammation: The Contribution of Microglia and Cytokines……….…127
3.5.1.6 Cell Death………………………………………………………..……..129
3.5.1.7 DA Agonists & Antioxidants………………………………………..….132
3.5.2 Rotenone & In Vitro Studies……………...………………………………135
3.5.2.1 DAergic Toxicity…………………………………………………….....136
3.5.2.2 Oxidative Stress (OS)……………………………………………….….138
3.5.2.3 α-Synuclein………………………………………………………….….140
3.5.2.4 Complex I Inhibitor……………………………………………………..141
3.5.2.5 Inflammation……………………………………………………………142
3.5.2.6 Cell death via Apoptosis………………………………………………..144
3.5.2.7 DA Agonists and Antioxidants………………………………………....147
3.6 ORGANOTYPIC CELL CULTURE: IN VITRO MODEL FOR PD………….147
3.6.1 MPTP application in organotypic cell cultures………………………………..148
3.6.2 Rotenone and Organotypic Cell Culture…………...……………………….…149
4.0 AIMS:………………………………………………………………........................151
CHAPTER 5: ………………………………………………………….........................152
MATERIAL & METHODS……………………………………………........................152
5.1 Animal care……………………………………………………………...................153
5.1.1 Ethics……………………………………………………………………...153
5.1.2 General………………………………………………………………..…..153
5.2 Organotypic slice cultures……………………………………………………...…153
5.2.1 Dissection of the ventral mesencephalon and striatum…………...…….153
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5.2.2 Mounting the slices……………………………………………………..155
5.2.3 Slice culture by the roller-tube technique………………………………156
5.3 Fixation of cultures…………………………………………………...…………...158
5.3.1 Immunoperoxidase antigen retrieval …………………………………...158
5.3.2 Immunohistochemistry………………………………………………...159
5.3.3 Co-localisation of TH and active Caspase-3………………………..…….160
5.4 Lactate Dehydrogenase (LDH) ELISA…………………………...……………...160
5.6 Quantification and statistical analyses of cell counts…………………….……..161
CHAPTER 6: ……………………………………………………………………….....170
DOPAMINERGIC NEURONS GROWN IN ORGANOTYPIC SLICE CULTURES,
TH-ir CELLS ARE DEPENDENT ON TARGET REGION-
STRIATUM……………………………………………………………………...……..170
6.0 INTRODUCTION……………………………………………………………...…171
6.1 AIMS………………………………………………………………………...……..171
6.2 METHODS………………………………………………………………………...171
6.3 RESULTS
6.3.1 TH-ir neurons of the ventral mesencephalon in culture…………………..172
6.3.2 TH-ir cell growth in the Co-cultures……………………………………...173
6.3.3 The influence of trophic ST target on VM TH-ir cells…………………...174
6.3.4 Glial cells contribute to the regulation of DAergic outgrowth…………...174
6.4 DISCUSSION……………………………………………………………………...186
6.4.1 General findings…………………………………………………………..186
6.4.2 Advantage and disadvantages of the culture system-organotypic slice culture
system…………………………………………………………………………..186
6.4.3 The influence of target region-Striatum on TH-ir neurons……………….188
6.4.4 The growth of TH-ir neurons with increasing culture duration…………..189
6.4.5 Possible influences on TH-ir cell development…………………………..189
6.4.6 Role of GFAP-ir cells in TH-ir neuronal development…………………..190
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CHAPTER 7: ……………………………………………………………………...…..193
THE EFFECTS OF NEUROTOXINS: MPTP & ROTENONE ON TH-ir CELLS
GROWTH IN ORGANOTYPIC SLICE CULTURES……………………………...…193
7.0 INTRODUCTION………………………………………………………………..194
7.1 AIMS………………………………………………………………………………194
7.2 METHODS………………………………………………………………………..194
7.2.1.1 MPTP Effects on 7 Day old VM & ST co-cultures…………………….195
7.2.1.2 Rotenone Effects on 7 Day old VM & ST co-cultures…………………196
7.2.2.1 MPTP Effects on 14 Day old VM & ST co-cultures………………..….197
7.2.2.2 Rotenone Effects on 14 Day old VM & ST co-cultures……………..…198
7.2.2.3 Effects of MPP+ on 14 Day old VM & ST co-cultures……….……….199
7.2.4 Cytotoxicity of Neurotoxins…………………………………………………….199
7.2.5 Neuronal Degeneration: Fluoro Jade C (FJ-C)……………………….……….200
7.2.6 Apoptotic Cell Death: Caspase-3……………………………………………….200
7.3 RESULTS………………………………………………………………………...201
7.3.1 Neurotoxin treatment of VM and ST co-cultures……………….………..202
7.3.2 Dose-dependent death of TH-ir cells-MPTP……………………………..202
7.3.3 MPTP toxicity varies with the time of co-culture exposure……………...214
7.3.4 Dose-dependent death of TH-ir cells-Rotenone………………………….215
7.3.5 Rotenone toxicity varies with the time of co-culture exposure…………..216
7.3.6 TH-IR CELL DEATH BY MPTP AND ROTENONE……………………….226
7.3.6.1 Cytotoxicity of MPTP and rotenone by LDH assay……………………226
7.3.6.2 Fluoro Jade C (FJ-C)……………………………………………………226
7.3.6.3 MPTP and rotenone induced Caspase-3 activation…………………….229
7.4 DISCUSSION……………………………………………………………………...235
7.4.1 Dose and time dependent toxicity of neurotoxins………………………...235
7.4.2 Toxicity of neurotoxins on Dopaminergic co-cultures varies with the age of
co-cultures……………………………………………………………………....238
7.4.3 Glial cells play an important role in DAergic degeneration……………...240
7.4.3 Rotenone vs MPTP neurotoxicity………………………………...……....243
7.4.4 MPTP and rotenone activate caspase-3……………………………..........244
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CHAPTER 8: ………………………………………………………………………….246
THE NEUROPROTECTIVE EFFECT OF GLIAL CELL-LINE DERIVED
NEUROTROPHIC FACTOR (GDNF) ON TH-ir CELLS IN CO-
CULTURES…………………………………………………………………………….246
8.0 INTRODUCTION………………………………………………………………...247
8.1 AIMS:……………………………………………………………………………...247
8.2 METHODS………………………………………………………………..............248
8.2.1 Dose-response of GDNF on TH-ir cells……………………..…………...248
8.2.2 TH-ir cells response to GDNF following neurotoxin treatment…….........250
8.2.3 TH-ir cell response to pre-treatment with GDNF followed by neurotoxin
insult……………………………………………………………………….……250
8.2 Analysis of cell death………………………………………………..……...254
8.3 RESULTS……………………………………………………………….................254
8.3.1 GDNF at varying doses did not induce a significant increase in TH-ir
cells……………………………………………………………………………..254
8.3.2 GDNF treatment promotes increase in cell size and branching……….…254
8.3.3 Post GDNF treatment following MPTP and rotenone exposure………....254
8.3.4 Pre-treatment with GDNF followed by MPTP and rotenone
exposure…………………………………………………………………….…..255
8.4 DISCUSSION………………………………………………………………...……270
8.4.1 The neurotrophic effects of GDNF on DAergic neurons survival and
development………………………………………………………………….…270
8.4.2 Neuroprotection and regeneration of DAergic neurons by GDNF…….....271
8.4.3 GDNF neuroprotection against neurotoxins in slice cultures…………….272
CHAPTER 9: ………………………………………………………………………….273
GENERAL DISCUSSION…………………………………………………………….273
9.1 INTRODUCTION………………………………………………………………...274
9.2 ORGANOTYPIC SLICE CULTURE OF VM & ST…………………………..274
9.2.1 The advantages of using organotypic slice cultures……………………...274
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9.2.2 Disadvantages of using organotypic slice cultures………………….……275
9.2.2 Summary of experimental findings…………………………………….…275
9.3 THE EFFECTS OF MPTP AND ROTENONE ON VM AND ST SLICE
CULTURES………………………………………………………………….………...278
9.3.1 MPTP and rotenone………………………………………………………278
9.3.2 Summary of experimental findings……………………………………….278
9.4 THE NEUROTROPHIC ROLE OF GDNF ON TH-ir NEURONS IN SLICE
CULTURES……………………………………………………………………….…...282
9.4.1 GDNF and DAergic neurons……………………………………………..282
9.4.2 Summary of experimental findings……………………………………….283
9.5 Summary…………………………………………………………………………...288
9.6 Conclusion ………………………………………………………………………...289
REFERENCES……………………………………………………………………...…290
APPENDICES………………………………………………………………………....400
xxv
LIST OF FIGURES AND TABLES
Figure 1.1: Diagrammatic representation of the age predilections for typical, sporadic Parkinson’s disease and genetically determined Parkinsonism.…………………………13 Figure 1.2: Action sites of pharmacological therapies currently available for PD………32 Figure 1.3: The current model of the basal ganglia………………………………..…47-48 Figure 1.4: Stages of MDN development……………………………………………..…57 Figure 1.5: Signaling cascades in the MDN development………………………...……..58 Figure 2.1: Schematic Representation of MPTP Metabolism………………………..….77 Figure 2.2: Schematic representation of MPP+ intracellular pathways………………….87 Figure 2.3: Pathways implicated in MPTP-mediated toxicity………………………..93-94 Figure. 2.4: Chemical structure of rotenone, paraquat, MPP+, and maneb…………….111 Figure 5.1: Schematic illustration of the preparation protocol of slice cultures from 4 to 5-day-old rats………………………………………………………………………...163-164 Figure 5.2: Schematic diagram of the culturing process………………………………..165 Figure 5.3: Schematic representation of the biotin-avidin peroxidase procedure for the identification of TH-ir neurons…………………………………………………….166-167 Figure 5.4: LDH ELISA…………………………………………………………...168-169 Figure 6.1: Dopaminergic (TH-ir) controls………………………………………...…..175 Figure 6.2: Dopaminergic (TH-ir) neuronal growth in VM & ST co-cultures…………176 Figure 6.3: Dopaminergic (TH-ir) neuronal growth in VM and ST co-cultures……….177 Figure 6.4: TH-ir cell growth in co-cultures vs single cultures (SVM)………………...178 Figure 6.5: Dopaminergic (TH-ir) neuronal growth in single VM cultures (SVM)……179 Figure 6.6: Dopaminergic (TH-ir) neuronal growth in co-cultures vs SVM…………...180 Figure 6.7: GFAP growths in co-cultures vs single cultures……………………….......181
xxvi
Figure 6.8: Glial fibrillary acidic protein (GFAP-ir) growth in VM & ST co-cultures………………………………………………………………………………….182 Figure 6.9: Glial fibrillary acidic protein (GFAP-ir) growth in single VM (SVM) cultures………………………………………………………………………………….183 Figure 6.10: Dopaminergic (TH-ir) & Glial fibrillary acidic protein (GFAP-ir) growth in VM & ST co-cultures. ………………………………………………………………….184 Figure 6.11: Glial fibrillary acidic protein (GFAP-ir) growth in VM & ST co-cultures and SVM…………………………………………………………………………………….185 Figure 7.1: Schematical diagram of the experiments………………………...…………201 Figure 7.2: MPTP treated co-cultures at 7 days……………………………………….203 Figure 7.3.1: Dopaminergic (TH-ir) neuronal growth following 1 week post MPTP treatment of 7 day co-cultures…………………………………………………………..204 Figure 7.3.2: Dopaminergic (TH-ir) neuronal growth following 2 week post MPTP treatment of 7 day co-cultures…………………………………………………………..205 . Figure 7.3.3: Dopaminergic (TH-ir) neuronal growth following 3 week post MPTP treatment of 7 day co-cultures…………………………………………………………..206 Figure 7.4: MPTP treated co-cultures at 14 days……………………………………...207 Figure 7.5.1: Dopaminergic (TH-ir) neuronal growth following 1 week post MPTP treatment of 14 day co-cultures………………………………………………………....208 Figure 7.5.2: Dopaminergic (TH-ir) neuronal growth following 2 week post MPTP treatment of 14 day co-cultures………………………………………………………....209 Figure 7.5.3: Dopaminergic (TH-ir) neuronal growth following 3 week post MPTP treatment of 14 day co-cultures………………………………………………...……….210 Figure 7.5.4: Dopaminergic (TH-ir) neuronal growth following 1 week post MPTP & MPP+ treatment of 14 day co-cultures………………………………………………….211 Figure 7.6: 7 day & 14 day old co-cultures MPTP treated………………….……212-213 Figure 7.7: Rotenone treated co-cultures at 7 days………………………………….…217 Figure 7.8.1: Dopaminergic (TH-ir) neuronal growth following 1 week post rotenone treatment of 7 day co-cultures…………………………………………………….…….218
E
*
xxvii
Figure 7.8.2: Dopaminergic (TH-ir) neuronal growth following 2 week post rotenone treatment of 7 day co-cultures…………………………………………………………..219 Figure 7.8.3: Dopaminergic (TH-ir) neuronal growth following 3 week post rotenone treatment of 7 day co-cultures………………………………………………………….220 Figure 7.9: Rotenone treated co-cultures at 14 days…………………………………..221 Figure 7.10.1: Dopaminergic (TH-ir) neuronal growth following 1 week post rotenone treatment of 14 day co-cultures…………………………………………………………222 Figure 7.10.2: Dopaminergic (TH-ir) neuronal growth following 2 week post rotenone treatment of 14 day co-cultures…………………………………………………………223 Figure 7.10.3: Dopaminergic (TH-ir) neuronal growth following 3 week post rotenone treatment of 14 day co-cultures…………………………………………………………224 Figure 7.11: 7 day & 14 day old co-cultures rotenone treated…………………….225-226 Figure 7.12: Lactate Dehydrogenase Cytotoxicity Assay (LDH) of MPTP & Rotenone treated co-cultures………………………………………………………………….228-229 Figure 7.13: MPTP & rotenone treated co-cultures 1 week post treatment- Degenerating neurons-FJC…………………………………………………………………………….230 Figure 7.14: Co-localisation of TH-ir and caspase-3 in co-cultures…………………...231 Figure 7.15: Co-localisation of TH-ir and caspase-3 in co-cultures following MPTP treatment at one week recovery period…………………………………………………232 Figure 7.16: Co-localisation of TH-ir and caspase-3 in co-cultures following rotenone treatment at one week recovery period…………………………………………………233 Figure 7.17: Co-localisation of TH-ir and caspase-3 in co-cultures following MPTP at one week recovery period…………………….………………………………………...234 Figure 7.18: Co-localisation of TH-ir and caspase-3 in co-cultures following rotenone treatment at one week recovery period………………………………………………....235 Figure 8.1: Schematical diagram of the experiments…………………………………..256 Figure 8.2: TH-ir cells in co-cultures treated with GDNF………………………...256-257 Figure 8.3: TH-ir analysis of 7 day co-cultures treated at varying doses of GDNF…………………………………………………………………………...………258
xxviii
Figure 8.4: 7 day co-cultures treated at varying doses of GDNF……………………259 Figure 8.5: The neuroprotective effects of GDNF on TH-ir cells following MPTP exposure…………………………………………………………………………….….260 Figure 8.6: The effects of GDNF post treatment following MPTP exposure…………261 Figure 8.7: The neuroprotective effects of GDNF on TH-ir cells following rotenone exposure………………………………………………………………………………...262 Figure 8.8: The effects of GDNF post treatment following rotenone exposure………………………………………………………………………………..263 Figure 8.9: LDH ELISA of GDNF post treatment following MPTP exposure……………………………………………………….……………………….264 Figure 8.10: LDH ELISA of GDNF post treatment following rotenone exposure………………………………………………………………………………...265 Figure 8.11: FJC-staining of GDNF Post-treatment on co-cultures following MPTP and
rotenone exposure………………………………………………………………………266
Figure 8.12: The neuroprotective effects of GDNF Pre-treatment on TH-ir cells following MPTP and rotenone exposure……………………………………………………….…267 Figure 8.13: The effects of GDNF Pre-treatment following MPTP exposure…………268 Figure 8.14: The effects of GDNF Pre-treatment following rotenone exposure………………………………………………………………………………...269 Table 1.1: Genes involved in Parkinsons Disease……………………...………………..11 Table 1.2: Summary of increased Oxidative stress (OS) in Idiopathic Parkinsons’s disease (IPD)……………………………………………………………………………………..15 Table 1.3: Summary of current PD treatment…………………………………………....33 Table 2.1: Characteristics of animal models of Parkinson’s disease………………….…75 Table 2.2: Relative toxicity of MPTP in different animals…………………………...….79 Table 2.3: MPTP-induced effects upon the DAergic system in different mouse strains…………………………………………………………………………………….84
xxix
Table 3 : Summary of Experimental PD In Vitro models…..…………………………..136
Table 7.1: 7 day old co-cultures treated with MPTP…………………………………...195 Table 7.2: 7 day old co-cultures treated with rotenone…………………………………196 Table 7.3: 14 day old co-cultures treated with MPTP……………………………….....197 Table 7.4: 14 day old co-cultures treated with rotenone………………………………..198 Table 7.5: 14 day old co-cultures treated with MPTP & MPP+………………………..199 Table 8.1: Dose-response of GDNF on TH-ir cells……………………………………249 Table 8.2: Post treatment of Co-cultures with GDNF following MPTP/rotenone exposure………………………………………………………………………………...251 Table 8.3: Pre-treatment of Co-cultures with GDNF prior to MPTP/rotenone exposure………………………………………………………………………………...252
xxx
ABSTRACT
In this study we aimed to investigate the effects of 1-methyl-4-phenyl-1,2,3,6-
tetrahydropyridine (MPTP) and rotenone neurotoxins on dopaminergic (DAergic)
neuronal survival using ventral mesencephalic (VM) organotypic cell culture derived
from postnatal rat pups (P4-5) and immunocytochemistry for tyrosine hydroxylase (TH)
as a marker of DAergic cells. In addition, we examined the neuroprotective effects of
glial cell line-derived neurotrophic factor (GDNF) on TH-ir cells exposed to MPTP and
rotenone as a possible treatment for PD.
The TH-ir cells in co-cultures with striatum (ST) as a target grew better then when VM
was cultured alone and that TH-ir cells in co-cultures could be maintained without using
conditioned and trophic media. We treated 7 day and 14 day co-cultures at different
times with varying MPTP and rotenone concentrations and found 14 day old cultures
were more vulnerable than 7 day old co-cultures to the effects of either neurotoxin with
TH-ir cell numbers significantly lower in 14 day cultures compared to 7 day cultures.
Both neurotoxins induced a dose-dependent TH-ir cell reduction in the co-cultures. In
addition we compared the toxicity of MPTP and its active metabolite 1-methyl-4-
phenylpyridinium (MPP+) as the neurotoxic effects of MPTP on DAergic cells depends
on its conversion to MPP+ by astrocytes. We found no significant difference in TH-ir cell
reduction in co-cultures treated with MPTP and MPP+. Rotenone was more toxic than
MPTP with less TH-ir cell survival in the weeks post treatment. GDNF exposure
produced increased cell size and significant increases in TH-ir cell branching in co-
cultures in a dose-dependent manner. Post treatment of GDNF against MPTP and
xxxi
rotenone provided significant neuroprotection as TH-ir cell survival was at the lower
neurotoxin doses and not at the higher doses.
1
CHAPTER 1:
OVERVIEW OF PARKINSON’S DISEASE
2
1.0 INTRODUCTION
Parkinson's disease (PD) is a progressive neurodegenerative movement disorder that is
estimated to affect approximately 1% of the population older than 65 years of age (Lang
and Lozano 1998a, b). Clinically the cardinal symptoms was first described by James
Parkinson (1817) (Parkinson 2002) with most patients presenting bradykinesia, resting
tremor, rigidity, and postural instability. A number of patients also suffer from
autonomic, cognitive, and psychiatric disturbances (Aarsland et al. 2007). The major
symptoms of PD result from the profound and selective loss of dopaminergic (DAergic)
neurons in the substantia nigra pars compacta (SNc), but there is widespread
neuropathology with the SNc becoming involved later toward the middle stages of the
disease (Braak et al. 2003). The pathological hallmarks of PD are round eosinophilic
intracytoplasmic proteinaceous inclusions termed Lewy bodies (LBs) and dystrophic
neurites (Lewy neurites) present in surviving neurons (Forno 1996). PD is primarily a
sporadic disorder and its specific etiology is incompletely understood, but with the recent
insights provided through studying the genetics, epidemiology, and neuropathology of
PD, and the development of new experimental models have improved our understanding.
1.1 INCIDENCE AND PREVALENCE OF PD
PD is a common neurodegenerative disorder, characterized by neuronal cell loss in the
substantia nigra (SN) and subsequently reduced secretion of dopamine (DA). PD is the
second most common neurodegenerative disease after Alzheimer’s disease, affecting up
to 1% of the elderly population over the age of 65. Epidemiological studies have
estimated a cumulative prevalence of PD of greater than one per thousand with the
3
prevalence being limited to senior populations; this proportion increases nearly 10-fold.
A family history of risk factors is a key factor for PD development. Indeed, the estimated
genetic risk ratio for PD is approximately 1.7 (70% increased risk for PD if a sibling has
PD) for all ages, and increases over 7-fold for those under 66 years of age (Langston
1998; Rocca et al. 2005; Levy 2007). The age-adjusted prevalence rate of PD revealed
from a pilot study of at least a 42.5% increase in the disease compared to 1966 (Chan et
al. 2001). The role for genes contributing to the risk of PD is also significant. Recently,
monogenically PD as familial PD (FPD) has been identified and seven causative genes
for FPD have been identified so far. Thus, the role of genetic factors and also increasing
age play an important risk factor for the development of PD.
1.2 CLINICAL CHARACTERISTICS OF PD
Resting tremor at a frequency of 4-6 Hz occurs at rest but decreases with voluntary
movement in about 70% of Parkinson’s patients. Rigidity refers to increased resistance
(stiffness) initially occur only to trunk region but also extends to passive movement of
limbs. Bradykinesia (slowness of movement), hypokinesia (reduction in amplitude, and
akinesia (absence of normal initiation of movement) manifest as a variety symptoms,
including paucity of normal facial expression, drooling, microphagia, and decreased
stride length during walking. PD patients also develop a stooped posture and possibly
lose postural reflexes. Freezing of gait, the inability to begin a voluntary movement such
as walking is a common symptom of Parkinsonism. Impairment in different cognitive
domains such as executive functions, language, memory, and visuospatial skills occurs
frequently in PD even in the early stages of the disease (Caballol et al. 2007).
4
1.3 NEUROCHEMICAL AND NEUROPATHOLOGICAL FEATURES OF PD
1.3.1 Cell loss in PD
Idiopathic Parkinsonism (IPD) is characterized by progressive and profound loss of
neuromelanin containing DAergic neurons in the substantia nigra pars compacta (SNpc)
(Forno 1996). Moreover, slight gliosis and neuronal loss in the locus coeruleus, dorsal
vagal nucleus with variable involvement of the nucleus basalis Meynert, and other
subcortical nuclei have been reported (Chung et al. 2003). Degeneration of pigmented
neuronal systems located in the brain stem, particularly in SNpc, is the most striking
pathological feature of PD. This causes striatal DA deficiency and all the major motor
PD symptoms.
The pattern of SNpc cell loss correlates with the level of expression of DA transporter
(DAT) mRNA (Uhl et al. 1994) and is consistent with the level of DA depletion being
most pronounced in the dorsolateral putamen (Jones 1999), the main site of projection for
these neurons. At onset of symptoms, putamental DA is depleted ~80%, and ~60% of
SNpc DAergic neurons have already been lost. The mesolimbic DAergic neurons, cell
bodies of which reside adjacent to the SNpc in the ventral tegmental area (VTA), are
much less affected in PD (Horn 1979) and significantly less depletion of DA in the
amygdala (Marsden 1987), the main site of projection for these neurons.
Neuropathological studies of PD-related neurodegeneration reveal DAergic neuronal loss
to have a characteristic topology, which is distinct from the pattern seen in the aging
process. In PD, cell loss is concentrated in ventrolateral and caudal portions of SNpc,
5
whereas during normal aging the dorsomedial aspect of SNpc is affected (Marsden 1987).
The degree of terminal loss in the striatum (ST) appears to be more pronounced than that
of SNpc DAergic neuronal loss, suggesting that ST DAergic nerve terminals are the
primary target of the degenerative process. The neuropathology of PD is characterized
solely by DAergic neuron loss which correlates with the progressive motor decline.
However the neurodegeneration extends well beyond DAergic neurons (Di Monte 2003),
with many “non-DAergic” PD features. Neurodegeneration and Lewy bodies (LB)
formation are found in noradrenergic (locus ceoruleus), serotonergic (raphe), and
cholinergic (nucleus basalis of Meynert, dorsal motor nucleus of vagus) systems, as well
as in the cerebral cortex (especially cingulated and entorhinal cortices), olfactory bulb,
and autonomic nervous system. Degeneration of hippocampal structures and cholinergic
cortical inputs contribute to high rate of dementia that accompanies PD, particularly in
older patients. The clinical correlation to lesions in serotonergic and noradrenergic
pathways is not clearly characterized as DAergic systems.
In about 95% of PD cases, there is no apparent genetic linkage (referred to as “sporadic”
PD), but in the remaining cases, the disease is inherited. The symptoms worsen and prior
to the introduction of levodopa, the mortality rate among PD patients were three times
that of normal age-matched subjects. Although levodopa has dramatically improved the
quality of life for PD patients, surveys revealed that these patients continue to display
decreased longevity compared to the general population (Katzenschlager and Lees 2002;
Marinus et al. 2003; Katsarou et al. 2004; Muller and Russ 2006). Furthermore, most PD
6
patients suffer considerable motor disability after 5-10 years of disease, even after
beneficial medications.
1.3.2 Lewy Bodies in PD
Apart from the loss of melanized (Mason 1984) nigrostriatal DAergic (DAergic) neurons
another major pathological hallmark of PD includes the presence of intraneuronal
proteinacious cytoplasmic inclusions “Lewy Bodies” (LBs). This results in the classical
neuropathological finding of SNpc depigmentation where the SNpc DAergic cell bodies
project primarily to the putamen. However LBs are not specific for PD as they are also
found in Alzheimer’s disease (AD), other α-synucleinopathies such as dementia with LB
disease (DLBD), and as “an incidental” pathologic finding in people of advanced age
than the prevalence of PD (Spillantini and Goedert 2000). The role of LB in neuronal
cell death is controversial, as the reasons for their increased frequency in AD and the
relationship of incidental LB to the occurrence of PD. LBs are spherical eosinophillic
protein aggregates composed of numerous proteins, including intracytoplasmic α-
synuclein, parkin, ubiquitin, and neurofilaments, which are found in all affected brain
regions (Chung, 2003; Fahn, 2004; Marsden, 1987; De Girolami, 1999). LBs are more
than 15μm in diameter with an organized structure containing a dense hyaline core
surrounded by a clear halo. Electron microscopy reveals a dense granulovesicular core
surrounded by radiating 8-10 nm fibrils (Marsden 1987).
1.4 ETIOLOGY OF PD
1.4.1 Environmental Factors
7
The specific etiology of PD is not known. It has been proposed to be multifactorial
because of its sporadic nature. Epidemiological studies show that a number of factors
may increase the risk of developing PD (Abbott et al. 2003; Baldereschi et al. 2003).
These include the exposure to well water, pesticides, herbicide, industrial chemicals,
farming and living in rural environment (Di Monte et al. 2002; Di Monte 2003). A
number of exogenous toxins have been associated with the development of parkinsonism,
including trace metals, cyanide, carbon monoxide, and carbon disulfide. In addition, the
possible role of endogenous toxins such as tetrahydro-isoquinolines and beta-carbolines
has also been implicated (McNaught et al. 1996; Storch et al. 2000; Maruyama and Naoi
2002). However, no specific toxin has been found in the human brain of PD patients, and
also in many instances of Parkinsonism associated with toxins is not that of typical LB
PD.
A study of monozygotic and dizygotic pairs of twins was conducted to assess the genetic
versus environmental factors in the etiology of PD (Tanner et al. 1999). Results show that
the contribution of environmental factors to both early and late onset forms can never be
completely excluded; however it is more evident in late onset forms (onset beyond age
50). Human exposure to chemical compounds of synthetic origin, including pesticides,
herbicides and insecticides has been the focus of several epidemiologic studies since the
first description of Parkinson-like symptoms among individuals who had taken drugs
contaminated with 1-methyl-4-phenyl-2,3,6-tetrahydropyridine (MPTP; (Langston et al.
1999)). MPTP is converted to 1-methylphenylpyridinium (MPP+) by MAO-B in glial
cells (Nicklas et al. 1985; Schmidt and Ferger 2001), and subsequently selectively
8
concentrated in mitochondria of DAergic neurons where it interacts with elements of the
mitochondrial respiratory chain (Langston et al. 1999), leading to ATP depletion, and
eventually cell death of the DAergic neurons (Fisher A. 1986; Schmidt and Ferger 2001;
Chung et al. 2003; Di Monte 2003). Paraquat is an herbicide structurally similar to
MPTP; human exposure to paraquat has also been associated with an increased risk of PD
(McCormack et al. 2002), and studies on animal models demonstrate that paraquat
induces a selective loss of DAergic neurons (Liu et al. 2003). Conflicting results have
been produced, in recent years, in association studies between exposure to pesticides,
fungicides and herbicides and PD risk (Gorell et al. 1998; McCann et al. 1998; Baldi et
al. 2003).
Human activities related to a possible pesticide exposure, including farming, living in
rural areas and drinking water, are among risk factors associated with PD, whereas
smoking and drinking coffee are protective factors (Hernan et al. 2002). A chronic
occupational exposure to manganese is the cause of manganism, a condition
characterized by tremor, rigidity and psychosis due to the accumulation of the metal in
the basal ganglia (Mergler and Baldwin 1997). Although no changes in manganese brain
concentration has been observed in PD, exposure to manganese has been linked to the
risk of PD in some epidemiological studies (Gorell et al. 1997; Gorell et al. 1998).
Copper exposure has been associated with PD, whereas iron exposure alone was not;
however exposure to combinations of iron and lead, iron and manganese and iron and
copper was associated with PD. Controversial or negative results have been obtained for
exposure to zinc, mercury and aluminium (Gorell et al. 1997; Gorell et al. 1998; Powers
9
et al. 2003). Inflammation of the brain in early life as a consequence of head injuries,
viral or bacterial infections or exposure to neurotoxicants, has been indicated as a
possible contributor to the development of PD later in life (Liu et al. 2003); moreover it
has been suggested that occupational exposure to viral (or other) respiratory infections
might be one of the risk factors for the observed increased incidence of PD cases among
teachers and healthcare workers (Tsui et al. 1999).
1.4.2 Genetic Factors
In a great majority of individuals with PD it is thought to occur sporadically, as the
family history does not indicate affected individuals in their first degree relatives.
However, a number of genetic forms of the disease have been recently discovered, and
research into these hereditary forms has helped to understand the pathophysiology of this
condition. To date, 10 loci (PARK1-10) have been mapped and found to be linked to
familial forms of PD (Riess et al. 2000; Riess et al. 2002; Steece-Collier et al. 2002;
Dawson and Dawson 2003; Fahn and Sulzer 2004; Pankratz and Foroud 2004) with
genes identified in five (see Table 1.1). They are presented either as autosomal
dominants or autosomal recessives.
The first gene discovered and mostly studied is the PARK-1 gene which encodes the
alpha synuclein (α-syn) protein (Polymeropoulos et al. 1996; Polymeropoulos 1998) and
is present in PD and dementia with LBs (DLB) but not in normal human brains. It is
normally densely accumulated within presynaptic axon terminals and beta (β)-positive
vesicles but gamma (γ)-negative are also present in the hippocampal dentate, hilar, and
10
CA2/3 regions (Galvin et al. 1999). Genetic variability in the α-syn gene has been shown
to be a risk factor for the development of PD (Conway et al. 2000; Hsu et al. 2000; Saha
et al. 2000; Lo Bianco et al. 2002; Lotharius and Brundin 2002). Immunohistochemical
studies have indicated that LBs of the brainstem and also cortical types in the brains of
PD patients with sporadic PD or DLB are strongly positive for α-syn (Hishikawa et al.
2001; Junn et al. 2003; Liani et al. 2004; Murakami et al. 2004) thus may be an important
component of LBs. α-syn knock out mice revealed a striking resistance to MPTP
induced degeneration of DA neurons and DA release. This resistance appeared to result
from the inability of the toxin to inhibit complex 1 (Dauer et al. 2002). The accumulation
of α-syn intracellulary at abnormally high quantities, in an aggregated form within the
neurons and sometimes glial cells, have been shown to contribute to the pathology of PD
and also to generate hydroxyl radicals (OH•) upon the addition of iron (Fe ii). The past
10 years has seen a shift in our etiological concepts of Parkinson's disease, moving from
a nearly exclusively environmentally mediated disease towards a complex disorder with
important genetic contributors. The identification of responsible mutations in certain
genes, particularly alpha-synuclein, Parkin, PINK1, DJ-1 and LRRK2, has increased our
understanding of the clinical and pathological changes underlying Parkinson's disease,
with implications for patient diagnosis, management and future research.
11
Table 1.1 Genes involved in Parkinsons Disease
Gene Inheritance Locus Clinical Features Histopathology α-synuclein AD PARK1/4 (4q21) Ala53Thr: early onset, rapid progression
Ala30Pro: typical parkinsonian phenotype Glu46Lys: cognitive decline, hallucinations Duplication: typical parkinsonian phenotype Triplication: early onset, rapid progression, dementia, early death
SN degeneration, Lewy body pathology in substantia nigra, cortex and hippocampus
Parkin AR PARK2 (6q25-27) Early disease onset, resembles idiopathic PD phenotype, slow disease progression, good response to levodopa, early dyskinesias, diurnal .uctuation and sleep relief
SN degeneration in the absence of Lewy bodies. Lewy body pathology reported in compound heterozygous carriers
Unknown AD PARK3 (2p13) Typical parkinsonian phenotype
Nigral Degeneration with Lewy bodies
UCH-L1 AD PARK5 (4p14) Typical parkinsonian phenotype N/A PINK1 AR PARK6 (1p35-36) Early disease onset, levodopa responsive,
slow progression, dyskinesias N/A
DJ-1 AR PARK7 (1p36) Early disease onset, levodopa responsive. Dystonia and psychiatric features reported
N/A
LRRK2 AD PARK8 (12q12) Predominantly levodopa responsive parkinsonism for all mutations. Supranuclear gaze palsy, dystonia, Supranuclear gaze palsy, dystonia, dementia and motor neuron disease is described in some individuals
All had SN degeneration. Variable additional pathology: novel ubiquitin positive inclusions and MND (Tyr1699Cys), tau pathology (Arg1441Cys), Lewy bodies (Gly2019Ser, Arg1441Cys), no additional pathology (Arg1441Cys)
Unknown N/A PARK10 (1p32) Typical parkinsonian phenotype N/A Unknown N/A PARK11 (2q36-37) Typical parkinsonian phenotype N/A
AD, autosomal dominant; AR, autosomal recessive; SN, substantia nigra; N/A, not available; MND, motor neuron disease
(amyotrophic lateral sclerosis) (adapted from Gosal et al., 2006).
11
12
1.4.3 Ageing and PD development
Age is an important factor in PD expression (Figure 1.1) (Fahn and Sulzer 2004). After
the age of 30-40, the incidence increases with the age reaching a maximum at the 8th
decade with a decline in prevalence thereafter (Marsden 1987). The progression of the
aged in the total population is also expected to rise from 12.2% in year 2000 to 15.5% in
the year 2020. Aging in individuals is affected to a great extent by genetic factors, diet,
social conditions, and the occurrence of age-related diseases. There is evidence that age-
induced alterations in cells are an important component of aging of the organism (Cotran
R.S 1999). A number of cell functions decline progressively with age, for example
oxidative phosphorylation in the mitochondria, the synthesis of nucleic acids and
structural enzymatic proteins are reduced. Oxidative damage to mitochondrial DNA
increases with ageing and can induce point mutations thus, contributing to mitochondrial
dysfunction and neuronal loss (Simon et al. 2004) [Simon, 2003]. Biochemical analysis
revealed a shift in the extractability of parkin upon aging in humans but not in mice thus
there is an effect of aging upon parkin in humans (Pawlyk et al. 2003).
1.5 MECHANISMS OF PARKINSONIAN CELL DEATH
1.5.1 Oxidative stress (OS)
OS and the consequent cell death in SNpc usually results from either an increased DA
turnover, resulting in excess peroxide formation, a deficiency in glutathione (GSH), thus
diminishing the brains capacity to clear H2O2 or an increase in reactive iron thereby,
promoting OH• formation (Olanow and Tatton 1999; Foley and Riederer 2000; Bharath et
al. 2002; Koutsilieri et al. 2002; Ischiropoulos and Beckman 2003; Jenner 2003; Klein
13
and Ackerman 2003). Most of the evidence suggesting OS is linked to the pathogenesis
of PD comes from postmortem studies (Jenner 1993; Yoritaka et al. 1996; Alam et al.
1997; Jenner 1998; Serra et al. 2001; Giasson et al. 2002; Hald and Lotharius 2005; Sofic
et al. 2006) with increased iron levels are increased in the SNpc in PD, (Gu et al. 1998;
Faucheux et al. 2003) and the reduced levels of GSH where excessive oxidative DAergic
cell burden occurs which disrupts mitochondrial complex I function (Shults et al. 1997;
Gu et al. 1998). Although ROS levels cannot be measured directly in living patients,
their reaction by products and the damage in postmortem tissue is used as an indirect
indicator (see Table 1.2).
Figure 1.1 Diagrammatic representation of the age predilections for typical,
sporadic Parkinson’s disease and genetically determined Parkinsonism.
Illustrating the concept that genetically determined Parkinsonism tends to start at a
younger age, but constitutes only a small proportion of all patients with typical,
progressive Parkinsonism (Langston, 2002).
14
1.5.2 Iron
Iron is richest in the basal ganglia with the highest levels being in the SNc, pallidus and
putamen (Dexter et al. 1989b; Foley and Riederer 2000). In IPD, the shift of iron Fe2+/
Fe3+ ratio is almost 2:1 to 1:2 (Riederer et al. 1989; Sofic et al. 1991; Gerlach et al. 1994;
Foley and Riederer 2000), this is consistent with the increased Fe2+-catalysed conversion
of H2O2 to the highly reactive OH• known as the Fenton reaction: H2O2 + Fe 2+ → OH•
+ OH• + Fe3+ (Foley and Riederer 2000). The reduction of Fe3+ by the superoxide
radical increases the reaction rate which drives the production of OH•. Iron levels are
increased in IPD by about 35% specifically in the SNc (Gu et al. 1998; Bartzokis et al.
1999; Graham et al. 2000; Berg et al. 2001); similar increases have also been noted in
other neurodegenerative diseases (Qian et al. 1997; Ke and Ming Qian 2003). Iron
participates in the free-radical generating reaction only in the free ferrous form (Rouault
and Cooperman 2006). Ferric iron (Fe 2+) in the SNc is normally bound either by ferritin
(about 90%) or neuromelanin (NM) and, is associated with both LB and NM in IPD
(Hirsch et al. 1991; Jellinger et al. 1992; Mann et al. 1994; Foley and Riederer 2000).
Infusion of iron into rat brains produces a model of PD characterized by a concentration-
dependent loss of striatal DA, and the degeneration of SNc neurons with the behavioral
changes (Fisher A. 1986; Berg et al. 1999; Santiago et al. 2000; Ponting 2001). How iron
accumulates within the SNc in PD is not clear but an increased lactoferrin receptors have
been detected on nigral neurons in PD patients which readily cross the BBB and are
synthesized in the brain. This may explain the iron accumulation in SN and subsequent
degeneration of DAergic neurons in PD (Berg et al. 2002).
15
Table 1.2 Summary of increased Oxidative stress (OS) in Idiopathic
Parkinsons’s disease (IPD)
Evidence for an increase of OS in IPD
brain
References
> membrane peroxidation in SNc
> ROS-induced protein modification
> protein carbonyls in all IPD brain
Ala,, 1997; Floor, 1998; Buhmann,
2004
> thiobarbituric acid-reactive substance
levels (measures the secondary
products of lipid peroxidation)
< polyunsaturated fatty acid levels
(peroxidant substrate)
Dexter, 1989
> 8-hydroxy-2’ deoxyguanosine levels
in the SNc + other brain regions
(indicates ROS-mediated DNA
damage)
Zhang, 1999
Several studies have suggested that transcranial ultrasound (TCS) may able to
demonstrate the presence of increased iron content in the SN of PD patients, even prior to
the development of the disease symptoms (Berg et al. 2002). The echogenicity in PD
patients substantially increases the echogenicity of the SNc (Berg et al. 2001). The
elevated tissue echogenicity has been examined in postmortem studies, revealing a close
correlation between echogenicity of the SN and tissue iron content. Iron accumulation in
16
the SN in rats confirmed that iron that is not protein-bound may lead to an increase of
tissue echogenicity (Berg et al. 1999). Exposures to welding fumes have been shown to
alter trace metal levels such as, manganese, iron, zinc and copper in the body. Serum
levels of iron in the welders were 1.9 fold higher than those of controls and the level of
erythrocytic superoxide dismutase activity was 2.4% less thus, occupational exposure to
these toxic fumes disturbs the homeostasis of trace elements in the systemic circulation
which induces OS (Li et al. 2004).
1.5.3 Neuromelanin (NM)
It’s been hypothesized that NM may act as an endogenous storage molecule for iron and
possibly influence the production of free radicals. Iron-binding studies have
demonstrated that both NM and synthetically-produced DA-melanin contained equivalent
numbers of high and low-affinity binding sites for iron but, that the affinity of NM for
iron was higher than that of synthetic melanin (Double et al. 2003). The iron-binding
capacity of NM is 10-fold greater than that of the synthetic melanin. This is consistent
with the hypothesis. The function or effect of NM on neuronal function is unclear; it is
generally regarded as a waste product of DA auto-oxidation, and accumulates with age in
nigral neurons (Zecca et al. 2004) but is dramatically decreased in PD patients (Zecca et
al. 2002). NM has been described as a ‘double-edged sword’ with respect to its
sequestering of redox-active metal ions such as iron [Zecca, 2002; Zecca, 2003] induced
by free radical reactions (Enochs et al. 1994), and inhibiting lipid peroxidation as an
oxidant (Korytowski et al. 1995). NM may also promote ROS-generating and act as a
depot for cellular toxins like MPP+ (D'Amato et al. 1987b, c; D'Amato et al. 1987a; Zecca
17
et al. 2003). In IPD, cell death occurs mainly in the DA-containing SN. It is suggested
that there is a selective vulnerability of neuromelanin-pigmented subpopulation of DA-
containing mesencephalic neurons in PD [(Hirsch et al. 1998). High levels of Cu/Zn
superoxide dismutase mRNA have been observed in the SN indicating high levels of
oxygen free radicals are being produced in PD (Hirsch 1992). Catecholaminergic
neurons surrounded by a low density of glutathione peroxidase cells were more
susceptible to degeneration in PD than those well protected against OS, indicating the
selective vulnerability of catecholaminergic neurons (Hirsch 1992).
1.5.4 Glutathione (GSH)
A defect in one or more of the naturally occurring antioxidants defences could lead to the
neurodegeneration in PD (Jenner 1996) although no basic defects have been detected in
levels of ascorbic acid, α-tocopherol, catalase, or glutathione peroxidase (Riederer et al.
1989; Dexter et al. 1994; Sian et al. 1994b; Sian et al. 1994a). The cellular radical
detoxification system (the most important being superoxide dismutase (SOD) and
glutathione peroxidase (GPX) activity unfortunately declines with the ageing process.
The activity of SOD is increased in discrete regions of the brain according to different
neurodegenerative diseases, in IPD it is characteristically increased in SNc (Marttila et al.
1988; Saggu et al. 1989). Of the two isozymes, mitochondral Mn-SOD and cytosolic
Cu/Zn-SOD, only levels of Mn-SOD are elevated in IPD (Yoshida et al. 1993; Yoshida et
al. 1994), suggesting increased OS levels in the mitochondria although there have been
report of high Cu/Zn-SOD levels occurring as well (Serra et al. 2001). This increase may
be a neuroprotective response to the elevated levels of OS however; an increase in H2O2
18
production might be a problem in the absence of corresponding catalase and GPX
activity. In genetically modified rodents, high SOD expression protected the rat nigral
cells against MPTP and 6-OHDA (Asanuma and Cadet 1998; Asanuma et al. 1998).
Similar results have been observed in the postnatal midbrain cultures from mice with
increased amounts of WT SOD enzyme, promoting cell survival and providing protection
against L-dopa-induced DAergic neurotoxicity whereas, increased amounts of mutated
Cu/Zn-SOD enzyme had inverse effects (Mena et al. 1997).
Most attention has been directed to the finding of a selective decrease in the reduced form
of GSH in the SNc in PD (Sofic et al. 1992; Sian et al. 1994b; Sian et al. 1994a).
Reduced levels of GSH have not been detected in other brain areas in PD and have not
been reported in any other neurodegenerative disorder. A reduction in GSH impairs
H2O2 clearance thus, promoting OH• formation, particularly in the presence of high levels
of iron. The cause of GSH reduction in PD is still unknown and no defects have been
detected in the enzymes associated with GSH synthesis. However, a significant increase
in γ-glutamyltranspeptidase (γ-GTT), the enzyme responsible for the translocation of
GSH precursor and the metabolism of the oxidised form of glutathione (GSSG) (Sian et
al. 1994b; Sian et al. 1994a; Jenner 1996; Foley and Riederer 2000), could reflect the
attempt of surviving cells to recruit and replenish diminished levels of GSH or act as a
compensatory mechanism to remove the toxic GSSG. In the normal human brain, nigral
GSH levels are low in comparison to with other brain regions (Perry et al. 1982; Riederer
et al. 1989; Sofic et al. 1992; Pearce et al. 1997; Bharath et al. 2002). However, there is a
further decline in IPD nigra of the total and reduced GSH levels (Dexter et al. 1994). The
19
GSH levels are similar in incidental LBs but are not accompanied by changes in iron
levels or mitochondrial function thus, a decline in GSH levels could possibly occur early
in the course of the disorder. An artificially induced 40-60% loss of GSH by the
administration of BSO (buthionine sulphoximine, a selective inhibitor of α-
glutamylcysteine synthetase) led to a similar loss as in IPD however, no nigral striatal cell
loss or changes in the mitochondrial SOD activity in rats were observed but the
sensitivity to both 6-OHDA and MPP+ were enhanced (Pileblad et al. 1989; Wullner et al.
1996; Toffa et al. 1997). This suggests a reduction in GSH itself may not damage
DAergic neurons but possibly rendering these vulnerable to other toxins.
1.5.5 Mitochondrial Dysfunction
There is a significant body of evidence which supports the hypothesis that the progressive
reduction in mitochondrial respiration (a feature of the ageing brain) is involved in a
number of neurodegenerative diseases including IPD (Schapira et al. 1989; Schapira et al.
1990; Di Monte 1991). A selective 38-49% decrease in complex 1 activity of the
mitochondrial respiratory chain has been found in the SNc of PD patients (Schapira et al.
1990; Reichmann et al. 1993a; Reichmann et al. 1993b). A complex 1 defect has also
been found in platelet and muscle tissues of PD patients (Schapira et al. 1990; DiMauro
1993; Reichmann et al. 1993a; Reichmann et al. 1993b) however, there have been no
reports on detectable structural or mitochondrial DNA changes. The other transport chain
complexes generally appear to be unaffected despite the report of a reduction in SNc
complex IV activity (Itoh et al. 1997). The cause of the reduced complex I activity in
IPD remains unknown. No MPTP-like toxin has been detected or any specific
20
abnormality in the subunits of complex I, DNA, nor nuclear gene which encodes complex
I protein.
A mitochondrial complex I defect could contribute to cell degeneration in PD through
decreased ATP synthesis and a bioenergetic defect. Complex I inhibition by MPTP or
MPP+ have been shown to lead to a depletion of cellular ATP in mouse brain
synaptosomes (Scotcher et al. 1990; Swerdlow et al. 1996). Studies in experimental
animals indicate that a decrease in complex I activity of 40% or less does not compromise
cellular ATP levels (Davey and Clark 1996). Complex I defect could also lead to cell
damage through the generation of free radicals or the compensatory mechanism of
increasing the respiration through complex II (Schulz et al. 1995a).
Factors that contribute to the progression of PD include brain defects in mitochondrial
respiration and the generation of H2O2 by MAO. One study revealed that these two
factors were linked (Cohen et al. 1997). Addition of glutathione disulfide (GSSG;
oxidized form of glutathione) to the mitochondria resulted in a similar reversible
inhibition of the electron transport (Cohen et al. 1997). A complex I defect may also
contribute to the development of apoptosis (Ozawa et al. 1997; Tatton et al. 1998), as
complex I defect is the major site of proton pumping; it is possible that this could lead to
cell death via neuronal vulnerability and contributing to the pathogenesis of PD (Liu and
Kane 1996; Susin et al. 1996).
1.5.6 Excitotoxicity
21
In the past decade, there has been wide acceptance of the theory of excitotoxicity as a
viable process underlying a variety of acute and chronic degeneration disorders (Plaitakis
and Shashidharan 2000). Activation of more than one ionotrophic glutamate receptor
subtype can produce neuronal death. The N-methly-D-aspartate (NMDA) receptor
mediated toxicity is so far the most characterised pathway (Dickie et al. 1996; Plaitakis
and Shashidharan 2000). It has been shown that excitatory amino acids given
systemically to immature animals can cause neuronal degeneration in areas of the brain
that lacks an intact BBB (Plaitakis and Shashidharan 2000). Unlike the classic
neuromodulators –DA, norepinephrine and acetylcholine, which show marked regional
distribution in the brain, amino acids is present at high concentration in all cells (Plaitakis
and Shashidharan 2000).
Postsynaptic transduction of excitatory transmission is mediated by several classes of
glutamate receptors. These include the NMDA, the AMPA (amino-3-[3-hydroxy-5-
methylisooxazol-4-1]), proprionic (acid)/kainite, and the metabotrophic receptor. SNpc
DAergic neurons are rich in glutamate receptor, receiving extensive glutamate
innervation from the cortex and subthalamic nuclei (STN) with a pattern of burst firing in
response to exogenous administration of glutamate (Johnson et al. 1992; Rothstein et al.
1994). DA lesions disinhibit the STN inducing an increase in the firing rate of its
excitatory output neuron (DeLong 1990). Glutamate has a dual function in synaptic
transmission being an excitatory neurotransmitter and mediating an inhibitory
postsynaptic potential in DAergic neurons (Fiorillo and Williams 1998). A reduction in
energy metabolism due to mitochondrial dysfunction is shown to result in a loss of ATP-
22
dependent Mg-blockade of NMDA receptors, allowing glutamate to mediate an influx of
Ca2+ into the cell (Greenamyre et al. 1999). The role of excitotoxicity in PD are
supported by reports of NMDA antagonists protecting DAergic cell loss against MPP+
infusion into the SNc of rats (Turski et al. 1991), and MPTP treatment in primates
(Greenamyre et al. 1994). NMDA receptor mediated neuroprotection has also been
shown to be dependent on nuclear factor-kβ (NF- kβ) and the release of brain derived
neurotrophic factor (BDNF) (Lipsky et al. 2001).
1.5.7 Nitric Oxide (NO)
Excitotoxic damage is thought to be mediated at least in part via NO. Increased NO
released from glial, microglia or macrophages could also be responsible for the elevation
of local OS (Dawson et al. 1992). Although NO is an effective free radical scavenger
(Rubanyi et al. 1991) it can also react with the superoxide radical to form the
peroxynitrite anion (••••ONOO-). The oxidative radical .ONOO- decomposes to form OH• +
NO2, which are potent initiators of lipid peroxidation (Beckman et al. 1990; Radi et al.
1991). NO also directly inhibits mitochondrial respiration at the level of complex IV and
I, by liberating ferritin-bound iron thus, promoting lipid peroxidation (Bolanos et al.
1996) with mitochondrial respiratory chain being damaged by sustained exposure to NO
and that GSH is an important defense. This has implications for PD where GSH levels are
decreased.
NO-mediated toxicity has been implicated in nigral damage induced by MPTP. The
neuronal NOS inhibitor 7-nitroindazole (7-NI), which blocks NO formation, protects
23
DAergic neurons from MPTP toxicity in both rats and baboons (Schulz et al. 1995b;
Hantraye et al. 1996). Similarly, MPTP toxicity is diminished in NOS knock-out mice
(Przedborski et al. 1996). A recent report noted that 7-NI inhibits MAO-B (Castagnoli et
al. 1997), raising the possibility that it may act by blocking the conversion of MPTP to
MPPC. Inducible NO synthetase activity (NI) has been shown to be greater in SNc of
IPD patients, possibly secondary to reactive gliosis (Hunot et al. 1996). Damage due to
NO can be estimated by mediating the formation of 3-Nitrotyrosine (3-NT), product of
the peroxynitrite induced nitration of tyrosine resides on cellular probes [Ischiropoulos et
al 1992]. Elevated levels of 3-NT have been reported in MPTP treated mice and
monkeys (Schulz et al. 1995b) 3-NT levels in LB core have also been demonstrated to be
increased in IPD, indicative of NO-induced damage (Jenner 1996; Good et al. 1998).
1.5.8 Inflammation
Inflammation plays a role in the pathogenesis of PD. Autopsy reports have shown a glial
response found in both ST and nigra of PD patients and MPTP-mice which may likely
exert deleterious effects on the remaining DAergic neurons (Przedborski et al. 2000;
Przedborski et al. 2001; McGeer et al. 2003; Teismann and Schulz 2004; McGeer and
McGeer 2008). This view has led many investigators to aggressively examine the
potential role of neuroinflammation in the pathogenesis of PD. The SN DAergic neurons
are vulnerable to a variety of insults due to their reduced antioxidant capacity with a high
number of microglia cells, compared to other areas of the brain (Liu et al. 2003). When
microglia becomes activated they release proinflammatory factors and such cytokines as
interleukin-1 (IL-1), IL-2, IL-6 and tumor necrosis factor alpha (TNFα ), which can be
24
cytotoxic (Blum et al. 2001). Autopsy reports from sporadic PD patients have shown at
early stages of the disease the activated microglia were mainly associated with tyrosine
hydroxylase positive (TH+ve) neurites in the putamen and at the advanced stage with
damaged neurons in the SN (Sawada et al. 2006). The number of activated microglia
was also TNFα-, and IL-6-positive. These increase in the SN during the progress of PD.
Not only is there an indication of cytokine elevation but a decrease in neurotrophins such
as BDNF and nerve growth factor (NGF), in the nigrostriatal DA regions and ventricular
and lumbar cerebral spinal fluid of PD patients (Nagatsu et al. 2000a). Furthermore,
levels of TNFα receptor R1 (TNFα-R1, p55), bcl-2, soluble Fas (sFas), and the activities
of caspase-1 and -3 were also elevated in the nigrostriatal DA regions. Activated
microglia and increased levels of inflammatory mediators have been detected in the ST of
PD patients, in vivo (Waters et al. 1987; Nagatsu et al. 2000a; Wu et al. 2002; Hald and
Lotharius 2005; Novikova et al. 2006; Sawada et al. 2006) and in vitro studies (Le et al.
2001; Gao et al. 2003b; Gao et al. 2003a; Wang et al. 2005; Shavali et al. 2006; Mount et
al. 2007) which all point to an inflammatory component of DAergic cell loss. Acute
toxin exposure in these models or decades after drug use in humans have shown that the
presence of activated microglia still persists, suggesting a brief insult can induce ongoing
inflammatory response (Shavali et al. 2006; Wilms et al. 2007). Moreover, not only do
activated microglia phagocytose damaged cell debris but also neighbouring intact cells,
further supporting their active participation in self-perpetuating neuronal damaging cycles
(Kim and Joh 2006).
25
1.6 CELL DEATH IN PD
1.6.1 Apoptosis
There has been much debate about the mode of cell death in PD. Apoptosis has been
suggested by some researchers (Agid and Blin 1987; Mochizuki et al. 1997; Tatton and
Olanow 1999; Tatton et al. 2003) and others who disagree (Kosel et al. 1997; Jellinger
2000). A recent study by Tatton (Tatton 2000) shows that there is an increased
occurrence of DNA fragmentation and chromatin condensation in the melanised cells of
SN of PD patients compared to controls. There was also an increase in caspase-3 and
Bax (a proapoptotic member of Bcl-2 protein family) immunoreactivity and nuclear
translocation of glyceraldehyde-3-phosphate dehydrogenase (GADPH) in PD nigral
DAergic neurons. This demonstrates the occurrence of ongoing apoptotic processes and
an involvement of mitochondrial processes.
1.6.2 Apoptotic markers in PD
Apart from caspase-3, Bax is another protein which plays a critical role in apoptosis is
p53 which has been found to be up-regulated in the midbrain and ST of PD patients (de la
Monte et al. 1998). This supports the role of intrinsic apoptotic pathways involving the
mitochondria. Interestingly, tumor necrosis factor receptor 1 (TNFR1, a death receptor)-
positive DAergic neurons have also been found in the SN of controls and PD patients
(Boka et al. 1994). The level of tumor necrosis factor (TNF-α)-positive glial cells were
markedly elevated (Hunot et al. 1999), and that of melanized neurons expressing death
the receptor adaptor protein Fas-associated death domain (FADD) and caspase-8 were
reduced in this area of PD patients (Hartmann et al. 2000; Hartmann et al. 2001). This
26
implicates the existence of an extrinsic mechanism known as the TNF receptor ligand
system.
Caspase-8 is also activated early on in mice treated with MPTP, possibly as result of of a
downstream consequence of cytochrome c release from mitochondria as inhibitopn of
caspase-8 inhibition resulted in necrosis rather than rescue of the DAergic cells
(Hartmann et al. 2001). Thus intervention as this point may be too late in the cascade of
events. Different toxins and models induce different outcomes, as comprehensively
reviewed by Blum (2001), 6-hydroxydopamine (6-OHDA) induces strong reactive
oxygen species (ROS) production, weakening cellular antioxidant defenses, lipid
peroxidation, DNA strand breaking, cytoskeletal changes, and some evidence of
mitochondrial impairment. Both in vivo and in vitro models, 6-OHDA induced both
apoptosis and necrosis and an upregulation of p53 and caspase activations. Although
overexpression of Bcl-2 and caspase inhibitors provide some protection from death it did
not maintain functionality of the cultured DA cells, thus these models provide useful
insights only into the aim of slowing disease progression.
MPP+ exposure in cell culture and in animal models also showed evidence of apoptosis
with p53 involvement, Bax upregulation, and caspase-3 activation (Blum et al. 2001). It
is well received that the common final pathway in DAergic degeneration involves
oxidative stress (OS) and/or mitochondrial impairment (Mattson 2000). In addition,
some neurons that die due to energy loss thus can not complete apoptosis die by necrosis,
resulting in gliosis and microgila activation inducing overproduction of nitric oxide (NO),
27
reinforcing the damages (Hirsch et al. 2000). Another complex I inhibitor is rotenone, a
widely used model of apoptosis in cell culture models. Chronic rotenone administration
leads to complex I inhibition in the brain, resulting in selective DAergic nigrostriatal
degeneration and typical parkinsonian symptoms and cytoplasmic inclusions reminiscent
of LBs (Bezard et al. 1997).
1.7 CURRENT TREATMENTS AND POTENTIAL THERAPIES FOR PD
1.7.1 Symptomatic Treatment
There is no cure for PD, since currently available therapies can neither arrest or reverse
the progression of the disease nor provide neuroprotection. However, the motor
symptoms can be initially managed with several different drugs. The drugs used to treat
PD either boost the levels of DA in the brain or mimic the effects of DA. The use of L-
DOPA which is still considered the gold standard aims to relieve PD motor symptoms by
replacing the deficient neurotransmitter, DA. However, the therapeutic efficacy of L-
DOPA tends to fade with time and the development of motor and/or psychiatric side
effects. An issue that is currently debated is the hypothesized toxicity of L-DOPA.
Experimental data suggest that prolonged use of the drug may contribute to the
degenerative process of PD. Various in vivo and in vitro studies have shown that L-
DOPA can be toxic, as a result of its pro-oxidant properties (Ziv et al. 1997; Melamed et
al. 1998). Whether L-DOPA is toxic per se or because it converts to DA whose
metabolism is associated with formation of ROS, as well is unclear (Coyle and
Puttfarcken 1993).
28
These considerations have led to the introduction of DA agonists in PD therapy. This
class of drugs, that includes compounds such as pergolide, bromocriptine, lisuride or the
more recent ropinirole and pramipexole, acts directly on the DA receptors mostly of the
D2 type mimicking the effect of DA. DA agonists reduce the occurrence of motor
complications, in both experimental and clinical studies and are used as an adjunct to L-
DOPA in patients with dyskinetic movements and fluctuations in the motor response to
the drug (Olanow 1992a, b) by restoring the normal efficiency of the DAergic pathways.
Evidence suggests that DA agonists may have also neuroprotective properties. Such
properties would be related to the ascertained efficacy of DA agonists in antagonizing
OS, at various levels (Olanow et al. 1998). Apart from levodopa, drugs that are currently
prescribed for the management of PD include DA receptor agonists, selegiline,
amantadine, catechol-O-methyl transferase (COMT) inhibitors and anticholinergics
(Figure 1.2, Table 1.3). Most current drugs are still in development and/or clinical phase
and continous research is employed to produce the most effective treatments which will
have all desired effects such as neuroprotection, less severe side effects, optimal delivery
of drugs to site of problems, relieve/reduce motor symptoms and delayment of the
neuronal degeneration.
1.7.2 Cell Transplantation
Transplantation of DA-producing neurons to replace those degenerated during the
pathogenesis of PD is a promising approach to treatment. Thus far this is the only
advancement that has shown the capacity to allow patients to achieve full restoration of
their functional capacity. The grafts have shown minimal immunological rejection in
29
recipients and in the most successful trials have even allowed patients to withdraw from
levodopa therapy (Lindvall and Hagell 2001). However, the majority of studies have
derived allogenic ventral mesencephalic (VM) tissue from human foetuses (Subramanian
2001; Levy et al. 2004). This presents, apart from the ethical issues, the implication that
sources of such tissues will be limited and cannot provide a consistent and reproducible
response. Furthermore, substantial quantities are required to be adequately efficacious.
Stem cells as alternative neuronal sources are receiving a great deal of focus as they can
provide an ‘unlimited’ source of tissue that can be employed to generate mature DAergic
neurons to innervate the affected ST (Sonntag et al. 2005). The most promising is the
bone marrow stem cells (Lu et al. 2004) and ordinary epithelial skin cells (Chunmeng and
Tianmin 2004) may possess the potential to be sources of inducible stem cells. Although
studies have shown promising results in preclinical trials, generated neurons have
survived poorly after transplantation in animal subjects (Lindvall and Hagell 2001; Levy
et al. 2004)}. One of the most difficult hurdles to overcome is the poor rate of graft cell
survival as 90% of the transplanted cells fail to survive following intracerebral grafts
(Sortwell et al. 2000; Emgard et al. 2003). Furthermore, few studies have been able to
investigate the long term implications of transplantation; thus the safety of such treatment
at this stage remains questionable.
1.7.3 Gene therapy
Transplanted neurons in PD patients are a potential target for the development of gene
therapy procedures. A variety of different viral and non-viral methods for achieving gene
delivery have been described (Latchman and Coffin 2001). Among the numerous
30
potential genes that have been evaluated for therapeutic efficacy for PD, those encoding
tyrosine hydroxylase (TH), guanosine triphosphate cyclohydrolase I and aromatic l-
amino acid decarboxylase all allow for an increase in the production of DA (Duan et al.
2005). The differentiation of these neurons is mediated by transforming growth factor
beta (TGF-�) and bone morphogenic factors (BMPs) (Sonntag et al. 2005). Glial cell
line-derived neurotrophic factors (GDNF) have thus far proven to be the most promising
option available to restore DAergic activity in the SN (Pahnke et al. 2004). Adenoviral
and lentiviral nigrastriatal implants to liposomes (Muramatsu et al. 2003; Azzouz et al.
2004; Muramatsu et al. 2005; Wong et al. 2006) have also been studied. However more
recent studies have indicated conflicting results regarding its effectiveness and the
numerous side effects (Nutt et al. 2003; Lang and Obeso 2004; Lang 2006).
Thus the development of a stem cell that has the capacity to differentiate into DAergic
neurons as well as produce such factors is much needed (Sonntag et al. 2005). Genes
encoding for the vesicular monoamine transporter-2 and glutamic acid decarboxylase
have also demonstrated some benefit. Kang et al. (Kang et al. 2001) investigated
potential genes that would be optimal for such therapy and found that the enzyme TH,
which has been used in earlier studies, functions only when the essential cofactor,
tetrahydrobiopterin is present. Thus, new developments into the delivery of genetically
modified cells that can convert levodopa to DA and store it for gradual release is
required. Alternatively, it has been proposed that nanomaterials like nanotubes may be
employed therapeutically as ligand carriers or vectors for drug, DNA or gene delivery for
PD (Singh et al. 2005). However, so far, this technology remains in the experimental
31
stages of development and far from clinical trials or commercial application (Pereira and
Aziz 2006).
1.7.4 Surgical methods
Prior to the commercial availability of L-DOPA, treatment for PD emphasized surgical
intervention (Kelly 1995). Such intervention such as thalamotomy focused on the
reduction of tremor, but failed to address the more debilitating symptom, bradykinesia
(Kelly 1995). However, pharmacological therapy for PD often becomes inadequate over
long-term use and during significantly progressed stages of the disease. The patient's
disability increases despite maximal drug management and many patients develop motor
fluctuations and dyskinesias. Surgical interventions for PD have been shown to be
beneficial for refractory symptoms. However their role is limited to being the ‘means of
last resort’ due to the high risk of potential complications and limited long-term efficacy.
Recent advances in this field have provided a greater range of surgical options.
Thalamotomy, pallidotomy are destructive lesions thought to improve motor deficient.
Deep brain stimulation (DBS) however offers subthalamic nucleus stimulation improves
levodopa-induced ‘off’ period function, decreases ‘off’ time, and reduces dyskinesia
(Zesiewicz and Hauser 2001). Surgery is considered for people with intolerable adverse
side effects from medication, and those patients who have significant cognitive capacity
(Olanow and Brin 2001; Olanow 2002; Betchen and Kaplitt 2003).
Possible adverse side effects of surgery include brain haemorrhage, infarction, seizures,
and even death (Beric et al. 2001; Seijo et al. 2007). Furthermore, successful surgical
32
DBS procedure can still lead to side effects that include worsening dyskinesia,
paraesthesias, speech and gait disturbances (Umemura et al. 2003). Developments in
nanotechnology have indicated the potential for implantable carbon nanotubes and
nanochips to be employed in this arena (Linazasoro 2008). These systems promise to
allow greater safety and precision for the delivery of impulses in the SN, and therefore
reduce side effects the aforementioned problems with regard to surgery and equipment
malfunctions. Studies are still in preliminary phases of development for eliminating
repeated surgeries; however animal studies have yielded exceedingly promising results.
Figure 1.2. Action sites of pharmacological therapies currently available for PD
(Singh et al. 2007).
33
Table 1.3 Summary of current PD treatment
Antiparkinsonian drugs Neuroprotection Effects or comments A. Anticholinergics Trihexyphenidyl biperiden piroheptine profenamine metixene
-
Classical parkinsonian drugs
B. DA replacements L-DOPA (levodopa) Amantadine Carbidopa Benserazide Selegiline(deprenyl) Lazabemide Tolcapone Entacapone
- + - - + - - -
DA precursor DA releaser, NMDA receptor antagonist Inhibits peripheral AADC (DCI) Inhibits peripheral AADC (DCI Inhibits MAO-B Inhibits MAO-B Inhibits COMT Inhibits peripheral COMT
C. DA receptor agonists Ergot-derivatives Bromocriptine Pergolide Cabergoline Non-ergot-derivatives Ropinirole Pramipexole Talipexole
+ + +
+
+ +
D2-R agonist, scavenger of ROS D2/D1-R agonist, scavenger of ROS D2/D1-R agonist, reduction of lipid peroxidation D2/D3-R agonist, scavenger of ROS, increases GSH with an induction of regulating enzymes D2/D3-R agonist, induction of Bcl-2 (at low conc.), inhibits α -synuclein aggregation (at high conc.) D2/α 2-R agonist, induction of Bcl-2 (at low conc.), inhibits α -synuclein aggregation (at high conc.)
D. Others L-threo-DOPS (droxidopa) GDNF GPI-1046 Estrogens KW6002 SIB1508Y, ABT418 Surgical therapy
_ + +
+ - + +
Precursor of noradrenaline Neurotrophic factor, induces Bcl-2 and Bcl-x Immunophilin ligand Neurite outgrowth and regenerative sprouting Female sex steroid hormone, induces Bcl-2 Adenosine A2A receptor antagonist, inhibits indirect pathway Nicotine receptor agonists Transplantation of DA-releasing cells, stem cells, etc. Gene therapy
+, exhibits neuroprotective effects; —, nothing or unknown (adapted from (Kitamura et
al. 2002)).
33
34
1.8 BASAL GANGLIA
1.8.1 INTRODUCTION
The basal ganglia (BG) circuit plays a key role in the regulation of voluntary movements
as well as in behavioural control and cognitive functions (Graybiel et al. 1994). In the
BG circuit, cortical inputs reach separate subpopulations of striatal �-aminobutyric acid
(GABA)-containing medium sized spiny neurons. From the striatum (ST), cortical
information are transmitted to SNpr through parallel routes named “direct” and “indirect”
pathways (Chesselet and Delfs 1996) and to the others output structures of the BG
(globus pallidus, subthalamic nucleus and thalamus) and finally link back to the frontal
cortex (see Figure 1.3). This circuit plays a key role in the regulation of voluntary and
purposive movements as well as in behavioural control and cognitive functions (Wise et
al. 1996). The selective neuronal loss in one of the structures within the circuit results in
clinical syndromes characterized by motor and cognitive dysfunctions such as Parkinson's
disease and Huntington's disease (Albin et al. 1995). PD’s core pathological features are
represented by the heterogenous loss of pigmented DAergic neurons in the SNpc and of
their projecting fibers in the ST (Lang and Lozano 1998a, b), whereas in Huntington's
disease the neurodegenerative disorder primary involves the degeneration of striatal spiny
neurons with sparing of striatal large cholinergic interneurons (Ferrante et al. 1985; Price
et al. 1998).
1.8.2 STRIATUM (ST)
The ST, which is the major component of the BG is situated in the forebrain and consists
of the caudate nucleus (CN), putamen (Put) and ventral ST (VST). The major neuronal
35
population in the ST is represented by spiny projection neurons, accounting for almost
95% of total striatal cells (Kemp and Powell 1971) with �-amino-butyric acid (GABA) as
the main neurotransmitter (Kita and Kitai 1988). Within the projection neurons, GABA
can be co-localized, alternatively, with enkephalin or substance P/dynorphin (Beckstead
and Kersey 1985). The remaining 5% of striatal cells consists of aspiny interneurons
containing, alternatively, acetylcholine, somatostatin, NADPH-diaphorase or GABA
associated with parvalbumin or calretin (Kawaguchi et al. 1995). Recently, the presence
of DAergic neurons intrinsic to the ST has also been suggested (Betarbet et al. 1997).
The main targets of striatal projections are the medial and lateral segments of the globus
pallidus and the SNpr (Parent and Hazrati 1995a, b). Neurons containing enkephalin are
believed to project to the lateral globus pallidus, while neurons containing substance
P/Dynorphin project to the medial globus pallidus and SNpr. This striatal output is known
as the direct and indirect pathway model of BG functional organization (figure 1.3a).
The ST is the main input structure of the BG circuit. The major neural input to the ST is
excitatory. Glutamatergic projections from all cortical areas (McGeorge and Faull 1989)
converge onto striatal neurons with other excitatory inputs to the ST arise from the
thalamus (Berendse and Groenewegen 1990), and from limbic structures, particularly the
amygdala (Kelley et al. 1982). Another input to the ST originates from DAergic neurons
located in the SNpc and in the ventral tegmental area (VTA) (Nieuwenhuys et al. 1982).
The ST also receives serotonergic afferent projections from the nucleus of the raphe
(Halliday et al. 1990) and a sparse noradrenergic innervation from the locus coeruleus
(Aston-Jones et al. 1986). The ST exhibits a variety of neurotransmitter receptors, which
36
also show a considerably higher density at the striatal level, compared to the other BG
nuclei.
1.8.2.1 Dopamine receptors (D1R & D2R)
Striatal neurons express both D1 and D2 DA receptors, which mediate the modulatory
effect of DA released from nigrostriatal terminal. D1 and D2 receptors are functionally
segregated to different subsets of striatal neurons. D1 receptors are expressed by neurons
projecting to the SNpr and medial globus pallidus, while D2 receptors are expressed by
neurons projecting to the lateral globus pallidus (Gerfen and Keefe 1994). A small
population of projection neurons expresses both D1 and D2 receptors ((Surmeier et al.
1996). The role played by DA at the striatal level has been extensively studied.
Electrophysiological studies suggest that DAergic transmission modulate the striatal
responses to incoming inputs, particularly those mediated by glutamate (Calabresi et al.
1993a; Cepeda and Levine 1998). Release of glutamate in the ST is modulated partly by
nigrostriatal DAergic projections.
Chronic blockade of D2 dopamine receptors causes an increase in the levels of both basal
extracellular and potassium-releasable glutamate in ST (Yamamoto and Cooperman
1994). In addition striatal DA depletion increases spontaneous glutamate release in ST
(Calabresi et al. 1993b). Behavioral studies conducted in freely moving animals show
that intra-striatal administration of DA attenuates neuronal excitation elicited by cortical
activation in both rats (Kiyatkin and Rebec 1996) and monkeys (Rolls et al. 1984).
Studies carried out in anaesthetized animals (Ohno et al. 1987; Johansen et al. 1991) or
37
using in vitro preparations (Calabresi et al. 1987; Nicola et al. 1996) have suggested that
the inhibitory modulation might be mediated primarily by D1 receptors (Kiyatkin and
Rebec 1999a, b).
1.8.3 PALLIDAL COMPLEX
In primates, the pallidal complex is comprised of two segments the medial globus
pallidus and lateral globus pallidus. In rodents, medial and lateral segments correspond to
the entopeduncular nucleus and globus pallidus, respectively. Both pallidal segments are
populated by GABAergic neurons (Oertel and Mugnaini 1984; Oertel et al. 1984).
1.8.3.1 Entopeduncular nucleus (medial globus pallidus)
The entopeduncular nucleus is the smallest nucleus of the BG circuit. It plays a central
role in the transmission of the BG output to the thalamus and to the motor cortex. Along
with the SNpr which shares many histologic and functional properties with the medial
globus pallidus, this area is the main output nucleus of the BG circuitry. The
entopeduncular nucleus projects primarily to the motor thalamus. In particular, to the
ventral anterior and ventral lateral thalamic nuclei which in turn, project diffusely to the
motor cortex (Carter and Fibiger 1978; Nauta 1979; Parent and Hazrati 1995b).
Entopeduncular neurons receive a combination of inhibitory (GABAergic) and excitatory
(glutamatergic) projections. The balance between the two opposite systems determines
the functional activity of the nucleus. The main source of GABAergic fibers is the ST
(Parent and Hazrati 1995b) with other GABAergic projections to the nucleus arising from
38
the adjacent globus pallidus (Parent and Hazrati 1995a). The excitatory innervation of
the entopeduncular neurons is provided by the subthalamic nucleus (STN) (Parent and
Hazrati 1995a), and the frontal cortex (Naito and Kita 1994a).
1.8.3.2 Globus pallidus (lateral globus pallidus)
Pallidal neurons use mainly GABA (co-localized with enkephalin) as neurotransmitter
(Fonnum et al. 1978) and project to structures localized within the BG circuit. These are
mainly the STN, SNpc, entopeduncular nucleus, pedunculopontine nucleus and reticular
thalamic nucleus (Parent and Hazrati 1995a). There have been reports of the existence of
pallidal, GABAergic projections to the SNpr (Smith et al. 1998) and, cholinergic and
non-cholinergic projections to the cortex have also been described. The main sources of
afferent projections to the globus pallidus are the ST-sending GABAergic fibers and the
subthalamic nucleus-sending glutamatergic fibers (Parent and Hazrati 1995a). Fèger
(Feger 1997) recently proposed the existence of another excitatory input to the globus
pallidus, originating from the thalamus. In addition, the nucleus seems to receive a
DAergic innervation from fibers originating from the nigrostriatal pathway (Lindvall and
Bjorklund 1979). This finding has been recently confirmed by Cossette et al. (Cossette et
al. 1999), who showed that, in the human brain, nigrostriatal axons provide collaterals
that reach the pallidal complex.
Like the entopeduncular nucleus, the globus pallidus is particularly enriched with GABA
receptors �1 and �2 subunits (Wisden and Seeburg 1992; Boyes and Bolam 2007). D1
39
and D2 dopamine receptors are also present in the globus pallidus (Richfield et al. 1987),
which further supports the potential role of DA at this level.
1.8.4 SUBSTANTIA NIGRA (SN)
Two distinct structures can be recognized within the SN: a densely populated, pigmented
area called substantia nigra pars compacta (SNpc) and an adjacent cell-sparse portion,
located ventrally, called substantia nigra pars reticulate (SNpr).
1.8.4.1 Substantia nigra pars compacta (SNpc)
Neurons in the SNpc contain neuromelanin and use DA as neurotransmitter. The
recipients of nigral projections are the striatum, subthalamic nucleus and globus pallidus.
Another group of DAergic neurons, homogeneous to nigral neurons, is located more in
the ventral tegmental area (VTA). These neurons project to the ventral ST, amygdala and
cerebral cortex (Hauber 1998). Recently, electrophysiological and morphological studies
have suggested the existence of a small percentage (5–8%) of nigrostriatal neurons
containing GABA instead of DA (Rodriguez and Gonzalez-Hernandez 1999).
Afferent projections to the SNc originate from various structures. Both the ST (Ribak et
al. 1980) and the globus pallidus (Smith et al. 1989) send GABAergic projections to
nigral DAergic neurons. On the basis of electrophysiological evidence, the existence of
GABAergic afferent projections from the SNpr has been suggested (Tepper et al. 1995).
The SNc receives glutamatergic projections from the prefrontal cortex, subthalamic
nucleus and pedunculopontine tegmental nucleus (which also sends cholinergic
40
projections) (Naito and Kita 1994a, b; Bezard et al. 1997). Nigral DAergic neurons also
receive serotonergic projections from the medial and dorsal raphe nuclei (Hauber 1998).
Among the diverse neurotransmitters that affect nigral activity, it has been pointed out the
importance of the glutamatergic input. Both NMDA and AMPA receptors are located on
the soma and dendrites of DAergic neurons and regulate their electrical activity (Meltzer
et al. 1997a). Iontophoretic administration of NMDA, AMPA and glutamate increase the
firing rate of nigral DAergic neurons. These effects are prevented by previous
administration of selective antagonists, such as the NMDA antagonists or the AMPA
antagonist (Christoffersen and Meltzer 1995; Meltzer et al. 1997b). Recent evidence
shows that nigral DAergic neurons express primarily GluR1, GluR2/3 and NR1 subunits
(Albers et al. 1999). The presence of metabotropic receptors mGluR1 has been
demonstrated within the cell body, axons and dendrites of SNpc neurons (Kosinski et al.
1998).
1.8.4.2 Substantia nigra pars reticulate (SNpr)
The SNpr is located ventral and adjacent to the SNpc, and is populated by GABAergic
neurons (Oertel and Mugnaini 1984; Oertel et al. 1984). The SNpc sends its inhibitory
projections to the ventral anterior and ventral lateral nuclei of the thalamus. Other targets
of nigral projection include the superior colliculus and the pedunculopontine nucleus
(Parent and Hazrati 1995b). The similarity with the entopeduncular nucleus extends to
the afferent projections, thus the neurons of the SNpc receive a combination of inhibitory
(GABAergic) and excitatory (glutamatergic) inputs from diverse structures. The main
41
source of inhibitory projections is the ST (Chevalier et al. 1985). Other sources of
GABAergic afferents include the GP (Smith and Bolam 1989; Smith et al. 1989), nucleus
accumbens (Deniau et al. 1994) and ventral pallidum (Groenewegen et al. 1993).
Glutamatergic projections to the SNpc originate in the subthalamic nucleus (Kita and
Kitai 1987), playing an important role in the regulation of nigral activity. Selective
subthalamic lesion reduces the activity of mitochondrial enzymes complex I, II and IV in
the SNpc. This reflects the reduced activity of nigral neurons resulting from the abolition
of the subthalamic excitatory input (Blandini 2001; Blandini et al. 2001b; Blandini et al.
2001a). The pars reticulata shares many histologic and functional properties ncluding the
input/output connections with the entopeduncular nucleus. The two nuclei are considered
the major output structures of the BG circuitry and are often referred to as the BG output
nuclei.
1.8.5 SUBTHALAMIC NUCLEUS (STN)
The STN is the only glutamatergic nucleus of the BG circuit (Smith et al. 1998) that
sends excitatory projections primarily to the BG output nuclei -SNpr and medial globus
pallidus (entopeduncular nucleus), and to the lateral GP. Other targets include the ST,
SNpc and motor cortex (Kita 2007). Subthalamic neurons receive an important inhibitory
innervation from GABAergic neurons of the lateral GP. Other inhibitory projections arise
from the ventral pallidum and ventral ST (Groenewegen and Berendse 1990). The STN
also receives excitatory projections from the sensory-motor cortex (Fujimoto and Kita
1993), thalamic parafascicular nucleus, and pedunculopontine nucleus (Canteras et al.
1990).
42
DAergic innervation of the STN, originating from the SNpc has also been described in
rats (Hassani et al. 1996) and recently has been demonstrated in the human STN, as well
(Cossette et al. 1999; Cossette et al. 2005b; Cossette et al. 2005a). Both GABA and
glutamate receptors are abundantly expressed in the STN (Wisden et al. 1992; Wisden
and Seeburg 1992). As for the glutamate receptors, both the ionotropic and the
metabotropic families have been described (Tallaksen-Greene and Albin 1996; Tallaksen-
Greene et al. 1998). DA receptors are present at the subthalamic level, where they play
an important role in the context of the BG functional organization. Both D1 and D2
receptors have been repeatedly described in the nucleus (Dawson et al. 1990; Bouthenet
et al. 1991; Johnson et al. 1992). Recently, Flores et al. (Flores et al. 1999) reported the
presence of mRNAs and binding sites for D1, D2 and D3 receptors.
1.8.6 THE BASAL GANGLIA CIRCUITRY
1.8.6.1 The direct and indirect pathway model
The functional architecture of BG has attracted numerous researchers in the last decade
leading to the formulation of a model of BG functioning (Gerfen et al. 1987b; Gerfen et
al. 1987a; Albin et al. 1989; Alexander and Crutcher 1990; Graybiel 1990; Gerfen 1992;
Albin et al. 1995). According to the model, the ST is the main input nucleus of the
circuit, transmitting the flow of information received from the cortex to the BG output
nuclei, SNpr and medial globus pallidus, via a direct and an indirect pathway. The two
pathways originate from different subsets of striatal neurons and, remain functionally
segregated. In the direct pathway, striatal GABAergic neurons, containing dynorphin as a
co-transmitter and expressing D1 DA receptors, project mono-synaptically to the SNpr
43
and medial globus pallidus. In the indirect pathway, the striatal output reaches the target
nuclei via a more complicated route, with different subset of GABAergic neurons
containing enkephalin and expressing D2 receptors, these projects to the lateral globus
pallidus, which sends GABAergic projections to the subthalamic nucleus. The STN, in
turn, sends its glutamatergic efferents to the output nuclei and to the lateral globus
pallidus. From the output nuclei, inhibitory, GABAergic projections reach the ventral
lateral and ventral anterior nuclei of the motor thalamus. Thalamic nuclei then send
glutamatergic projections to the motor cortex, thus closing the loop (Figure 1.3).
The functional consequence of such organization is that the activation of the direct or the
indirect pathway leads to opposite changes in the net output of the BG circuitry. The
activation of the striatal GABAergic neurons giving rise to the direct pathway causes
inhibition of GABAergic neurons of the output nuclei, leading to the disinhibition of
thalamic nuclei, which are under the inhibitory control of the output nuclei projections.
Conversely, activation of the striatal neurons that project to the lateral globus pallidus, in
the indirect pathway, causes inhibition of the lateral globus pallidus and subsequently
disinhibits the STN. The activation of the subthalamic nucleus which is glutamatergic
increases the activity of the output nuclei. Consequently, their inhibitory control over the
motor thalamus results is enhanced (Alexander and DeLong 1985a, b).
Subthalamic excitatory projections modulate the neuronal activity in the SNpr and medial
globus pallidus. Electrical subthalamic stimulation increases metabolic rates for glucose
in these nuclei (Tzagournissakis et al. 1994) with unilateral ablation of the STN, causes
44
an ipsilateral reduction in the activity of mitochondrial enzymes complex I, complex II
and complex IV in the recipients of subthalamic projections (Blandini et al. 1997;
Blandini 2001; Blandini et al. 2001b). Thus, the final output of the basal ganglia
processing is under the direct control of the STN.
1.8.6.2 BG changes in PD
The neurodegenerative process in PD causes a functional re-arrangement of the BG
circuitary. The current model (Graybiel et al. 1990; Gerfen and Engber 1992; Albin et al.
1995) states that the DAergic denervation of the ST induces a cascade of events leading
ultimately to the increased activity of the BG output nuclei. The enhanced activity of the
output nuclei would be a result of the enhanced glutamatergic drive fom the STN. This
model also predicts that the enhanced activity from the output nuclei results in an
increased inhibitory control over the motor thalamus and thus the reduction of the
thalamic glutamatergic output to the motor cortex. These changes are believed to be the
main reason for the parkinsonian symptoms (Figure 1.3b).
These functional changes occurring in PD has been mainly provided by animal studies in
particular rodent and non-human primates. The most significant alterations in rodent
models include increases in neuronal firing rate (Bergman et al. 1994), glucose
metabolism (Mitchell et al. 1992), and mitochondrial enzyme activity (Porter et al. 1994;
Vila et al. 1997) in the STN or it sprojection nuclei. In rats, lesion of nigrostriatal
pathway causes down-regulation of glutamate receptors in projection nuclei of STN this
is consistent with the current hypothesized hyperactivity of subthalamic glutamatergic
45
projections (Porter et al. 1994). Subthalamic ablation leads to increases in mitochondrial
enzyme activity in the entopeduncular nucleus and SNpr of 6-OHDA lesioned rats
(Blandini et al. 1997) and abolishes rotational response to apomorphine (Burbaud et al.
1995; Blandini et al. 1997). Also it normalizes the firing rate and discharge pattern of
pars reticulate neurons in addition preventing the changes in gene expression in the ST,
entopeduncular nucleus and GP (Delfs et al. 1995) of rat with unilateral nigrostriatal
lesion.
Human studies also support the view that subthalamic hyperactivity plays a central role in
PD pathophysiology. Not only reports of PD symptoms disappearance in a PD patient
after the occurrence of a subthalamic hematoma (Sellal et al. 1992) but also a down-
regulation of NMDA receptors in the medial GP as result of increased activity of
subthalamic projections to the medial GP (Lange et al. 1997). Thus the introduction of
electrophysiological techniques in the therapy of PD was rationaled by blocking the
hyperactivity of subthalamic neurons resulting in relievement of PD symptoms.
The STN is also a key player in the compensatory mechanisms that sustain the DAergic
function in the pre-symptomatic phase of PD. The gradual loss of in the SNpc is initially
accompanied by an increased DA at the ST level by efficiency of residual DAergic
neurons (Zigmond et al. 1990b; Zigmond et al. 1990a). Monkeys rendered parkinsonian
by MPTP have increased glutamatergic inputs to SNpc as temporary blockade of these
inputs reveals the PD motor abnormalities (Bezard et al. 1997). This compensatory
mechanism masks the PD symptoms in an early phase of PD but as it progresses and the
46
degenerative process reaches a threshold at which the functional changes leads to PD
symptoms are triggered.
47
Figure 1.3 The current model of the basal ganglia.
The diagrams illustrate the functional organization of the basal ganglia in (a) normal
conditions, (b) Parkinson's disease. Thick and broken lines indicate pathways that are
believed to be respectively hyperactive or hypoactive in the pathological conditions. The
blue and red indicate inhibitory (Glutamate) and excitatory (GABA) projections,
respectively. D1 = Direct pathway, D2 = Indirect pathway. Substance P (SP) + GABA
(excitatory) = D1 pathway, Enkephalin + GABA (excitatory) = D2 pathway. Dopamine
on D2 receptor (inhibitory), dopamine on D1 (excitatory) (Parent A et al., 2000). With the
current model it is thought that dopamine inhibits neuronal activity in the indirect
pathway and to excite neurons in the direct pathway. Whereas in the parkinsonian state,
dopamine depletion leads to disinhibition of D2-receptor-bearing striatal neurons in the
GPi/SNr and overinhibition of thalamo-cortical and brainstem motor centers resulting in
parkinsonism.
48
D2 D2 D1 D1
49
1.9 THE NIGROSTRIATAL PATHWAY
1.9.1 INTRODUCTION
Nigrostriatal DAergic neurons are critically involved in the control of voluntary
movements (DeLong 1990; Grillner and Mercuri 2002). As a result, their loss in PD
leads to profoundly disabling motor symptoms (Agid 1991). The development of the
mDA cell type during mammalian embryogenesis can be subdivided into three distinct
processes: (1) the induction of a progenitor field within the neuroepithelium competent to
generate mDA precursors at early stages of neural development (approx. from embryonic
day (E) 8.5 to E10.5 in the mouse); (2) the specification of a mDA neuronal fate in these
precursors at intermediate stages (approx. from mouse E10.5 to E12.5); and (3) the
acquisition of the mature phenotype or terminal differentiation of mDA neurons at
relatively late stages of neural development (i.e. from E12.5 onwards).
1.9.2 Ontogeny of Midbrain DAergic neurons
During the development of the rat, the cells of the SN and VTA are produced between
embryonic day 11 and 15 (E11 and E15). This occurs in the ventricular zone where the
greatest numbers of cells are undergoing division between E13 to E15 (Noisin and
Thomas 1988; Voorn et al. 1988; Santana et al. 1992; Aubert et al. 1997; Park et al. 2000;
Gates et al. 2006). It has been demonstrated that rat mDN are specified by the floor plate
(FP-specialised group of cells at the ventral midline of the neural tube) which is mediated
by contact-dependent mechanism (Hynes 1995). The FP, secretes the lipid-modified
glycoprotein Sonic hedgehog (Shh), which is known for its pivotal role in the
specification of the different ventral populations in the hindbrain and spinal cord by
50
controlling the expression of different transcriptional regulators (Placzek and Briscoe
2005) in addition mediates the DAergic induction.
Following induction of DAergic neurons, migration occurs. Initially the neurons migrate
ventrally in a radial pattern (Marchand and Poirier 1983) along glia paths (Shults et al.
1990) which extends to the ventral surface of the mesencephalon and in a rostral
direction. During the migration process the cells differentiate and become
immunoreactive for TH from E12.5 (Specht et al. 1981b). The D1 (excitatory receptors)
are intially present in striatal patches then gradually the homogenous distribution
characteristic of the adult ST emerges. D2 (inhibitory) receptor expression within the ST
is a homogeneous population which finally forms clusters.
1.9.3 In vivo development of the nigro-striatal pathway
TH-ir terminals are first detected in the caudal putamen from E14.5 (Specht et al. 1981a,
b). Thereafter, the innervation of the rostral striatum shows a ventrolateral to
dorsomedial gradient and the innervation of caudal regions is from a ventromedial
position (Voorn et al. 1988). As they reach the ventral mesencephalon the dopaminergic
neurons then project axons to the ST. At E14, DAergic neuron bundles move towards
the ST and by E17 the bundles are dense and are present in the dorsolateral ST.
The DAergic neurons become more closely packed togther within each ventral
mesencephalon during stages E15-16 along each side of the midline. These two groups
in the more caudal regions merge along the midline to form the origin of the VTA, at E17
51
the dendrites can be clearly seen and from E19 onwards the SNpc, SNpr and VTA can be
distiguished throughout the whole VM. During the first postnatal week, the VM
develops the typical adult organiztion with DAeric neurons of the SN projecting dendrites
both within the pars compacta and reticulata (Voorn et al. 1988). There is a functional
and anatomical diversity among mDNs (Carlsson 1959; Dahlstroem and Fuxe 1964a, b).
mDN located within the SNpc (SN or A9) form the nigrostriatal pathway to the
dorsolateral ST and are essential for the control of voluntary movement; mDNs within the
ventral tegmental area (VTA or A10) form the mesolimbocortical system and controls
brain mechanisms of novelty, reward, and addiction; and mDNs within the retrorubral
field (RRF or A8) are involved in both local midbrain circuitry and the SN pathway.
1.9.4 In vivo development of patch/matrix innervation
The adult ST is functionally and structurally separated into “patch” and “matrix” regions.
These differences arise from the development of DAergic innervation of the ST, as
innervation to patch regions of the ST occurs before formation of the matrix (Voorn et al.
1988). By E19, the DAergic axons produce ramifications in “patch” regions throughout
the ST. Following birth a second group of neurons project axons to the matrix regions of
the ST, by third week postnatal week the characteristic adult pattern of innervation has
formed. Studies by Gerfen (Gerfen et al. 1987a) confirm these observations by 6-OHDA
injection into nigro-striatal pathway into newborn rats destroyed DAergic innervation of
patch regions whereas the matrix region which develops later, remained intact.
52
The subdivision of the ST into two major compartment-the striosomes (or patches) and
the extrastriosomal matrix, was first proposed by Graybiel and Ragsdale in 1978
(Graybiel and Ragsdale 1978). These investigators noted small AChE-poor striosomes
that represented approximately 10–20% of the total striatal volume and formed complex
three-dimensional labyrinths embedded in an AChE-rich matrix (Graybiel and Ragsdale
1978; Penney and Young 1986; Johnston et al. 1990). The notion of striosome/matrix
compartmentalization of the ST was further supported by studies of the distribution of a
wide variety of transmitter-related substances and by the organization of striatal afferent
and efferent connections (Pickel et al. 1981a; Pickel et al. 1981b; Ferrante et al. 1986;
Gerfen et al. 1987b; Gerfen et al. 1987a; Graybiel et al. 1987; Holt et al. 1997; Prensa et
al. 1999).
1.9.5 In Vitro modeling of nigrostriatal pathway
In vitro studies have been utilized to study the development of the nigrostriatal pathway.
Both dissociated neuronal cultures of DAergic nigro-striatal neurons and their target
region the striatum (Prochiantz et al. 1979; di Porzio et al. 1980; Hemmendinger et al.
1981; Prochiantz et al. 1981; Kotake et al. 1982; Denis-Donini et al. 1983, 1984) and
organotypic co-cultures (Whetsell et al. 1981; Jaeger et al. 1989; Ostergaard et al. 1990)
have been used to investigate the development of nigrostriatal neurons.
DAergic neurons in both organotypic (Ostergaard et al. 1990) and dissociated neuronal
cultures (Prochiantz et al. 1979) can survive in the absence of their striatal target region.
The TH-ir neurons of the VTA and SN derived from rats aged between E21 and postnatal
53
day 7 retain their basic morphological characteristics in organotypic cultures (Ostergaard
et al. 1990) however, the maturation and development of DAergic nigro-striatal neurons
measured by studies of DA synthesis and uptake, is enhanced by the presence of striatal
cells (Prochiantz et al. 1979; di Porzio et al. 1980; Shalaby et al. 1983) (or striatal
membranes (Prochiantz et al. 1981). Also, the DAergic neurons do not develop axons in
the absence of target cells but in their presence DAergic neurons form axons
characteristic of those formed in vivo (Hemmendinger et al. 1981).
The growth of these DAergic neurons is also shown to be target specific by innervation of
the ST. Total neurite outgrowth of the neurons is significantly reduced when DA neurons
are cultured with striatal cells but not with the predominant mesencephalic non-DA cell
population or with cerebellar cells at any concentration tested (Denis-Donini et al. 1983).
Direct neuro-neuronal interactions between DA neurons and striatal cells regulate the
development or maturation of the afferent DA cells is in agreement on the basis of
biochemical data (di Porzio et al. 1980). Further support that co-cultures containing the
VTA/SN-complex and the PFC or the striatum, dopaminergic mesencephalic neurons
able to innervate the target areas in the PFC and the STR but no TH+ve fibers and cells
were found in the prefrontal as well as in the striatal tissue cultured alone and also in co-
cultures which had not developed a bridge (Franke et al. 2003).
Co-cultures of dissociated striatal and mesencephalic cells have been used to demonstrate
the influence of target cells on the development of DA arbors (Prochiantz et al. 1979;
Manier et al. 1997). However, tissue organization and spatial cues related to this
54
organization are lost in dissociated cultures, and mechanisms relying on these aspects,
such as selectivity of the innervation, can hardly be studied in such cultures. Organotypic
slice cultures might represent a convenient compromise between in vivo models and the
use of dissociated cocultures (Bahr 1995; Gahwiler et al. 1997).
1.10 FACTORS INFLUENCING THE DEVELOPMENT OF MESENCEPHALIC
DOPAMINE NEURONS (mDNs)
Many factors have been identified that may be involved in the formation of the nigrostital
pathway by promoting survial, differentiation and maturation of nigrostriatal neurons in
vitro. The development of mDNs can be divided into 4 stages (Figure 1.4). First,
progenitors at the ventral mesencephalic surface that have self-renewing properties and
give rise to multiple cell types (‘stem-like’) arise at embryonic day 7.5 (E7.5). In a
second step, these are specified to a DAergic neuronal precursor cell fate, and several
molecular markers are associated with this population (see below). In a third stage, the
DAergic precursors exit the cell cycle and begin to display early mDN markers. Finally,
the early mDNs functionally mature, express mature mDN markers establishing
appropriate connectivity.
1.10.1 Early development
Mesencephalic organization is initiated by the positioning of key signaling centers such
as the floor plate, present at the ventral midline, and the midbrain–hindbrain junction
(MHJ, also known as the isthmic organizer) at the caudal extreme of the mesencephalon.
In addition, early patterning events also position the dividing stem-like precursors that are
55
competent to become mDNs in the context of instructive cues from signaling centers.
Genetic loss-of-function studies have identified a network of transcription regulators and
other signaling factors that underlie patterning of the midbrain and hindbrain precursors
(Joyner et al. 2000). Expression of two homeodomain transcription factors, Otx2 in the
midbrain and Gbx2 in the hindbrain, initially form a boundary at the MHJ at E7.5 in the
mouse and becomes well-defined by E9.5. Multiple signaling and regulatory molecules
function as effectors downstream of Otx2 and Gbx2 at the MHJ, including fibroblast
growth factor-8 (FGF8), Engrailed (En)-1 and -2 homeobox transcription factors, the
secreted factor Wnt1, the Lim-domain transcription factors Lmx1a and Lmx1b, Foxa2,
and the paired-like homeodomain proteins Pax-2, -5, and -8 (Liu and Joyner 2001).
1.10.2 Specification of mitotic mesencephalic precursors to the mDN fate
In the early embryonic midbrain (E7.5–E9.5 in the mouse), the floor plate organizer at the
ventral surface of the neural tube (Hynes et al. 1995) and the midbrain/hindbrain junction
at the caudal extreme of the midbrain (MHJ or isthmus) (Alvarado-Mallart 1993) instruct
the fate of adjacent neural progenitors. A number of secreted proteins encode the action
of these signaling centers such as Sonic hedgehog (SHH), secreted by the floor plate
organizer, is necessary to instruct the ventral cell fate of progenitors in vivo and in vitro
and fibroblast growth factor-8 (FGF8) a key MHJ-derived signal that is required for
specification of the mesencephalic progenitors (Ye et al. 1998). Wnt1 is an additional
secreted factor that promotes mDN development (Schulte et al. 2005). Transgenic
markers demonstrate Wnt1 expression (lateral to the floor plate) at the region of DAergic
56
neuronal progenitors at the time point when postmitotic mDNs are born (murine E9.5–
E11) (Zervas et al. 2004)
1.10.3 Postmitotic development of mDNs
As midbrain precursor cells exit the cell cycle (E10.5–E11.5 in mice), they migrate away
from the ventricular surface. Early phenotypic markers of mDNs, such as TH and other
enzymes in the dopamine biosynthesis pathway are induced at this time (E11.5 in the
mouse; (Lauder and Bloom 1974; Wallen et al. 1999). Subsequent maturation of mDNs
(E12–15) is characterized by the expression of common synaptic markers and the DAT.
DAT expression is relatively specific to the mDN population, unlike TH, which is also
expressed in other catecholaminergic cell types. Several transcription factors have been
implicated in the postmitotic development of mDNs (Figure 1.5). Within the postmitotic
mesencephalic cell population, expression of these factors is restricted to the early mDNs
but, none of these transcription factors (TF) appear sufficient individually to instruct the
mDN phenotype, suggesting a network model. One important TF is Nurr1 an orphan
nuclear receptor/transcription factor which is induced in postmitotic mDN precursors at
approximately E10.5, just preceding TH expression (Wallen and Perlmann 2003). Nurr1
is necessary for mDN specification as Nurr1-deficient animals fail to express early mDN
phenotypic markers including TH and cRet, a component of the Glial cell derived
neurotrophic factor (GDNF) receptor.
57
Figure 1.4 Stages of MDN development.
Initially mature stem cells are patterened to a ventral cell fate and away from a dorsal fate
(in grey). Subsequently ventral dividng mesencephalic precursors are specified towards a
MDN fate away from other fates, such as serotonergic neurons (in grey). A network of
transcription regulatory factors promotoes (blue) or inhibits (green) the process. An array
of secreted factors is implicated at each step of the developmental program (red) (adapted
from (Abeliovich and Hammond 2007)).
58
Figure 1.5 Signaling cascades in the MDN development.
Soluble factors (green) and transcription factors (blue). (A) At the dividing
mesencephalic progenitor stage, SHH induces Gli1 activation and Foxa2 induction
(which can induce SHH expression ie autocrine loop). (B) SHH also induce transcription
cascade involving activation of Lmx1a, Msx1, and Ngn2, and inhibition of Nkx6.1. Otx2
induces Ngn2 expression of Nkx2.2. Nurr1, Lmx1b, and En1/2 function in parallel to
induce aspects of the postmitotic mDN fate. Nurr1 and Pitx3 act cooperatively to induce
late markers of the mDN phenotype (adapted from (Abeliovich and Hammond 2007)).
59
1.10.4 Development of postmitotic mDNs
Several secreted factors have been implicated in postmitotic mDN development including
Wnt family members such as Wnt5a (Schulte et al. 2005) and FGF family members such
as FGF-20 (Ohmachi et al. 2003). It is unclear whether secreted factors that act at an
earlier stage in the development of mitotic precursors, such as SHH or FGF8, serve an
additional function in postmitotic mDN development. Overexpression of Nurr1 in
neuronal cell lines (Wagner et al. 1999), hippocampal neural progenitor cultures
(Sakurada et al. 1999), midbrain precursors (Kim et al. 2003), and embryonic stem cells
(Sonntag et al. 2004; Kim et al. 2006) appears to promote the generation of mDNs, but
this may reflect a broad proneural activity of Nurr1 (Kim et al. 2003). Furthermore, only
a subset of mDN markers appears to be induced by Nurr1 alone. Pitx3 overexpression in
undifferentiated ES cultures (Chung et al. 2005) or in neuron progenitor cells (Sakurada
et al. 1999) leads to either no changes or the induction of a limited set of nigral mDN
markers suggesting Pitx3 drives the expression of markers specific to the SN but not the
VTA, as VTA neurons are relatively spared in Pitx3 mutant mice (Chung et al. 2005).
These transcription factors in mDN maturation may function within a network as Nurr1
alone is sufficient to induce early mDN markers such as TH, and late events in mDN
maturation can only be induced by the combination of Nurr1 and Pitx3 (Martinat et al.
2006).
1.10.4 Functional maturation and axonal pathfinding
Functional maturation of mDNs is distinguished by axonal pathfinding, synaptogenesis,
and depolarization-induced vesicular DA release. Several studies have suggested that
60
interaction with ST target tissue promotes mDN maturation events (Perrone-Capano and
Di Porzio 2000). Newly born mDNs extend axonal and dendritic processes that must be
correctly targeted to their destinations. mDN axons project rostrally at E11 in the mouse
(Nakamura et al. 2000b) by E14.5, many of these axons have reached their targets. mDN
axonal pathfinding is likely governed by some of the same guidance cues that regulate
CNS projections (Tessier-Lavigne 2002). Expression of Neuropilin-1, a co-receptor for
the semaphorin family ligands, appears to be induced by Nurr1 (Hermanson et al. 2006),
but its role for semaphorin signaling in the guidance of mDN axons has not been clearly
defined. In addition, the homeodomain proteins En-1 and En-2, which are implicated in
mDN maturation, can induce the expression of the guidance molecule receptors ephrin-
A2 and ephrin-A5 (Logan et al. 1996; Kimura et al. 2004).
Slits and Netrins may also be involved in target recognition by nigrostriatal mDNs as
they express the Slit receptors, Robo1 and Robo2, and the Netrin receptor DCC (Lin et al.
2005). The Slit family of guidance cues has also been implicated in the guidance of
mDN projections, as mice deficient in Slit2 or both Slit1 and Slit2 show DAergic
projections that are displaced ventrally in the diencephalon (Bagri et al. 2002).
Furthermore, in the double mutants some axons aberrantly cross the ventral midline.
mDNs are repelled by Slit2-expressing cells in culture, whereas they are attracted by
Netrin-1-expressing cells, suggesting that Slit2 and Netrin-1 act in concert to guide these
axons to the lateral ganglionic eminence (LGE), the precursor to the ST (Lin et al. 2005).
61
Once the nigrostriatal mDNs reach the ST their targeting may be regulated by ephrin
signaling, as these mDNs express the EphB1 receptor, while ephrin-B2 ligand is
expressed in the ST. Cultures of SN mDNs, but not VTA mDNs, show reduced axonal
outgrowth on monolayers of ephrin-B2-expressing cells (Yue et al. 1999). Such
signaling could ensure accurate (topographical) innervation of the ST by the nigrostriatal
mDNs. Interestingly application of ephrin-B2 to midbrain neuronal cultures results in an
upregulation of Nurr1, suggesting that correct target recognition is required for the
maintenance of DAergic identity and better survival (Calo et al. 2005).
Upon target innervation, mDNs axons likely compete to establish synapses and survive.
In support of this model, a wave of apoptotic mDN cell death is observed perinatally in
rodents (Burke 2003). Growth factors, notably GDNF and BDNF, have long been
implicated in this process, as this increases the survival (Beck et al. 1995) and
arborization of mDNs (Costantini and Isacson 2000a, b). In vivo these data have been
corroborated by the observation that transplantation of GDNF-expressing cells into mice
lesioned with the mDN toxin 6-hydroxydopamine (6-OHDA) results in increased survival
of mDNs and sprouting of DAergic axons into the graft site (Akerud et al. 1999).
Notably Nurr1 mutant mice lack expression of the GDNF receptor cRet (Wallen and
Perlmann 2003), suggesting that defective GDNF signaling may partially account for the
migration and target innervation defects seen in such mice.
Finally, there is surprising functional and anatomical diversity among mDNs (Carlsson
1959; Dahlstroem and Fuxe 1964b). mDN located within the SNpc (SN or A9) form the
62
nigrostriatal pathway to the dorsolateral ST and are essential for the control of voluntary
movement; mDNs within the ventral tegmental area (VTA or A10) form the
mesolimbocortical system and controls brain mechanisms of novelty, reward, and
addiction; and mDNs within the retrorubral field (RRF or A8) are involved in both local
midbrain circuitry and the SN pathway. The guidance cues that underlie this diversity
remain to be identified. This may also relate to the apparent cell-intrinsic differences
among types of mDNs, as described above (Maxwell et al. 2005).
1.11 NEUROTROPHINS AND THE DEVELOPMENT OF mDNs
The experimental evidence on which the neurotrophic factor concept was built came
mainly from studies of the first-discovered and purified neurotrophic factor (NTF), nerve
growth factor (NGF), and its capacity to increase survival, neurite growth, and
neurotransmitter synthesis of paravertebral sympathetic neurons (Levi-Montalcini 1987).
NGF is synthesized by the target cells of these neurons, is released, binds to high affinity
binding receptors of the nerve terminals, is subsequently internalized, and is retrogradely
transported together with its receptors to the neuronal cell soma where it exerts its action
(Lewin 1996; Lewin and Barde 1996; Campenot and MacInnis 2004). NGF has been
shown to act in vitro and in vivo (Levi-Montalcini 1987; Snider 1994; Klein 1994), and
has become more clear during the last few years that cell death from Caenorhabditis
elegans to mammals is a tightly regulated process that requires both specific death-
promoting external signals and the coordination and synergies of several intracellular
signaling networks (Frade and Barde 1998; Krieglstein et al. 2000; Raoul et al. 2000;
Barker et al. 2001). This is demonstrated by glial-cell-line-derived neurotrophic factor
63
(GDNF) promoting the survival of impaired mDN more efficiently when it is co-applied
with anti-apoptotic agents (Eberhardt et al. 2000; Perrelet et al. 2002).
Five established growth factors may be considered to act as target-derived neurotrophic
factors for nigrostriatal DAergic neurons: GDNF, neurturin (NRT), brain-derived
neurotrophic factor (BDNF), neurotrophin-4 (NT-4), and fibroblast-growth factor-2
(FGF-2).
1.11.1 Glial-cell-line-derived neurotrophic factor (GDNF)
GDNF was identified and purified from a glioma cell line, B49, by its ability to promote
survival of mesencephalic DAergic neurons (Unsicker et al. 1991; Lin et al. 1993;
Airaksinen and Saarma 2002). GDNF is expressed throughout the CNS, with the highest
levels occurring in the P0 and P10 rat, in the human ST (Springer et al. 1994; Choi-
Lundberg and Bohn 1995), and at lower levels, in the SN (Schaar et al. 1993; Choi-
Lundberg and Bohn 1995) suggesting that GDNF may act both as a local and a target-
derived NTF for nigrostriatal DAergic neurons. The developmental pattern of expression
correlates with striatal target innervation and with ontogenetic cell death (Burke 2004).
The selectivity of GDNF for nigrostriatal DAergic neurons is still debatable as other
neuron populations have been shown to be responsive to GDNF (Henderson et al. 1994;
Arenas et al. 1995; Buj-Bello et al. 1995; Trupp et al. 1995; Farkas et al. 1997;
Krieglstein et al. 1998a), the relevance of and excitement for GDNF as the prime
candidate for a NTF-based Parkinson’s therapy are however still unaltered (Kordower
2003).
64
In addition to promotion of survival, GDNF has also been shown to induce neurite
growth, sprouting (Akerud et al. 1999), synaptic efficacy (Bourque and Trudeau 2000),
and TH expression (Xiao et al. 2002). GDNF has also been shown to enhance long-
lasting changes on DAergic afferents, including increased DA turnover in vivo (Hudson
et al. 1995). In animal models of PD, the effect of GDNF has a major impact on the
metabolic strengthening and increased DA synthesis of the remaining neurons (Gash et
al. 1996). However, there are also reports of aberrant effects of GDNF in certain
experimental situations. GDNF has been shown to induce aberrant sprouting and
downregulation of TH in 6-OHDA-lesioned nigrostriatal DAergic neurons (Georgievska
et al. 2002a).
1.11.2 GDNF family- ligands
GDNF is part of a family of growth factors which includes neurturin, artemin, and
persephin (Baloh et al. 2000; Saarma 2000) which share the transmembrane tyrosine
kinase receptor with its intracellular tyrosine kinase, c-Ret. Their specificity, however, is
through their individual ligand-binding receptor, which is a GPI-anchored receptor
interacting with c-Ret upon ligand binding (Baloh et al. 2000; Sariola and Meng 2003).
They all have survival-promoting activity of GDNF on MDN under many in vitro
conditions (Lin et al. 1993) such as preventing apoptosis in vitro (Burke 1998) and in
vivo (Oo et al. 2003). Administration of GDNF into the ST during the biphasic period of
ontogenetic cell death suppresses apoptosis of nigral DAergic neurons. Neutralizing
65
endogenous GDNF in the ST diminishes cell death, although only during the first period
of cell death (Kholodilov et al. 2004); Baloh, 2000 #1300}.
1.11.2.1 Neurturin
The other GDNF family ligands NRT, artemin, and persephin, are also expressed in the
nigrostriatal system. NRT is of interest as it is also expressed in the ST (Widenfalk et al.
1997; Horger et al. 1998; Akerud et al. 1999). NRT mRNA has been found in the mouse
VMB and ST during development with peaks in the MB floor at E11.5–E13.5 as DAergic
progenitors differentiate. As DAergic neurons extend their axonal projections to targeted
areas, NRT expression decreases in the SN and gradually increases in the ST with a peak
at P10. NRT expression levels are relatively low compared with those of GDNF and do
not precisely match the time course of ontogenetic cell death (Cho et al. 2004a; Cho et al.
2004b) suggesting NRT’s role in regulating ontogenetic cell death in DAergic neurons,
either as a target-derived or as a local paracrine NTF. Its ability to promote the survival
of DAergic neurons in vitro is comparable to GDNF (Horger et al. 1998; Akerud et al.
1999) but does not induce sprouting and hypertrophy of developing DAergic neurons like
GDNF. In vivo, NRT protects 6-OHDA-lesioned nigrostriatal neurons (Horger et al.
1998; Akerud et al. 1999), again does not induce TH staining and sprouting (Akerud et al.
1999). There are also regionally distinct effects of NRT in vivo beeing reported with
infusion into the lateral ventricle in rats’ leads to higher DA levels in the mediodorsal ST,
relative to the ventrolateral ST, whereas such a distinct regional distribution could not be
identified upon GDNF infusion (Hoane et al. 1999).
66
1.11.2.2 Neublastin/artemin and persephin
In addition to GDNF and NRT, neublastin/artemin (ART) and persephin (PSP) also
promote the survival of DAergic neurons in vitro and in vivo (Baloh et al. 1998;
Milbrandt et al. 1998; Rosenblad et al. 2000b). PSP is expressed in relatively high levels
in the MB floor and in the ST (Jaszai et al. 1998; Akerud et al. 2002). DAergic neurons
also express the preferred ligand-binding receptor for PSP, GFR-4 (Enokido et al. 1998;
Akerud et al. 2002). However, there are too few data available to discuss their
physiological relevance in the context of promoting DAergic neuron survival.
1.11.3 Neurotrophins
Neurotrophins represent a family of growth factors that are closely related to NGF,
namely brain-derived neurotrophic factor (BDNF), neurotrophin-3 (NT-3), and NT-4/5
(Lindsay et al. 1993; Seroogy and Gall 1993; Unsicker 1994; Lykissas et al. 2007; Skaper
2008). BDNF and NT-4 signal via the tyrosine kinase receptor trkB, and NT-3 does so
via trkC (Huang and Reichardt 2003; Segal 2003). Neurotrophins and NGF have been
shown to promote survival of MDN (Chaturvedi et al. 2006; Hirata et al. 2006; Jiang et
al. 2006). Nigral DAergic neurons express the high affinity receptors trkB and trkC
(Merlio et al. 1992; Numan and Seroogy 1999), suggesting that DAergic neurons direct
response to the corresponding trk ligands. BDNF mRNA has been detected in the ST, and
BDNF and NT-3 occur in the VMB (Hofer et al. 1990; Friedman et al. 1991; Gall et al.
1991; Seroogy et al. 1994). In contrast to other neurotrophins, the expression of NT-4 is
relatively low in the CNS (Berkemeier et al. 1991). BDNF and NT-3 have been shown to
be retrogradely transported by DAergic neurons when injected into the ST (DiStefano et
67
al. 1992; Mufson et al. 1994), whether BDNF acts as a target-derived factor for
nigrostriatal DAergic neurons is unknown; none of the mice carrying null mutations of
the neurotrophins nor their receptors provide supportive evidence (Yan et al. 1993; Yan
and Miller 1993; Klein 1994).
Chronic BDNF infusion results in an increased number of spontaneously active DAergic
neurons, showing an increased firing rate and an increased number of action potentials
within bursts (Shen et al. 1994) which may be the basis for the increased locomotor
behavior and striatal DA turnover as observed during supranigral BDNF infusions.
Similarly, the reduction of endogenous BDNF by the intranigral and intrastriatal
application of anti-sense BDNF in rats modulates nigrostriatal neurotransmission (Lau et
al. 1998), and homozygous BDNF mutations produce severe catecholamine depletion
within the nigrostriatal DAergic system, whereas heterozygous mice exhibit a slight but
significant elevation of striatal DA content (Dluzen et al. 1999). Rite et al. (Rite et al.
2003) have shown that the expression of BDNF in the SN is dependent on target integrity
but is independent of neuronal activation. Suggesting BDNF is probably not a target-
derived NTF for nigrostriatal DAergic neurons, but that it possibly acts in the SN in an
autocrine or paracrine fashion to modulate nigrostriatal function anterogradely.
1.11.4 Local factors
Several other growth factors have been shown to promote the survival of DAergic
neurons. These include fibroblast-growth factor-2 (FGF-2) (Otto and Unsicker 1990;
Timmer et al. 2007; Peng et al. 2008), transforming growth factor beta (TGF-�)
68
(Krieglstein et al. 1995b; Farkas et al. 2003; Roussa et al. 2004; Roussa and Krieglstein
2004; Roussa et al. 2006), growth/differentiation factor 5 (GDF5) (Krieglstein et al.
1995c), GDF15 (Strelau et al. 2000), bone morphogenetic proteins (BMPs) (Jordan et al.
1997), TGF-α (Alexi and Hefti 1993), heparin-binding epidermal growth factor (HB-
EGF) (Farkas and Krieglstein 2002), and sonic hedgehog (Shh) (Miao et al. 1997). FGF-
2, BMPs, and HB-EGF survival-promoting capacity may be indirect and is possibly
mediated via astrogial cells and represent a relevant endogenous source for
dopaminotrophic growth factors (Nakayama et al. 2003). Their complete spectrum of
survival-promoting molecules has not been analyzed (Schaar et al. 1994; Engele and
Franke 1996; Engele et al. 1996; Engele and Schilling 1996).
1.11.4.1 TGF-�
This is a multifunctional growth factor (Bottner et al. 2000; Unsicker 2000; Unsicker and
Strelau 2000). TGF-�2 and TGF-�3 are expressed in the developing and adult SN and in
the ST (Flanders et al. 1991; Unsicker et al. 1991). In vivo, TGF-�’s role as endogenous
promoter of DAergic neuron survival has been shown (Farkas et al. 2003; Roussa et al.
2006) and is supported with GDNF (Krieglstein et al. 1998b), Shh and FGF-8 (Roussa
and Krieglstein 2004). Similar to BDNF, TGF-� acts directly on DAergic neurons, as
demonstrated by TGF-� dependent translocation to the nucleus of TH+ve neurons, but
may act additionally on other mesencephalic cells (Hyman et al. 1994; Roussa et al.
2004).
1.11.5 Low molecular weight compounds
69
The delivery of proteins into the brain is a problem (Fricker and Miller 2004; Pan and
Kastin 2004) thus, there is an ongoing search to find small compounds that either mimic
neurotrophic factors, enhance neurotrophic effects, or induce neurotrophic factors in the
substantia nigra. Several low molecular weight compounds have been indentified that
increases the production of BDNF in nigral DAergic neurons: salicylic acid, cGMP
analogs, okadaic acid, IBMX, dipyridamole and glutamate (Chun et al. 2000) AMPA
receptors have been found to serve as potential targets as their stimulation increases
neuronal activation and activity-dependent signalling (Zafra et al. 1990). A potent and
selective AMPA receptor potentiator, LY404187 protects against unilateral infusion of 6-
OHDA into the SN or ST of rats (ONeill et al. 2004). The protective effect may be
attributable to the increased expression of BDNF in DAergic neurons in vivo (O'Neill et
al. 2004), as these compounds have previously demonstrated their capacity to increase
BDNF expression in primary neuron culture (Legutko et al. 2001).
An alternative approach to enhance effects of endogenous BDNF is to block
dephosphorylation of the activated tyrosine kinase receptor trkB. Inhibition of protein
tyrosine phosphatases has been shown to protect axotomized nigral DAergic neurons in
vivo (Lu et al. 2002). Application of an ineffective dose of BDNF synergistically
increases the protective capacity of such nhibitors, suggests that tyrosine phosphatase
inhibition prevents neuronal death by enhancing neurotrophic signalling.
1.11.6 Immunophilin ligands
70
The immunophilin ligands has been introduced as potential neurotrophic molecules (Guo
et al. 2001a; Guo et al. 2001b) acting via their protein receptors and are known
immunosuppressants. The first immunophilin ligand GPI-1046 came of interest due to its
neurotrophic activities (Steiner et al. 1997). GPI-1046 bound to FK506-binding protein-
12 induces neurite outgrowth in vitro similarly to NGF, and regenerative sprouting from
spared nigrostriatal neurons following MPTP or 6-OHDA lesioning in rats. Systemically,
GPI-1046 in MPTP treated rhesus monkeys failed to produce neuroprotective effects
(Emborg et al. 2001). As the immunosuppressant effect of immunophilin ligands makes
them unsuitable for neurological application, nonimmunosuppressant immunophilin
ligands have been introduced. Novel compounds include V-10,367 (Costantini et al.
1998); and V-13,661 (Costantini et al. 2001) (Constantini et al. 2001). Both ligands
prevent loss of striatal DA innervation in the MPTP mouse model following oral
administration. However, only orally applied V-10,367, which binds to FKBP12 protects
against a slow and progressive intrastriatal 6-OHDA lesion.
1.12 CHANGES IN CYTOKINE LEVELS, DECLINE OF ESSENTIAL NTFs AND
LINKS WITH PD
1.12.1 Human evidence
Degeneration of the DAergic neurons of the SNpc and the resulting loss of nerve
terminals accompanied by DA deficiency in the ST are responsible for most of the
movement disturbances observed in PD. One hypothesis of the cause of degeneration of
the nigrostriatal DA neurons is that PD is caused by programmed cell death (apoptosis)
due to increased levels of cytokines and/or decreased ones of neurotrophins. There is a
71
marked increase in levels of cytokines (Nagatsu et al. 2000a; Nagatsu and Sawada 2005;
Sawada et al. 2006), such as tumor necrosis factor (TNF)-alpha, interleukin (IL)-1beta,
IL-2, IL-4, IL-6, transforming growth factor (TFG)-alpha, TGF-beta1, and TGF-beta2,
and decreased levels of neurotrophins (Siegel and Chauhan 2000; Chauhan et al. 2001),
such as brain-derived neurotrophic factor (BDNF) and nerve growth factor (NGF), in the
nigrostriatal DA regions and ventricular and lumbar cerebrospinal fluid of PD patients.
Furthermore, the levels of TNF-alpha receptor R1 (TNF-R1, p55), bcl-2, soluble Fas
(sFas), and the activities of caspase-1 and caspase-3 were also elevated in the nigrostriatal
DA regions in PD (Dragunow et al. 1997; Tatton et al. 2003; Nagatsu and Sawada 2007).
1.12.2 Evidence from experimental PD models
In experimental animal models of PD, IL-1 beta level is increased and NGF decreased in
the ST of 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP)-induced parkinsonian
mice, and TNF-alpha level increased in the SN and ST of the 6-hydroxydopamine (6-
OHDA)-injected side of hemiparkinsonian rats (Grunblatt et al. 2000a, b). L-DOPA
alone or together with 6-OHDA does not increase the level of TNF-alpha in the brain in
vivo (Mogi et al. 1999). Increased levels of proinflammatory cytokines, cytokine
receptors and caspase activities, and reduced levels of neurotrophins in the nigrostriatal
region in PD patients, and in MPTP and 6-OHDA-produced parkinsonian animals
suggest increased immune reactivity and programmed cell death (apoptosis) of neuronal
and/or glial cells (Nagatsu et al. 2000a, b). These data indicate the presence of such
proapoptotic environment in the SN in PD that may induce increased vulnerability of
neuronal or glial cells towards a variety of neurotoxic factors. The probable causative
linkage among the increased levels of proinflammatory cytokines and the decreased
72
levels of neurotrophins, candidate parkinsonism-producing neurotoxins such as
isoquinoline neurotoxins (Nagatsu 1997), and the genetic susceptibility to toxic factors,
remains for further investigation in the molecular mechanism of PD. The increased
cytokine levels, decreased neurotrophin ones, and the immune response in the
nigrostriatal region in PD indicate new neuroprotective therapy such as nonsteroidal anti-
inflammatory drugs (NSAIDs) such as aspirin, immunosuppressive or immunophilin-
binding drugs such as FK-506, and drugs increasing neurotrophins (Gold and Nutt 2002;
McGeer and McGeer 2002; Tansey et al. 2008).
73
CHAPTER 2:
IN VIVO PARKINSONIAN MODELS: MPTP & ROTENONE
74
2.0 INTRODUCTION
As discussed in chapter 1 the etiology of PD is not completely understood, but it is likely
to involve both genetic and environmental factors (Sherer et al. 2002a; Allam et al. 2005).
Epidemiological studies suggest that exposure to environmental agents containing
neurotoxins such as pesticides, may increase PD risk (Gorell et al. 1998; Menegon et al.
1998). Mitochondrial dysfunction has also been linked to PD. Specifically; there are
systemic reductions in the activity of complex I of the mitochondrial electron transfer
chain (ETC) in PD brain, muscle, and platelets (Mizuno et al. 1989; Parker et al. 1989;
Schapira et al. 1989; Cardellach et al. 1993; Haas et al. 1995). Additional evidence for
mitochondrial impairment in PD comes from the finding that MPP+ (1-methyl-4-phenyl-
2,3-dihydropyridinium), the active metabolite of the parkinsonism toxin N-methyl-4-
phenyl-1,2,3,6-tetrahydropyridine (MPTP), acts as a complex I inhibitor (Nicklas et al.
1985). There are three major toxin models of PD: 6-hydroxydopamine (6-OHDA),
MPTP, and rotenone (see Table 2.1), in which there is DAergic neuronal cell death
associated with Parkinsonism. Since two of these well known selective DAergic cell
neurotoxins are employed in the experiments discussed in this thesis, they are reviewed in
more depth in this chapter.
2.1 MPTP
In the early 1980s the DAergic neurotoxin 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine
(MPTP) a meperidine analogue 1-methyl-phenyl-propion-oxypiperidine (MPPP) was
discovered by accident during synthetic heroin production (Davis et al. 1979; Langston
et al. 1983). It is of interest to note the biochemical, pathological and clinical features
75
induced in these young addicts corresponded to the hallmarks of PD. A follow up of this
study presented evidence that MPTP induced severe and unremitting Parkinsonism
caused by an active nerve cell degeneration with characteristic features of
neuroinflammation (Langston et al. 1999)
Table 2.1 Characteristics of animal models of Parkinson’s disease
Model
Gradual cell loss of DA neurons in adulthood
Easily detectable motor deficits
Development of Lewy bodies
Short timecourse
6-OHDA
No
Yes (quantifiable rotation deficit)
No
Yes
Rotenone Yes, but variable Individual susceptibility
Yes Yes Yes
Acute MPP+ No Yes No Yes Drosophila α-synuclein overexpression
Yes Yes Yes Yes
Mouse α-synucelin overexpression
Yes, but not in SN Yes Nuclear as well as cytoplasmic inclusions
No (1 year)
* Chronic MPTP administration induces slow Parkinsonian syndrome and might produce
Lewy bodies (Adapted from (Beal 2001)).
2.2 BIOCHEMISTRY OF MPTP
MPTP itself is not toxic in the brain but requires a two–step biotransformation process to
form the metabolite MPP+ which is the ultimate toxin (See Figure 2.1). The lipophilic
MPTP rapidly enters the blood brain barrier after systemic administration where MPTP is
converted via MAO-B to form the 1-methyl-4-phenyl-1,2-dihydroxypyridinium ion
(MPDP+) (Markey et al. 1984; Chiba et al. 1985). MAO-B is mainly located in
76
astrocytes (Takada et al. 1990; Di Monte et al. 1991) but not in DAergic neurons
(Westlund et al. 1985). MPDP+ is spontaneously oxidized to MPP+. Theoretically both
MAO-A and MAO-B isoforms are able to convert MPTP but MAO-B is the likely key
enzyme as inhibition of MPTP metabolism by MAO-B inhibitors, but not MAO-A
inhibitors, was effective in preventing MPTP neurotoxicity (Heikkila et al. 1984a;
Nagatsu and Sawada 2006). MPP+ is released by glial cells into the extracellular space
by an active mechanism using the extracellular monoamine transporter (Russ et al. 1996).
Subsequently, MPP+ can be specifically taken up into DAergic nerve terminals via
plasma membrane DA transporter (DAT) (Chiba et al. 1985; Javitch et al. 1985). It has
been demonstrated that mice with a deficiency of the DAT are resistant against MPTP
toxicity (Gainetdinov et al. 1997; Takahashi et al. 1997) thus overexpression of DAT may
result in enhanced MPTP neurotoxicity. MPP+ is then taken up by then intraneuronal
vesicular monoamine transporters (VMATs) and sequestrated into synaptosomal vesicles
or accumulated in mitochondria via energy-driven uptake (Singer 1987). Intravesicular
storage of MPP+ via VMAT suppressed its toxicity in vitro (Liu 1992). This was
confirmed in vivo using VMAT-2 deficient mice which demonstrated an increased MPTP
toxicity (Takahashi et al. 1997; Gainetdinov et al. 1998).
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Figure 2.1 Schematic Representation of MPTP Metabolism.
After systemic administration, MPTP crosses the blood-brain barrier. Once in the brain,
MPTP is converted to MPDP+ by MAO-B within nondopaminergic cells, such as glial
cells and serotonergic neurons (not shown), and then to MPP+ by an unknown mechanism
(?). Thereafter, MPP+ is released, again by an unknown mechanism (?), into the
extracellular space. MPP+ is concentrated into dopaminergic neurons via the dopamine
transporter (DAT) (Dauer and Przedborski 2003).
78
2.3 ANIMAL MODELS OF PD USING MPTP
2.3.1 Factors influencing the neurotoxic action of MPTP
Models of MPTP have been widely used and investigated (Heikkila and Sonsalla 1987;
Gerlach et al. 1991; Heikkila and Sonsalla 1992; Tipton and Singer 1993). Neurological
effects following systemic application of MPTP have been found in a variety of animals,
including monkeys, mice, dogs, cats, sheep and goldfish (Heikkila and Sonsalla 1987;
Zigmond and Stricker 1989; Gerlach et al. 1991; Heikkila and Sonsalla 1992; Tipton and
Singer 1993). There are marked differences with regard to sensitivity to the neurotoxic
action of MPTP (Table 2.2). Even large doses of MPTP elicit only slight neurotoxic
effects in rats and guineapigs. To produce a DA loss similar to that observed and
elsewhere in the monkeys in even the most sensitive strain of mice, the C57/Black
mouse, a 50-fold dose of MPTP is required. The reasons for the differential sensitivities
are still not completely understood, however the pharmacokinetics of MPTP, and the
distribution and excretion rate of its main metabolite MPP+, may be responsible. Other
factors that may influence these species differences include neuromelanin, distribution
and localisation of MAO-subtypes in the brain, DA metabolism and the anti-oxidant
content of the nigrostriatal system.
2.3.2 Behavioural changes
Monkeys treated with MPTP develop motor disturbances comparable to those in man
(Stern 1990; Gerlach et al. 1991). The most prominent of these are akinesia and rigidity;
resting tremor is only seen in isolated instances. The precise appearance of these
neurologic abnormalities varies depending on the age, species and dosage. MPTP-
79
syndromes have been described in rhesus monkeys ((Burns et al. 1983; Smith et al.
1993), squirrel monkeys (Irwin et al. 1990), macaques (Crossman et al. 1989), baboons
(Hantraye et al. 1993) and marmosets (Ueki et al. 1989; Russ et al. 1991). The
quantitation of the neurologic abnormalities in monkeys is achieved using the modified
PD scales.
Table 2.2 Relative toxicity of MPTP in different animals
Animal Family Cumulative dose Dopamine concentration
(mg/kg) (% of normal values)
Rodents
- Rat
Sprague Dawley 151 77 (Caudate nucleus)
- Guineapigs 105 50 (Striatum)
- Mouse
C57/Black 90 24 (Caudate nucleus)
CF/1 80 60 (Striatum)
Swiss-Webster 410 35 (Striatum)
Non-Human primates
- Common marmoset 6.9-9.2 15 (Caudate nucleus)
- Rhesus monkey 1.5 3 (Striatum)
2.1-6.5 0.4 (Caudate nucleus)
0.5 (Putamen)
- Squirrel monkey 2 30 (Caudate nucleus)
15 (Putamen)
See (Gerlach and Riederer 1996)
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MPTP-treated mice show only transient behavioral changes as an initial short-term toxic
effect (Sundstrom et al. 1990; Zuddas et al. 1992b). They exhibit hypersalivation,
piloerection, seizures and hypokinesia (Weihmuller et al. 1989; Zuddas et al. 1992b) and
recover within 24–48h, although decreased locomotor activity (-66%) has been described.
Moderate DA depletion does not produce overt motor symptoms, but performance in
spatial working memory tasks was impaired when more difficult tasks were applied
(Tanila et al. 1998). Long-term effects as a result of dopamine denervation including
including postural abnormalities, bradykinesia and tremor were observed by some groups
(Teismann et al. 2001). The mode of MPTP administration and the strain of animals used
may lead to apparently different observations (Schmidt and Ferger 2001).
2.3.3 Histopathological changes
A selective destruction of SN neurons has been described in MPTP-treated non-human
primates (Burns et al. 1983; Ueki et al. 1989; Mavridis et al. 1991; Bernocchi et al. 1993;
Hantraye et al. 1993). However, detailed examination shows that other regions of the
brain are also affected (Mitchell et al. 1985; German et al. 1988; Gibb et al. 1989). This
mainly occurs in older animals. A loss of neurons is also found in the locus ceruleus
(Mitchell et al. 1985; Forno et al. 1993). In younger animals, losses of TH-ir neurons
occur in the VTA and hypothalamus (German et al. 1988); quantitative evaluation
revealed that MPTP-treated macaques observed cell loss between 46-93% in the SNpc
leading to a stable Parkinsonian syndrome with more than 99% loss of DA in the ST
compared to the 28-57% in VTA. Additionally, a decreased density of DA receptors was
found after MPTP treatment (Mitsumoto et al. 1998). MPP+ acutely displaces DA from
81
synaptosomal vesicles and gives rise to DA-induced hydroxyl radical formation (Chiueh
et al. 1992a; Chiueh et al. 1992b). Using the reverse microdialysis a 60-fold increase of
DA release after striatal application of MPP+, and significantly increased hydroxyl
radical formation was observed (Obata et al. 1992; Ferger et al. 1998b; Ferger et al.
1998a; Obata et al. 2001).
The presence of LBs has not been established in MPTP-treated non-human primates. In
old monkeys eosinophil inclusion-bodies have been found in the same areas of where
LBs are found in humans (Forno et al. 1993). In MPTP-treated C57/Black mice an
average 40% loss of TH-ir neurons in the SN and 17% loss in the VTA are observed
(Date et al. 1990b; Kupsch et al. 1992). This degree of damage led to a substantial
depletion (-85%) of DA in the ST. This pattern of neuronal loss was further confirmed
by Nissl-staining methods (Seniuk et al. 1990) and showed that the locus ceruleus and the
hypothalamus are also affected.
2.3.4 Neurochemical changes
MPTP produces a series of neurochemical changes in primates and rodents. Decreased
concentration of DA and it metabolites, diminished TH activity and fewer DA-uptake
sites in the ST and GP indicate damage to the nigrostriatal DAergic system. MPTP
treatment of monkeys’ results in striatal DA levels being reduced, followed by the
reduction of DOPAC and HVA levels, whereas the noradrenergic and serotonergic
system were less affected (Ueki et al. 1989; Pifl et al. 1991; Russ et al. 1991). In
contrast, noradrenaline, serotonin and neuropeptides are less affected by MPTP.
82
Concentrations of enkephalins, cholecystokinin, substance P and neurotensin are
unchanged in the BG of MPTP-treated marmosets (Taylor et al. 1991).
In marmosets (Russ et al. 1991) and squirrel monkeys (Moratalla et al. 1992) the putamen
is more severely affected than the caudate nucleus, but in the macaque (Alexander et al.
1991) the opposite was found. This can be explained by the supersensitivity of the post-
synaptic DA D2-receptor (Alexander et al. 1991). The essential features of the
neurochemical effects observed in primates with MPTP-treatment, such as depletion of
DA in the nigrostriatal and mesolimbic systems and depletion of adrenaline are also
demonstrated in various strains of mice (Heikkila et al. 1989; Weihmuller et al. 1989;
Date et al. 1990a; Seniuk et al. 1990; Sundstrom et al. 1990; Gerlach et al. 1993).
2.3.5 Invertebrates
Studies have shown that MPTP toxicity is lower in animals such as frogs and leeches.
The frog Rana pipiens, showed a decrease in DA content and motor dysfunction with
MPTP and MPP+ (Barbeau et al. 1985a; Barbeau et al. 1985b). However, MPP+ was
more toxic and effective at lower doses than MPTP in frogs. The medicinal leech
(Hirudo medicinalis), also showed decreased DA content and motor dysfunction with
MPTP, but Macrobdella decora showed no MPTP sensitivity (Langston and Irwin 1986;
Kopin and Schoenberg 1988). The planaria (Dugesia japonica) is the most primitive
species to have a centralized nervous system (Kitamura et al. 1998) and is considered an
ancestor of the mammalian brains (Sarnat and Netsky 1985). This flatworm contains
neurotransmitters such as DA, NA and 5-HT. Treatment with L-dopa or reserpine
83
induces an increase or decrease DA content respectively, and DA agonists or antagonist
influence the planarias behaviour (Sarnat and Netsky 1985). Not only does it have high
capacity for regeneration and reorganization (Agata and Watanabe 1999; Kato et al.
1999; Kobayashi et al. 1999), MPTP induced degeneration is also completely rescued by
anti-parkinsonian drugs such as talipexole and pramipexole (Kitamura et al. 1998).
2.3.6 Genetic mouse models and MPTP
Several gene polymorphsims have been identified which contribute to the development of
PD in humans (See Table 2.3). This includes the different mutations in the α-synuclein
gene (Polymeropoulos et al. 1997; Kruger et al. 1998; Zarranz et al. 2004). Mice with a
targeted deletion of the α-synuclein gene are viable and develop normally (Abeliovich et
al. 2000). Although they show normal brain architecture and DAergic neurons, they
display a reduction in ST DA content and attenuated DA-dependent locomotor response
following amphetamine treatment. In addition, the reserve pool of neurotransmitter
vesicles in the hippocampus was reduced (Cabin et al. 2002).
2.4 MECHANISMS OF MPTP NEUROTOXICITY
2.4.1 Mitochondrial impairment
Within mitochondria, MPP+ acts by inhibiting the electron transport system of the
mitochondrial complex I (NADPH-ubiquinone oxidoreductase I; see Figure 2.2) (Nicklas
et al. 1985; Ramsay et al. 1991b). This leads to impairment in ATP production, to an
elevation of the intracellular calcium concentration and to the generation of free radicals,
84
resulting in cellular energy failure (Nicklas et al. 1987) and in the formation of
superoxide anions (Dawson 2000).
Table 2.3 MPTP-induced effects upon the DAergic system in different
mouse strains
Mouse lines Neurodegeneration ST denervation
in the SNpc
Normal mice Yes Yes
α-synuclein KO mice No No
α-synuclein (A30P) transgenic mice > >
α-synuclein (A53T) transgenic mice like in normal mice like in normal mice
α-synuclein overexpressing mice > >
DJ-1 KO mice > >
> as compared to normal mice, KO knockout mice, see (von Bohlen Und Halbach
2005; von Bohlen und Halbach et al. 2005).
There is a correlation between the time course of ATP changes with MPP+ brain levels
(Chan et al. 1991, 1992). The selective toxicity of MPP+ to DAergic neurons derives, at
least in part, from its high affinity for the dopamine transporter (Javitch and Snyder 1984;
Javitch et al. 1984). In line with this, mice lacking this transporter are protected from
MPTP toxicity (Bezard et al. 1999). MPP+ is also sequestered into synaptic vesicles by
the vesicular monoamine transporter (VMAT), and sequestration into vesicles decreases
85
MPP+ toxicity by preventing its interaction with mitochondria (Reinhard et al. 1987; Liu
et al. 1992). Thus, mice with a 50% depletion of VMAT2 show increased vulnerability
to MPTP (Takahashi et al. 1997).
2.4.2 Energy Failure
MPP+ inhibits complex I and impairs ATP formation which in turn disables or reduces
energy-dependent processes such as maintenance of calcium homeostasis, cellular
membrane potential, and ion- and transmitter transport (Di Monte, 1991; Royland &
Langston, 1998). The energy failure due to MPTP-induced ATP depletion is aggravated
by secondary steps due to energy consuming repair processes. In particular enzymes
such as poly (ADP-ribose) polymerase (PARP) which require ATP for DNA repair are
critically involved in MPTP toxicity. This is supported by PARP inhibitors or PARP KO
mice showing protection against MPTP toxicity (Cosi et al. 1996). Additionally MPP+ is
thought to also inhibit the α-ketoglutarate dehydrogenase of the tricarboxyclic acid cycle
(complex II of the respiratory chain) (Mizuno et al. 1987b). This mechanism acts
synergistically enhancing the MPTP-induced disruption of cellular energy metabolism.
2.4.3 Calcium homeostasis
As a consequence of severe energy impairment there is decreased activity of dependent
calcium-ATPase which leads to intraneuronal calcium-overload. Elevated intracellular
levels of calcium have been shown to activate degradative enzymes such as proteases and
phosphatases. These results in cell membrane and cytoskeleton degradation, disruption
of cell function, loss of cell membrane potential and ultimately neuronal death. Binding
86
of excess calcium and calcium channel blockers have shown protection against MPTP-
induced nigral degeneration (German et al. 1992; Kupsch et al. 1995; Kupsch et al.
1996).
2.4.4 Glutamate release
Excessive MPP+ concentrations may promote excitotoxicity since an enhanced glutamate
release has been shown (Carboni et al. 1990b; Carboni et al. 1990a). Glutamate has been
shown to be neurotoxic despite having a physiological role as the most abundant
excitatory amino acid. Excessive glutamate release activates NMDA receptors leading to
a massive influx of calcium thereby inducing the formation of reactive oxygen species
(ROS) as well as a reducing intracellular glutathione synthesis (Murphy et al. 1989; Beal
et al. 1993).
2.4.5 Reactive oxygen (ROS) and nitrogen species (NOS)
ROS are continuously formed in the body as byproducts from numerous biochemical
reactions. Compared to other brain regions SNpc is exposed to higher levels of oxidative
stress (OS) caused by the catabolism of DA via MAO-B mediated deamination, DA
autoxidation and high levels of iron all result in a high degree of ROS formation (Coyle
and Puttfarcken 1993). In addition, low levels of glutathione peroxidise diminish the
ability of the SNpc to cope with OS (Sian et al. 1994b; Sian et al. 1994a). MPTP
87
Figure 2.2 Schematic representation of MPP+ intracellular pathways.
Inside dopaminergic neurons, MPP+ can follow one of three routes: (1) concentration into
mitochondria through an active process (toxic); (2) interaction with cytosolic enzymes
(toxic); (3) sequestration into synaptic vesicles via the vesicular monoamine transporters
(VMAT; protective). Within the mitochondria, MPP+ blocks complex I (X), which
interrupts the transfer of electrons from complex I to ubiquinone (Q). This perturbation
enhances the production of reactive oxygen species (not shown) and decreases the
synthesis of ATP (Dauer and Przedborski 2003).
X
88
administration increases DA turnover (Teismann et al. 2001) resembling the enhanced
DA turnover in PD. In early stages of PD the remaining DAergic neurons try to
compensate for the reduced number of DA neurons by producing more DA (Fahn and
Cohen 1992).
Transgenic mice which overexpressed the superoxide detoxifying enzyme superoxide
dismutase (SOD) showed significant protection against MPTP toxicity (Przedborski et al.
1992). Nitric oxide (NO) also seems to be directly and indirectly involved in MPTP
neurotoxicity (Castagnoli et al. 1997; Di Monte et al. 1997). The reaction of NO with
superoxide results in the formation of peroxynitrite (Beckman et al. 1990), which is
neurotoxic by oxidation and nitration of biomolecules (Beckman 1996; Beckman and
Koppenol 1996; Beckman et al. 1996). It has been shown that peroxynitrite inhibits
complex I, II and III of the mitochondrial respiratory chain and irreversibly enhances
energy depletion (Radi et al. 1991). NO synthase inhibitors and mice with a deficiency in
NO synthesizing enzymes showed protection against MPTP toxicity (Schulz et al. 1995c;
Schulz et al. 1995b; Hantraye et al. 1996; Przedborski et al. 1996; Liberatore et al. 1999).
This has been confirmed by studies with 3-nitrotyrosine, which is a marker for nitration
reactions and has been found to be elevated after MPTP administration (Pennathur et al.
1999).
2.4.6 Cytokines and inflammatory processes
ROS and NO production are enhanced by microglia infiltration and activation as well
inflammatory reactions in the MPTP mouse model (Czlonkowska et al. 1996;
Kurkowska-Jastrzebska et al. 1999a, b). Reactive microglia release cytotoxic species like
89
hydroxyl radicals, NO, glutamate and cytokines such as interleukin (IL)-1β, IL-6 and
tumor necrosis factor-α (TNF-α) (Grunblatt et al. 2000a, b). MPTP-treated mice showed
elevated levels of proinflammatory cytokines (Mogi et al. 1998; Kaku et al. 1999) which
are similar to the post mortem analysis of PD brains (Mogi et al. 1994b; Mogi et al.
1994a). Furthermore, humans who ingested MPTP up to 14 years later still showed
evidence of chronic inflammation (Langston et al. 1999). Anti-inflammatory drugs such
as meloxicam and acetlysalicyclic acid showed a pronounced neuroprotection against
MPTP toxicity (Ferger et al. 1999; Teismann and Ferger 2001).
Further support for the implication of microglia in the loss of DAergic neurons in PD is
the increased number of major histocompatibility complex class II-positive microglial
cells in the SNpc of patients who developed PD (McGeer et al. 1988) or suffered from
parkinsonism due to accidental administration of MPTP (Langston et al. 1999). The
same observation was also made in the SNpc of MPTP-treated mice (Liberatore et al.
1999) and monkeys (Hurley et al. 2003; McGeer et al. 2003; Barcia et al. 2004),
suggesting that microglial cells might be involved in a putative antigen-specific immune
response. A marked increase in cytokine levels such as the pro-inflammatory cytokines
TNF-�, and IL-1� has been observed in the SNpc of PD patients compared to control
subjects (Hunot et al. 1999). Anti-inflammatory drugs like pioglitazone and minocycline
(Du et al. 2001; Breidert et al. 2002; Wu et al. 2002), as well as gene deletions targeting
pro-inflammatory molecules such as iNOS and cyclooxygenase type-2 (Liberatore et al.
1999; Dehmer et al. 2000; Teismann et al. 2003b; Teismann et al. 2003a; Hunot et al.
2004) have been shown to protect the nigral DA neurones in animal models of PD.
90
2.4.7 MPTP and apoptosis
In PD, estimations of apoptotic cells by terminal deoxynucleotidyl transferase-mediated
2'-deoxy-uridine-5'-triphosphate nick end labelling (TUNEL)-positive DAergic neurons
vary between 2 and 12% with great interindividual variability (Mochizuki et al. 1997;
Tompkins et al. 1997; Kingsbury et al. 1998; Tatton 2000). Additionally, apoptotic cell
profiles have been demonstrated ultrastructurally (Anglade et al. 1997). In other small
series, no neuronal TUNEL reactivity was observed (Kosel et al. 1997; Banati et al. 1998;
Wullner et al. 1999; Jellinger 2000). Sometimes, solely apoptotic glial profiles were
noted in SN (Kosel et al. 1997). Considering the slow evolvement of the disease, it is to
be expected that just a small percentage of cells will show signs of active cell death at any
given time-point (Blum et al. 2001). The contradictory results regarding apoptosis may be
due to differences in patient age, disease duration, postmortem time, and experimental
protocol.
TUNEL staining cannot be considered entirely specific for apoptotic cell death as DNA
strand breaks might also occur randomly in necrotic cell death (Andersen 2001) and may
reflect deficient DNA repair (Jellinger 2000). Thus, TUNEL staining should be
complemented by analysis of morphological markers of apoptosis (Burke 1998).
Currently, it is unknown how long apoptotic profiles may persist in vivo and this is an
area of ongoing research (Banati et al. 1998; Hirsch et al. 1999). Less well-known is the
contribution of autophagic cell death to Parkinson's disease, characterized by prominent
early cytoplasmic changes and vacuolization.
91
In vivo, apoptotic nuclei were detected in mice undergoing a 5-day treatment with MPTP,
peaking on day 1 after the final injection (Tatton and Kish 1997), whereas this could not
be demonstrated after a 1-day regimen of MPTP exposure (Jackson-Lewis et al. 1995).
This may be related to the degree of ATP depletion, as apoptosis might shift to necrosis
under conditions of extreme lack of energy substrates (Leist et al. 1997). Thus, only with
prolonged cellular damage more in accordance with the human situation may apoptotic
cell death be observed.
Another approach to showing apoptotic cell death relies on the demonstration of caspase
cleavage (Figure 2.3). Caspases are a family of cysteine proteases with substrate
specificity for aspartic acid. Currently 14 members are known. Ced-3 is the structural
homolog in Caenorhabditis elegans, testifying to their marked conservation through
phylogeny. Each caspase is comprised of two large and two small subunits. Activation
requires a caspase-dependent cleavage of a precursor protein into a prodomain, and a
large and a small subunit, the latter two then forming the heterotetramer. According to
their differential substrate specificities, they may be divided into three major groups,
which also provide insight into their biological roles in inflammation and apoptosis
(Thornberry et al. 1997; Thornberry and Lazebnik 1998; Nicholson 1999).
Caspase-3 activation has been demonstrated in vivo after MPTP treatment (Eberhardt et
al. 2000; Hartmann et al. 2000) (Viswanath et al. 2001). Activated caspase-3 has also
been demonstrated in SNpc of PD (Hartmann et al. 2000; Tatton 2000), while this has
been refuted by others (Jellinger 2000). Interestingly, caspase-mediated cleavage of both
92
wild-type and mutant parkin has been demonstrated (Kahns et al. 2002). The functional
significance of this finding in relation to pathogenesis is presently unknown. Using
electron microscopy, either apoptotic (Sheehan et al. 1997) or early necrotic and late
apoptotic changes (Mukherjee et al. 1997) were observed after MPP+ in cell culture.
Unfortunately, it is not known from the few pathological reports on patients with MPTP-
related Parkinsonism whether apoptotic markers were present, although an ongoing
neuronal degeneration was assumed (Langston et al. 1999).
2.5 DA AGONISTS AND MPTP
From a therapeutic perspective both humans and monkeys who received MPTP respond
very well to anti-PD treatments such as L-DOPA/carbidopa. However, as in PD patients
(Kostic et al. 1991), long-term treatment with L-DOPA leads to hyperkinetic motor
complications called dyskinesia, which can be as disabling as the parkinsonian symptoms
themselves. For instance, among the seven indexed MPTP human cases, five developed
dyskinesia within the first year of treatment with L- DOPA/carbidopa (Langston and
Ballard 1984). As of yet, the occurrence of L-DOPA-induced motor complications
remains a major impediment to the proper management of PD patients. The MPTP
monkey model has emerged as an invaluable tool to investigate the molecular basis of
these drug-induced abnormal movements and to test therapeutic strategies to control these
(Blanchet et al. 2004). This is elegantly illustrated in the study by Bézard (Bezard et al.
2003) in which the administration of a D3-dopamine partial agonist markedly improved
L-DOPA-induced dyskinesia in MPTP monkeys, without exacerbating the parkinsonian
symptoms.
93
Figure 2.3 Pathways implicated in MPTP-mediated toxicity.
Two major pathways leading to caspase activation have been characterized (Kaufmann
and Hengartner, 2001). One involves the activation of death receptors on the cell surface,
e.g. Fas/CD95 or TNF�. Both share a common motif, the death domain, necessary for
the binding of cytoplasmic adaptor molecules, e.g. FADD or TRADD. Through
involvement of a common death effector domain, the prodomain of procaspase-8 is
bound, bringing about the autocatalytic activation of the enzyme. This, in turn, leads to
the activation of downstream effector caspases, directly or via a mitochondrial pathway
as an amplification step. The mitochondrial pathway involves the release of cytochrome
c from mitochondria, forming a ternary complex with procaspase-9 and with the
scaffolding protein Apaf-1 and procaspase-9. Alongside with cytochrome c, other
proapoptotic effectors, such as Smac/Diablo or apoptosis-inducing factor (AIF), are
released from mitochondria into the cytosol. In the ternary complex, the so-called
apoptosome, caspase-9 activation can take place. Again, in further steps effector
caspases, such as caspase-3, are activated and are responsible for the cleavage of at least
200 protein substrates involved in the degenerative process (Eberhardt and Schulz 2003).
94
95
2.6 ROTENONE
2.6.1 INTRODUCTION
Several epidemiological studies suggest that pesticides and other environmental toxins
may be involved in the pathogenesis of PD (Beal 2001). Rotenone, an agricultural toxin
used as an insecticide and fish poison, produces Parkinsonism in rats when administered
intravenously (Betarbet et al. 2000). Rotenone is a mitochondrial complex I inhibitor, but
differs from MPTP in that rotenone acts uniformly throughout the brain. This model is
unique as the rats develop progressive degeneration of nigrostriatal DAergic neurons and
inclusions that stain with antibodies against ubiquitin and �-synuclein. These inclusions
contain a dense core surrounded by fibrils, similar to LBs, supporting the link between
dysfunction in complex I with alterations in �-synuclein. Furthermore, the rats have
clinical signs suggestive of PD including bradykinesia, postural instability, unsteady gait,
and tremor that respond to the DA agonist apomorphine. Although this model satisfies
most of the criteria for an excellent model of PD, its utility may be limited since there is a
high mortality rate, and less then 50% of surviving rats treated with rotenone develop
consistent lesions.
2.6.2 Biochemistry of rotenone
Rotenone is widely used around the world as an insecticide and pesticide. Rotenone
readily breaks down by exposure to sunlight with most of the toxicity of the compound
lost in 2–3 days. It is also rapidly broken down in soil and in water with the half-life in
both of these environments being between 1 and 3 days (Caboni et al. 2004). Because of
its short half-life and because it does not readily leach from soil, it is not considered to be
96
a groundwater pollutant. Consequently, the likelihood of PD being caused by
environmental exposure to rotenone is low; however epidemiological data show that the
risk of PD increases with exposure to pesticides (Butterfield et al. 1993; Gorell et al.
1998). The most common way that rotenone exposure to humans would take place is
through ingestion. However, absorption in the stomach and intestines is slow and
incomplete, and the liver breaks down the compound effectively, thus enormous
quantities would need to be ingested. Consistent with this view is the fact that chronic
ingestion of rotenone by rats for 24 months at doses 30 times greater than that used to
model PD by systemic infusion (Betarbet et al. 2000) failed to cause any behavioral or
neuropathological features of the disease (Marking 1988). In a single reported case of
fatal rotenone poisoning after acute ingestion (De Wilde et al. 1986) rotenone was not
found in the brain although present in blood, liver and kidney.
2.7 ROTENONE-TREATED ANIMAL PD MODELS
2.7.1 Rodents
Rotenone has been used extensively as a prototypic mitochondrial poison in cell cultures,
but less frequently in living animals. In animals, rotenone has been administered by
different routes. As stated above, oral delivery of rotenone appears to cause little
neurotoxicity in animals (Gorell et al. 1998). Systemic administration, on the other hand,
often causes toxicity and lethality, the degree of which is related to the dose used.
Stereotaxic injection of rotenone into the median forebrain bundle depletes striatal DA
and serotonin (Gao et al. 2002). Rats treated for a week with rotenone by intravenous
infusion show bilateral lesions of the striatum and the globus pallidus, characterized by
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neuronal loss and gliosis (Heikkila et al. 1985). In that study, the nigrostriatal DAergic
pathway remained unaffected. Similarly, subcutaneous injection of rotenone once or
multiple times, although causing fatality, failed to affect striatal DAergic content in mice
(Thiffault et al. 2000). Conversely, Greenamyre and collaborators (Betarbet et al. 2000;
Sherer et al. 2003b) found that intravenous and subcutaneous infusion of rotenone for
about 3 weeks to rats does produce nigrostriatal DAergic neurodegeneration. By
quantitative analysis, SN DAergic neuron numbers are reduced by about 30% in
rotenone-infused rats compared with vehicle controls (Hoglinger et al. 2003b). This
study also shows that the numbers of mesolimbic DAergic neurons, the cell bodies of
which reside adjacent to the SN in the ventral tegmental area (VTA), are unchanged by
rotenone administration. In the ST, the average loss of DAergic fibers was estimated to
be 55% after rotenone infusion in rats, similar in PD, was greater than the loss of SN
DAergic neurons. Despite the use of the exact same regimen of rotenone, the severity of
the striatal DAergic damage in rats within a given experiment appears highly variable,
ranging from none to near complete (Betarbet et al. 2000; Hoglinger et al. 2003b; Sherer
et al. 2003a; Lapointe et al. 2004; Zhu et al. 2004). After the infusion of rotenone, the
loss of tyrosine hydroxylase-positive (TH+ve) fibers in the ST is either focal, showing a
zone of maximal loss at the center, or diffuse; whether the latter represents a more severe
lesion of the striatal DAergic fiber network than the former remains to be demonstrated.
Of note, the focal loss in the center of the ST seen in some of the lesioned rats is a pattern
that differs from that of PD in which the dorsolateral quadrant of the ST is typically the
most affected.
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2.7.2 Invertebrates
Rotenone has also been shown to produce behavioural impairments related to the
DAergic system in invertebrates. In Drosophila, application of rotenone (25-500 μM)
produces a reduction in locomotion and selective loss of TH (Coulom and Birman 2004;
Meulener et al. 2005). Drosophila lacking DJ-1 function (Meulener et al. 2005), despite
being viable, fertile and with a normal lifespan, show striking selective sensitivity to
environmental toxins, such as paraquat and rotenone, that are linked epidemiologically to
sporadic PD in humans (Hertzman C. 1990; Semchuk et al. 1992; Liou et al. 1997; Gorell
et al. 1998; Fall et al. 1999; Uversky 2004). In flatworms, rotenone evokes neuronal
degeneration which is prevented by antiparkinsonian drugs (Kitamura et al. 2003). The
pond snail (Lymnaea stagnalis) is widely used in invertebrate neurobiology (Vehovszky
et al. 2007). Both acute and chronic rotenone treatment inhibited spontaneous
locomotion and feeding, both behaviours being dependent on muscle movement.
Biochemical assays revealed a significantly lower DA content in addition to TH loss in
the cell bodies and axons (Beal 2001; Vehovszky et al. 2007). Snails have advantages
over Drosophila that their neurons in the CNS are large and individually identifiable from
preparation to preparation (Syed and Winlow 1991; Skingsley et al. 1993; Sakharov and
Tsyganov 2000; Tsyganov and Sakharov 2000; Zhu et al. 2004; Haque et al. 2006).
DAergic neurons have also been identified in the buccal ganglia of snails (Kyriakides et
al. 1989; Kyriakides and McCrohan 1989; Elekes et al. 1991; Rosen et al. 1991; Quinlan
et al. 1997; Kabotyanski et al. 2000). This suggests that vertebrates and invertebrates
share some common mechanisms in rotenone-triggered changes leading to syndromes of
Parkinsonism.
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2.8 ROTENONE AND PD PATHOLOGY
In contrast to the 6-OHDA and MPTP models, rotenone-infused rats show proteinaceous
inclusions in some of the remaining SN DAergic neurons. Like LBs in PD, these
inclusions are immunoreactive for both ubiquitin and �-synuclein (Betarbet et al. 2000),
and by electron microscopy they appear composed of a dense core with fibrillar
peripheral elements. Like PD in which neurodegeneration extends beyond the DAergic
system (Agid and Blin 1987), rotenone infusion is associated with 35% reduction in
serotonin transporter density in the striatum, 26% reduction of noradrenergic neurons in
the locus coeruleus, and 29% reduction in cholinergic neurons in the pedunculopontine
nucleus (Hoglinger et al. 2003a).
Although the initial descriptive studies did not report any striatal lesion (Betarbet et al.
2000), the number of DA-regulated phosphoprotein-32 projecting neurons, cholinergic
interneurons and nicotinamide adenine dinucleotide phosphate (NADPH) diaphorase-
positive neurons in the ST were all significantly reduced by the infusion of rotenone in
rats (Hoglinger et al. 2003a; Lapointe et al. 2004). Unexpectedly, even at doses of
rotenone that did not damage the nigrostriatal DAergic pathway in rats, despite the
significant loss of intrinsic striatal neurons. Remarkably, Zhu and collaborators (Zhu et
al. 2004) found that the rotenone-induced intrinsic striatal neuronal loss occurs especially
in those rats exhibiting the central striatal loss of tyrosine hydroxylase immunoreactivity
(TH-ir) mentioned above. These results indicate that rotenone exerts a much more
widespread neurotoxicity than initially thought and, contrary to the initial contention that,
it does not consistently spare striatal postsynaptic DAergic neurons.
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2.9 ROTENONE AND L-DOPA
Behaviorally, rotenone-infused rats exhibit reduced mobility, flexed posture, and in some
cases rigidity (Sherer et al. 2003a) and even catalepsy (Alam and Schmidt 2002). Four
weeks after the infusion of rotenone, rats show more than 70% reduction in spontaneous
motor activity (Hoglinger et al. 2003b). These motor abnormalities appear to be reversed
by L-DOPA administration (Alam and Schmidt 2004). However, some rotenone-infused
rats without a nigrostriatal DAergic lesion have been reported to exhibit a similar set of
motor abnormalities (Sherer et al. 2003a). In addition, indices of DAergic damage
across different doses of rotenone did not correlate with motor behavior in individual rats
(Fleming et al. 2004). Therefore, whereas the rotenone-related motor abnormalities are
dramatic, it is still questionable that they result from a loss of nigrostriatal DAergic
neurons and thus the use of these behavioral alterations as an experimental correlate of
PD symptoms must be done with caution.
2.10 MECHANISMS OF ROTENONE TOXICITY
2.10.1 The Complex 1 Inhibitor
Like MPTP, rotenone is highly lipophilic and thus readily gains access to all organs
including the brain. After a single intravenous injection, rotenone reaches maximal
concentration in the CNS within 15 min and decays to about half of this level in less than
2 hours (Talpade et al. 2000). Its brain distribution is heterogeneous, paralleling regional
differences in oxidative metabolism. It also freely crosses all cellular membranes and can
accumulate in subcellular organelles such as the mitochondria where it impairs oxidative
phosphorylation by inhibiting reduced NADH-ubiquinone reductase activity through its
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binding to the PSST subunit of the multipolypeptide enzyme complex I of the electron
transport chain (Schuler and Casida 2001a). Aside from its action on mitochondrial
respiration, rotenone also inhibits the formation of microtubules from tubulin (Brinkley et
al. 1974; Marshall and Himes 1978). This effect may be quite relevant to the mechanism
of DAergic neurodegeneration because excess of tubulin monomers may be toxic to cells
(Burke et al. 1989; Weinstein and Solomon 1990). Interestingly, a protein implicated in
some familial forms of PD, parkin, appears to bind to tubulin, thereby enhancing the
ubiquitination and degradation of misfolded tubulins, an effect that is lacking with the
PD-linked parkin mutants (Ren et al. 2003).
2.10.2 Rotenone and OS
Brains of PD patients show evidence of OS, including decreased levels of reduced
glutathione and oxidative modifications to DNA, lipids, and proteins (Dexter et al. 1989a;
Alam et al. 1997; Pearce et al. 1997; Floor and Wetzel 1998), and oxidative damage is
hypothesized to contribute to the neurodegenerative process in PD (Jenner 1998). The
source of this oxidative damage is unknown. ROS are generated during dopamine
metabolism and by mitochondrial respiration. Within complex I, upstream of the
rotenone-binding site, is a site of electron leakage that produces ROS (Hensley et al.
1998; Kushnareva et al. 2002) and impaired complex I activity enhances ROS formation
(Cassarino et al. 1997; Barrientos and Moraes 1999; Kushnareva et al. 2002).
As in vivo rotenone infusion resulted in uniform complex I inhibition across brain regions
(Betarbet et al. 2000), ROS production by complex I dysfunction cannot fully explain the
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selectivity of oxidative damage resulting from rotenone exposure. Nevertheless, rotenone
infusion caused relatively selective oxidative damage to certain DA-rich brain regions
known to degenerate in PD. It has been hypothesized that there may be an interaction
between ROS synthesized during rotenone-induced mitochondrial inhibition and ROS
produced during DA metabolism. DAergic neurons are believed to exist normally in a
state of compensated OS because they contain DA. Normal metabolism of DA by
monoamine oxidase produces H2O2 (Maker et al. 1981), which is sufficient to increase
levels of oxidized glutathione {Maker, 1981 #533; Spina and Cohen 1988), and which
suggests ongoing oxidative stress. In addition, in the presence of iron (abundant in the
SN), H2O2 is converted to the reactive and highly damaging hydroxyl radical.
Furthermore, DA may be oxidized nonenzymatically to form reactive DA semiquinones
(Hastings et al. 1996). For these reasons, the greatest oxidative damage after rotenone
exposure may occur in DAergic neurons. Thus, the extent of oxidative damage in
DAergic neurons may be much greater than that observed for the entire homogenate
because protein from DAergic neurons comprises only a small percentage of the total
midbrain homogenate. Oxidative damage after rotenone exposure may explain, in part,
the aggregation of α-synuclein caused by rotenone (Betarbet et al. 2000). In the PD
brain, α-synuclein becomes oxidatively modified and insoluble, leading to aggregation
(Giasson et al. 2000).
2.10.3 Rotenone and Inflammation
The rotenone-induced degeneration of DAergic neurons may not be solely attributable to
an impairment of neuronal mitochondrial complex I activity but may also involve the
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activation of microglia (Gao et al. 2003a; Sherer et al. 2003b). Microglial cells, which
are the resident macrophages in the brain, respond to many insults by rapid proliferation,
hypertrophy, and expression of a number of cytokines (Floyd et al. 1999; Raivich et al.
1999). SN has the highest density of microglia in the brain ((Kim et al. 2000).
Importantly, there is an increase in reactive microglia in the ST and SN of patients with
idiopathic PD (McGeer et al. 1988; Beal 2003). In vitro, rotenone-induced microglial
activation before apparent neurodegeneration (Gao et al. 2002). Activated microglia
upregulate cell surface markers such as the macrophage antigen complex I and produce a
variety of proinflammatory cytokines. One of the consequences of microglia activation is
the production of ROS.
The reaction of O2• with NO• generates peroxynitrite ONOO–, which is strongly
implicated in PD pathogenesis (Beal 2003). Prior studies have shown that the LB-
positive SN neurons in PD stain with antibodies to 3-nitrotyrosine, which is thought to be
a specific marker for peroxynitrite-mediated damage (Good et al. 1998). Furthermore,
LBs are known to react with antibodies to nitrated α-synuclein, the major protein
component of LBs and LNs (Duda et al. 2000; Giasson et al. 2000).
2.10.4 Rotenone and Apoptosis
Rotenone is assumed to exert its cytotoxicity through the induction of apoptosis (Duan
and Mattson 1999; Duan et al. 1999; Isenberg and Klaunig 2000; Armstrong et al. 2001),
a form of programmed cell death that normally occurs during the development of the
nervous system. The term apoptosis is applied to a group of characteristic structural and
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molecular events that separate this type of cell death from necrosis. In contrast to
necrosis, which involves a group of cells simultaneously, apoptosis may occur in a single
cell surrounded by a group of viable cells. There is a distinct and precisely localized
control over the fate of specific cells in a mixed cell population that undergo apoptosis.
The biochemical cascades that lead to apoptotic cell death seem to involve the activation
of one or more members of a family of cysteine proteases, called caspases, and the
release of factors, such as cytochrome c from mitochondria, that ultimately induce
nuclear DNA condensation and fragmentation (Kiechle and Zhang 2002). There are two
major pathways through which apoptosis are induced; one involves death receptors and is
exemplified by Fas-mediated caspase-8 activation, and the other is the stress-mediated or
mitochondria-mediated caspase-9 activation pathway. Both pathways converge on
caspase-3 activation, resulting in nuclear degradation and cellular morphological change
(Ueda et al. 2002). Most evidence has stemmed from in vitro models (see Chapter 3).
2.11 OTHER PARKINSONIAN-INDUCING NEUROTOXINS
2.11.1 6-hydroxydopamine (6-OHDA)
6-Hydroxydopamine (6-OHDA), a hydroxylated derivative of DA, was introduced as the
first animal model of PD (Porter et al. 1963; Ungerstedt 1968; Blum et al. 2001). In
contrast to MPTP and rotenone, 6-OHDA does not cross the rat BBB, therefore it is
administered by stereotaxic injection along the pathway of the DAergic fibers into the
SNc, the median forebrain bundle, or the ST. The stereotaxic injection allows a unilateral
lesion avoiding generalized symptoms and providing a contralateral control (Perese et al.
1989; Cenci et al. 2002). The extent of the lesion can be quantified in vivo by measuring
105
the asymmetric turning behaviour of the rats induced by systemic administration of DA
receptor agonists, L-DOPA or DA- releasing drugs (Ungerstedt 1968; Hefti et al. 1980).
However, the turning behaviour due to unilateral lesions may also cause contralateral
modifications at striatal level or in the STN (Salin et al. 1996; Perier et al. 2000). In
rodents, cats and non-human primates, 6-OHDA-induced toxicity is relatively selective
for monoaminergic neurons, as a result of its uptake by DA and noradrenaline
transporters (Luthman et al. 1989). Once inside the neurons 6-OHDA produces toxic
effects by inhibiting mitochondrial complex I and causing OS (Sachs and Jonsson 1975a,
b; Glinka et al. 1997).
The 6-OHDA model has provided considerable amount of information. DA denervation
is known to affect corticostriatal glutamatergic neurotransmission (Calabresi et al. 1993a;
Schwarting and Huston 1996a, b; Dunah et al. 2000). In the parkinsonian rat, L-DOPA
therapy is able to reverse the alterations observed in glutamatergic neurotransmission
(Calabresi et al. 2000; Picconi et al. 2004). However, similar to humans after prolonged
L-DOPA treatment dyskinesias appear. The 6-OHDA model does not reproduce all the
pathological and clinical features of human Parkinsonism. It induces DAergic neuron
death with preservation of non-DAergic neurons, whereas the formation of cytoplasmatic
inclusions (Lewy bodies) does not occur. 6-OHDA does not affect other brain areas
involved in PD, such as the anterior olfactory structures, lower brain stem areas or the
locus coeruleus (Betarbet et al. 2002b; Del Tredici et al. 2002). Reports of Parkinsonian-
like tremor are rare in studies of 6-OHDA-lesioned rodents however occasional akinesia,
rigidity and tremor have been described (Lindner et al. 1999; Cenci et al. 2002). Finally,
the regimen of the 6-OHDA model with intrastriatal injections may be more useful for
106
neuroprotective studies, whereas the regimen with 6-OHDA injections into the SN–VTA
complex appears to be a more useful approach for testing new pharmacological or cell
replacement therapies (Hirsch et al. 2003). In general, this model exclusively induces
acute effects, which differs significantly from the slowly progressive pathology of human
PD (Betarbet et al. 2002b).
2.11.2 Maneb
The mechanism of maneb toxicity is not well known (Thiruchelvam et al. 2002). Maneb
contains a major active fungicidal component, manganese ethylene-bis-dithiocarbamate
(Mn–EBDC), and belongs to the dithiocarbamate (DTC) fungicide family. Chronic
exposures of humans to maneb have been linked to the development of Parkinsonism
(Ferraz et al. 1988; Meco et al. 1994). Furthermore, in experimental models, maneb
appears to decrease locomotor activity (Morato et al. 1989), potentiate MPTP effects on
locomotor activity and catalepsy (Takahashi et al. 1989), and modulates the toxicity of
paraquat (see above).
Commercial maneb contains not only Mn–EBDC but also many other minor reagents
without clearly defined functions. Other constituents of maneb, rather than the fungicidal
Mn–EBDC, may be responsible for maneb-mediated neurotoxicity, as is the case of
MPTP-contaminated synthetic heroin. Alternatively, the neurotoxic effect can be
attenuated by the combined action of several constituents of maneb. Furthermore, Mn–
EBDC, being relatively stable in vitro, could potentially degenerate to manganese and
EBDC in vivo, both of these compounds being potentially neurotoxic. Indeed, manganese
is known to be relatively non-toxic to the adult organism except for the brain, where it
107
causes PD-like symptoms when inhaled, even at moderate amounts over longer periods of
time (Carpenter 2001; Gerber et al. 2002).
Occupational exposure to manganese occur mainly in mining, alloy production,
processing, ferro-manganese operations, welding, and work with agrochemicals. Among
the neurologic effects is an irreversible parkinsonian-like syndrome, manganism (Levy
and Nassetta 2003; Takeda 2003). However, although the neurological signs of
manganism have received close attention because they resemble several clinical disorders
collectively described as extrapyramidal motor system dysfunction and in particular,
idiopathic PD (IPD) and dystonia, there are well-established distinct dissimilarities
between IPD and manganism. Therefore, whether manganese plays an etiologic role in
idiopathic PD remains to be determined (Aschner 2000).
On the other hand, the EBDC component (and not the manganese moiety of maneb) has
been suggested to contribute to toxicity. This conclusion follows from the finding that
both MANCOZEB (Mn–Zinc–EBDC) and ZINEB (Zinc–EBDC) produce neurotoxicity
in cell cultures (Soleo et al. 1996). Indeed, EBDC per se enhances MPTP-induced
neurotoxicity (McGrew et al. 2000). The direct involvement of manganese ethylene-bis-
dithiocarbamate (Mn–EBDC) in selective DAergic neurodegeneration was recently
demonstrated in a rat model (adult male Sprague-Dawley rats), in which Mn–EBDC was
directly delivered to the lateral ventricles (Zhang et al. 2003). Use of this model has
shown that Mn–EBDC is able to induce extensive striatal dopamine efflux comparable
with that induced by MPP+. Furthermore, Mn–EBDC preferentially inhibits
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mitochondrial complex III. As mitochondrial dysfunction is pivotal in the pathogenesis of
PD, these results support the proposal that exposure to pesticides such as maneb, or other
naturally occurring compounds that inhibit mitochondrial function, may contribute to PD
development (Zhang et al. 2003).
2.11.3 Paraquat
Acute poisoning by high levels of paraquat is known to cause lung, liver, kidney, and
brain injury. This herbicide is poorly absorbed when inhaled but causes severe illness
when ingested orally, usually causing death within 2 days of ingestion. At lower doses,
death may be delayed for several weeks. Paraquat has been shown to accumulate in lung
tissue, where free radicals are formed, lipid peroxidation is induced, and NADPH is
depleted, producing diffuse alveolitis, followed by extensive pulmonary fibrosis (Bismuth
et al. 1990). Free radical production and the activation of cholinergic and glutamatergic
transmission are related events palying a crucial role in paraquat-induced neurotoxicity
(Corasaniti et al. 1998). Furthermore, paraquat is known to be a potent redox cycler,
being readily converted to a free radical, which, in reaction with molecular oxygen,
generates superoxide anions and subsequently other redox products (Jones and Vale
2000; Yumino et al. 2002).
Paraquat induces NADPH-dependent lipid peroxidation (Hara et al. 1991a, b, c, d; Jones
and Vale 2000; Yumino et al. 2002). Subcutaneous injections in rats resulted in a
glutamate efflux initiating excitotoxicity of nitric-oxide-synthase-containing neurons and
eliciting the depolarization of N-methyl-d-aspartate (NMDA) receptor channels and Ca2+
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penetration into cells by activation of non-NMDA receptor channels (Shimizu et al.
2003b). The influx of Ca2+ into cells stimulates nitric oxide synthase release and
diffusion into DAergic terminals inducing mitochondrial dysfunction and DA overflow
(Shimizu et al. 2003b). Importantly, paraquat dose-dependently reduces the number of
DAergic neurons in cultured rat organotypic midbrain slices (Shimizu et al. 2003a).
Since this damage is prevented by GBR-12909, a selective DAT inhibitor, DAT has been
assumed to be the initial step in paraquat-induced DAergic neurotoxicity. The death of
DAergic neurons is prevented by inhibitors of NMDA, nitric oxide synthase,
cycloheximide, and the caspase cascade (Shimizu et al. 2003a). Neurodegeneration is
also inhibited by l-deprenyl and dopamine D2/3 agonists. These results strongly support
that constant exposure to low levels of paraquat leads to the vulnerability of DAergic
neurons in the nigrostriatal system via excitotoxic pathways (Shimizu et al. 2003a).
Another line of evidence emphasizes the pathological importance of the direct interaction
between paraquat and α-synuclein. Paraquat markedly accelerates the in vitro rate of α-
synuclein fibril formation (Uversky et al. 2001; Uversky et al. 2002), with the
accelerating effects being dose-dependent (Manning-Bog et al. 2002). Furthermore,
when mice are exposed to the herbicide, brain levels of α-synuclein are significantly
increased. This up-regulation follows a consistent pattern, with higher α-synuclein levels
at 2 days after each of three weekly paraquat injections and with protein levels returning
to control values by day 7 post-treatment (Manning-Bog et al. 2002). This is
accompanied by the formation of intraneuronal aggregates with histological properties of
amyloid fibrils (thioflavine S staining), to which α-synuclein co-localizes (Manning-Bog
110
et al. 2002). Thus, paraquat exposure triggers α-synuclein fibrillation in the mouse brain.
This suggests that the up-regulation of α-synuclein as a consequence of toxicant insult
and the direct interaction between the protein and environmental agents are potential
mechanisms leading to α-synuclein pathology in neurodegenerative disorders.
2.12 SUMMARY
It is clear that certain environmental neurotoxins can promote and accelerate the
development of PD, whereas other environmental agents may be neuroprotective; for
example, caffeine consumption and cigarette smoking may decrease the risk of PD
{Manning-Bog, 2002 #631; Ross, 2000 #659; Ross, 2000 #660; Ragonese, 2003 #662;
Ross, 2001 #661; Tan, 2003 #663; Schwarzschild, 2003 #665; Ascherio, 2001 #664;
Gale, 2003 #1537; Paganini-Hill, 2001 #1538}. PD-promoting environmental toxins are
metals (Tanner et al. 1989; Hirsch et al. 1991; Good et al. 1992; Yasui et al. 1992;
Hertzman et al. 1994; Hellenbrand et al. 1996b; Hellenbrand et al. 1996a; Altschuler
1999; Gorell et al. 1999), solvents (Uitti et al. 1994; Pezzoli et al. 1996; Seidler et al.
1996; Davis and Adair 1999; Hageman et al. 1999), carbon monoxide (Klawans et al.
1982), and MPTP (Langston et al. 1983). Epidemiological studies have shown an the
association between increased PD risk and such environmental factors as rural residence
(Morano et al. 1994; Liou et al. 1997; Marder et al. 1998), farming (Semchuk et al. 1992;
Liou et al. 1997; Gorell et al. 1998; Fall et al. 1999), the drinking of water from wells
(Morano et al. 1994; Marder et al. 1998), and exposure to agricultural chemicals,
pesticides, and/or herbicides (Semchuk et al. 1992; Liou et al. 1997; Gorell et al. 1998;
Fall et al. 1999; Vanacore et al. 2002). The ability of different environmental factors to
111
increase PD risk and produce PD characteristics in some experimentally has resulted in
the development of valuable models for the analysis of mechanisms of neurodegeneration
targeting the nigrostriatal system (Di Monte 2003).
Figure. 2.4. A–C Chemical structure of rotenone (A), paraquat and MPP+ (B), and
maneb (C) (Uversky 2004).
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CHAPTER 3:
IN VITRO MODELS OF PD
113
3.0 INTRODUCTION
Neural tissue grown and maintained in vitro provides an alternate way of studying in situ
properties of neurons and glial cells under simple and controlled conditions. A given
neural tissue culture preparation retains its physiological and biochemical properties
representing those exhibited by the intact brain making tissue culture a useful model for
studying the nervous system. There are four broad classes in which in vitro cultivation
and maintenance of living biological units from multicellular organisms techniques fall
into:
1) Organotypic (explant) cultures are explanted from specific neuroanatomical loci
to substrates as small tissue fragments.
2) Dissociated cell cultures involve the seeding of enzymatically or mechanically
dispersed single cells on various substrates.
3) Reaggregate cultures require reassociation of dissociated single cells into small
aggregates.
4) Cultures of purified cell populations or monotypic cultures are prepared by the
isolation of particular cell types by gradient centrifugation or other separation
technique.
3.1 DISSOCIATED CULTURES
This form of cell culture involves the isolation of the brain region containing the cells of
interest mainly from embryonic tissue, and the dissociation of the tissue into a single cell
suspension. These cells are then plated onto a surface that they will adhere to such as
laminin or poly-D-lysine coated coverslips in multiwell dishes. Alternatively, neurons
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can be cultured within the culture medium without attachment to a surface, thus allowing
aggregation to take place (Prochiantz et al. 1979; di Porzio et al. 1980; Prochiantz et al.
1981; Denis-Donini et al. 1983).
Many neuronal populations have been studied using this form of cell culture including the
DAergic neurons of the VM (Prochiantz et al. 1979; Daguet et al. 1980; Prochiantz et al.
1981; Berger et al. 1982; Dal Toso et al. 1988; Soderback et al. 1989; Niijima et al. 1990;
O'Malley et al. 1991; Masuko et al. 1992; Fawcett et al. 1995). These dissociated
cultures of DAergic cells develop in vivo morphological characteristics such as the
release and synthesis of DA (Prochiantz et al. 1979; Daguet et al. 1980; Prochiantz et al.
1981; Dal Toso et al. 1988), respond to a variety of neurotrophic factors (Ferrari et al.
1989; Knusel et al. 1990b; Knusel et al. 1990a; Ferrari et al. 1991; Kushima et al. 1992;
Kushima and Hatanaka 1992; Lin et al. 1993; Nikkhah et al. 1993; Hyman et al. 1994), in
addition express many of the electrophysiological characteristics that have been reported
for midbrain DAergic neurons in vivo and in slice cultures (Ort et al. 1988; Masuko et al.
1992; Cardozo 1993).
Disadvantages of this technique come from the mechanical destruction of all components
of the tissue matrix and alterations off cell/cell interactions during the culture process.
Cell plating density has been shown to influence the survival of DAergic neurons in
several studies such as increasing cell density enhances the specific high affinity DA
uptake per cell as well as cell survival in dissociated cultures of E13 mouse
mesencephalic cells (Dal Toso et al. 1988). Others such as O’ Malley et al., (O'Malley et
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al. 1991) have demonstrated that high plating conditions of embryonic rat mesencephalic
cells selectively enhanced the survival of the DAergic subpopulation; Fawcett et al.,
(Fawcett et al. 1995) found that 87% of DAergic neurons in a dissociated monolayer
culture system died which was far greater than cell death seen in a 3-dimensional culture
system where cell/cell contacts were increased.���
3.2 REAGGREGATE CULTURES
Reaggregate cultures involve the dissociation of neural tissues into single cells and
subsequently the dispersion of cells resembling themselves under controlled conditions.
The dissociated cell suspensions are maintained in rotating flasks. These single cell
suspensions can be cultured alone or in combination with single cells from other areas of
the embryonic brain. Over a period of 12-24 hours these cells form spherical
reaggregates of typical 300 μm in diameter (Heller et al. 1988). It was observed that
dissociated DAergic neurons from the rostral mesencephalic tegmentum of embryonic
day 14 mouse brain, co-aggregated with dissociated cells from the ST, developed an
increasing ability to synthesise, accumulate and store DA with time (Kotake et al. 1982;
Shalaby et al. 1983). This temporal development resembled the in vivo development of
DAergic neurons. Furthermore, an increased survival of mouse embryonic DAergic
neurons when co-cultured with striatal target neurons (Hoffmann et al. 1983) and a
selective association of DA axons with their ST target cells have been shown and also the
formation of synaptic contacts between embryonic mouse DAergic and striatal neurons
within reaggregate cultures (Won et al. 1989).
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A major advantage is that these cells develop into 3-dimensions. This technique has also
proven useful in allowing studying of transition from unicellular to a multicellular system
and be maintained long term (Choi et al. 1993). There is electron microscopic evidence
of synapse and myelin formation in these aggregate cultures (Seeds 1975). In
reaggregate cultures, a disadvantage is that cells cannot be visualized under direct
observation, and the morphological examination requires time-consuming and tedious
histological methods.
3.3 ORGANOTYPIC EXPLANT CULTURES
The goal for an explant culture system is to culture small fragments of tissue that will
maintain the cytoarchitecture, simulate differentiation and maturation patterns, and
preserve the functions of the original tissue. Explants maybe maintained in special
Maximow chambers (Bornstein 1958) or in the culture dishes. The development of
DAergic neurons within explant cultures of postnatal DAergic neurons co-cultured with
ST tissue have been described (Whetsell et al. 1981; Mytilineou et al. 1983). These
nigro-striatal explant cultures survived for up to 15 weeks were capable of DA synthesis,
uptake and release.
The explant culture system has the advantage that cells may be maintained for months in
vitro, unlike dissociated and reaggregate cultures; the cytoarchitecture of the culture is
maintained and preserved within the explants. Most of the important structural and
functional attributes of neurons and glial cells have been found to be maintained in
culture. These include the formation of synapses, the appearance of normal myelin, rise
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in enzyme activity associated with neurotransmitter metabolism, the increase in myelin-
specific enzymes, and the development of the bioelectric activity characteristic of
interneuronal communication (Burton and Bunge 1975; Crain and Peterson 1975; Crain
et al. 1975). The explant technique also enables the study of the formation of connections
between two co-culture explants. However, despite these advantages of explant cultures
due to the thickness of the tissue sections (1 mm), cells within the explant are almost as
inaccessible as they are in situ. Furthermore, also as a consequence of the thickness of
the culture, examination by light microscopy has poor results.
3.4 ORGANOTYPIC SLICE CULTURES
This technique will be employed throughout this thesis and was developed by Gahwiler
(Gahwiler 1981, 1988). Organotypic tissue culture techniques allow neuronal tissue to be
maintained for weeks in vitro, and have thus expanded the utility of in vitro preparations
for neurological study. When prepared from thick (200–400 �m) sections of neuronal
tissue, organotypic cultures permit the maintenance of the three-dimensional tissue
architecture observed in vivo. These cultures maintain fundamental neuronal circuitries,
neuronal and glial interactions and have the ability to form new neuronal synapses with
other appropriate brain regions when cultured together.
Two organotypic culture techniques are predominately utilized. The first is the roller-tube
method (Gahwiler 1988; Ostergaard et al. 1990), in which tissue is continuously rotated
within a bath of medium. The second is the static membrane model developed by
Stoppini and colleagues (Stoppini et al. 1991); in this technique, tissue rests on a porous
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membrane exposed to culture medium below while the upper surface is exposed to air so
that tissue oxygenation can occur. Regardless of the culture method used, the organotypic
preparation allows the study of an in vivo-like anatomical system with the ease of in vitro
sample manipulations, such as administration of controlled levels of growth factors
(Hoglinger et al. 1998; Schatz et al. 1999; Meyer et al. 2001; Jaumotte and Zigmond
2005) or neurotoxicants (Kotake et al. 2003; Shimizu et al. 2003a; Kress and Reynolds
2005; Testa et al. 2005).
The organotypic slice culture technique is an adaptation of the explant culture technique
that permits the long-term preservation of tissue structure within cultures that are highly
accessible to observation and experimental manipulation. This approach involves
culturing slices of neuronal tissue attached to glass coverslips, and are rotated in tissue
culture tubes. The main advantage over dissociated neuronal cultures is that the
organotypic cultures retain a high degree of their original morphology and characteristics
(including electrophysiological characteristics and the formation of functional synaptic
networks) and have been described using source tissue from various brain regions. These
include the VM (Jaeger et al. 1989; Ostergaard et al. 1990; Sorensen et al. 1994; Holmes
et al. 1995; Jones et al. 1995; Steensen et al. 1995; Dickie et al. 1996) hippocampus
(Zimmer and Gahwiler 1984; del Rio et al. 1991), cerebellum (Calvet and Calvet 1988a,
b; Mouginot and Gahwiler 1995), visual cortex (Caeser et al. 1989), retina (Feigenspan et
al. 1993), and ST (Ostergaard 1993; Ostergaard et al. 1995). Organotypic co-cutlures of
slices from two (and sometimes three) brain regions may also be prepared. Other regions
of the brain have also been employed such as co-cultures of locus coeruleus with
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hippocampus (Knopfel et al. 1989), cortex with thalamus (Bolz et al. 1992), hippocampus
and medial septum (Gahwiler and Brown 1985), ST with neocortex (Ostergaard 1993),
visual cortex and lateral geniculate nucleus or superior colliculus (Bolz et al. 1992;
Novak and Bolz 1993) and VM with ST (Jaeger et al. 1989; Ostergaard et al. 1990) have
been described. The co-culture technique is used in experiments described in Chapter 5
to study the development of the nigrostriatal pathway in organotypic cultures.
The roller-tube technique for slice cultures were introduced by Hogue (1947) and
modified by Costero (Costero and Pomerat 1951). Since then Gawhiler (Gahwiler 1981)
has developed this technique culturing postnatal rat tissue from several brain regions
including the cerebellum, hippocampus and hypothalamus (Gahwiler 1984a, b). This
technique also allows the culturing of two different regions of the brain together such as
the ST and neocortex (Knopfel et al. 1989; Ostergaard et al. 1991; Bolz et al. 1992) and
more importantly the ventral mesencephalon with the striatum (Whetsell et al. 1981;
Jaeger et al. 1989; Ostergaard et al. 1990).
The organotypic slice culture system has been used to study the development and survival
of DAergic neurons of the VM employing different donor ages (E21- P12) and different
species, including human (Ostergaard and Rasmussen 1991) have been shown to survive
for up to nine months. The morphology of DAergic neurons within organotypic slice
cultures resemble that of the postnatal rat in vivo: they exhibit several shapes including
bipolar, pyramidal and multipolar (Ostergaard et al. 1990). Electrophysiological
characteristics of DAergic neurons in organotypic cultures of VM have been shown to
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resemble those seen in other in vitro preparations and in vivo (Steensen et al. 1995).
However, one fundamental difference between organotypic slice cultures and other in
vitro preparations is the presence of spontaneous burst firing activity in organotypic
cultures (Steensen et al. 1995), which resembles a firing pattern characteristically only
seen in the intact animal. It has been suggested that burst firing results from afferent
inputs (Yung et al. 1991) thus these afferents are likely to be present in organotypic
cultures and so are a more realistic representation of the intact animal.
Cultures of purified cell populations have been recognized as important development in
the field of neural tissue culture. Reports of cultures enriched for neurons (Mains and
Patterson 1973a, b; Wood 1976), astrocytes (McCarthy and de Vellis 1980; Kim et al.
1983a), oligodendrocytes (McCarthy and de Vellis 1980; Lisak et al. 1981; Kim et al.
1983b), and Schwann cells (Brockes et al. 1979; Brockes and Raff 1979). These culture
systems have contributed to a better understanding of morphological, functional, and
metabolic properties of individual cell types and neuron-glia interactions.
All of these culture systems are considered to be primary cultures as they started from
material obtained directly from the brain tissue of origin without any subculturing. The
inability of normal neurons to divide continually not only makes the maintenance of
neural cultures challenging but those cultures that do survive will grow, mature,
differentiate, and eventually die within the course of 3 to 6 months and no longer than a
year. Primary cultures can not divide indefinitely and die thus clonal cell lines are
defined as “immortal”. Their source usually being neoplastic tissue either explanted from
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in situ neural tumors and subcultured or taken cell lines derived from cloning of
chemically, virally or transformed cells.
3.5 In Vitro SYSTEMS AND THEIR RELEVANCE TO PD
At present, all forms of pharmacological treatment are mainly aimed at increasing
DAergic neurotransmission which only control the symptoms. The underlying cause of
neuronal loss remains unknown. It has been speculated that the etiology of PD is
concerned with the four pathomechanisms:
1) Direct effect of neurotoxins MPTP, 6-OHDA and rotenone on SN DAergic
neurons, resulting in neuronal loss in SN (Davis et al. 1979; Burns et al. 1983;
Calne and Langston 1983; Langston et al. 1983; Bywood and Johnson 2003; Testa
et al. 2005)
2) Toxicity of excitatory amino acids acting on DAergic neurons in SN (Spencer et
al. 1987b; Spencer et al. 1987a; Choi 1988).
3) The involvement of free radical-induced OS leading to degeneration of SN
neurons (Ambani et al. 1975; Dexter et al. 1986).
4) Deficiency in target-derived neurotrophic factor that is responsible for the
maintenance and survival of SN DAergic neurons (Prochiantz et al. 1979; Hyman
et al. 1991).
3.5.1 MPTP & IN VITRO STUDIES
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As previously mentioned in chapter 2, MPP+ the toxic metabolite of MPTP has been used
numerously in animal models and other in vitro systems. In vitro models of PD have
been developed and employed for decades in an attempt to better understand the
mechanisms underlying the causes of DAergic cell death and the PD pathology. These
include primary cell lines derived from rat or mouse mesencephalon either from fetal or
postnatal rodents (Hartikka et al. 1992; Chen et al. 1995; Koutsilieri et al. 1996; Marini
and Nowak 2000; Pong et al. 2000; Shinpo et al. 2000; Bilsland et al. 2002; Callier et al.
2002; Gille et al. 2002; Han et al. 2003; Hoglinger et al. 2003a; Andres et al. 2005; Chu
et al. 2005; Du et al. 2005; Mercer et al. 2005; Li et al. 2006; Bains et al. 2007;
Grammatopoulos et al. 2007; Radad et al. 2007a; Radad et al. 2007b), human
neuroblastoma cell line, SH-SY5Y (Fang et al. 1995; Fall and Bennett 1999; Conn et al.
2002; Gomez-Santos et al. 2002; Ba et al. 2004; Mathiasen et al. 2004), PC12 cells, a cell
line established from rat adrenal pheochromocytoma cell (Greene and Tischler 1976),
possess intracellular substrates for DA synthesis, metabolism and transport (Rebois et al.
1980; Hatanaka 1981; Tuler et al. 1989). The cells contain the enzymes required for
synthesis and decomposition of DA such as TH and MAO. The membrane receptors and
synthesized transmitters in PC12 cells are similar to DAergic neurons located in
midbrain. Therefore, PC12 cell line has been used as a cellular model in PD studies
(Denton and Howard 1987; Marongiu et al. 1988; Basma et al. 1990; Basma et al. 1992;
Itano et al. 1994; Cappelletti et al. 1995; Desole et al. 1996; McNaught et al. 1996; Wei et
al. 1996; Desole et al. 1997b; Desole et al. 1997a; Hom et al. 1997; Ara et al. 1998;
Kohda et al. 1998; Soldner et al. 1999; Lee et al. 2000; Seyfried et al. 2000; Fonck and
Baudry 2001; Guo et al. 2001a; Bai et al. 2002; Gelinas and Martinoli 2002; Quigney et
123
al. 2003; Shimoke et al. 2003; Noda et al. 2004; Virmani et al. 2004; Xu et al. 2005;
Chalimoniuk et al. 2006; Chiasson et al. 2006; Gesi et al. 2006; Meuer et al. 2006;
Weinreb et al. 2006; Iuvone et al. 2007; Kang et al. 2007), DAergic cell line MN9D has
features such as the transport activity for DA transporter (DAT) and VMAT2, and
secretory vesicle properties makes this in vitro model advantageous for studies in
vesicular monoamine transporters (VMAT2) neuroprotection and molecular mechanisms
involved in the protection and its regulation (Chen et al. 2005), and slice cultures of SN
(Mytilineou et al. 1983; Mytilineou and Cohen 1984; Schmidt et al. 1997; Liu et al. 2001;
Madsen et al. 2003).
3.5.1.2 DAergic Neurons-Target Specific Toxicity
Exposure to MPP+ not only significantly reduces the number of TH+ve cells and neurites
(Sanchez-Ramos et al. 1986; Sanchez-Ramos et al. 1988b; Michel et al. 1989; Michel et
al. 1990; Dickie and Greenfield 1997; Goto et al. 1997; Madsen et al. 2003) but also a
significant decrement in TH, aromatic L-amino acid decarboxylase (AAAD) activities,
reduced DA and DOPAC content and, DA uptake. In addition, TH mRNA was
decreased, whereas AAAD mRNA was no different from control cultures (Michel et al.
1990; Dalia et al. 1993, 1996). In vivo, high doses of MPTP induces a significant
reduction in both striatal DAconcentrations (95%), and midbrain DAergic cells; 69% loss
of nucleus A8 cells, 75% loss of nucleus A9 cells, and in nucleus A10 subnuclei there
was 42% loss of VTA cells, 55% loss of interfascicular nucleus cells, and no loss of cells
in the central linear nucleus. This provides evidence for differential susceptibility to
MPTP toxicity among different mouse strains and provides us the precise locations of
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midbrain DAergic cells that are vulnerable to MPTP (Zuddas et al. 1992a; Del Zompo et
al. 1993; German et al. 1996). The strong belief by which-MPTP selectively destroys
only certain midbrain DAergic neurons is further supported by in vitro studies (Sanchez-
Ramos et al. 1986; Sanchez-Ramos et al. 1988a; Sanchez-Ramos et al. 1988b; Michel et
al. 1990; Saporito et al. 1992; Zuddas et al. 1992a; Del Zompo et al. 1993; German et al.
1996).
3.5.1.3 Complex 1 Inhibitor
Despite intensive investigation, the molecular mechanism of MPTP has not been
definitively demonstrated. MPP+, the active metabolite of MPTP, has been shown to
inhibit complex I of the mitochondrial electron transport chain, which was thought to be
the mechanism by which MPTP induced parkinsonism (Nicklas et al. 1985; Cheeseman
and Clark 1987; Pai and Ravindranath 1991; Sayre et al. 1991; Basma et al. 1992;
Rollema et al. 1994; Sriram et al. 1995; Sriram et al. 1997; Stephans et al. 2002;
Richardson et al. 2007). A study by Soldner (Soldner et al. 1999) reported otherwise. It
was found that treatment of undifferentiated PC12 cells with MPP+ primarily inhibited
proliferation of PC12 cells and secondarily led to cell death after the depletion of all
energy substrates by glycolysis. This cell death showed no morphological characteristics
of apoptosis and was not blocked by treatment with caspase inhibitors. It would seem that
the inhibition of cell growth was not dependent on an inhibition of complex I activity
since MPP+ also inhibited cell proliferation in SH-SY5Y cells lacking mitochondrial
DNA and complex I activity (Soldner et al. 1999).
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However, the weak inhibitory ability of MPP+ at complex I (Ramsay et al. 1991b;
Ramsay et al. 1991a; Richardson et al. 2005) has led some to question the role of
complex I inhibition in MPTP toxicity (Lotharius and O'Malley 2000; Nakamura et al.
2000a). It has been demonstrated that the single-subunit nicotinamide adenine
dinucleotide (reduced) dehydrogenase of Saccharomyces cerevisiae (NDI1) can serve as
a replacement for complex I in mammalian cells and restore electron transfer in cell
devoid of mitochondrial DNA (Seo et al. 1998; Seo et al. 2000; Bai et al. 2001) and that
NDI1 expression in neuroblastoma cells abolishes the toxicity of rotenone, a complex I
inhibitor that has been demonstrated to reproduce features of PD in rats (Betarbet et al.
2000; Sherer et al. 2003b; Sherer et al. 2003a). Recently, it has been shown that NDI1
can be expressed in vivo using viral-mediated techniques and that unilateral expression of
NDI1 partially protected against reduced TH-ir resulting from an acute high-dose
regimen of MPTP (4 x 15 mg/kg over 1 day; (Seo et al. 2006; Richardson et al. 2007).
These findings suggested that NDI1 could provide some protection from MPTP
neurotoxicity. Others have hypothesized alternate mechanisms of action for MPTP
including release of vesicular stored DA and subsequent oxidative damage, release of
stored iron deposits, increased cytoplasmic calcium and intraneuronal calcium release,
and redox cycling of MPP+ (Trevor et al. 1987b; Trevor et al. 1987a; Obata et al. 1992;
Chen et al. 1995; Lotharius and O'Malley 2000; Obata et al. 2001; Obata 2002; Obata et
al. 2002; Kooncumchoo et al. 2006; Obata 2006). Thus, the molecular mechanism of
MPTP remains to be established.
3.5.1.4 Oxidative Stress (OS)
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There is growing evidence indicating that OS and mitochondrial dysfunction play a role
in the pathogenesis and progression of PD. Toxic build up of superoxide and NOS are
major OS indicators such as lipid peroxidation and protein oxidation, are increased in the
cell membrane and/or mitochondria in the ventral MB and the ST may trigger
pathogenesis in PD (Fahn and Cohen 1992; Przedborski and Jackson-Lewis 1998; Reiter
1998; Heales et al. 1999; Munoz et al. 2006; Zeng et al. 2006).
Exposure of mouse brain to MPTP resulted in significant increases of ROS and
malondialdehyde (MDA, the product of lipid peroxidation) and decreases in GSH content
(Okawara et al. 2007). Pretreatment of mouse brain slices with GSH or GSH isopropyl
ester attenuated MPTP toxicity as assessed by the tissue activity of the mitochondrial
enzyme, NADH-dehydrogenase (NADH-DH), and by leakage of the cytosolic enzyme,
lactate dehydrogenase (LDH), from the slice into the medium. This supports that MPTP
exposure generates ROS resulting in OS (Sriram et al. 1997). In addition transgenic mice
with the overexpression of copper/zinc containing SOD were shown to be resistant to
MPTP-induced DAergic toxicity (Przedborski et al. 1992).
While it is generally acknowledged that exposure to MPTP can possibly lead to the
generation of oxygen free radicals, it is debatable whether this is a primary mechanism of
neurotoxicity of MPTP or a secondary event caused by the inhibition of mitochondrial
respiration and resultant energy depletion (Tipton et al. 1993; Tipton and Singer 1993).
The latter hypothesis is supported by the evidence that inhibition of NADH-DH by
rotenone (Turrens and Boveris 1980) or MPP+ (Hasegawa et al. 1990) results in the
127
generation of superoxide. The observation that the irreversible inhibition of complex I by
MPP+ in a non-neuronal system such as bovine heart mitochondria is totally prevented by
the addition of GSH, ascorbate or catalase in the incubation medium, suggests that free
radical generation may be a primary event (Cleeter et al. 1992).
However, the role of OS in MPTP-induced neurotoxicity is seemingly controversial. The
generation of oxygen free radicals by MPTP has been demonstrated in non-neuronal
systems such as cerebellar granule cells (CGC) (Oyama et al. 1993). The addition of
MPP+ induced an increased in the percentage of ROS-positive cells (Oyama et al. 1993).
Interestingly the neuronal non-calcium antagonistic 1,4-dihydropyridine (DHP)
derivatives cerebrocrast, glutapyrone and tauropyrone was shown to decrease the
generation of ROS and the loss of mitochondrial membrane potential which provided
protection against MPP+-induced deterioration of mitochondrial bioenergetics
(Klimaviciusa et al. 2007). The formation of free radicals through redox cycling of MPP+
has been demonstrated in purified hepatic microsomal NADPH cytochrome P-450
reductase (Sinha et al. 1986; Frank et al. 1987; Lambert and Bondy 1989).
3.5.1.5 Inflammation: The Contribution of Microglia and Cytokines
The evidence so far suggests that chronic inflammation, mitochondrial dysfunction, and
OS play significant and perhaps synergistic roles in PD, where the primary pathology is
significant loss of the DAergic neurons in the SN. Mesencephalic neuron-glia cultures
treated with MPTP and an inflammogen lipopolysaccharide (LPS) synergistically,
induced a progressive and selective degeneration of DAergic neurons. The synergistic
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neurotoxicity was observed when both agents were applied either simultaneously or in
tandem. The synergistic neurotoxicity was more prominent when lower doses of both
agents were applied for a longer period of time. In addition, MPTP and LPS
synergistically stimulated the NADPH oxidase-mediated release of superoxide free
radical in which pharmacological inhibition and genetic inactivation of NADPH oxidase
prevented superoxide production and the synergistic neurotoxicity. Furthermore,
inhibition of NOS significantly provided neuroprotection suggesting the involvement of
nitric oxide in the synergistic neurotoxicity (Gao et al. 2003b; Gao et al. 2003a).
There is increasing evidence that suggests an involvement of glia in MPTP neurotoxicity,
the nature of this involvement remains unclear. In particular microglia, but not astroglia
is significantly involved in enhancing the progression of MPTP-induced DAergic
neurodegeneration through the release of NADPH oxidase-derived ROS, microglia
contribute to the progressive neuronal damage. Among the factors measured, the
production of extracellular superoxide was the most prominent. NADPH oxidase
inhibitor, apocynin were shown to attenuate MPTP-induced DAergic neurodegeneration
but only in the presence of glia. More importantly, DAergic neurons from mice lacking
NADPH oxidase, a key enzyme for superoxide production, were significantly more
resistant to MPTP neurotoxicity than those from wild-type controls. This suggests that
NADPH oxidase may also be a promising target for therapeutic interventions in PD (Gao
et al. 2003b; Gao et al. 2003a).
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Microglial cells also produce cytokines which are important in the inflammation process.
Studies of neuronal/glial mesencephalic cell cultures treated with MPP+ not only induced
the proliferation of microglial cells, the inflammatory cytokine granulocyte macrophage
colony-stimulating factor (GM-CSF), which could be produced by glial cells contributing
to microgliosis (Henze et al. 2005). On the other hand, some cytokines have been shown
to be protective such as transforming growth factor-beta 1 (TGF-beta) (Krieglstein and
Unsicker 1994; Akaneya et al. 1995a, b; Krieglstein et al. 1995b; Krieglstein et al. 1995c;
Henze et al. 2005) in DAergic neurons against MPP+ toxicity. IL-6 is a major
immunomodulatory cytokine with neuroprotective activity. The absence of IL-6 resulted
in increased vulnerability of DAergic neurons to MPTP, and a compromised reactive
microgliosis with significantly greater amounts of the pro-inflammatory cytokines IL-1α,
IL-1β and TNF-α, IL-10 and IL-12 (Bolin et al. 2005). Taken together, these data
strongly suggest that microglial cells can participate in the death of DA neurons via the
production of pro-inflammatory cytokines.
3.5.1.6 Cell Death
Apoptosis has been suggested as a mechanism of cell death in MPTP-induced
parkinsonism, since MPP+ induces apoptosis in cultured cerebellar granular neurons
(Dipasquale et al. 1991), in PC12 cells (Hartley et al. 1994) and in rat ventral
mesencephalic-striatal co-culture (Mochizuki et al. 1994). The syntheses of 1-amino-4-
phenyl-1,2,3,6-tetrahydropyridine (APTP) and 1-amino-4-phenyl-pyridinium salt
(APP+), the 1-amino analogues of the DAergic neurotoxins, MPTP and MPP+,
respectively, have also demonstrated that APTP and APP+ are both cytotoxic to PC12
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cells inducing apoptotic cell death (Noda et al. 2004). The major contributors to DAergic
cell death mainly involve ROS cell death pathways such as caspase-3 activation post
MPP+ treatment (Du et al. 1997; Leist et al. 1998; Shimoke et al. 2003). Another regular
marker for apoptosis is TUNEL. Evidence for TUNEL-positive DAergic cells after
application of MPP+ in cell culture has been demonstrated repeatedly (Dodel et al. 1998)
while denied by other researchers (Mochizuki et al. 1997).
Apoptosis is the mode of cell death induced in SH-SY5Y cells by MPTP, and this was
confirmed with nick end labeling and bisbenzimide staining (Sheehan et al. 1997). The
human neuroblastoma cell line SH-SY5Y possesses many of the qualities of human
neurons and, as such, has served as a model. Analysis by electronic microscopy revealed
ultrastructural changes consistent with the morphological characteristics of apoptosis
(Sheehan et al. 1997). These changes included plasma membrane blebbing, altered
cytosolic density, nuclear condensation and fragmentation, pronounced vacuole
formation, ribosomal dispersion, and the disappearance of the Golgi complex,
microtubules, and smooth endoplasmic reticulum. The in vitro induction of apoptosis by
a parkinsonian neurotoxin might be reflective of the mechanisms of in vivo nigral
degeneration occurring during PD (Ba et al. 2003).
Caspase-3 activation following DAergic cell death there are also evidence of time-
dependent increases in ROS generation, cytochrome c release, and also caspase-9
activation (Kaul et al. 2003). The caspase-3 inhibitor Z-DEVD-fmk effectively blocked
MPP+-induced PKC delta cleavage and kinase activity, suggesting that the proteolytic
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activation is caspase-3 mediated. Similar results were seen in MPP+-treated rat midbrain
slices. Z-DEVD-fmk and the PKC delta specific inhibitor rottlerin almost completely
blocked MPP+-induced DNA fragmentation. The SOD mimetic, MnTBAP also
effectively attenuated MPP+-induced caspase-3 activation, PKC delta cleavage, and DNA
fragmentation. Intracellular delivery of catalytically active recombinant PKC delta
significantly increased caspase-3 activity, further indicating that PKC delta regulates
caspase-3 activity. Finally, over-expression of a kinase inactive PKC delta K376R mutant
prevents MPP+-induced caspase activation and DNA fragmentation, confirming the pro-
apoptotic function of PKC delta in DAergic cell death (Kaul et al. 2003).
Another pathway involved in DAergic cell death is the stress-activated protein kinase
(SAPK) cascade. Recent in vitro and in vivo evidence has directly implicated this kinase
in the death of DAergic nigral neurons in the MPTP model of PD. MPP+ specifically
induces apoptotic changes in nuclear morphology in TH+ve (DAergic) cells and a nuclear
phospho-c-Jun-ir (a key transcription factor substrate of SAPK). This indicates a role for
the SAPK/JNK pathway including its c-Jun transcriptional target in the apoptotic death of
DAergic nigral neurons in the MPTP model of PD (Chun et al. 2001; Gearan et al. 2001;
Xia et al. 2001). Inhibition of the proapoptotic c-Jun N-terminal kinase (JNK) signaling
cascade blocked JNK, c-Jun, and caspase activation, the death of DAergic neurons, and
the loss of catecholamines in the ST in mice (Xia et al. 2001).
3.5.1.7 DA Agonists & Antioxidants
Anti-parkinsonian agents, pramipexole (PPX) and ropinirole (ROP) have been reported to
possess neuroprotective properties, both in vitro and in vivo. Incubation of primary
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mesencephalic cultures with PPX and ROP or the conditioned medium from PPX- or
ROP-treated primary cultures induced a marked increase in the number of DAergic
neurons in the cultures (Du et al. 2005). Similar effects can be observed after incubating
with the conditioned medium derived from PPX- and ROP-treated SN astroglia.
Meanwhile, PPX and ROP can protect the primary cells from the MPP+ insult.
Furthermore, the neurotrophic effects of PPX and ROP on mesencephalic DAergic
neurons could be significantly blocked by D3 receptor antagonist, but not by D2 receptor
antagonist (Du et al. 2005).
Most antioxidants and DA agonists provide neuroprotection by blocking DA depletion in
the ST such as minocycline, a semisynthetic tetracycline (Du et al. 2001) and/or
reduce/inhibit ROS production. OS has been implicated in the selective degeneration of
DAergic neurons in PD in particular as a mechanism of MPTP induced neuronal death.
However not all antioxidants and DA agonists can provide such protection. Apoptosis
induced by MPP+ in PC12 cells was unable to be rescued by the antioxidant drugs N-
acetylcysteine (NAC) and ascorbic acid (AA) or DA agonist deprenyl (Desole et al. 1996;
Desole et al. 1997a) again confirming that apoptosis is an important mechanism of cell
death in MPTP. There are still potential therapeutic agents such as the bioenergetic
coenzyme Q (10) (CoQ (10)) which has beneficial effects on TH and complexes I and II
of the respiratory chain in striatal slice cultures induced by MPP+ (Gille et al. 2004).
The neuroprotective effects of anti-parkinsonian agents are suggested to be associated
with marked reductions in inducible NO synthase (iNOS) and caspase 1 expression (Du
133
et al. 2001) since Peroxynitrite anion, an active metabolite of NO has been shown to
cause significant cytotoxic effects against DAergic neurons (Sawada et al. 1996). But
MPP+-induced neurotoxicity by NO is inhibited only in the presence of glia when treated
with minocycline (Du et al. 2001) pointing out the involvement of glial/astrocytic cells
presence contributing to cell death (McNaught and Jenner 1999).
MPP+ and other mitochondrial inhibitors (e.g., rotenone) induce apoptosis in vitro is
believed to be via stimulation of autocrine excitotoxicity (Leist et al. 1998). Cell death,
increase in intracellular Ca2+ concentration, release of cytochrome c, and all biochemical
and morphological signs of apoptosis were prevented by blockade of the N-methyl-D-
aspartate receptor (NMDA) with noncompetitive, glycine-site or glutamate-site
inhibitors. In addition, MPP+-induced apoptosis was reduced by high Mg2+
concentrations in the medium or by exocytosis with clostridial neurotoxins (Leist et al.
1998). Many anti-excitotoxic agents have been shown to be neuroprotective by inhibiting
blocking NMDA receptors and/or release of excessive glutamate such as memantine
(Volbracht et al. 2006), IV cAMP phosphodiesterase inhibitor, Ro20-1724 (Dickie and
Greenfield 1997), NMDA/glutamate antagonist MK-801 (McNaught and Jenner 1999;
Shimizu et al. 2003b).
Antioxidant enzymes such as SOD and glutathione peroxidase (GSHPx) have also been
shown to be affected and the target area for therapeutic intervention. Following MPTP
treatment, a decreased GSHPx activity was observed in both SN and ST, while
erythropoietin (Epo) restored nigral GSHPx activity decreased by MPTP, SOD enzyme
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activity was not significantly changed by MPTP and Epo treatment. Furthermore, Epo
stimulated astroglial GSHPx production in astroglial cell culture (Genc et al. 2002). SOD
treatment has been shown to be neuroprotective against MPP+ exposure (Gonzalez-Polo
et al. 2004), pretreatment of cultures with EUK-134 (a SOD and catalase mimetic)
completely protected DAergic neurons against MPP+-induced neurotoxicity and
prevented MPP+-induced nitration of tyrosine residues in TH (Pong et al. 2000).
Tetrahydrobiopterin, an essential cofactor for TH, may also act as an antioxidant in
DAergic neurons and protect against the toxic consequences of glutathione depletion. The
increasing intracellular tetrahydrobiopterin levels may protect against OS by complex-I
inhibition (Madsen et al. 2003). Dextromethorphan (DM), a widely used anti-tussive
agent, attenuated endotoxin-induced DAergic neurodegeneration in vitro. The
neuroprotective effect of DM in mesencephalic neuron-glia cultures significantly reduced
the MPTP-induced production of both extracellular superoxide free radicals and
intracellular ROS. More importantly, due to its proven safety record of long-term clinical
use in humans, DM may be a promising agent for the treatment of degenerative
neurological disorders such as PD (Zhang et al. 2004).
Studies have also focused on the potential of growth factors as neuroprotective and/or
neuroregenerative therapeutic agents for PD. Treatment with nerve growth factor (NGF)
promote acute cell proliferation when exposed with MPP+ via the sustained extracellular
signal-regulated kinases (ERKs) and the p38 MAPK pathway in PC12 cells (Shimoke
and Chiba 2001; Shimoke and Kudo 2002). Another growth factor that has been focused
lately is glial cell line-derived neurotrophic factor (GDNF). Exposure to MPP+ was
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found to be also toxic for hESC-derived DAergic neurons (Human embryonic stem cells
can proliferate indefinitely yet also differentiate in vitro, allowing normal human neurons
to be generated in unlimited numbers). Treatment with (GDNF) protected TH+ve neurons
against MPP+-induced apoptotic cell death and loss of neuronal processes as well as
against the formation of intracellular ROS (Zeng et al. 2006). Moreover, we found that
the levels of glial cell line-derived neurotrophic factor (GDNF) and brain-derived
neurotrophic factor (BDNF) in the conditioned medium of mesencephalic cultures treated
with PPX and ROP were significantly increased. Blocking GDNF and BDNF with the
neutralizing antibodies, the neurotrophic effects of PPX and ROP were greatly
diminished. These results suggest that D3 dopamine receptor-preferring agonists, PPX
and ROP, exert neurotrophic effects on cultured DA neurons by modulating the
production of endogenous GDNF and BDNF, which may participate in their
neuroprotection (Du et al. 2005).
3.5.2 ROTENONE & IN VITRO STUDIES
Similar to MPP+, rotenone uptake into brain synaptosomes is similar among primates and
rodents, and hence is used in neuronal cultures. Cultures of dissociated mesencephalic
neurons from fetal rats and DA-neuron-derived cell lines such as human SH-SY5Y and
rat PC12 (see Table 3) is suitable for studying mechanisms of DAergic neuronal
degeneration and for screening new pharmacological agents. Rotenone toxicity is belived
to be a result from OS generated during DA metabolism and by mitochondrial
respiration. In addition, progressive depletion of glutathione, oxidative damage to
proteins and DNA, release of cytochrome c from mitochondria to cytosol, activation of
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caspase 3, mitochondrial depolarization, and eventually apoptosis (Greenamyre et al.
2001).
Table 3 Summary of Experimental PD In Vitro models
In Vitro model
References
Primary Embryonic/ Postnatal mouse mesencephalon
(Danias et al. 1989; Lotharius and O'Malley 2001; Gao et al. 2002; Ahmadi et al. 2003; Gao et al. 2003b; Gao et al. 2003a; Gille et al. 2004; Diaz-Corrales et al. 2005; Grammatopoulos et al. 2005; Radad et al. 2006)
PC12 cell lines (Basma et al. 1992; Hartley et al. 1994; Seaton et al. 1997, 1998; Bal-Price and Brown 2000; Lamensdorf et al. 2000b; Seyfried et al. 2000; Taylor et al. 2000; Ishiguro et al. 2001; Dargusch and Schubert 2002; Zheng et al. 2002; Lee et al. 2005; Wang et al. 2005; Bal-Price et al. 2006; Hirata et al. 2006; Hirata and Kiuchi 2007; Marella et al. 2007; Tan et al. 2007; Yim et al. 2007)
Rat VM slices (Seo et al. 2002; Bywood and Johnson 2003; Sherer et al. 2003b; Sherer et al. 2003a; Tai et al. 2003; Xu et al. 2003; Gille et al. 2004; Yang et al. 2004b; Dukes et al. 2005; Hirata and Nagatsu 2005; Kress and Reynolds 2005; Liu et al. 2005; Sousa and Castilho 2005; Testa et al. 2005; Centonze et al. 2006)
Mouse MN9D cell line (Seo et al. 2002; Kweon et al. 2004; Cao et al. 2007) SH-SY-5Y cells (Seaton et al. 1997; Lamensdorf et al. 2000b; Lamensdorf et al.
2000a; Imamura et al. 2006; Klintworth et al. 2007)
Synaptosomes (Bougria et al. 1995; Fonck and Baudry 2003)
3.5.2.1 DAergic Toxicity
Rotenone treatment induces a dose- and time-dependent destruction of SNpc neuron
processes, morphologic changes, some neuronal loss, and decreased TH protein levels.
Chronic complex I inhibition also caused oxidative damage to proteins, measured by
protein carbonyl levels. This oxidative damage was blocked by the antioxidant alpha-
tocopherol (vitamin E). At the same time, alpha-tocopherol also blocked rotenone-
induced reductions in TH protein and TH immunohistochemical changes (Testa et al.
2005). The number of TH+ve neurons was shown to be reducted by 50 +/- 6%
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[Grammatopoulos TN], along with a reduction in DA, dihydroxyphenyl acetic acid
(DOPAC) and homovanillic acid (HVA) levels in PC12 cells, DOPA formation with an
accompanying decrease in ATP and increase in lactate in rat striatal slices (Hirata and
Nagatsu 2005).
Decreases in ATP levels, changes in catechol levels, and increased DA oxidation is the
general outcome of rotenone treated neurons. Whether endogenous DA makes PC12 cells
more susceptible to rotenone is still unknown. Cells treated with the TH inhibitor alpha-
methyl-p-tyrosine (AMPT) to reduce DA levels prior to rotenone exposure revealed no
changes in rotenone-induced toxicity with or without AMPT treatment. However, a
potentiation of toxicity was observed following coexposure of PC12 cells to rotenone and
methamphetamine. PC12 cells were depleted of DA prior to methamphetamine and
rotenone cotreatment to determine whether this effect was due to DA, resulted in a large
attenuation in toxicity. These findings suggest that DA plays a role in rotenone-induced
toxicity and possibly the vulnerability of DA neurons in PD (Dukes et al. 2005).
Cell death and reduced DA release has been observed in PC12 cells induced by rotenone
(Yang et al. 2004b), this response was abolished by removal of extracellular Ca2+.
Mitochondrial inhibitors and uncouplers evoke catecholamine secretion from PC12 cells
which is wholly dependent on Ca2+ influx through voltage-gated Ca2+ channels (Taylor et
al. 2000). Rotenone also induces a rapid accumulation of DOPAL and DOPET in the
medium of cultured PC12 cells but is decreased by MPP+. 3,4-
Dihydroxyphenylacetaldehyde (DOPAL) is a toxic metabolite formed by the oxidative
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deamination of DA. This aldehyde is mainly oxidized to 3,4-dihydroxyphenylacetic acid
(DOPAC) by aldehyde dehydrogenase (ALDH), but is also partly reduced to 3, 4-
dihydroxyphenylethanol (DOPET) by aldehyde or aldose reductase (ARs) combined
inhibition of ALDH and ARs potentiated rotenone-induced toxicity (Lamensdorf et al.
2000a).
The neurotoxic vulnerabilities between DAergic subgroups have also been studied
Rotenone induced severe dendrite loss among SN DAergic neurons, whereas
hypothalamic A11 DAergic neurons were spared. Adjacent sections that were
immunolabeled for calbindin or stained with cresyl violet also revealed a striking
dendritic degeneration of SN neurons in rotenone-exposed slices, whereas
noncatecholamine neurons, such as those in the perifornical nucleus were more resistant
(Bywood and Johnson 2003).
3.5.2.2 Oxidative Stress (OS)
One possible result of complex I inhibition is increased formation of ROS, creating
oxidative damage within the cell. Oxidative damage, rather than a bioenergetic defect, is
also seen in the in vivo rotenone model (Sherer et al. 2003b; Sherer et al. 2003a;
Bashkatova et al. 2004). PC12 cells treated with rotenone causes an apoptotic cell death
and elevated intracellular ROS and lactic acid accumulation (Lotharius and O'Malley
2001; Dargusch and Schubert 2002; Liu et al. 2005; Radad et al. 2006), the formation of
ROS and the depletion of GSH in differentiated PC12 cells (Sousa and Castilho 2005;
Kim et al. 2007).
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Mitochondrial complex I inhibitor rotenone induces apoptosis through enhancing
mitochondrial ROS production. Recently, it has been shown that fraxetin (coumarin) and
myricetin (flavonoid) have significant neuroprotective effects against apoptosis induced
by rotenone, increases the total glutathione levels in vitro, and inhibit lipid peroxidation.,
such as Mn and CuZn superoxide dismutase (MnSOD, CuZnSOD), catalase, glutathione
reductase (GR), and glutathione peroxidase (GPx) on rotenone neurotoxicity in
neuroblastoma cells. N-acetylcysteine (NAC), a potent antioxidant, was employed as a
comparative agent. SH-SY5Y-rotenone treated cells significantly increased the
expression and activity of MnSOD, GPx and catalase (endogenous antioxidant defense
system). Cells preincubated with fraxetin resulted in a decrease in the protein levels and
activity of both MnSOD and catalase, in comparison with the rotenone treatment.
Activity and expression of GPx were increased by rotenone and pre-treatment with
fraxetin did not modify those levels significantly. This suggests that rotenone-induced
neurotoxicity is partially mediated by free radical formation and OS, and that it can be
partially protected by fraxetin providing the main protection system of the cells against
oxidative injury (Molina-Jimenez et al. 2005).
The effect of the complex I-inhibitor rotenone on glutathione redox status and the
generation of reactive oxygen intermediates (ROI) in rat pheochromocytoma PC12 cells
induce a time-dependent loss of GSH, whereas treatment with lower concentrations of
rotenone increases cellular GSH. Both MPP+ and rotenone increase cellular levels of
oxidised glutathione (GSSG) and the higher concentrations of both compounds lead to an
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elevated ratio of oxidised glutathione (GSSG) vs total glutathione (GSH + GSSG)
indicating a shift in cellular redox balance. However MPP+ or rotenone does not induce
the generation of ROI or significant elevation of intracellular levels of thiobabituric acid
reactive substances (TBARS) for up to 48 h (Seyfried et al. 2000).
3.5.2.3 αααα-Synuclein
In vitro, rotenone-induced α-synuclein aggregation (Lee et al. 2002) is correlated with
elevations in insoluble protein carbonyls (Lee et al. 2002; Sherer et al. 2002b). In
addition, in vivo rotenone infusion results in selective upregulation of α-synuclein in the
substantia nigra and the increased occurrence of higher-molecular-weight, aggregated
forms of α-synuclein (Betarbet et al. 2000; Betarbet et al. 2002a; Hoglinger et al. 2003a).
An elevation in insoluble protein carbonyls in the midbrains of rotenone-treated animals
suggests that rotenone treatment may cause oxidative modifications to α-synuclein that
make it more likely to aggregate (Giasson et al. 2000; Krishnan et al. 2003). On the other
hand, elevated α-synuclein expression can itself cause oxidative damage and render cells
more vulnerable to oxidative insults (Hsu et al. 2000; Kanda et al. 2000; Ko et al. 2000).
Aggregation of gamma-tubulin protein, which is a component of the centrosome matrix
and recently identified in LBs of PD, was observed in primary cultures of mesencephalic
cells treated with rotenone. Rotenone-treated neurons and astrocytes showed enlarged and
multiple centrosomes. These centrosomes also displayed multiple aggregates of α-
synuclein protein. Neurons with disorganized centrosomes exhibited neurite retraction
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and microtubule destabilization, and astrocytes showed disturbances of mitotic spindles
(Diaz-Corrales et al. 2005).
3.5.2.4 Complex I Inhibitor
Complex I inhibition has several potential functional consequences, including decreased
ATP production, altered calcium handling, and oxidative damage. Complex I is the first
of four multisubunit protein complexes in the mitochondrial electron transport chain
responsible for converting energy from cellular metabolism into the proton gradient used
by complex V to generate ATP. Electron transport chain dsyfunction could lead to loss of
the proton gradient, impaired ATP production, and a bioenergetic defect. However, in
dissociated culture systems, rotenone does not kill cells via ATP depletion instead
rotenone increases OS (Zhang et al. 2001; Sherer et al. 2003b).
Complex I inhibition provokes the following events: 1) activation of specific kinase
pathways; 2) release of mitochondrial proapoptotic factors, apoptosis inducing factor, and
endonuclease G. AS601245, a kinase inhibitor, exhibited significant protection against
these apoptotic events. The traditional caspase pathway does not seem to be involved
because caspase 3activation was not observed. This suggests that overproduction of ROS
caused by complex I inhibition is responsible for triggering the kinase activation
(Ishiguro et al. 2001; Lee et al. 2005; Bal-Price et al. 2006; Hirata et al. 2006; Ito et al.
2006; Radad et al. 2006; Hirata and Kiuchi 2007; Marella et al. 2007; Tan et al. 2007;
Yim et al. 2007).
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Rotenone causes the loss of mitochondrial membrane potential, released cytochrome c
into the cytosol, and reduced cytochrome c content in mitochondria addition of bHB
blocked this toxic effect and also attenuated the rotenone-induced activation of caspase-9
and caspase-3 (Imamura et al. 2006). Numerous studies also suggest that dysfunction of
mitochondrial proton-translocating NADH-ubiquinone oxidoreductase (complex I) is
associated with neurodegenerative disorders, such as PD and Huntington's disease. It was
previously shown that the single-subunit NADH dehydrogenase of Saccharomyces
cerevisiae (Ndi1P) can work as a replacement for complex I in mammalian cells such as
DAergic cell lines rat PC12 and mouse MN9D. The cells expressing the Ndi1 protein
were resistant to rotenone (Seo et al. 2002).
3.5.2.5 Inflammation
Increasing evidence suggests an important role for environmental toxins such as
pesticides in the pathogenesis of PD. Rotenone-induced DAergic neurodegeneration has
been associated with both its inhibition of neuronal mitochondrial complex I and the
enhancement of activated microglia. Previous studies with NADPH oxidase inhibitors,
diphenylene iodonium and apocynin suggest that NADPH oxidase-derived superoxide
might be a major factor in mediating the microglia-enhanced rotenone neurotoxicity.
However, because of the relatively low specificity of these inhibitors, the exact source of
superoxide induced by rotenone remains to be determined. Primary mesencephalic
cultures from NADPH oxidase--null (gp91phox-/-) or wild-type (gp91phox+/+) mice,
demonstrate the critical role for microglial NADPH oxidase in mediating microglia-
enhanced rotenone neurotoxicity. In neuron--glia cultures, DAergic neurons from
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gp91phox-/- mice were more resistant to rotenone neurotoxicity than those from
gp91phox+/+ mice. However, in neuron-enriched cultures, the neurotoxicity of rotenone
was not different between the two types of mice. These results show that the greatly
enhanced neurotoxicity of rotenone was attributed to the release of NADPH oxidase-
derived superoxide from activated microglia (Gao et al. 2003a).
Observations of rotenone and the inflammogen lipopolysaccharide (LPS) synergistically
induced DAergic neurodegeneration. The synergistic neurotoxicity of rotenone and LPS
is observed when the two agents were applied either simultaneously or in tandem.
Mechanistically, microglial NADPH oxidase-mediated generation of ROS appeared to be
a key contributor to the synergistic DAergic neurotoxicity. This conclusion is based on
the facts that inhibition of NADPH oxidase or scavenging of free radicals induced
significant neuroprotection. Finally, rotenone and LPS failed to induce the synergistic
neurotoxicity as well as the production of superoxide in cultures from NADPH oxidase-
deficient animals. This is the first demonstration that low concentrations of pesticide and
an inflammogen work in synergy to induce a selective degeneration of DAergic neurons
(Gao et al. 2003b).
Neuron-enriched and neuron/glia cultures from the rat mesencephalon, has been used to
study the role of microglia in rotenone-induced neurodegeneration. Significant and
selective DAergic neurodegeneration was observed in neuron/glia cultures 2d after
treatment with rotenone. The greatly enhanced neurodegenerative ability of rotenone was
attributed to the presence of glia, especially microglia, because the addition of microglia
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to neuron-enriched cultures markedly increased their susceptibility to rotenone.
Mechanistically, rotenone stimulated the generation of superoxide from microglia that
was attenuated by inhibitors of NADPH oxidase. Furthermore, inhibition of NADPH
oxidase or scavenging of superoxide significantly reduced the rotenone-induced
neurotoxicity (Gao et al. 2002).
3.5.2.6 Cell death via Apoptosis
Two cell models, human cultured cells HL-60 and BJAB, have shown that the exposure
of cells to rotenone induces the generation of H2O2, which leads to significant changes in
the mitochondrial membrane potential, and which is accompanied by the fragmentation
of internucleosomal DNA and the formation of DNA-ladders (Tada-Oikawa et al. 2003).
Furthermore, caspase-3 activity increases in rotenone-treated HL-60 cells in a time-
dependent manner. These apoptotic events are delayed in HP100 cells, an H2O2-resistant
clone of HL-60, confirming the involvement of H2O2 in apoptosis. In the same study, the
expression of anti-apoptotic protein, Bcl-2, in BJAB cells, was shown dramatically to
inhibit rotenone-induced changes in the mitochondrial membrane potential and the
formation of DNA ladders, confirming the involvement of mitochondrial dysfunction in
apoptosis (Tada-Oikawa et al. 2003).
Nanomolar concentrations of rotenone are also able to induce caspase-3-mediated
apoptosis in primary DA neurons in VM cultures from E15 rats (Ahmadi et al. 2003;
Hirata et al. 2006). After 11h of exposure to 30 nM rotenone, the number of DA neurons
identified by TH immunostaining decreases rapidly, and only 20% of the neurons
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survive. On the other hand, more than 70% of total neurons survive rotenone treatment,
implying that DAergic neurons (TH+-neurons) are more sensitive to rotenone (Ahmadi et
al. 2003). Rotenone significantly increases the number of apoptotic TH+-neurons
(Hartley et al. 1994; Seaton et al. 1997, 1998; Radad et al. 2006; Hirata and Kiuchi 2007;
Tan et al. 2007), which is correlated with an increase in immunoreactivity for active
caspase-3 in DAergic neurons (Seaton et al. 1997; Lee et al. 2005; Wang et al. 2005;
Hirata et al. 2006; Ito et al. 2006). Furthermore, the caspase-3 inhibitor, DEVD, rescues
a significant number of TH+-neurons from rotenone-induced death. Thus, at low
(nanomolar) concentrations, rotenone is able to induce caspase-3-mediated apoptosis in
primary DA neuron cultures, with TH+-neurons being more sensitive to rotenone-induced
toxicity than the total VM cell population (Ahmadi et al. 2003). This immediately raises
a question concerning the mechanisms involved in the selectivity of dopaminergic
neuronal death by rotenone.
Nitric oxide (NO) can also trigger either necrotic or apoptotic cell death. A 24-h
incubation of PC12 cells with NO donors or specific inhibitors of mitochondrial
respiration rotenone, in the absence of glucose, caused ATP depletion and resulted in 80-
100% necrosis. Glucose addition almost completely prevented the decrease in ATP level
and the increase in necrosis induced by the NO donors or mitochondrial inhibitors. This
suggests that the NO-induced necrosis in the absence of glucose was due to the inhibition
of mitochondrial respiration and subsequent ATP depletion. However, in the presence of
glucose, NO donors and mitochondrial inhibitors induced apoptosis of PC12 cells as
determined by nuclear morphology. The presence of apoptotic cells is prevented
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completely by benzyloxycarbonyl-Val-Ala-fluoromethyl ketone (a nonspecific caspase
inhibitor), this shows apoptosis was mediated by caspase activation. Indeed, both NO
donors and mitochondrial inhibitors in PC12 cells caused the activation of caspase-3- and
caspase-3-processing-like proteases. Caspase-1 activity was not activated. Cyclosporin A
(an inhibitor of the mitochondrial permeability transition pore) decreased the activity of
caspase-3- and caspase-3-processing-like proteases after treatment with NO donors, but
was not effective in the case of the mitochondrial inhibitors. The activation of caspases
was accompanied by the release of cytochrome c from mitochondria into the cytosol,
which was partially prevented by cyclosporin A in the case of NO donors. These results
show that NO donors may trigger apoptosis in PC12 cells partially mediated by opening
the mitochondrial permeability transition pores, release of cytochrome c, and subsequent
caspase activation (Bal-Price and Brown 2000).
Rotenone exposure caused apoptosis in SH-SY5Y cells but not in PC12 cells. This
selective ability of paraquat and rotenone to induce apoptosis in different cell lines
correlates with their ability to activate c-Jun N-terminal protein kinase (JNK) and p38
mitogen-activated protein kinases. Furthermore, JNK and p38 are required for rotenone-
induced apoptosis in SH-SY5Y cells (Newhouse et al. 2004) as well as primary neurons
(Klintworth et al. 2007). Along with nuclear damage, the changes in the mitochondrial
membrane permeability, leading to the cytochrome c release and caspase-3 activation, the
formation of reactive oxygen species and the depletion of GSH in differentiated PC12
cells was induced by rotenone (Sousa and Castilho 2005; Kim et al. 2007).
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3.5.2.7 DA Agonists and Antioxidants
Markers of DAergic neurons are more altered than those of �-aminobutyric acid (GABA)
neurons in PD-toxin models, suggesting greater susceptibility of DAergic neurons. Thus
DA agonists are employed to replace the DA lost. Intracellular DA depletion with
reserpine significantly attenuated rotenone-induced ROS accumulation and apoptotic cell
death but not with deprenyl (Lotharius and O'Malley 2001; Dargusch and Schubert 2002;
Liu et al. 2005; Radad et al. 2006). Another DA agonist which has been shown to
possess anti-apoptotic actions is pramipexole, which also reduces caspase-3 activation,
decreases the release of cytochrome c and prevents the fall in the mitochondrial
membrane potential induced by rotenone (Schapira 2002a, b; Gu et al. 2004). The
oxidative damage and DAergic neuronal loss caused by rotenone were shown to be
blocked by alpha-tocopherol (Seyfried et al. 2000; Sherer et al. 2003b).
3.6 ORGANOTYPIC CELL CULTURE: IN VITRO MODEL FOR PD
The ability to conduct morphological assessments of ventral mesencephalic (VM) and
striatum co-cultures provides support for the use of the organotypic co-culture model of
the basal ganglia, given that DA neurons present in the VM will project axons to an
adjacent cultured striatal tissue slice (Snyder-Keller et al. 2001; Gates et al. 2004) and
form functional synapses (Tseng et al. 2007). Further analyses have shown that even the
complex patch/matrix organization of the striatal tissue is maintained when donor tissue
of VM and ST is age-appropriate (Snyder-Keller et al. 2001). These studies suggest that
it is important to use VM tissue in which the dopamine (DA) neurons have differentiated,
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yet have not begun to project DA axons/terminals to the striatum; in this way, the likely
degeneration incurred with axotomy during tissue dissection can be limited.
PD research has utilized VM organotypic cultures, containing the DA neurons affected in
PD, in the study of tissue explants for transplantation purposes, as well as to assess
potential DA neurotoxicants. Researchers have used the VM cultured alone (Dickie et al.
1996; Hoglinger et al. 1998; Meyer et al. 2001; Shimizu et al. 2003a), the VM in co-
culture with ST (the target of VM DA neuron terminals) (Schatz et al. 1999; Katsuki et
al. 2001; Snyder-Keller et al. 2001; Kotake et al. 2003; Gates et al. 2004), and the VM,
striatum, and cortex in triple cultures to model the BG DAergic system (Plenz and Kitai
1996; Snyder-Keller et al. 2001). However, to date, studies have not adequately
described the DA development, in terms of neurochemical and protein analyses, of the
VM and striatum in co-culture (Lyng et al. 2007). Here, we have used organotypic co-
cultures of P5-6 VM and P5-6 rat ST in the roller tube system with slight modifications
(Stoppini et al. 1991; Snyder-Keller et al. 2001) to model the developing nigrostriatal DA
system.
3.6.1 MPTP application in organotypic cell cultures
Using midbrain slice cultures a few studies have demonstrated MPP+ to be a complex I
inhibitor (Madsen et al. 2003; Xu et al. 2003; Gille et al. 2004), specifically toxic to
DAergic neurons by reduction in DA levels, its metabolites and tyrosine hydroxylase-
positive cells in a dose-dependent manner (Reinhardt 1993; Madsen et al. 2003; Shimizu
et al. 2003b; Shimizu et al. 2003a; Kress and Reynolds 2005). MPP+’s toxic action was
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attenuated by NMDA inhibitor MK801 (Kress and Reynolds 2005), DA agonists such as
L-Dopa, antioxidant and bioenergetic coenzyme Q(10) (CoQ(10)) had preserving
potential on the activities of TH, complexes I and II of the respiratory chain along wit the
survival rate of DAergic neurons (Gille et al. 2004). Cell death process is believed to
occur via apoptosis evident by the loss of cell bodies and the fragmentation of processes
along with nucleus shrinkage (Kress and Reynolds 2005). This cell death was prevented
by the inhibitors of NMDA, nitric oxide synthase (NOS), cycloheximide and caspase
cascade and also rescued by L-deprenyl and dopamine D2/3 agonists (Shimizu et al.
2003b; Shimizu et al. 2003a). The influence of target tissue was also examined on co-
cultures of DAergic neurons forming dense innervation to the striatal tissue. DAergic
neurons in midbrain--striatum co-cultures were more resistant to the cytotoxic actions of
NMDA and a nitric oxide donor NOC-18, than the same neuronal population in single
midbrain cultures. On the other hand, the toxicity of MPP+ was more prominent in
midbrain-ST co-cultures than that in single midbrain cultures (Katsuki et al. 1999). The
neurotrophic factor GDNF was also examined in this in vitro model showing that pre-
treatment and post-treatment with GDNF is important to obtain maximal protection
against MPP+ toxicity (Jakobsen et al. 2005).
3.6.2 Rotenone and Organotypic Cell Culture
To a lesser extent, the use of rotenone in both animal and cell cultures is fewer than
MPP+ and has mainly been studied in primary cell cultures deriving from fetal
mesencephalon. Only a few studies have employed rotenone in this culture system.
Similar to MPP+, it is observed to be a complex I inhibitor and specific DAergic toxin
150
leading to a dose- and time-dependent destruction of SNpc neuron processes,
morphologic changes, neuronal loss, and decreased TH protein levels (Xu et al. 2003;
Kress and Reynolds 2005; Testa et al. 2005). Chronic complex I inhibition also caused
oxidative damage to proteins, measured by protein carbonyl levels. This oxidative
damage was blocked by the antioxidant alpha-tocopherol (vitamin E). At the same time,
alpha-tocopherol also blocked rotenone-induced reductions in TH protein and TH
immunohistochemical changes (Testa et al. 2005). Addition of L-Dopa, DA agonists and
antioxidants such as CoenzymeQ were able to counteract rotenone’s toxicity (Gille et al.
2004).
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4.0 AIMS OF STUDY
1) To grow and maintain the development of dopaminergic neurons in vitro using the
organotypic slice culture technique.
2) To establish a period of optimal dopaminergic neuronal growth for application of
neurotoxins.
2) To assess the effects of neurotoxins MPTP and rotenone on the survival of
dopaminergic neurons.
3) To examine the neuroprotective effects of neutrophic factor GDNF on dopaminergic
neurons with or without the toxic stress from the neurotoxins.
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CHAPTER 5:
GENERAL MATERIAL AND METHODS
153
5.1 ANIMAL CARE
5.1.1 Ethics
The experimental studies presented in this thesis were performed within the guidelines
established by the National Health and Medical Research Council (NH&MRC) and were
approved by the ethics committees of the Institue of Medical and Veterinary Science
(IMVS; 64A/04, 152A/07) and the University of Adelaide (M-64-2004) to minimize
animal suffering and to reduce the number of animals used.
5.1.2 General
Sprague-Dawley rats from postnatal day of 4 to 5 (P 4-5) were kept in a heated pad
insulated container until required.
5.2 ORGANOTYPIC SLICE CULTURES
All the culture procedures were carried out under sterile conditions, within a laminar flow
hood. Similarly, all equipment was sterilized in an oven for four hours at 180°C and
solutions were either sterile filtered or obtained pre-sterilized or autoclaved.
5.2.1 Dissection of the ventral mesencephalon and striatum
Organotypic slices were prepared from brains of pups 4-5 d of either sex according to the
procedure described by Stoppini (Gahwiler 1981; Stoppini et al. 1991) and Gahwiler
{Gahwiler, 1988 #27) with slight modifications. Briefly, pups were quickly decapitated;
small dissecting scissors were used to make a cut along the midline of the skull before the
skull was opened to expose the dorsal surface of the brain. A spatula was used to cut
154
through cranial nerves and ease the brain, dorsal surface downwards onto a sterile chilled
Petri dish. Following the rapid removal of the brain, placed on its dorsal side with the
ventral side up on a sterile glass cold Petri dish, under a dissecting microscope.
In order to dissect the striatum (ST), two cuts in the coronal plane were made; one
through the forebrain anterior to the level of the optic chiasm, the second through the
optic chiasm itself. The block was placed on its anterior cut surface and another two
horizontal cuts separated the striatum from the cerebral cortex and basal forebrain, whilst
two vertical cuts removed the septum and any remaining cortex from the striatum. Any
meninges attached to the tissue were removed with tweezers. The tissue block was
transferred to a McIlwain tissue sectioner where serial coronal sections of 300μm
thickness were cut. After transferring the ST to a petri dish containing cooled ringers
balanced salt solution (BSS; see appendices), the sections were gently teased apart. Up to
10 slices were obtained from each tissue block.
A similar procedure was used to dissect the ventral mesencephalon (VM). A coronal cut
was made through the posterior thalamus at the level of the mammilary bodies. A second
cut was made through the brain at the level of the pons in order to isolate the midbrain. A
horizontal cut removed the dorsal midbrain and cortical tissue from the ventral midbrain.
The tissue was then turned onto the dorsal cut surface and any attached meninges were
removed with fine tweezers before making a vertical midline cut to separate the two VM.
Then 200μm thick slices were cut. Up to ten cultures could be produced from each tissue
block. Finally, the sections were transferred to a solution of cooled ringers and the
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sections were teased apart under the dissecting microscope. Petri dishes with ringer’s
solution containing either SN or ST were then transferred into another sterile Petri dish
containing chilled Gey’s balanced salt solution (GBSS; JRH biosciences) and 0.1 % D-
glucose and 0.1% potassium chloride. The Petri dishes containing either ST or SN placed
in GBSS were placed at 4°C for 90min in a fridge before mounting.
5.2.2 Mounting the slices
The slices were mounted on glass coverslips (11cm x 22cm). The coverslips were
cleaned for fifteen minutes in a 1% solution of sodium thiosulphate and potassium
hexacyano ferrate, then boiled in distilled water for two hours, immersed in ethanol for
two days and sterilised.
A plasma clot was used to mount the tissue onto the cover slips. This involved
embedding the tissue in a plasma clot which was formed by mixing a solution of chicken
plasma with thrombin. A stock solution of chicken plasma was prepared from
lyophilised chicken plasma to which 5ml of sterile purified water was added. The
solution was centrifuged at 2500 revolutions per minute (RPM) for thirty minutes and the
supernatant was sterile filtered before being frozen in 1ml aliquots. The thrombin
solution was prepared from 0.1g of bovine thrombin by addition of 5ml of sterile purified
water. The solution was centrifuged for thirty minutes at 2500 RPM and the supernatant
was sterile filtered before being frozen in 200μl aliquots.
156
On the day of preparation of cultures, 150μl of the prepared thrombin solution was
diluted with 5ml of sterile GBSS. Both the chicken plasma (Sigma P-3266-5mL) and the
thrombin (Sigma T-4648-10KU) solutions were kept on ice. A 20μl drop of chicken
plasma solution was placed on one end of the pre-cleaned coverslip. A fine spatula was
then used to gently lift a section of ventral mesencephalon followed by the ST, from the
GBSS and into the drop. A 20μl drop of thrombin was then placed on the cover slip and
a spatula was used to mix the two solutions around the sections. Finally, the tissues were
carefully positioned approximately 1mm apart within the plasma clot which had formed.
5.2.3 Slice culture by the roller-tube technique
Each coverslip with the mounted cultures was placed in screw cap, diagonal bottomed
culture tube (NUNC TC tube; NC-156758, SI 110 X 6 mm) and 0.75ml of culture
medium was added to each tube. The culture medium contained 25% horse serum which
had been inactivated by heating at 56oC for thirty minutes, 50% Eagle minimum essential
medium and 25% Hanks BSS ([10 mM HEPES] IMVS; Infectious disease lab media
unit). The medium was also supplemented with d-glucose to a final concentration of
6.5mg/ml and L-glutamine to a final concentration of 0.15 mg/ml. New culture medium
was made each week. Penicillin and gentamycin were also added to the medium.
The culture tubes were placed in a roller drum which was tilted at an angle of 5o from the
horizontal axis to ensure that the cultures were immersed by the medium for about half of
the rotating cycle. The culture tubes were rotated at a rate of approximately ten
revolutions per hour. The roller drum was positioned within an incubator which kept the
157
cultures at a temperature of between 35.5 and 36.5oC. The culture medium was changed
twice a week by pouring away the old medium and adding 0.75ml fresh medium to each
tube. In order to control the number of non-neuronal cells, anti-mitotic drugs were added
to the medium for 24 hours on the third or fourth day in culture. The anti-mitotic
substances used were uridine and cytosine-β-D-arabino-furanoside, both at
concentrations of 10-3 M. Cultures were maintained for variable lengths of time, most
cases for 7 weeks.
Co-cultures included whole intact hemispheres of ventral mesencephalon containing SN
were plated 1mm apart from the ST on glass coverslips (LOMB Scientific) with 25 μl of
chicken plasma and 20 μl thrombin. Placed in a humidified chamber for 15 min to allow
the chicken plasma and thrombin to clot then into a flat-bottom culture tubes with screw
caps containing 1 ml medium [50% DMEM (IMVS; Infectious disease lab media unit),
25% Hanks balanced salt solution with 10 nM HEPES (IMVS; Infectious disease lab
media unit), 25% heat inactivated horse serum (JRH biosciences; 50120-100 M),
supplemented with glucose (1 ml 50% D-glucose to 100 ml of medium) and glutamine
(500 μl of 1 mM L-glutamine to 100 ml of medium), and 0.5% gentamycin and penicillin
(IMVS; Infectious disease lab media unit). The medium was changed twice a week.
After 4 to 6 days in vitro (DIV), co-cultures were treated with antimitotic agents 1 ml of
cytosine-β-D-arabinofuranoside (Sigma C-6645) and 1 ml of uridine (Sigma U-3750) to
100 ml of medium for 24 h to retard glial and fibroblast growth, then discarded and fresh
media was added. Cultures were maintained for variable lengths of time, most cases for 7
weeks, in a cell culture incubator at 35-36.5 °C in a roller drum at 5 revolutions per hour.
158
5.3 FIXATION OF CULTURES
Cultures were fixed at 1 week intervals with 4% paraformaldehye (pH 7.4) overnight or
minimum 4 hours at room tempertature. Then endogenous peroxidases were inactivated
with treatment with 0.5 % H2O2 (8.3 mL) plus 100% of methanol (500 mL) for 30 min.
Sections were the rinsed twice (5 min) in fresh PBS (pH 7.4) 0.3 % Triton X-100 (Sigma
T9284-500 mL).
5.3.1 Immunoperoxidase antigen retrieval
For antigen retrieval, the cultures on coverslips or 7μM sections of paraffin embedded
tissue were placed in coplin glass staining jars or slide rack (respectively) (Proscitech;
L056) containing citrate solution (see appendices) with lid screwed slightly, the coplin
jars are then placed within a kartel slide dish filled with distilled water (approximately
250 ml) and lid placed on, then microwave as follows. Place side dishes in Panasonic
microwave oven (Model NE-1037) press 0 the start button. Microwave the coverslips
until they start to boil. Once boiled remove the boiled dishes and place in a second
microwave oven (NEC). If there is 2 dishes select timer fro 10 min and power level at 2,
for 3 dishes select timer for 10 min and power level at 3. Once microwaving is complete
remove dishes from microwave take lids off and allow 30 min to cool (approximately
below 40°C). Cultures are then washed twice (5 min) fresh 0.3% Triton X-100 PBS.
5.3.2 Immunohistochemistry
Antisera: The following mono- and polyclonal antibodies were used: rabbit polyclonal
anti-tyrosine hydroxylase (TH) antibody (1:2000; Chemicon AB151), rabbit monoclonal
159
anti-glial fibrillary acidic protein (GFAP) antibody (DAKO, Z0334; 1:20 000), mouse
polyclonal anti- Neu-N (1:4000; Chemicon MAB 377). Antigen retrieval was mainly
citrate unless primary antibodies were Caspase -3 in which EDTA solution was used
instead. Secondary antibodies were Vector Biotinylated anti-mouse IgG (BA-2000) or
anti-rabbit IgG (BA-1000) at 1/250. Followed by the 3° step where the Pierce peroxidase
conjugated streptavidin tertiary (21127) at 1/1000 (is also referred to as SPC).
Paraffin sections: Following immunoperoxidase antigen retrieval procedure the cultures
were blocked with normal donor horse serum [1 ml to 30 ml 0.3% Triton X-100 PBS
(CSL Biosciences 09532301)] for 30 min at room temperature then incubated with
primary antibodies overnight at RT. Rinsed twice in Triton X-100 PBS and incubated
with 30 min secondary antibodies (1/250) at room temperature, rinsed twice then
incubated with tertiary antibodies SPC (1/1000) at room temperature for 1 hour. Cultures
were rinsed twice before antigen-antibody complexes were revealed using 3,3’ –
diaminobenzidine-Tetrahydrochloride (DAB; Sigma, D8001) as chromogen for 7 min
until a dark brown colour can be seen. The cultures were then brought to tap water for 10
min to wash off any excess DAB, and were further counterstained with Lillie Mayers
alum haematoxylin (see appendices for procedure). 5 μm sections of paraffin sections
from P5-7 rat pups were de-waxed in heater for 10 min then placed in 2 changes of
xylene (2 min) and alcohol (2 min) before treatment with H2O2 and methanol as
described with the cultures.
160
Controls and cultures were dehydrated with two changes of alcohol and cleared with two
lots of histolene and mounted onto microscope glass slides (coverslipped for controls)
with mounting glue (D.P.X).
Cultures: Procedure is the same as paraffin sections however no need for antigen
retrieval procedure and incubation of primary antibodies is overnight at 4 °C.
5.3.3 Co-localisation of TH and active Caspase-3
Cultures were treated as above however the secondary antibodies were conjugated with a
flourochrome. TH (Chemicon; AB152) polyclonal.
5.4 LACTATE DEHYDROGENASE (LDH) ELISA
LDH is a cytoplasmic enzyme retained by viable cells with intact plasma membranes and
released from necrotic cells with damaged membranes. Briefly, LDH catalyzes the
conversion of lactate to pyruvate upon reduction of NAD+ to NADH/H+, the added TMB
(yellow) is formed, the amount of yellow correlates with LDH activity. Culture medium
samples were collected and snapped frozen (- 80° C) and thawed in 60° C water bath
when required. The bovine serum albumin (BSA) standard was used as an internal
control (1 mg/1 mL) in tris buffered saline (TBS) and also as a blocking solution.
Samples (20 μL) were pre-mixed with TBS (80 μL) with a total volume of 100 μL per
well in a 96-well plate (Maxicorp) in triplicates per treatment group, overnight at 4° C.
Following day plates were aspirated (tipped out) and blocked for 1 hr at RT with 3%
BSA on orbital shaker at low. Washing in TBS x 2 then incubated with the 1° Ab LDH
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(AB2101-1; AbCam) 1/1000 min 1 hr in humidifier. Another TBS wash (2 x) and
incubation with 2° Ab HRP-conjugated anti-goat (Jomar Diagnostics; 611-701-127) in
the humidifier for an 1 hr. Final TBS x3 wash then revealed with TMB (Sigma-Aldrigde;
T-8665 100 ml) 100 μL per well) forming a yellow by-product and reaction terminated
with addition of H2SO4 (50 μL). Colour intensity was assessed using a plate reader
(Ascent Multiwell Plate reader) at 450 nm (see Figure 5. 4).
5.5 QUANTIFICATION AND STATISTICAL ANALYSES OF CELL COUNTS
Cultures were examined and photographed by light microscopy at 400 x with camera
attached to microscope (Olympus) and a monitor. Cell counts were estimated by using
unbiased, blind fashion, uniform and systematic random samples of sections throughout
the cultures. A minimum 3 cultures per group and 4 or more images per cultures were
taken and TH-ir cells with clear nucleus and neuritic or axonal projections observed were
counted. Areas of where the tissues were still thick were not counted as single cell
labeling was not defined.
For experimental Chapters 6, 7 and 8 an assessment of the survival and growth of the TH-
ir neurons within the organotypic slice culture was required. Therefore, using a light
microscope the number of TH-ir cells surviving in each culture was determined by
counting the cells under high power (400 x) magnification. This process was facilitated
by the fact that the cultures were approximately only one cell layer thick.
162
Data are shown as mean ± standard error of measurement (SEM). Repeated measures
analysis of variance (ANOVA) followed by Bonferroni t-tests were used to analyse the
growth of TH-ir cells. 1-way ANOVA was used to compare cell counts between groups
and weeks in vitro and a 2-way ANOVA was used when comparing treated groups of
various concentrations and following weeks. A P value of < 0.05 was considered
significant. Prism (Graphpad TM Software, SanDiego, CA) statistics computer program
was used for all analyses.
163
Figure 5.1. Schematic illustration of the preparation protocol of slice cultures from
4 to 5-day-old rats.
Tissue blocks were dissected at the position A and B to obtain the ventral tegmental
area/substantia nigra-complex (A), the striatum (B). Blocks were trimmed along the
dashed lines and coronal sections of the regions of interest (250-300 �m; asterisks) were
selected and mounted on a glass coverslip by means of a plasma clot. The coverslip was
then placed in a flat-bottom culture tube containing culture medium. Continuous rotation
of the culture was achieved by placing the culture tube in a roller within an incubator
(36ºC). STR, striatum; VTA/SN-complex, ventral tegmental area/substantia nigra-
complex. (Adapted from Heike Franke (Franke et al. 2003)).
164
165
thrombin
coverslip
chick plasma
plasma clot
culture tube
medium
roller drum 5 rev./min
Figure 5.2 Schematic diagram of the culturing process.
Each coverslip contains a piece of VM and ST tissue placed under a chicken plasma cloth
and thrombin; it is then inserted into a flat-bottom screw cap tube with 1 mL medium the
finally placed into a roller drum in a 36 º C incubator rotating at 5 rev/min.
166
Figure 5.3 Schematic representation of the biotin-avidin peroxidase procedure for
the identification of TH-ir neurons.
Each arrow between the steps represents 3 x 5 minute washes with Triton X-100 with
PBS.
167
CULTURE FIXED
4% paraformaldehyde (30mins)
↓↓↓↓
ENDOGENOUS PEROXIDASE ACTIVITY BLOCKED
0.2% H2O2 in methanol (30mins)
↓↓↓↓
CELL MEMBRANES PERMEABILISED
1% Triton X-100 in PBS (3x 5mins)
↓↓↓↓
NON-SPECIFIC BINDING BLOCKED
10% normal horse serum in Triton X-100 in PBS (30mins)
TYROSINE HYDROXYLASE LABELLED
Rabbit anti-TH, 1:2000 dilution with 10% NHS Triton X-100 in PBS (O/N cold room)
↓↓↓↓
SECONDARY ANTIBODY
Vector Biotinylated anti-rabbit IgG (BA-1000) at 1/250 dilution with 10% NHS Triton
X-100 in PBS (30mins at room temp.)
↓↓↓↓
PEROXIDASE LABEL ATTACHED
Pierce peroxidase conjugated streptavidin tertiary (# 21127) at 1/1000 (referred to as
SPC) with 10% NHS Triton X-100 in PBS (1hr at room temp.)
↓↓↓↓
ANTIGEN-ANTIBODY COMPLEXES REACTION VISUALISED
0.05% DAB with 0.01% in PBS (<10mins)
↓↓↓↓
COUNTERSTAIN
Lillie Mayers haematoxylin (1min)
↓↓↓↓
DEHYDRATE TO HISTOLENE
2x100% ethanol then 2x100% histolene (30mins)
168
Figure 5.4 LDH ELISA.
(A) Samples loaded into wells with buffer (TBS) and incubated overnight (4° C). (B)
Samples become attached to the well surface excess sample is aspirated. (C) Incubation
with BSA to block unoccupied sites on well surface with neutral protein on orbital shaker
for 1 hr at RT (low). (D) Add 1° Ab and incubate in humidifier (1 hr). (E) Unattached
Abs is removed followed by incubation of 2° Ab (min 1 hr) in humidifier. (F) Add
substrate and incubate-peroxidase enzyme breaks down substrate and turns yellow,
reaction terminated by H2SO4 (blue).
169
A
B
C
TBS x2 wash
170
CHAPTER 6:
DOPAMINERGIC NEURONS GROWN IN ORGANOTYPIC
SLICE CULTURES: TH-ir CELLS ARE DEPENDENT ON
TARGET REGION-STRIATUM
171
6.0 INTRODUCTION
There are two main populations of DA neurons that innervate the forebrain, the substantia
nigra (SN; A9) cell group, and the ventral tegmental area (VTA; A10) cell group
(Ungerstedt 1971; Beckstead et al. 1979). The SN confines its outgrowth to the dorsal
striatum (ST; or caudate-putamen) (Huang 1990), whereas the VTA also innervates
several other structures, including the nucleus accumbens (Chronister et al. 1980) and
cerebral cortex (Kalsbeek et al. 1988). The determinants of these innervation patterns are
not known, however, evidence for specific factors that influence the survival and
outgrowth of dopaminergic (DAergic) neurons has been accumulating since the 1970s.
Prochiantz and colleagues showed that DAergic neurons from the ventral mesencephalon
(VM) grew toward and formed terminals with neurons from the ST, and they further
demonstrated that this resulted at least in part from a diffusible factor released from
striatal glia {Barbin, 1985 #1642; Prochiantz, 1981 #120; {Prochiantz, 1982 #1643;
Denis-Donini, 1983 #75}. Bohn and co-workers then showed that glial cell lines released
one or more substances that promoted the survival and outgrowth of DAergic neurons
(Engele et al. 1991).
6.1 AIMS
Using postnatal SD rats, we aim to grow and maintain a DAergic system and study the
trophic effects of ST target, which could be used as a model for PD in vitro.
6.2 METHOD
The slice cultures were prepared according to the method described in Chapter 5.
172
Co-cultures of ventral mesencephalon (VM) and striatum (ST) were prepared from 4-5
day old (P4-5) Sprague-Dawley rats. Cultures of VM with and without ST were prepared.
The cultures were incubated by the roller tube technique according to Gahwiler (1981)
for 49 days. The live cultures were inspected twice a week. Cultures producing yellow
colour medium indicated healthy live cells whereas a pink to orange buffer is indicative
of poor growth. Once every week a minimum of 3 tubes were removed, fixed in
paraformaldehyde and stained for tyrosine hydroxylase immunoreactivity (TH-ir), a
marker of DAergic neurons and counter-stained with haematoxylin to show the cell
cytology. The cultures were then analyzed in a blind fashion. The numbers of TH-ir
cells in a minimum of 3 fields under 400x magnification were counted using specific
criteria to include only cells with neurites and a nucleus. These were photographed and
independently checked by a second researcher.
Analysis of the number of TH-ir cells was performed by using One-way ANOVA and
Two-way ANOVA, followed by Bonferroni's Multiple Comparison Test. A p value of
<0.05 was considered significant.
6.3 RESULTS
6.3.1 TH-ir neurons of the ventral mesencephalon in culture
The primary TH antibody was tested for its specificity as a DAergic marker. In all our
cultures only whole VM slices were used instead of halves to minimize the variability as
both sides of the VM do not have the same amount of DAergic neurons (Figure 6.1A &
B), thus if VM halves were cultured this would result in some co-cultures with good
173
growth and others with poor growth of DAergic neurons. Culture controls showed no
staining in the negative control (Figure 6.1C) and brown DAB staining with no
background staining in positive control (Figure 6.1D).
The characteristic organization of the VM disappeared with progressive cultures and
neither the substantia nigra pars compacta (SNpc), nor the VTA, could be detected
separately. The cultures consisted of overlaying layers of neurons upon non-neuronal
cells. A monolayer was formed after 14-28 days incubation with both non-TH-ir cells
and TH-ir neurons extended fibres from the centre of the tissue towards the surrounding
plasma clot (Figure 6.2A). Some of the TH-ir cells were able to direct their axons, to
innervate nearby striatal tissue, forming an axon bridge that between VM-ST (Figure
6.2B). Different morphological types of TH-ir cells were present (Figure 6.2C),
projecting their axons and dendrites in all directions within the tissue forming neuronal
networks (Figure 6.2D). .
6.3.2 TH-ir cell growth in the Co-cultures
Between 7-14 days, the co-cultures were thick with many layers of non-neuronal and
neuronal cells emerging from the tissue surfaces. Individual TH-ir cells could be clearly
seen as they grew from the edge of the tissue outwards into the plasma clot. After 21
days, the co-cultures were reduced to a 1-2 cells thick layer and by the 28th day single
cells formed distinct cell bodies, axons and dendrites. These TH-ir cells formed networks
with neighbouring cells.
174
6.3.3 The influence of trophic ST target on VM TH-ir cells
Single cultures of VM (SVM) grew poorly compared to the co-cultures. These cultures
did not flatten to a monolayer and neuronal network formation was sparse (Figure 6.3A)
compared to co-cultures (Figure 6.3B). Generally TH-ir cells in SVM were smaller and
had fewer processes (Figure 6.3C) compared to co-cultures (Figure 6.3D). SVM cultures
grown for 42 days had significantly lower TH-ir cells (Figure 6.4) when compared to co-
cultures (Figure 6.5 & Figure 6.6).
6.3.4 Glial cells contribute to the regulation of DAergic outgrowth
Cultures were also stained for the astrocytic marker glial fibrillary acidic protein (GFAP).
GFAP-ir cells in SVM cultures were large with numerous projections (Figure 6.7B)
compared to the co-cultures (Figure 6.7C). GFAP-ir cell numbers were initially lower in
the first weeks in co-cultures (Figure 6.8) and increased to similar numbers compared to
the SVM cultures (Figure 6.9). In SVM cultures GFAP-ir cell numbers were initially
higher than co-cultures and remained stable in the following weeks (Figure 6.9). The
GFAP-ir cell numbers were not siginificantly higher than the TH-ir cell numbers in both
co-cultures and SVM cultures (Figure 6.10 & Figure 6.11).
175
Figure 6.1 Dopaminergic (TH-ir) controls
A) TH-positive control [VM rat pup 4-5d, paraffin section 5-7 μm] (x200). (B) Whole
VM hemisphere of figure (A) at x40. Note: left side has less TH-ir neurons than the
right side (C) TH-negative control from co-cultures (x400). (D) TH-positive control
from co-cultures (x400). Bar = 10 μm (C & D), bar = 20 μm (A), bar = 100 μm (B).
A B
C D
176
Figure 6.2 Dopaminergic (TH-ir) neuronal growth in VM & ST co-cultures.
(A) 7 -14 days, TH-ir cells emerge from tissue (x100). (B) Bridge between VM & ST
targets (x40). (C) Subtypes of DAergic neurons (x400). (D) 4 weeks, formation of axons
and dendrites and networks with neighbouring cells (x 200). Bar = 10 μm (C), bar = 20
μm (A), bar = 50 μm (B), bar = 100 μm (D).
A
B
C D
177
0
2
4
6
8
10
12
14
1 2 3 4 5 6 7
Weeks in vitro
Nu
mb
er o
f ce
lls/
0.22
90m
m2
TH-ir
Figure 6.3 Dopaminergic (TH-ir) neuronal growth in VM and ST co-cultures.
The number of cells per field (0.2290mm2) viewed at 400x for each week (wk). The
Means are calculated from the raw data and data expressed as mean ± SEM followed by
One-way ANOVA which was calculated using STDEV, ** p<0.01 [1 wk compared to 3-
6 wk; 2 wk compared to 3-6 wk], *** p<0.001 [1 and 2 wk compared to 7 wk].
**
***
**
178
Figure 6.4 TH-ir cell growth in co-cultures vs single cultures (SVM).
(A) Poor TH-ir cell growth in SVM slice culture at 3 weeks (x 40). (B) TH-ir cells from
SVM cultures at 2 weeks (x400). (C) TH-ir cells in co-cultures SVM at 2 weeks (x 400).
Bar = 10 μm (B & C), bar = 100 μm (A).
A
B C
179
0
1
2
3
4
5
6
7
8
1 2 3 4 5 6
Weeks in vitro
Nu
mb
er o
f ce
lls/
0.22
90m
m2
TH-ir
Figure 6.5 Dopaminergic (TH-ir) neuronal growth in single VM cultures (SVM).
The number of cells per field (0.2290mm2) viewed at 400x for each week (wk). The
Means are calculated from the raw data and data expressed as mean ± SEM followed by
One-way ANOVA which was calculated using STDEV followed by Bonferroni's
Multiple Comparison Test. * p<0.05 [1 wk compared 2 wk], *** p<0.001 [2 wk
compared to 3-6 wk].
*
***
180
0
2
4
6
8
10
12
14
1 2 3 4 5 6
Weeks in vitro
Nu
mb
er o
f T
H-i
r ce
lls/
0.22
90m
m2
VM & ST
VM
Figure 6.6 Dopaminergic (TH-ir) neuronal growth in co-cultures vs SVM.
The number of cells per field (0.2290mm2) viewed at 400x for each week (wk). The
Means are calculated from the raw data and data expressed as mean ± SEM followed by
Two-way ANOVA which was calculated using STDEV followed by Bonferroni post-
tests, *** p<0.001.
*** ***
*** *** ******
181
Figure 6.7 GFAP growths in co-cultures vs single cultures.
(A) GFAP-positive control (VM rat pup 4-5d, paraffin section 5-7μm; x 200). (B)
GFAP-ir cells from co-cultures at 7 DIV (x 400). (C) GFAP-ir cells in SVM at 7 DIV (x
400). Bar = 10 μm (B & C), bar = 20 μm (A).
A
* *
* * *
*
*
*
C B
182
0
1
2
3
4
5
6
7
8
9
10
1 2 3 4 5 6
Weeks in vitro
Nu
mb
er o
f ce
lls/
0.22
90m
m2
GFAP-ir
Figure 6.8 Glial fibrillary acidic protein (GFAP-ir) growth in VM & ST co-cultures.
The number of cells per field (0.2290mm2) viewed at 400x for each week (wk). The
Means are calculated from the raw data and data expressed as mean ± SEM followed by
One-way ANOVA which was calculated using STDEV followed by Bonferroni's
Multiple Comparison Test, * p<0.05, ** p<0.01, *** p<0.001.
****
***
**
183
0
1
2
3
4
5
6
7
8
9
1 2 3 4 5 6
Weeks in vitro
Nu
mb
er o
f ce
lls/
0.22
90m
m2
GFAP-ir
Figure 6.9 Glial fibrillary acidic protein (GFAP-ir) growth in single VM (SVM)
cultures.
The number of cells per field (0.2290mm2) viewed at 400x for each week (wk). The
Means are calculated from the raw data and data expressed as mean ± SEM followed by
One-way ANOVA which was calculated using STDEV followed by Bonferroni's
Multiple Comparison Test, * p<0.05, ** p<0.01.
***
**
184
0
2
4
6
8
10
12
14
1 2 3 4 5 6
Weeks in vitro
Nu
mb
er o
f c
ells
/0.2
290m
m2
TH-ir cells
GFAP-ir cells
Figure 6.10 Dopaminergic (TH-ir) & Glial fibrillary acidic protein (GFAP-ir)
growth in VM & ST co-cultures.
The number of cells per field (0.2290mm2) viewed at 400x for each week (wk). The
Means are calculated from the raw data and data expressed as mean ± SEM followed by
Two-way ANOVA which was calculated using STDEV and Bonferroni's Multiple
Comparison Test, *** p<0.001.
*** ***
******
*** ***
185
0
1
2
3
4
5
6
7
8
9
10
1 2 3 4 5 6
Weeks in vitro
Nu
mb
er o
f ce
lls/
0.22
90m
m2
VM & ST
SVM
Figure 6.11 Glial fibrillary acidic protein (GFAP-ir) growth in VM & ST co-cultures
and SVM.
The number of cells per field (0.2290mm2) viewed at 400x for each week (wk) from co-
cultures & SVM. The Means are calculated from the raw data and data expressed as
mean ± SEM followed by Two-way ANOVA which was calculated using STDEV and
Bonferroni's Multiple Comparison Test, *** p<0.001.
***
186
6.4 DISCUSSION
6.4.1 General findings
In this study we were able to maintain the growth of DAergic cells without any special
conditioned medium using the organtotypic culture technique. In particular, we
demonstrated that growth and maturation of VM DAergic neurons was in some way
regulated and dependent on its specific trophic target- namely the ST. The DAergic
neurons were able to innervate the ST and that glial cells played a supportive role in this
process.
Ostergaard (Ostergaard et al. 1990) described 5 distinct groups of TH-ir cell types which
we were able to identify in our slice cultures. The multipolar neurons were the most
commonly observed throughout the period of culture which mainly is comprised of three
primary dendrites and a single unbranched axon from the cell body with a small number
had spines on the dendrites. Pyramidal shaped neurons have a triangular cell body shape
with both smooth axons and dendrites that are spineless and have three dendrites
emerging from the cell body. The bipolar neurons commonly have two dendrites and like
the pyramidal neurons, their axons and dendrites are spineless whereas the small,
multiform neurons have either round, pyramidal or bipolar cell body shape with two or
three dendrites.
6.4.2 Advantage and disadvantages of the culture system-organotypic slice culture
system
187
This DAergic in vitro system has been employed widely over the last 5 decades and
shown to be reliable and reproducible. Slice cultures are usually derived from early
postnatal rodents. Early postnatal periods (p0 to p7) are ideally suited for culturing
because cytoarchitecture is already established in most brain networks, the brain is larger
and easier to dissect, and the nerve cells survive explantation readily. Tissue that is
derived from embryonic animals survives in slice cultures (Steensen et al. 1995;
Ostergaard et al. 1996; Becq et al. 1999; Berglof et al. 2008) but the organotypic
organization often becomes distorted since in early development, the majority of nerve
cells are still in their migratory phase from the periaquaductal area, where they originate,
to the SNpc. Few attempts have been made to culture adult tissue (Sorensen et al. 1994;
Wilhelmi et al. 2002). The organotypic slice culture system provides a highly accessible
system for studying the development of neuronal tissues. The TH-ir cells in organotypic
cultures of VM showed morphological characteristics similar to those described in intact
animals (Ostergaard et al. 1990; Tepper et al. 1994), consistent with their in vivo
morphology.
Variability was the main disadvantage of this culture technique. The numbers of TH-ir
cells and their neuritic outgrowth varied widely between the cultures. This variability
impeded quantitative analysis. However, we overcame this problem by only selecting
fields containing single DA cell growth, with a clearly identified cell body and processes
with a nucleus. Areas which had dense growth were avoided because of overlapping
DAergic neurons originating from the VM. One explanation for the increased variability
of slice cultures is that the numbers of cells present in each slice at the start of the culture
188
period is unknown compared to dissociated cultures where the plating density can be
calculated. Another factor is that each VM slice contains DAergic neurons from a
different VM level (rostral to caudal nigral) and since the whole rostral-caudal extent of
SN and ST were dissected in all experiments and then combined randomly to form co-
cultures; it is possible that the specific topographical projection of DAergic neurons to the
ST resulted in good growth in certain cultures and poor in others. To reduce the
variability only whole VM slices were used and large numbers of cultures were prepared
for each experimental condition to reduce the standard error of deviation (SD) number of
TH-ir cells in each culture. Cultures which detached from the coverslips (overall < 10%
of plated VM-ST cultures) and cultures that did not grow were discarded. To obtain
unbiased counting, slides were coded and cells were scored blindly.
6.4.3 The influence of target region-Striatum on TH-ir neurons
As discussed in chapter 1 and 3 during development, DA neurons follow a specific
temporal pattern by which their axons project from the VM and innervate the ST, a
process that has been well documented both in vivo (Voorn et al. 1988; Gates et al. 2004)
and in vitro in VM/ST co-cultures (Voorn et al. 1988; Schatz et al. 1999; Snyder-Keller et
al. 2001; Gates et al. 2004). The number of DAergic neurons grown in SVM cultures in
the absence of the ST target started low and remained low for the whole culture period
compared to co-cultures with the ST. Although there were no significant differences in
the morphology of the TH-ir neurons between SVM and co-cultures it was apparent that
SVM cultures yielded poor TH-ir cell growth compared to co-cultures. These results are
supported by previous studies in which the ST exerted a trophic influence on the
189
development of the DAergic neurons in dissociated cell cultures (Prochiantz et al. 1979;
Prochiantz et al. 1981). Further studies also demonstrated that ST extracts exerted
neurotrophic effects on mesencephalic DAergic cells (Dal Toso et al. 1988). In addition,
the development of dendritic and axonal patterns of TH-ir aggregates was also influenced
by the presence of ST cells (Hemmendinger et al. 1981).
6.4.4 The growth of TH-ir neurons with increasing culture duration
Significant differences between SVM and VM-ST cultures were observed regarding the
numbers of TH-ir cells. There is a an increase in TH-ir cell number from week two to
three in the co-cultures in agreement with Ostergaard (Ostergaard et al. 1990) who found
that following six days in co-culture no TH-ir neurons had innervated the ST but by day
twenty five innervation had occurred in the majority of co-cultures. Following the third
and fourth week, growth of TH-ir cells were at its maximum in VM-ST cultures. In
contrast in the SVM cultures, the numbers of TH-ir cells remained low as they had no
target tissue to innervate. Once the TH-ir cells from the VM had innervated the ST in the
co-cultures, the number of TH-ir cells did not significantly increase in the remaining
weeks of culturing. One possibility is that the target region could regulate this
innervation by producing a limited amount of neurotrophic factor (Barde 1989) as BDNF
and GDNF are produced by the ST and enhance survival of TH-ir neurons in culture
(Knusel et al. 1990b; Hyman et al. 1991; Oo et al. 2003; Jaumotte and Zigmond 2005)
controlling innervation by TH-ir neurons. Thus, axons which are deprived of any
neurotrophic factor would die.
190
6.4.5 Possible influences on TH-ir cell development
The prenatal development of DAergic neurons of the SN is characterized by their birth,
specification and migration to their final positions. Their postnatal development is
characterized by the establishment of contact and interactions between the SN and other
neural nuclei, particularly the ST target, by extension of axons, terminal differentiation,
and synapse formation. In the postnatal process there is natural cell death, which is
apoptotic and biphasic in time course, initially peaking on postnatal day (PND) 2, and a
second on PND14 (Mahalik et al. 1994; Jackson-Lewis et al. 2000; Burke 2003; Oo et al.
2003) and by PND20 apoptotic events have largely subsided. This natural cell death
event is regulated in vivo by interaction with ST target. This target dependence is present
largely within the first two postnatal weeks (Burke 2003). Since our co-cultures derived
from P4-5 day old rat pups the DAergic cells would have gone through the initial
apoptotic phase however due to the dissection of the VM and ST regions resulting in
severed afferent/efferent connections this may have augmented the natural cell death
process. After the tissue slices were plated next to each other the regenerative process
would have taken a while to establish and to allow the TH-ir cells to migrate and project
axons innervating the ST. This may explain the low levels of TH-ir cells in the first two
weeks of culture and then increased in the following third week. It is well accepted that
the ST regulates DAergic cell death as suggested by in vitro (Prochiantz et al. 1979;
Hemmendinger et al. 1981; Hoffmann et al. 1983; Tomozawa and Appel 1986) and in
vivo studies (Coyle and Schwarcz 1976; Burke et al. 1992; Macaya et al. 1994).
6.4.6 Role of GFAP-ir cells in TH-ir neuronal development
191
Glial cells play an important role in CNS development (Hatten 1993). In our co-cultures
the numbers of GFAP-ir cells were always less than TH-ir cell numbers. However as the
TH-ir cell numbers increased so did the number of GFAP-ir cells. In contrast in the SVM
cultures GFAP-ir cell numbers were significantly higher than the TH-ir cell numbers in
the first week of incubation and remained similar numbers as the TH-ir cells throughout
the culturing period. Studies have shown that DAergic neurons in culture may be guided
by astroglial cells (Johansson and Stromberg 2002; Johansson et al. 2005). The long-
term surviving outgrowth of DAergic nerve fibers were shown to be guided by astrocytes
DAergic. Little is known about how cell types other than astrocytes may influence nerve
fiber formation from DA neurons. It is known that conditioned medium from
oligodendrocyte-type-2 astrocyte progenitors and microglia may enhance survival of
cultured DAergic neurons (Nagata et al. 1993b; Nagata et al. 1993a; Takeshima et al.
1994a, b; Takeshima et al. 1994c; Zietlow et al. 1999; Sortwell et al. 2000), although the
presence of both oligodendrocytes and microglia is known to exert inhibitory effects
(Ling et al. 1998b; Wang et al. 2002; Park et al. 2007).
The GFAP-ir cells in SVM may provide support since no ST target tissue is available to
innervate and in turn regulate development as astroglias are thought to be involved in the
guidance of neurons and axonal growth cones (Jacobs and Goodman 1989; Hidalgo et al.
1995; Hidalgo and Booth 2000). It is likely that the interaction between glia and neurons
are mutual, where astroglia and neurons are active team players and thus regulate mutual
survival (Gates and Dunnett 2001; Chotard and Salecker 2004). It has been shown that
astroglia-associated TH-positive neurite outgrowth development only innervates fetal ST,
192
the innervation pattern overlapping astrocytic-dense areas (Johansson and Stromberg
2003). Although it is not known how the astrocytes might attract the TH-positive axons,
in situ, studies have shown that extracellular matrix molecules, which are produced by
astrocytes, are concentrated in striatal patches when DA innervation occurs (O'Brien et al.
1992; Charvet et al. 1998a, b). In addition, enhanced migration of astrocytes increases
formation of TH-positive nerve fibers (Johansson and Stromberg 2002). However, it
cannot be determined from our study as to what factors influence the astrocytes to
terminate their migration.
In vitro, DAergic neurones from the mesencephalon plated on glial monolayers either
from the striatal or the mesencephalic region on mesencephalic glial cells, the DAergic
neurones develop highly branched and varicose neurites, whereas on striatal glia they
only exhibit one long, thin and rather linear neurite (Denis-Donini et al. 1984). This
demonstrates that glial cells from two different brain regions have distinct properties.
Studies have also shown that neurotrophic factors are produced by glial cells which
protect DAergic neurons against oxidative stress (Hou et al. 1997). Furthermore, an
astrocyte conditioned medium promotes the survival of DAergic cells (Engele et al. 1991;
O'Malley et al. 1994) and protects these cells against L-DOPA toxicity (Mena et al.
1996).
193
CHAPTER 7:
THE EFFECTS OF NEUROTOXINS: MPTP & ROTENONE
ON TH-ir CELLS GROWTH IN ORGANOTYPIC SLICE
CULTURES
194
7.0 INTRODUCTION
As discussed in chapter 2 the neurotoxin MPTP has been studied extensively in both in
vivo and in vitro PD models (Uversky 2004; Bove et al. 2005; Smeyne et al. 2005;
Smeyne and Jackson-Lewis 2005). Most in vitro studies have employed MPP+ compared
to rotenone with its application mainly in specific types of primary cell lines or derived
from embryonic mice (Fonck and Baudry 2001; Gao et al. 2003b; Gao et al. 2003a;
Hirata and Nagatsu 2005; Zeng et al. 2006). Rotenone has been shown to be more toxic
to DAergic neurons (Sherer et al. 2003b; Sherer et al. 2003a) compared to MPTP. Both
toxins have been reported to act through mitochondrial dysfunction causing reactive
oxygen species (ROS) and subsequent cell death (Storch et al. 2000; Fonck and Baudry
2001; Richardson et al. 2007). Astrocytes have been recognized as the key player in
converting MPTP to it active metabolite MPP+ (Brooks et al. 1989; Di Monte et al. 1991;
Di Monte et al. 1992) and also contribute to ROS production (Block and Hong 2007;
Block et al. 2007). Rotenone on the other hand is highly lipophilic and does not require
astrocytes for its conversion into an active metabolite to be toxic.
7.1 AIMS
This study aims to characterize the effects of both MPTP and rotenone on TH-ir cell
growth and survival in organotypic slice cultures derived from 4-5d postnatal rats.
7.2 METHOD
The slice co-cultures were prepared according to the method described in Chapter 5.
195
7.2.1.1 MPTP Effects on 7 Day old VM & ST co-cultures
Co-cultures of ventral mesencephalon and striatum were prepared from P4-5 Sprague
Dawley rats, cytostatics were added on day 4 post co-culture and media changed twice a
week. Co-cultures were then divided into groups which were incubated in different toxin
doses (see Table 7.1). 24 hours post treatment with MPTP the media was changed with
fresh medium, and thereafter changed twice a week. The following 1, 2, and 3 weeks
post-treatment, a minimum of 3 tubes per group were immunocytochemically stained for
tyrosine hydroxylase (TH-ir). Thereafter, the number of TH-ir cells were counted in a
blinded fashion and analysed as previously described (chapter 5).
Table 7.1
7 day old co-cultures treated with MPTP
GROUPS TREATMENTS
1
Naïve controls (normal media)
2 Vehicle controls (PBS + media)
3 100 μM
4 50 μM
5 10 μM
6 5 μM
7 1 μM
8 0.5 μM
196
7.2.1.2 Rotenone Effects on 7 Day old VM & ST co-cultures
Co-cultures were divided into the following groups which were incubated in different
doses (see Table 7.2). 24 hours post treatment with rotenone the media was changed with
fresh medium, and thereafter changed twice a week. The following 1, 2, and 3 weeks
post-treatment, a minimum of 3 tubes per group were immunocytochemically stained for
tyrosine hydroxylase (TH-ir). Thereafter, the number of TH-ir cells were counted in a
blinded fashion and analysed as previously described (chapter 5).
Table 7.2
7 day old co-cultures treated with rotenone
GROUPS TREATMENTS
1 Naïve controls (normal media)
2 Vehicle controls (DMSO + media)
3 100 nM
4 50 nM
5 10 nM
6 5 nM
7 1 nM
197
7.2.2.1 MPTP Effects on 14 Day old VM & ST co-cultures
Co-cultures of ventral mesencephalon and striatum were prepared from P4-5 Sprague
Dawley rats, cytostatics were added on day 4 post co-culture and media changed twice a
week. Co-cultures were then divided into groups which were incubated in different doses
(see Table 7.3). 24 hours post treatment with MPTP the media was changed with fresh
medium, and thereafter changed twice a week. The following 1, 2, and 3 weeks post-
treatment, a minimum of 3 tubes per group were immunocytochemically stained for
tyrosine hydroxylase (TH-ir). Thereafter, the number of TH-ir cells were counted in a
blinded fashion and analysed as before.
Table 7.3
14 day old co-cultures treated with MPTP
GROUPS TREATMENTS
1
Naïve controls (normal media)
2 Vehicle controls (PBS + media)
3 100 μM
4 50 μM
5 10 μM
6 5 μM
198
7.2.2.2 Rotenone Effects on 14 Day old VM & ST co-cultures
Co-cultures were divided into the following groups which were incubated in different
doses (see Table 7.4). 24 hours post treatment with rotenone the media was changed with
fresh medium, and thereafter changed twice a week. The following 1, 2, and 3 weeks
post-treatment, a minimum of 3 tubes per group were immunocytochemically stained for
tyrosine hydroxylase (TH-ir). Thereafter, the number of TH-ir cells were counted in a
blinded fashion and analysed as before.
Table 7.4
14 day old co-cultures treated with rotenone
GROUPS TREATMENTS
1 Naïve controls (normal media)
2 Vehicle controls (DMSO + media)
3 100 nM
4 50 nM
5 10 nM
6 5 nM
199
7.2.2.3 Effects of MPP+ on 14 Day old VM & ST co-cultures
Co-cultures were divided into 2 groups which were incubated in different doses and
neurotoxin (see Table 7.5). 24 hours post treatment with MPTP or MPP+ the media was
changed with fresh medium, and thereafter changed twice a week. The following first
week of post-treatment, a minimum of 3 tubes per group were immunocytochemically
stained for tyrosine hydroxylase (TH-ir). Thereafter, the number of TH-ir cells were
counted in a blinded fashion and analysed as before.
Table 7.5
14 day old co-cultures treated with MPTP & MPP+
MPTP MPP+
Control Control
5 μM 5 μM
10 μM 10 μM
50 μM 50 μM
100 μM 100 μM
7.2.4 Cytotoxicity of Neurotoxins
Lactate dehydrogenase (LDH) is released from dead or degenerating cells with
permeabilized membranes into the culture medium, the medium was analysed at 7 days
post toxin treatment by a fully automatic spectrophotometer (Ascent Multiwell plate
reader). The medium was collected for LDH determination and frozen immediately with
200
storage at −20°C until analysed. Two independent series of LDH measurements were
performed.
7.2.5 Neuronal Degeneration: Fluoro-Jade C (FJC)
Neuronal cell death was assessed using the neuron-specific cell death marker FJC using a
standard protocol (Schmued et al. 2005), with modification. (1) cultures were transferred
to 0.06% postassium permanganate solution for 10 min, then rinsed in DH2O for 2 min;
(2) incubation with 0.0001% solution of FJC (Chemicon, Tememcula, CA) for 20 min;
(3) FJC was immediately before use bt diluting a stock solution of 0.01% FJC by 100-
fold in 0.1% acetic acid; (4) followed by washes x3 time for 1 min in DH2O, dried in a 45
°C oven, cleared in histolene and coverslipped using DPX; (5) cultures were examined
using epifluorescence microscope with an excitation wavelength of 488 nm.
7.2.6 Apoptotic Cell Death: Caspase-3
Cultures were treated as previously described in chapter 5, incubated overnight with TH
and active caspase-3 at 4 °C, following day incubation with appropriate biotinylated
secondary antibody for 30 min, followed by streptavidin-conjugated AlexaFluor 488, 546
or 660.
201
A
DIV 0 4 7 14 21 28
B
DIV 0 4 7 14 21 28 35
Figure 7.1 Schematical diagram of the experiments.
A) 7 day old co-cultures were treated with MPTP or rotenone for 24 hours then fixed at 1,
2 & 3 weeks post treatment. B) 14 day old co-cultures were treated with MPTP or
rotenone. * = Normal media change x2/wk.
cytostatics@24hrs
MPTP/Rotenone
1 wk post 2 wk post 3 wk post
* * * *
cytostatics@24hrs
MPTP/Rotenone
* * * *
1 wk post 2 wk post 3 wk post
202
7.3 RESULTS
7.3.1 Neurotoxin treatment of VM and ST co-cultures
All experiments included both a naïve control and vehicle control and were compared at
the same time as toxin treated co-cultures. All naïve and vehicle controls had TH-ir cell
numbers that were similar to each other and correlated to their respective weeks in
culture. There was no significant difference between controls and vehicles thus,
suggesting neither had any effect on TH-ir cell growth during the treatments. Both
MPTP and rotenone induced dose-dependent cell death and 14 day old co-cultures were
more vulnerable than 7 day co-cultures.
7.3.2 Dose-dependent death of TH-ir cells-MPTP
MPTP treatment of 7 day co-cultures induced a reduction in TH-ir cell numbers and
neuritic outgrowth. The greatest reductions were seen at the highest dose (100 μM)
(Figure 7.2). All MPTP concentrations induced a significant reduction in numbers of
TH-ir cells compared to naïve and vehicle controls. One week post treatment at lower
doses (1 μM) induced a 13 % significant decrease in TH-ir cell number compared to
naïve controls with further significant decreases at highest dose (100 μM) of 35 %
compared to naïve controls. At two weeks post treatment (Figure 7.3.2) the dose-
response was similar to one week post treatment. TH-ir cell numbers were lower than
one week post at 1 μM (11 %) but the reduction continued to increase at the highest dose
(100 μM) 47 % compared to naïve controls. The reduction in TH-ir cells continued
steeply during third week of post treatment (Figure 7.3.3) as 1 μM caused a significant
reduction of 32 % and 61 % at 100 μM compared to naïve controls.
203
Figure 7.2. MPTP treated co-cultures at 7 days.
(A), control; (B), 5 μM; (C), 10 μM; (D), 50 μM; (E), 100 μM, all 1 week post. *
Degenerating cells. Bar = 10 μm.
A
B C
D E
**
*
*
204
0
2
4
6
8
10
12
14
control vehicle 0.5uM 1uM 5uM 10uM 50uM 100uM
Treatments
Nu
mb
er o
f ce
lls/
0.22
90m
m2
TH-ir
Figure 7.3.1 Dopaminergic (TH-ir) neuronal growth following 1 week post MPTP
treatment of 7 day co-cultures.
The number of cells per field (0.2290mm2) viewed at 400x following different doses.
The Means are calculated from raw data and expressed as mean ± SEM followed by
Two-way ANOVA which was calculated using STDEV, * p<0.05, ** p<0.01, ***
p<0.001 compared to controls.
* ** *****
******
205
0
2
4
6
8
10
12
14
16
control vehicle 0.5uM 1uM 5uM 10uM 50uM 100uM
Treatments
Nu
mb
er o
f ce
lls/
0.22
90m
m2
TH-ir
Figure 7.3.2 Dopaminergic (TH-ir) neuronal growth following 2 week post MPTP
treatment of 7 day co-cultures.
The number of cells per field (0.2290mm2) viewed at 400x following different doses.
The Means are calculated from raw data and expressed as mean ± SEM followed by
Two-way ANOVA which was calculated using STDEV, * p<0.05, ** p<0.01, ***
p<0.001 compared to controls.
** ** ******
***
206
0
2
4
6
8
10
12
14
16
control vehicle 0.5uM 1uM 5uM 10uM 50uM 100uM
Treatments
Nu
mb
er o
f ce
lls/
0.22
90m
m2
TH-ir
Figure 7.3.3 Dopaminergic (TH-ir) neuronal growth following 3 week post MPTP
treatment of 7 day co-cultures.
The number of cells per field (0.2290mm2) viewed at 400x following different doses.
The Means are calculated from raw data and expressed as mean ± SEM followed by
Two-way ANOVA which was calculated using STDEV, * p<0.05, ** p<0.01, ***
p<0.001 compared to controls.
***
*** ******
******
207
Figure 7.4. MPTP treated co-cultures at 14 days.
(A), control; (B), 5 μM; (C), 10 μM; (D), 50 μM; (E), 100 μM, all 3 week post. Bar =
10 μm.
A
E D
B C
D E
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0
2
4
6
8
10
12
14
control vehicle 5uM 10uM 50uM 100uM
Treatments
Nu
mb
er o
f ce
lls/
0.22
90m
m2
TH-ir
Figure 7.5.1 Dopaminergic (TH-ir) neuronal growth following 1 week post MPTP
treatment of 14 day co-cultures.
The number of cells per field (0.2290mm2) viewed at 400x following different doses.
The Means are calculated from raw data and expressed as mean ± SEM followed by
Two-way ANOVA which was calculated using STDEV, * p<0.05, ** p<0.01, ***
p<0.001 compared to controls.
*** ***
******
209
0
2
4
6
8
10
12
14
control vehicle 5uM 10uM 50uM 100uM
Treatments
Nu
mb
er o
f ce
lls/
0.22
90m
m2
TH-ir
Figure 7.5.2 Dopaminergic (TH-ir) neuronal growth following 2 week post MPTP
treatment of 14 day co-cultures.
The number of cells per field (0.2290mm2) viewed at 400x following different doses.
The Means are calculated from raw data and expressed as mean ± SEM followed by
Two-way ANOVA which was calculated using STDEV, * p<0.05, ** p<0.01, ***
p<0.001 compared to controls.
****** ***
***
210
0
2
4
6
8
10
12
14
control vehicle 5uM 10uM 50uM 100uM
Treatments
Nu
mb
er o
f ce
lls/
0.22
90m
m2
TH-ir
Figure 7.5.3 Dopaminergic (TH-ir) neuronal growth following 3 week post MPTP
treatment of 14 day co-cultures.
The number of cells per field (0.2290mm2) viewed at 400x following different doses.
The Means are calculated from raw data and expressed as mean ± SEM followed by
Two-way ANOVA which was calculated using STDEV, * p<0.05, ** p<0.01, ***
p<0.001 compared to controls.
******
***
***
211
0
2
4
6
8
10
12
cont
rol 5 10 50 100
Treatments (uM)
Nu
mb
er o
f T
H-i
r ce
lls/
0.22
90m
m2
naïve control
MPP+
MPTP
Figure 7.5.4 Dopaminergic (TH-ir) neuronal growth following 1 week post MPTP &
MPP+ treatment of 14 day co-cultures.
The number of cells per field (0.2290mm2) viewed at 400x following different doses and.
The Means are calculated from raw data and expressed as mean ± SEM followed by
Two-way ANOVA which was calculated using STDEV, * p<0.05, ** p<0.01, ***
p<0.001 compared to controls.
212
Figure 7.6. 7 day & 14 day old co-cultures MPTP treated. The number of cells per
field (0.2290mm2) viewed at 400x following different doses and at all weeks of recovery.
The Means are calculated from raw data and expressed as mean ± SEM followed by
Two-way ANOVA which was calculated using STDEV, *** p<0.001 7 days compared
to their respective 14 day groups. (A), 1 week post; (B), 2 week post; (C), 3 week post.
213
0
2
4
6
8
10
12
14
control vehicle 5uM 10uM 50uM 100uM
Treatments
Nu
mb
er o
f T
H-i
r ce
lls/
0.22
90m
m2
7DIV
14DIV
0
2
4
6
8
10
12
14
16
control vehicle 5uM 10uM 50uM 100uM
Treatments
Nu
mb
er o
f T
H-i
r ce
lls/
0.22
90m
m2
7DIV
14DIV
0
2
4
6
8
10
12
14
16
control vehicle 5uM 10uM 50uM 100uM
Treatments
Nu
mb
ers
of
TH
-ir
cell
s/0.
2290
mm
2
7DIV
14DIV
***
******
******
***
***
***
***
*** ******
A
B
C
214
7.3.3 MPTP toxicity varies with the time of co-culture exposure
14 day old co-cultures treated at lower doses were more affected than 7 day old treated
co-cultures. TH-ir cells showed less intense immunostaining as well as loss of neural
processes (Figure 7.4). MPTP treated 14 day old co-cultures produced a more consistent
dose-response cell count than 7 day old co-cultures. One week post treatment at 5 μM
induced a significant reduction in TH-ir cell numbers (38%) whereas 7 day old co-
cultures treated at the same dose reduced TH-ir cells by 22 % (Figure 7.5.1). At the 100
μM a significant decrease of 72 % was observed in 14 day old co-cultures. Reductions in
TH-ir cell number during second week of post treatment at low doses (5 μM) 46 % (7 day
cultures at same dose and week; 12.5 %) continued to increase but at higher doses (100
μM) were slowed down to 64 % compared to naïve controls (Figure 7.5.2). By the third
week reduction in TH-ir cell numbers at low doses (5 μM ) were also slowing down to
41% (7 day old cultures treated at same dose and week; 32 %). However the higher
doses of 100 μM continued to cause significant TH-ir cell reductions (79 %) compared to
naïve controls (Figure 7.5.3). To compare whether MPTP or its by product MPP+ was
more toxic we treated 14 day old co-cultures with varying concentrations and analysed
TH-ir cells at one week post treatment. There was no significant difference in toxicity
between the two toxins but a dose-response was observed for both (Figure 7.5.4).
We compared dose effect at each point for 7 and 14 day old co-cultures (Figure 7.6). 7
day co-cultures was observed to be more resistant to MPTP compared to 14 day co-
cultures. The number of TH-ir cells was significantly reduced at all concentrations
compared to naïve controls but recovered much faster in all three weeks of post treatment
215
in 7 day old co-cultures. In contrast in 14 day co-cultures even at the lower doses MPTP
induced significant reductions in TH-ir cell numbers and continued to decrease at all
weeks post treatment compared to naïve controls.
7.3.4 Dose-dependent death of TH-ir cells-Rotenone
Rotenone co-cultures treated at 7 days old induced reductions in TH-ir cell numbers and a
few projections compared to naïve controls (Figure 7.7). The intensity of TH-
immunostaining was clearly less intense compared to naïve controls. A dose-response
was observed with rotenone treated co-cultures. 7 day old co-cultures treated with
rotenone at the lowest dose (1 nM) induced a 3 % reduction in TH-ir cell numbers in the
first week of post treatment compared to naïve controls (Figure 7.8.1). Significant TH-ir
cell reduction was observed at the higher doses 100 nM (67 %) compared to naïve
controls. TH-ir cell reductions continued to increase significantly at 1 nM (38 %) and at
100 n M (89 %) during the second week compared to naïve control (Figure 7.8.2). Third
week of recovery TH-ir cell numbers were reduced to 31 % (1 nM) and 51 % (100 nM)
compared naïve controls (Figure 7.8.3).
14 day old co-cultures treated with rotenone induced swelling of TH-ir cell bodies and
reduction in TH-ir neurites (Figure 7.9). The dose response was similar to MPTP treated
co-cultures. First week of recovery resulted in a significant reduction in TH-ir cell
numbers by 31 % at 5 nM whereas at 100 nM induced a greater reduction by 93 %
compared to naïve controls (Figure 7.10.1). This percentage is much larger than 7 day
old co-cultures where 5 nM resulted in 3.5 % and 100 n M was 67 % in the first week.
216
Second week of recovery TH-ir cell number reduction continued to increase to 53 % at 5
nM and 92 % at 100 nM compared to naïve controls (Figure 7.10.2). Again these
numbers were much higher compared to 7 day old co-cultures (5 nM = 43 % & 100 nM =
67 %). Finally third week of recovery TH-ir cell number decreased to to 26 % at 5 nM
and 86 % at 100 nM compared naïve controls (Figure 7.10.3). Compared to 7 day old co-
cultures treated at 5 nM which resulted in 44 % TH-ir cell number reduction this
percentage were much higher than 14 day old co-cultures.
7.3.5 Rotenone toxicity varies with the time of co-culture exposure
Similar to MPTP treatment, rotenone was observed to be more toxic to 14 day co-cultures
than 7 day co-cultures (Figure 7.11). There was less variability between the treatment
groups and a more consistent dose-response effect in the weeks post treatment. Lower
doses of rotenone treated in 7 day co-cultures resulted in low TH-ir cell reductions but
progressively increased in the weeks of recovery whereas the higher doses varied
between 67 % in the first week, 89 % in the second and 51 % in the last week. 14 day old
co-cultures on the other hand when treated with low doses of rotenone a 31 % reduction
in TH-ir cell numbers in the first week was observed which continued to increase to 53 %
in second week and reduce to 26 % in the final week. At the highest concentration of 100
nM 14 day treated co-cultures induced a substantial reduction in TH-ir cell numbers by
93 % in the first week and were stable in the second week (92 %) and declined slightly to
86 % in the third week. Thus, 14 day co-cultures were observed to be more prone to
rotenone at all doses whereas 7 day old co-cultures were more vulnerable at the higher
doses as indicated in weeks of post treatment.
217
Figure 7.7. Rotenone treated co-cultures at 7 days.
(A), control; (B), 5 nM; (C), 10 nM; (D), 50 nM; (E), 100 nM, all 1 week post treatment.
* Degenerating neuron. Bar = 10 μm.
B
C
A
*
*
B C
D E
*
218
0
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4
6
8
10
12
14
control vehicle 1nM 5nM 10nM 50nM 100nM
Treatments
Nu
mb
er o
f ce
lls/
0.22
90m
m2
TH-ir
Figure 7.8.1 Dopaminergic (TH-ir) neuronal growth following 1 week post rotenone
treatment of 7 day co-cultures.
The number of cells per field (0.2290mm2) viewed at 400x following different doses.
The Means are calculated from raw data and expressed as mean ± SEM followed by
Two-way ANOVA which was calculated using STDEV, * p<0.05, ** p<0.01, ***
p<0.001 compared to controls.
***
***
***
219
0
2
4
6
8
10
12
14
16
control vehicle 1nM 5nM 10nM 50nM 100nM
Treatments
Nu
mb
er o
f ce
lls/
0.22
90m
m2
TH-ir
Figure 7.8.2 Dopaminergic (TH-ir) neuronal growth following 2 week post rotenone
treatment of 7 day co-cultures.
The number of cells per field (0.2290mm2) viewed at 400x following different doses and
2 weeks of recovery. The Means are calculated from raw data and expressed as mean ±
SEM followed by Two-way ANOVA which was calculated using STDEV, * p<0.05, **
p<0.01, *** p<0.001 compared to controls.
******
*** ***
***
220
0
2
4
6
8
10
12
14
16
control vehicle 1nM 5nM 10nM 50nM 100nM
Treatments
Nu
mb
er o
f ce
lls/
0.22
90m
2
TH-ir
Figure 7.8.3 Dopaminergic (TH-ir) neuronal growth following 3 week post rotenone
treatment of 7 day co-cultures.
The number of cells per field (0.2290mm2) viewed at 400x following different doses.
The Means are calculated from raw data and expressed as mean ± SEM followed by
Two-way ANOVA which was calculated using STDEV, * p<0.05, ** p<0.01, ***
p<0.001 compared to controls.
***
******
*** ***
221
Figure 7.9. Rotenone treated co-cultures at 14 days.
(A) control; (B), 5 nM; (C), 10 nM; (D), 50 nM; (E), 100 nM, all 1 week post. Bar = 10 μm
A
E
B C
D E
222
0
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4
6
8
10
12
control vehicle 5nM 10nM 50nM 100nM
Treatments
Nu
mb
er o
f ce
lls/
0.22
90m
m2
TH-ir
Figure 7.10.1 Dopaminergic (TH-ir) neuronal growth following 1 week post
rotenone treatment of 14 day co-cultures.
The number of cells per field (0.2290mm2) viewed at 400x following different doses.
The Means are calculated from raw data and expressed as mean ± SEM followed by
Two-way ANOVA which was calculated using STDEV, * p<0.05, ** p<0.01, ***
p<0.001 compared to controls.
***
******
***
223
0
2
4
6
8
10
12
14
control vehicle 5nM 10nM 50nM 100nM
Treatments
Nu
mb
er o
f ce
lls/
0.22
90m
m2
TH-ir
Figure 7.10.2 Dopaminergic (TH-ir) neuronal growth following 2 week post
rotenone treatment of 14 day co-cultures.
The number of cells per field (0.2290mm2) viewed at 400x following different doses.
The Means are calculated from raw data and expressed as mean ± SEM followed by
Two-way ANOVA which was calculated using STDEV, * p<0.05, ** p<0.01, ***
p<0.001 compared to controls.
*** ***
*** ***
224
0
2
4
6
8
10
12
control vehicle 5nM 10nM 50nM 100nM
Treatments
Nu
mb
er o
f ce
lls/
0.22
90m
m2
TH-ir
Figure 7.10.3 Dopaminergic (TH-ir) neuronal growth following 3 week post
rotenone treatment of 14 day co-cultures.
The number of cells per field (0.2290mm2) viewed at 400x following different doses.
The Means are calculated from raw data and expressed as mean ± SEM followed by
Two-way ANOVA which was calculated using STDEV, * p<0.05, ** p<0.01, ***
p<0.001 compared to controls.
***
***
******
225
Figure 7.11. 7 day & 14 day old co-cultures rotenone treated.
The number of cells per field (0.2290mm2) viewed at 400x following different doses and
at all weeks of recovery. The Means are calculated from raw data and expressed as
mean ± SEM followed by Two-way ANOVA which was calculated using STDEV, **
p<0.01, *** p<0.001 7 days compared to their respective 14 day groups. (A), 1 week
post; (B), 2 week post; (C), 3 week post.
226
0
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6
8
10
12
14
control vehicle 5nM 10nM 50nM 100nM
Treatments
Nu
mb
ers
of
TH
-ir
cell
s/0.
2290
mm
2
7DIV
14DIV
0
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8
10
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14
16
control vehicle 5nM 10nM 50nM 100nM
Treatments
Nu
mb
ers
of
TH
-ir
cell
s/0.
2290
mm
2
7DIV
14DIV
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16
control vehicle 5nM 10nM 50nM 100nM
Treatments
Nu
mb
er o
f T
H-i
r ce
lls/
0.22
90m
m2
7DIV
14DIV
***
***
***
***
**
***
***
******
***
A
B
C
227
7.3.6 TH-IR CELL DEATH BY MPTP AND ROTENONE.
7.3.6.1 Cytotoxicity of MPTP and rotenone by LDH assay.
To evaluate the degree of cell death, we used an LDH cytotoxicity assay, apoptotic
marker caspase-3 and neuronal degenerating marker fluoro Jade-C (FJ-C) which are
different methods. The LDH cytotoxicity assay detects LDH released from the cytosol of
dead cells and indicates destruction of the cell membrane (Korzeniewski and Callewaert,
[1983]; Decker and Lohmann-Matthes, [1988]). Exposure of co-cultures to 100 nM of
rotenone increased LDH release from the cytosol by 58 % compared to controls. This
was greater than at the lower toxic doses 31 % (5 nM) and 42 % (10 nM). MPTP on the
other hand resulted in 52 % release of LDH compared to controls at 100 μM. This was
lower than with rotenone but greater when compared to relative lower doses (34 % at 5
μM, 42 % at 10 μM) (Figure 7.12).
7.3.6.2 Fluoro Jade C (FJ-C)
The Fluoro Jade C antibody was used to detect degenerating neurons following MPTP
and rotenone treatment. Both high and low doses of MPTP and rotenone induced
degeneration of neurons despite their intact processes and cell bodies compared to control
(Figure 7.13A). The loss of cell body integrity and processes however, were clearly
marked by FJ-C. The toxicity of rotenone and MPTP can be shown by the loss of cell
body and processes (Figure 7.13 B, C, D, E).
228
Figure 7.12. Lactate Dehydrogenase Cytotoxicity Assay (LDH) of MPTP &
Rotenone treated co-cultures.
The lactate dehydrogenase (LDH) cytotoxicity assay was used to detect LDH in the
culture medium after its release from the cytosol of dead cells. 14 day cultures were
treated with either rotenone or MPTP at 5, 10 and 100 (nM/μM) for 24 hrs then allowed
to recover for 7 days in normal culture medium. On 7th day medium was collected and
snap frozen in – 80 °C and thawed when analysis was carried out. Data is represented
as the mean (absorbance) calculated from raw data and expressed as mean ± SEM
followed by Two-way ANOVA which was calculated using STDEV, # p<0.01 BSA vs.
control; *** p<0.001 BSA vs. all doses both MPTP & rotenone; control vs all doses both
MPTP & rotenone; *** p<0.001 5 nM vs 100 nM; * p<0.05 5 μM vs. 100 μM. n = 5
per group.
229
7.3.6.3 MPTP and rotenone induced Caspase-3 activation
Treatment of co-cultures with MPTP and rotenone at high doses (100 μM; 100 nM,
respectively) induced caspase-3 activation at one week of recovery. However only a
small number were TH+ve compared to controls (Figure 7.14, 7.15, 7.16). Caspase-3
was more prominent at the lower doses of MPTP and rotenone (5 μM; 5 nM,
respectively) compared to controls (Figure 7.17, 7.18).
0
0.2
0.4
0.6
0.8
1
1.2
1.4
BSA Control 5 10 100
Treatments
Ab
sorb
ance
lev
els
(450
nm
)
MPTP (uM)
Rotenone (nM)
#***
***
****
230
Figure 7.13. MPTP & rotenone treated co-cultures 1 week post treatment-
Degenerating neurons-FJC.
(A) No degenerating neurons were seen in control co-cultures; 3 weeks. (B) MPTP: 100
μM; (C) 10 μM; (D) rotenone: 100 nM; (E) 10 nM. Bar = 10 μm.
A
B C
D E
231
Figure 7.14. Co-localisation of TH-ir and caspase-3 in co-cultures.
Control 3 weeks, no neurotoxin treatment (A) TH-ir (Red); (B) caspase-3 (green); (C)
merge; (x 400). Bar = 10 μm.
A B
C
>
>
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>
>
> >
>
> >
>
232
Figure 7.15. Co-localisation of TH-ir and caspase-3 in co-cultures following MPTP
treatment at one week recovery period.
MPTP treatment at 100 μM (A) TH-ir (Red); (B) caspase-3 (green); (C) merge; (x 200).
Bar = 10 μm.
A B
C
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233
Figure 7.16. Co-localisation of TH-ir and caspase-3 in co-cultures following
rotenone treatment at one week recovery period.
Rotenone treatment at 100 nM (A) TH-ir (Red); (B) caspase-3 (green); (C) merge; (x
200). Bar = 10 μm.
A B
C
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234
Figure 7.17. Co-localisation of TH-ir and caspase-3 in co-cultures following
MPTP at one week recovery period.
MPTP treatment at 10 μM (A) TH-ir (Red); (B) caspase-3 (green); (C) merge; (x
200). Bar = 10 μm.
>
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235
Figure 7.18. Co-localisation of TH-ir and caspase-3 in co-cultures following
rotenone treatment at one week recovery period.
Rotenone treatment at 10 nM (A) TH-ir (Red); (B) caspase-3 (green); (C) merge;
(x 200). Bar = 10 μm.
A B
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7.4 DISCUSSION
7.4.1 Dose and time dependent toxicity of neurotoxins
A dose and time dependent response was observed when the co-cultures were treated
with either neurotoxin. 7 day old co-cultures treated with MPTP were not affected as
much at the lower doses as compared with the higher doses, post treatment, in a dose
dependent manner. However 14 day old co-cultures demonstrated more vulnerability at
both low and high doses, as shown by the significant reductions in TH-ir cells at all
weeks post treatment. Our results confirm multiple other reports indicating a dose-
dependent toxicity by either MPTP (Chang and Ramirez 1986; Notter et al. 1988;
Christie-Pope et al. 1989; Michel and Agid 1992; Koutsilieri et al. 1993; Schmidt et al.
1997; Lannuzel et al. 2003; Madsen et al. 2003) and rotenone (Gao et al. 2002; Ahmadi
et al. 2003; Xu et al. 2003; Gille et al. 2004; Kress and Reynolds 2005; Testa et al. 2005;
Radad et al. 2006; Klintworth et al. 2007; Ren and Feng 2007) in different animal models
and organotypic cell cultures.
As shown in the 7 and 14 day co-cultures, toxin application at a critical time (or age of
cells) has different effects on the TH-ir cells in the following recovery weeks. This was
an unexpected finding. Previous studies applied toxins at different doses for a specific
time and analysed cell response within a week. This study has demonstrated that acute
exposure to the complex I inhibitors MPTP or rotenone that cell death and/or
neurodegeneration may occur within hours, days or weeks post toxin application.
Therefore, DAergic survival after neurotoxin treatment is dependent on the time and dose
at which the toxin is applied and when the cells are analysed. Studies have suggested that
237
rotenone causes cell death of TH-ir cells at a very high concentration of rotenone (2μM
vs. 100nM rotenone) or where cells have been treated for longer periods of time (48 vs.
24 h) (Hartley et al. 1994; Bal-Price and Brown 2000; Klintworth et al. 2007). Cultures
treated with rotenone at a low dose of 20 nM for 24 hrs resulted in a 50 % reduction of
TH-ir cells (Radad et al. 2006).
In vivo studies have shown that MPTP induces neuronal destruction in the SNpc and the
VTA with the active phase of degeneration beginning 12 h post-injection and lasting up
to 4 days. During this period, there was a greater decrease in TH-ir neurons than in Nissl-
stained neurons suggesting that MPTP can cause a functional synthetic loss in TH,
without necessarily destroying the neuron (Jackson-Lewis et al. 1995). In vitro, MPP+
treatment (30 μM MPP+ for 3, 6 or 24 hrs) can induce cell death, with increased caspase-
3 in western blot analysis and the DEVDase activity assay suggesting apoptotic death
(Chu et al. 2005). Whereas MPP+ has been shown to elicit a minimal amount of caspase-
3 activation (Hartmann et al. 2000) which was detectable only at the 24-h time point by
the DEVDase activity assays (2-fold increase vs. control). This may explain that even at
the lower doses of MPTP and rotenone, TH-ir cell reduction continued in the weeks post
treatment (Hartley et al. 1994). Apart from the effects observed in these cells there are
also functional changes. Following exposure of cells to 1 μM MPP+ 3H-DA uptake was
decreased to 38% compared to controls within the first two hours of incubation and to 8%
after 48 hours (Koutsilieri et al. 1993). Loss of TH-ir cells became evident at 0.1 μM
MPP+ (80% of control) leading to maximal toxicity at 10 μM (20% of control).
238
7.4.2 Toxicity of neurotoxins on Dopaminergic co-cultures varies with the age of co-
cultures
Although there was a significant dose dependent TH-ir cell loss following MPTP and
rotenone treatment in both 7 and 14 day co-cultures, the effects were most potent in 14
day co-cultures especially at the higher doses. It was unexpected that the TH-ir cells
were still affected after a single dose exposure and three weeks of recovery. The
vulnerability of 14 day compared to 7 day co-cultures to both toxins was confirmed by
cell counts and changes in the intensity of TH-ir and cytologic alterations. A possible
reason for this is that neurotoxins may have a greater effect on the cells that have already
migrated away from the thick tissue. By the second week in vitro there are more single
TH-ir cells to be exposed to the neurotoxins. This method while retaining organotypic
connections allows examination of a thinned out region of tissue where individual DA
cells can be morphologically examined and counted free of influence of overlying
DAergic cells in the thicker regions of tissue. Short exposure to both neurotoxins at low
and high doses were more deleterious in 14 day than 7 day co-cultures as indicated by
minimal TH-ir cell recovery at all time periods of post treatment. 7 day co-cultures were
able to maintain a greater number of TH-ir cells during the weeks of post treatment
especially after the lower doses. The mesencephalic DAergic cells are more vulnerable
to MPTP or rotenone mitochondrial insult and in addition rotenone is a more potent toxin
compared to MPTP or MPP+ in this DAergic cell model.
The vulnerability of 14 day to 7 day co-cultures is at variance with previous studies.
Jakobsen et al., (Jakobsen et al. 2005) observed that one week old cultures derived from
239
embryonic day 12 mice were more vulnerable to MPP+ than three week old cultures.
Others have demonstrated that immature or newly plated neurons were more sensitive to
MPP+ and as lengths of time in culture increased the neurotoxin effects were diminished
(Danias et al. 1989; Golde et al. 2002; Jakobsen et al. 2005). These studies however
employed embryonic dissociated cultures of the mesencephalon and compared one week
cultures with 4 week cultures. Thus, different experimental paradigms were used and
cannot be compared directly with this current study. In our study 14 day co-cultures were
more vulnerable and showed significant reduction in TH-ir cell numbers compared to 7
day old cultures. Thus, it would seem that the age of the DAergic neurons in organotypic
cultures is an important factor in toxin-induced degeneration. In vivo studies have
supported our finding where a low systemic dose of rotenone had no effect on young rats,
but led to a 20–30% reduction of TH-ir neurons in the SN of older rats (Phinney et al.
2006).
The establishment of axonal projections and innervation of ST by TH-ir cells of VM
plays an important role in the vulnerability to these neurotoxins. Some studies have
implicated an excitotoxic component to the injury induced by DAergic neurotoxins in
animal models because drugs such as the N-methyl-d-aspartate (NMDA) receptor
antagonist MK801 can protect against the loss of DAergic neurons caused by MPTP
(Turski et al. 1991). In addition, DAergic toxicity after chronic application of MPP+ or
rotenone does not occur unless an excitotoxic stimulus such as NMDA is co-applied
(Kress and Reynolds 2005). NMDA receptor blockade with MK801 lead to a rescue of
60-70% of mature neurons. The expression of NMDA receptors increases with
240
maturation in culture (Phinney et al. 2006). This could explain why 7 day co-cultures
were less vulnerable than 14 day co-cultures as TH-ir cell innervation of ST would be
greater in 14 day co-cultures. Interestingly Maeda et al., (Maeda et al. 1998) observed
that VM and ST slices co-cultured at a wide distance from each other neither resulted in
failure of outgrowth of DAergic fibres to the ST and no cytotoxic response to NMDA.
VM slices co-cultured at a close distance to the ST were affected even at the lowest
concentration of neurotoxins. Thiruchelvam (Thiruchelvam et al. 2003) and Li (Li et al.
2005) also report application of neurotoxins to be age-dependent as catecholamine
synthesis and uptake in immature cells renders these vulnerable to oxidative stress. In
addition, DAergic neurons innervate and develop synaptic connections in the ST over the
course of 2–3 weeks in vitro (Plenz and Kitai 1998) and as glutamatergic connections
increase so do their vulnerability to excitoxic stress.
7.4.3 Glial cells play an important role in DAergic degeneration
The role of nigral glial cells is important in PD and in this model of DA cell growth. As
previously discussed in chapter 2, MPTP is only toxic when it is converted to its
metabolite MPP+ and transported into the cell via the DAT (Chiba et al. 1985;
Gainetdinov et al. 1997; Miller et al. 1999)and concentrated in the mitochondria to inhibit
complex I (Ramsay et al. 1991b; Ramsay et al. 1991a; Russ et al. 1996). MPTP is taken
up by glial cells where inside monoamine oxidases convert it to the toxic metabolite.
Administration of MPTP to mice lacking MAO-B shows that SNpc neurons do not die
(Shih and Chen 1999) further demonstrating that MAO conversion of MPTP to MPP+ is
necessary to cause cell death. Several articles suggest that glial cells can contribute
241
directly to the toxic effects of MPTP, primarily through the action of free radical
mediators such as inducible nitrous oxide synthase (iNOS) (McGeer et al. 1988; Hirsch et
al. 1998; McNaught and Jenner 1999). MPTP administration leads to a rapid gliosis
(Schneider and Denaro 1988), followed by an increase in the production of iNOS
(Zietlow et al. 1999). iNos has been shown to interact with free radicals to produce
peroxynitrite, which can interact with tyrosine residues in cellular proteins and inhibit
neuronal signal transduction (Ferrante et al. 1999; Kuhn et al. 1999; Kuhn and Geddes
1999). In a model of iNOS action that extends the role of glia, Hirsch and Hunot (Hirsch
and Hunot 2000) suggest that MPTP acts directly to induce cytokines which in turn
activate iNOS. The iNOS is then released from the glial cells to act upon DAergic
neurons, inducing damage within these cells. Although differences in iNOS levels have
not been examined in different strains of mice, it is possible that this pathway could
underlie some of the differences in toxicity that occur after administration of MPTP or
MPP+ (Smeyne et al. 2001).
The organotypic cell culture system differs from primary cell lines and dissociated
cultures in that it does not just contain a single pure population of a specific cell type but
maintains the different cell types and original morphology and characteristics of the
tissue of interest. Neurons in slice culture are still associated with glial cells and more
likely to express their in vivo morphology and characteristics than dissociated neuronal
cultures although reaggregated (Hemmendinger et al. 1981) and plated (Berger et al.
1982) cultures have produced DAergic neurons retaining their ability to interact and form
morphologically characteristic pattern of fibres. MPTP was applied in our studies to slice
242
cultures containing glia. MAO-A and MAO-B are the major enzymes that catalyze the
oxidative deamination of monoamine neurotransmitters such as DA, noradrenaline, and
serotonin in the central and peripheral nervous systems. MAO B is mainly localized in
glial cells which convert MPTP to MPP+ (Bove et al. 2005; Nagatsu and Sawada 2006).
MAO-B inhibitors such as L-deprenyl (selegiline) and rasagiline have been shown to be
effective in the treatment of PD (Nicotra and Parvez 2000; Nagatsu and Sawada 2006),
preventing both PD behavioural manifestations and cell loss in the SN of monkey
(Heikkila et al. 1984a; Langston and Ballard 1984) and reduction in DA levels in rodents
(Heikkila et al. 1984b). Our results have shown that MPTP induced TH-ir cell loss in a
dose-dependent manner and maintained significant cell loss in weeks of post treatment.
It would be of interest to compare the actions between MPP+ and MPTP as other studies
have only used slice cultures with embryonic rat midbrain and MPP+ and/or excitotoxic
toxins (Ito et al. 1999; Katsuki et al. 1999).
Rotenone is highly lipophilic and freely crosses brain cellular membranes to accumulate
in subcellular organelles including the mitochondria. In mitochondria, rotenone impairs
oxidative phosphorylation by inhibiting reduced nicotinamide adenine dinucleotide
(NADH)-ubiquinone reductase activity through its binding to the subunit of the
multipolypeptide enzyme complex I of the electron transport chain (Syed and Winlow
1991; Schuler and Casida 2001a, b). The mechanism of cell death may also occur from
withdrawal of trophic support from glia and microglia. Little neurotoxicity was detected
in the mesencephalic neuron-enriched cultures after treatment for 8 day with up to 20 nm
rotenone compared to significant and selective DAergic neurodegeneration in co-cultures
243
with glia two days after treatment with 20 nm rotenone or 8 days after treatment with 1
nm rotenone (Gao et al. 2002). This is a result of glial presence especially microglia as it
markedly increases neuronal susceptibility to rotenone. Rotenone stimulated the release
of superoxide from microglia that was attenuated by inhibitors of NADPH oxidase.
Furthermore, inhibition of NADPH oxidase or scavenging of superoxide significantly
reduces the rotenone-induced neurotoxicity (Gao et al. 2002).
In vivo studies further support the notion that glial cell activation contributes to toxicity
as rotenone administration in young rats has no effect but produces a 20-30% reduction in
TH-ir cells in the SN of older rats (Pasinetti et al. 1999). GFAP-ir is an astrocytic marker
which has been observed to be less intense in young rats when compared to aged rats
(Pasinetti et al. 1999). It is recognised that there is an increase in the basal level of glial
cell activation with aging (Linnemann and Skarsfelt 1994; Rozovsky et al. 1998; Gao et
al. 2003b) and studies have also shown involvement of microglia and increased
inflammatory markers following rotenone toxicity (Hogue 1953; Gao et al. 2002; Gao et
al. 2003a). Several identified growth factors for DA neurons are present in diminished
quantity or absent in the adult and aging organism (Wilcox and Derynck 1988; Schaar et
al. 1993; Seroogy and Gall 1993; Stromberg et al. 1993; Choi-Lundberg and Bohn 1995;
Kaseloo et al. 1996; Nosrat et al. 1996; Kornblum et al. 1997; Widenfalk et al. 1997)
these include BDNF, NGF, NT-3, CNTF and GDNF which could contribute to the
diminished neurite-promoting activity and increased sensitivity to neurotoxins.
7.4.3 Rotenone vs MPTP neurotoxicity
244
Results from our current study show that rotenone is more toxic than MPTP. There is a
large unit difference (nM to μM) between the toxins and the 14 day old co-cultures
treated with rotenone at the higher doses (50 and 100 nM) induced a marked TH-ir cell
reduction at all three weeks of recovery compared to MPTP. In addition, the neurotoxic
effects of MPTP were variable. The observation that complex I was uniformly inhibited
throughout the brain (and the rest of the body) but that only the SN neurons degenerated
(Betarbet et al. 2000; Hoglinger et al. 2003b; Sherer et al. 2003a) indicates that the
DAergic neurons of the SN have a higher sensitivity to rotenone, presumably because of
the interplay between rotenone-induced defects in complex I and the oxidation of DA.
Rotenone is a high affinity inhibitor of complex I, and interacts specifically with the ND1
and PSST subunits of complex I (Nicolaou et al. 2000; Schuler and Casida 2001a, b)
compared to MPP+ which has a low affinity for complex I and typically produces 104- to
106-fold less complex I inhibition than rotenone (Mizuno et al. 1987b; Mizuno et al.
1987a; Hasegawa et al. 1990; Ramsay et al. 1991b; Ramsay et al. 1991a; Hasegawa et al.
1995).
7.4.4 MPTP and rotenone activate caspase-3
Both MPP+ and rotenone have been shown to activate caspase-3 in DAergic cells
(Ahmadi et al. 2003; Kaul et al. 2003; Yang et al. 2004a; Ramachandiran et al. 2007).
Co-cultures treated with high doses of MPTP and rotenone (100 μM; 100 nM) did not
induce high numbers of caspase-3 co-localised with TH-ir following one week of
recovery. However, at the low doses of both neurotoxins (5 and 10 μM; nM) there were
a greater number of apoptotic cells compared to controls. Other studies have shown
245
otherwise (Abbott et al. 2003; Chee et al. 2005). An explanation for this is that different
apoptotic pathways may be activated. As there are more than 10 caspases that have been
identified, caspase-3 is believed to be the final executor of apoptotic DNA damage and
activated caspase-3 has been detected (Cohen 1997; Naoi et al. 2000). Therefore, severity
of insult and duration of MPP+ treatment may be responsible for the inconsistent results.
In addition, the inhibition of caspase in primary cultures of DAergic neurons treated with
MPP+ can trigger a switch from apoptosis to necrosis (Hartmann et al. 2001). MPP+ and
rotenone may have the potential to induce caspase-dependent and -independent cell death,
and the final outcome may be determined by several extrinsic factors placed on cells in
defined situations (Han et al. 2003). Despite this, caspase-3 inhibitors such as DEVD has
been successful in preventing apoptosis occurring in DAergic neurons treated with either
rotenone or MPP+ (Dodel et al. 1998; Viswanath et al. 2001; Bilsland et al. 2002; Ahmadi
et al. 2003).
246
CHAPTER 8:
THE NEUROPROTECTIVE EFFECT OF GLIAL CELL-
LINE DERIVED NEUROTROPHIC FACTOR (GDNF) ON
TH-ir CELLS IN CO-CULTURES
247
8.0 INTRODUCTION
The first trophic factor for DAergic neurons was termed glial cell line-derived
neurotrophic factor (GDNF) (Lin et al. 1993). Subsequently, it was shown as a survival
factor in dissociated cultures of DA neurons, and promoted the survival and outgrowth of
DAergic neurons (Engele et al. 1991). GDNF protected DAergic neurons against the
toxic effects of 6-hydroxydopamine (6-OHDA) and 1-methyl-4-phenyl-1,2,3,6-
tetrahydropyridine (MPTP) (Lapchak et al. 1997c; Lapchak et al. 1997b; Lapchak et al.
1997a; Bohn et al. 2000). These observations have raised the possibility that GDNF can
serve as a therapeutic agent in PD, a condition associated with the premature loss of
DAergic neurons (Olanow and Tatton 1999; Airaksinen and Saarma 2002) as well as in
other neurological diseases, including amyotrophic lateral sclerosis (Manabe et al. 2002;
Manabe et al. 2003), and aging (Ai et al. 2003).
Mesencephalic DAergic neuronal development has been suggested to be dependent on
GDNF (Krieglstein et al. 1995a; Krieglstein 2004) thus we employed organotypic
cultures which develop nigrostriatal connections to examine the role of GDNF in
protecting and/restoring DAergic neurons from toxic stress. Current DAergic therapies
for PD specifically target the symptoms of PD and are not known to alter disease
progression. In addition, the effectiveness of these treatments wanes with time, and
patients develop significant side effects including dyskinesias that can become disabling.
For these reasons, the development of neuroprotective or neuroregenerative therapies for
PD would represent a tremendous advance in our ability to treat the disease.
Collectively, in vitro and in vivo data, in both the rodent and nonhuman primate, provide
248
strong support for a role for GDNF in treating PD, showing considerable promise for
continued investigation as a drug candidate for the treatment of PD. Not much work has
been performed using GDNF and neurotoxins-MPTP and rotenone in organotypical cell
culture derived from postnatal rats, thus we aim to further characterise this system.
8.1 AIMS
Assess:
i) Dose-response of GDNF on TH-ir cells.
ii) Neuroregenerative properties of GDNF post toxin exposure on TH-ir cells.
iii) Neuroprotective effects GDNF pre-toxin exposure on TH-ir cells.
8.2 METHOD
The slice co-cultures were prepared according to the method described in Chapter 4.
8.2.1 Dose-response of GDNF on TH-ir cells
Co-cultures of ventral mesencephalon (VM) and striatum (ST) were prepared from P4-5
Sprague-Dawley rats. Co-cultures received cytostatics between 4-5 days in culture for 24
hrs then fresh media was added and changed twice a week. The co-cultures were then
divided into four groups (see Table 8.1).
24 hrs after cytostatic exposure co-cultures received fresh media change, co-cultures that
were treated with GDNF (Promega; human glial cell-line derived neurotrophic factor;
249
G2781-5 mg) were added in the media according to their assigned concentrations at every
media change.
Table 8.1
Dose-response of GDNF on TH-ir cells
GROUP TREATMENTS
1 Control (normal medium)
2 10ng/mL GDNF + medium
3 20ng/mL GDNF + medium
4 100ng/mL GDNF + medium
At one, two, and three weeks during the treatment a minimum of three tubes were stained
for tyrosine hydroxylase-immunoreactivity (TH-ir), a marker of DAergic neurons, and
counter stained with haematoxylin. The cultures were then analyzed in a blind fashion.
The number of TH-ir cells was counted in a minimum of 3 fields under 400x
magnification with a light microscope. Analysis of number of TH-ir cells was performed
by a 2 way-ANOVA followed by Bonferroni's Multiple Comparison Test. P value of
<0.05 was considered significant.
250
8.2.2 TH-ir cells response to GDNF following neurotoxin treatment
Co-cultures were prepared as described in Chapter 4, and grown for two weeks before
receiving their treatments (see Table 8.2). Neurotoxins were incubated with the co-
cultures for 24 hrs after which the media was changed with fresh media; GDNF was
added to the media twice a week for a week in the designated co-cultures. One week post
recovery period, a minimum of 3 tubes per group were immunocytochemically stained
for tyrosine hydroxylase (TH-ir). The number of TH-ir cells was counted in a blind
fashion and analysed.
8.2.3 TH-ir cell response to pre-treatment with GDNF followed by neurotoxin insult.
Co-cultures were prepared as described in Chapter 4, and grown for two weeks with
GDNF supplement at 20ng/mL at each media change before receiving their neurotoxin
treatments (see Table 8.3). Co-cultures were incubated with neurotoxins for 24 hrs and
then the media was exchanged with fresh media. Media changes were carried out twice
a week for a week. One week post recovery period a minimum of 3 tubes per group were
immunocytochemically stained for tyrosine hydroxylase (TH-ir). The number of TH-ir
cells was counted in a blind fashion and analysed. See Figure 8.4.1A, 8.4.1B & 8.4.1C
for schematical diagram of experiments.
251
Table 8.2
Post treatment of Co-cultures with GDNF following MPTP/rotenone
exposure
GROUPS TREATMENT GDNF (20ng/mL) 1 Control/normal medium -
2
Control/supplement +
3 100 μM MPTP
-
4 100 μM MPTP
+
5 50 μM MPTP
-
6 50 μM MPTP
+
7 100 nM rotenone
-
8 100 nM rotenone
+
9 50 nM rotenone
-
10 50 nM rotenone
+
11 10 μM MPTP
-
12 10 μM MPTP
+
13 5 μM MPTP
-
14 5 μM MPTP
+
15 10 nM rotenone
-
16 10 nM rotenone
+
17 5 nM rotenone
-
18 5 nM rotenone
+
252
Table 8.3
Pre-treatment of Co-cultures with GDNF prior to MPTP/rotenone exposure
GROUPS TREATMENT PRE-TREATMENT
GDNF (20ng/mL)
1 Control (normal
medium)
-
2 Control (medium with
supplement)
+
3 10 μM MPTP
-
4 10 μM MPTP
+
5 5 μM MPTP
-
6 5 μM MPTP
+
7 10 nM rotenone
-
8 10 nM rotenone
+
9 5 nM rotenone
-
10 5 nM rotenone
+
253
A
DIV 0 4 7 14 21
B
DIV 0 4 7 14 21
C
DIV 0 4 7 14 21
Figure 8.1 Schematic diagram of the experiments.
A) 5 day old co-cultures were treated with GDNF at 10, 20 or 100 ng/mL for 3 weeks at
every media change, TH-ir cell analysis at 1 week intervals. B) 14 day old co-cultures
were treated with MPTP or rotenone, 24 hrs post toxin co-cultures were treated with
GDNF (20 ng/mL) for 1 week at media changes followed by TH-ir cell analysis. C) Co-
cultures were pre-treated with GDNF (20 ng/mL) straight after cytostatic treatment up
until toxin exposure, 1 week of recovery in normal media followed by TH-ir cell analysis.
* GDNF supplementation x2/wk; # normal media change x2/wk.
Cytostatics for
TH-ir analysis
Cytostatics for
*
MPTP/RotenoneTH-ir analysis
#
*
TH-ir analysis TH-ir analysis
* *
Cytostatics for
MPTP/RotenoneTH-ir analysis
* # *
254
8.2 Analysis of cell death
The medium from toxin treated at 7 day recovery and GDNF post treatment (same time at
toxin recovery day) will be collected for LDH determination as previously described in
chapter 7. FJC staining will also be carried out to assess whether GDNF post treatment
of co-cultures provided neuroprotection.
8.3 RESULTS
8.3.1 GDNF at varying doses did not induce a significant increase in TH-ir cells
Despite the increasing concentration of GDNF in the media, the number of TH-ir cells
did not significantly increase throughout the weeks compared to normal controls except
at the highest dose (100 ng/mL) during the first week. There was, however a trend
(statistically not significant) for the number of TH-ir cells to increase with increasing
GDNF dose (Figure 8.2A; B; C & E, 8.3).
8.3.2 GDNF treatment promotes increase in cell size and branching.
The main difference between cells treated with GDNF and those that were not is that the
size and number of branches of TH-ir cells increased markedly compared to normal
controls (Figure 8.2A; E, 8.4). The difference in intensity of TH-ir staining between the
normal controls and GDNF-treated cells was distinctive as they were much darker
(Figure 8.2A; B).
8.3.3 Post GDNF treatment following MPTP and rotenone exposure.
255
GDNF treatment at 20ng/mL does not provide any significant protection to the TH-ir
cells at the higher doses of MPTP (50 & 100 μM). However at the lower doses of 5 and
10 (μM) MPTP there is a significant increase of TH-ir cells when treated with GDNF
(Figure 8.5 & 8.6). An obvious change was in the morphology of the cells, where the cell
body and processes were intact compared to toxin treated cells. Similar results were seen
with rotenone (Figure 8.7 & 8.8). Further support of GDNF protection against MPTP and
rotenone was observed by LDH assays (Figure 8.9 & 8.10) showing protection is
significant only at the lower doses of toxins but not at the higher doses. FJC staining
shown GDNF post treatment was able to provide partial protection of TH-ir cells
indicated by the decreased number of degenerating neurons (Figure 8.11).
8.3.4 Pre-treatment with GDNF followed by MPTP and rotenone exposure.
Since treatment with GDNF of co-cultures exposed to higher doses of MPTP and
rotenone resulted in no protection, we decided to pre-treat co-cultures with GDNF in
conjunction with MPTP and rotenone at the lower doses. Unlike post-treatment with
GDNF, TH-ir cells were not significantly protected against MPTP and rotenone during
pre-treatment with GDNF (Figure 8.12, 8.13 & 8.14). However, a trend is observed with
co-cultures pre-treated with GDNF. These had higher numbers of TH-ir cells after MPTP
and rotenone exposure compared to co-cultures that did not receive any GDNF treatment
post-toxin exposure. In addition, the morphology of TH-ir cells improved with increased
neuritic branching and cell body size.
256
Figure 8.2. TH-ir cells in co-cultures treated with GDNF.
7 day co-cultures treated with GDNF (A) control, DAB; (B) 5 ng/mL; 10 ng/mL; (C)10
ng/mL; (D) 100 ng/mL for 1 week followed by 1 week of recovery, DAB + nickel
solution for dark grey colour. (E) 100 ng/mL treated TH-ir cell. Note: enlarged cell body
and long processes plus multiple branching, in addition boutons/nodules on them. Image
analysis at 400 x; bar = 10 μm.
257
B A
C D
E
258
0
2
4
6
8
10
12
14
16
1 2 3
weeks post treatment
Nu
mb
er o
f T
H-i
r ce
lls/
0.22
90m
m2
control
5 ng/mL
10 ngmL
100 ng/mL
Figure 8.3. TH-ir analysis of 7 day co-cultures treated at varying doses of GDNF.
Cultures were treated for two weeks and allowed to recover in fresh normal media;
following consecutive weeks, co-cultures were randomly fixed and processed for TH-ir
cell analysis. Data is presented as Mean ± SEM. Two-way ANOVA followed by
Bonferroni’s post tests was performed on Mean ± SD. * p < 0.05 (Control vs 5 ng at 3
weeks); *** p <0.001 (Control vs 100 ng at 1 week); * p < 0.05 (5 ng vs 100 ng at 1 & 3
weeks), n = 5 for each group.
*
****
*
259
0
1
2
3
4
5
6
7
1 2 3
weeks post treatment
NO
. o
f b
ran
chin
g/c
ell
control
5 ng/mL
10 ng/mL
100 ng/mL
Figure 8.4. 7 day co-cultures treated at varying doses of GDNF.
Cultures were treated for two weeks then allowed to recover in fresh normal media;
following consecutive weeks co-cultures were randomly fixed and processed for TH-ir
process analysis. Data is presented as Mean ± SEM. Two-way ANOVA followed by
Bonferroni’s post tests was performed on Mean ± SD. *** p <0.001 (Control vs 5, 10 &
100 ng/mL at 1, 2 & 3 weeks); α p < 0.01 (5 ng vs 10 ng at 3 weeks); # # # p <0.001 (5
ng vs 100 ng/mL at 1, 2 & 3 weeks); § p <0.001 (10 ng vs 100 ng/mL at 1, 2 & 3 weeks),
n = 50 for each group.
# # # # # #
# # #
αααα
****** ***
§ §
§
260
Figure 8.5. The neuroprotective effects of GDNF on TH-ir cells following MPTP
exposure.
14 day co-cultures treated with MPTP followed by GDNF treatment (A) control; (B) 5
μM; (C) 5 μM + GDNF (20 ng/mL); (D) 10 μM; (E) 10 μM + GDNF. Image analysis
at 400 x; bar = 10 μm.
A
B C
D E
261
0
2
4
6
8
10
12
naïve
contro
l
GDNF 5
uM10
uM50
uM
100u
M
Treatments
Nu
mb
er o
f T
H+
ve c
ells
/0.2
290m
m2
No Toxins
MPTP
MPTP+GDNF
Figure 8.6. The effects of GDNF post treatment following MPTP exposure.
14 day co-cultures treated with varying doses of MPTP for 24 hrs, followed by 7 days of
media changes with GDNF supplementation (20 ng/mL). Cultures were fixed and
processed for TH-ir cell anlaysis. Data are presented as Mean ± SEM, one-way ANOVA
was performed on Mean ± SD followed by Bonferroni’s Multiple comparison test. *** p
<0.001 (Naïve control vs 5, 10, 50 & 100 μM); # # # p <0.001 (GDNF vs 5, 10, 50 &
100 μM); * p < 0.05 (5 μM vs + 5 μM GDNF); * p <0.05 (10 μM vs 10 μM + GDNF).
***
* *
# # #
262
Figure 8.7. The neuroprotective effects of GDNF on TH-ir cells following rotenone
exposure.
14 day co-cultures treated with rotenone followed by GDNF treatment (A) control; (B) 5
nM; (C) 5 nM + GDNF (20 ng/mL); (D) 10 nM; (E) 10 nM + GDNF (20 ng/mL).
Image analysis at 400 x; bar = 10 μm.
A
B C
D E
263
0
2
4
6
8
10
12
naïve
contro
l
GDNF
5nM
10nM
50nM
100n
MTreatments
Nu
mb
er o
f T
H+
ve c
ells
/0.2
290m
m2
No toxins
Rotenone
Rotenone+GDNF
Figure 8.8. The effects of GDNF post treatment following rotenone exposure.
14 day co-cultures treated with varying doses of rotenone for 24 hrs, followed by 7 days
of media changes with GDNF supplementation (20 ng/mL). Cultures were fixed and
processed for TH-ir cell anlaysis. Data are presented as Mean ± SEM, Two-way
ANOVA was performed on Mean ± SD followed by Bonferroni’s Multiple comparison
test. *** p <0.001 (Naïve control vs 5, 10, 50 & 100 nM); # # # p < 0.001 (GDNF vs 5,
10, 50 & 100 nM); * p < 0.05 (5 nM vs 5nM +GDNF); * p <0.05 (10 nM vs 10 nM +
GDNF).
***
**
# # #
264
0
0.2
0.4
0.6
0.8
1
1.2
1.4
BSA
contr
ol
GDNF cont
rol
5uM
10uM
100u
M
Treatments
Ab
sorb
ance
lev
els
(450
nm
)
controls
GDNF control
- GDNF
+ GDNF
Figure 8.9. LDH ELISA of GDNF post treatment following MPTP exposure.
14 day co-cultures treated with varying doses of MPTP for 24 hrs, followed by 7 days of
media changes with GDNF supplementation (20 ng/mL). Culture media were collected
on final day of recovery and snapped frozen in -80 °C. Bovine serum albumin (BSA)
used an internal control and as blocking. Data are presented as Mean absorbance levels ±
SEM, one-way ANOVA was performed on Mean absorbance levels ± SD followed by
Bonferroni’s Multiple comparison test. *** p <0.01 (Control vs 5, 10, & 100 μM); ***
p < 0.001 (GDNF Control vs 5, 10 & 100 μM); * p <0.05 (10 μM vs 10 μM + GDNF);
** p <0.01 (5 μM vs 5 μM + GDNF).
***
** *
265
0
0.2
0.4
0.6
0.8
1
1.2
1.4
BSA
cont
rol
GDNF cont
rol
5nM
10nM
100n
M
Treatments
Ab
sorb
ance
lev
els
(450
nm
)
controls
GDNF control
- GDNF
+ GDNF
Figure 8.10. LDH ELISA of GDNF post treatment following rotenone exposure.
14 day co-cultures treated with varying doses of rotenone for 24 hrs, followed by 7 days
of media changes with GDNF supplementation (20 ng/mL). Culture media were
collected on final day of recovery and snapped frozen in -80 °C. Bovine serum albumin
(BSA) is used an internal control and as blocking. Data are presented as Mean
absorbance levels ± SEM, one-way ANOVA was performed on Mean absorbance levels
± SD followed by Bonferroni’s Multiple comparison test. *** p <0.001 (Control vs 5,
10 & 100 nM); *** p < 0.001 (GDNF control vs 5 , 10 & 100 nM); * p <0.05 (5 nM vs 5
nM + GDNF); * p <0.05 (10 nM vs 10 nM + GDNF).
****
**
***
266
Figure 8.11. FJC-staining of GDNF Post-treatment on co-cultures following MPTP
and rotenone exposure.
(A) control; (B) 14 day co-cultures treated with MPTP 5 μM + GDNF (20 ng/mL); (C) 5
μM; (D) rotenone 5 nM + GDNF (20 ng/mL); (E) 5 nM. Image analysis at 400 x; bar =
10 μm.
A
B C
D E
267
Figure 8.12. The neuroprotective effects of GDNF Pre-treatment on TH-ir cells
following MPTP and rotenone exposure.
(A) control; (B) 14 day co-cultures treated with MPTP 5 nM; (C) 5 nM + GDNF (20
ng/mL); (D) rotenone 5 nM; (E) 5 nM + GDNF (20 ng/mL). Image analysis at 400 x; bar
= 10 μm.
A
B C
C E
268
0
2
4
6
8
10
12
Naïve control GDNF control 5 uM 10 uM
Treatments
Nu
mb
er o
f T
H-i
r ce
lls/
0.22
90m
m2
Controls
- GDNF
+ GDNF
Figure 8.13. The effects of GDNF Pre-treatment following MPTP exposure.
GDNF (20 ng/mL) treatment followed cytostatics up to 14 day; co-cultures were then
treated with low doses of MPTP for 24 hrs, followed by 7 days of media changes with
normal media. Data are presented as Mean ± SEM, one-way ANOVA was performed on
Mean ± SD followed by Bonferroni’s Multiple comparison test. *** p < 0.001 naïve
control compared to all treatments excluding GDNF control; # # # p < 0.001 GDNF
control compared to all treatments excluding naïve control.
# # #
***
269
0
2
4
6
8
10
12
Naïve control GDNF control 5 nM 10 nM
Treatments
Nu
mb
er o
f T
H-i
r ce
lls/
0.22
90m
m2
Controls
- GDNF
+ GDNF
Figure 8.14. The effects of GDNF Pre-treatment following rotenone exposure.
GDNF (20 ng/mL) treatment followed cytostatics up to 14 day; co-cultures were
thentreated with low doses of rotenone for 24 hrs, followed by 7 days of media changes
with normal media. Data are presented as Mean ± SEM, one-way ANOVA was
performed on Mean ± SD followed by Bonferroni’s Multiple comparison test. *** p <
0.001 naïve control compared to all treatments excluding GDNF control; # # # p <
0.001 GDNF control compared to all treatments excluding naïve control.
# # #
***
270
8.4 DISCUSSION
8.4.1 The neurotrophic effects of GDNF on DAergic neurons survival and
development.
GDNF supports the development of embryonic DAergic neurons (Lin et al. 1993) and is
particularly important for postnatal survival of mesencephalic DAergic neurons
(Granholm et al. 2000). Despite being named glial-derived growth factor, it is mainly
present in the striatal medium spiny neurons that receive DAergic input from the SN (Oo
et al. 2005). GDNF levels in various parts of the brain are reported to be no different in
parkinsonian and control patient brains (Mogi et al. 2001). However, GDNF mRNA
expression is increased in the putamen of PD patients compared with controls (Backman
et al. 2006), and so there is no evidence that loss of GDNF is responsible for development
of PD.
Although GDNF’s neurotrophic effect is on a smaller scale than expected, it did
demonstrate a positive outcome on postnatal DAergic neuron development. The DAergic
neurons were larger with increased cell branching and lengthening of processes and
greater staining intensity which may be the result of an increase in tyrosine hydroxylase
and/or VMAT2 expression induced by GDNF (Emborg et al. 2008). In vivo studies have
shown similar results where GDNF injected into the ST promotes cell survival by
inhibiting cell death and induced regeneration by fiber outgrowth (Hou et al. 1996; Zeng
et al. 2006; Kowsky et al. 2007; Elsworth et al. 2008). It may be that a constant supply of
GDNF over a longer period of time is needed to produce a much greater effect as these
cultures were only exposed to GDNF for a short period of time. Previous studies have
271
used embryonic cultures (Meyer et al. 2000; Jakobsen et al. 2005; Zeng et al. 2006),
genetically modified cells expressing GDNF (Bjorklund et al. 2000; Park et al. 2001;
Kordower 2003) or implantation of a pump in vivo (Palfi et al. 2002) for continous supply
of GDNF.
8.4.2 Neuroprotection and regeneration of DAergic neurons by GDNF.
A variety of experiments in rodent models have shown that GDNF injected directly into
the SN or ST protects DAergic neurons from neurotoxins (Kearns and Gash 1995; Tomac
et al. 1995; Kirik et al. 2000a; Kirik et al. 2000b; Kirik et al. 2004). Elevating GDNF in
the ST by gene therapy is also protective (Kordower et al. 2000; Georgievska et al.
2002b; Eslamboli 2005; Eslamboli et al. 2005). Unlike the case with BDNF, there is also
evidence for restoration of function of injured neurons by GDNF after toxic insults
(Tomac et al. 1995; Kirik et al. 2000a; Kirik et al. 2001)—although restorative actions are
generally not as dramatic as the neuroprotective actions.
A series of influential studies from Gash, Gerhardt, and coworkers (Gash et al. 1996;
Zhang et al. 1997; Grondin et al. 2002; Grondin et al. 2003) have demonstrated GDNF-
induced improvement in bradykinesia, rigidity, and postural instability in monkeys with
stable MPTP-induced hemiparkinsonism. GDNF was effective when given by bolus or by
constant infusion, via three routes of administration: intranigral, intrastriatal, and
intracerebroventricular (ICV). These studies, showing clear improvement in monkeys
with established hemiparkinsonism, stimulated the clinical trials that followed. In
addition to neuroprotective and neurorestorative actions, GDNF also has direct effects on
272
DAergic neurons, modulating excitability via changes in A-type potassium channels
(Yang et al. 2001). This may be a mechanism by which GDNF acutely increases DA
release (Hebert et al. 1996).
8.4.3 GDNF neuroprotection against neurotoxins in slice cultures.
Studies so far have demonstrated protection by growth factors against MPP+ in
embryonic systems such as stem cells (Zeng et al. 2006); midbrain from mice (Son et al.
1999; Jakobsen et al. 2005) or primary DAergic neurons (Chu et al. 2005; Anantharam et
al. 2007a; Anantharam et al. 2007b) and only a few experiments have been carried out on
postnatal slice cultures exposed to MPP+ (Schmidt et al. 1997) and rotenone (Yang et al.
2008). A majority of current findings are based on in vivo studies using MPTP, MPP+ or
6-OHDA as the neurotoxin. Our current study employs MPTP and rotenone in slice
cultures derived from postnatal rats in conjunction with GDNF. We were able to show
neuroprotection from MPTP or rotenone but only at the low doses of neurotoxins. This
may be due to the fact that higher doses (50-100 μM, nM) induce a 75-90% decrease in
DAergic neurons and thus, not a lot of surviving cells are present for treatment with
GDNF. Although TH-ir cells were rescued by GDNF treatment it did not prevent these
cells from degenerating. It may be that the GDNF concentration was too low to provide
neuroprotection against the ongoing degenerating process.
273
CHAPTER 9:
GENERAL DISCUSSION
274
9.1 INTRODUCTION
Organotypic slice cultures are widely used in CNS research to investigate neuronal
functioning and cell-cell interactions in situ. The present study employed organotypic
slice cultures to investigate the effects of two neurotoxins on brain DAergic neurons
together with the neuroprotective role of GDNF against these toxins. Chapter 6 described
the characterization of DAergic neurons using this culture system. The effects of two
toxins, MPTP and rotenone, on DAergic neurons, were described in chapter 7 and the
neuroprotective effects of GDNF were outlined in chapter 8.
9.2 ORGANOTYPIC SLICE CULTURE OF VM AND ST
9.2.1 The advantages of using organotypic slice cultures.
The advantages of the organotypic slice culture model over an in vivo preparation include
its relative simplicity, and ease of access. These factors readily allow studies into the
DAergic innervation of the ST. Furthermore, unlike a typical in vitro system, the
physiological environment of the developing ST neurons is preserved. The slice culture
thus represents a system that is more anatomically and physiologically relevant than
cultures of cell lines (Cattaneo and McKay 1990; Evrard et al. 1990) or dissociated
primary cells (Panula et al. 1979; Bockaert et al. 1986). The maintenance of cell-cell
interactions in slice cultures allows the development and cell biology of VM neurons and
their connections with the ST to be analyzed as if in situ. These properties, and the
experimentally accessible condition of cells in the slice model, make the organotypic
culture of VM and ST a valuable compromise paradigm between the in vivo whole
animal and in vitro dissociated or cell culture systems. Additionally, it is possible with
275
this method to co-culture different brain regions and form neuronal connections between
them (Gahwiler 1988; Ostergaard et al. 1990; Yamamoto et al. 1992). Novel findings can
thus be obtained when the co-culture system is used in neurotoxicologic studies (Maeda
et al. 1998; Katsuki et al. 2001). Organotypic slice co-cultures of VM and ST are also
suitable for electrophysiological studies of the nigrostriatal pathway as the morphological
and electrophysiological characteristics of SN neurons in such preparations are similar to
those in other non organotypic in vitro and in vivo studies (Steensen et al. 1995;
Rohrbacher et al. 2000). Importantly, slice cultures have demonstrated that DAergic
axons do actually form functional synapses with neurons of the ST in vivo (Ostergaard et
al. 1990; Ostergaard et al. 1991; Ostergaard 1993; Holmes et al. 1995; Ostergaard et al.
1995; Ostergaard et al. 1996; Franke et al. 2003).
9.2.2 Disadvantages of using organotypic slice cultures
Unfortunately, organotypic slice cultures do not exactly replicate the in vivo environment
of the cells. This is due to unavoidable severance of existing cell-cell connections during
removal of the tissue. As the in vitro environment is not exactly the same as the in vivo
environment it is unlikely that all the factors which regulate the innervation of ST by
DAergic neurons, in vivo, are present in culture. Despite these drawbacks, the co-culture
system is still closer to representing in vivo conditions, in terms of preservation of the
basic structural organization of the tissue, than single-cell culture systems usually
employed for studies of DAergic neurodegeneration.
9.2.2 Summary of experimental findings
276
In this study we were able to maintain the growth of DAergic cells without any
conditioned medium using, the organotypic culture technique. Furthermore, the
morphology of TH-ir cells observed in the current study resembled that of those seen in
the postnatal rat in vivo (Mytilineou et al. 1983; Ostergaard et al. 1990; Tepper et al.
1994). Our study focused initially on the important role of the ST in the DAergic
development of the VM. This is because it has been shown to have a trophic influence on
the development of DAergic neurons of the SN in vitro (Prochiantz et al. 1979). It was
demonstrated that the growth and maturation of VM DAergic neurons was regulated by,
and dependent on, its specific trophic target (the ST). Growth of TH-ir cells in co-culture
remains regular throughout the 7 weeks of culture and such cells formed dense networks
of neuritic projections. Similar observations were reported earlier by Østergaard et al.
(1991). Single VM (SVM) cultures showed poor TH-ir cell growth with such cells being
smaller with fewer processes during an analogous 6 week period. Poor growth of SVM
DAergic neurons was likely to be due to the absence of the trophic target (ie the ST) and
the lack of contact-dependent mechanisms specific for DAergic fibers (Ostergaard et al.
1990). Striatal membranes or extracts have been demonstrated to have a neurotrophic
effect on mesencephalic DA cells (Prochiantz et al. 1981; Dal Toso et al. 1988).
Furthermore, the development of axonal and dendritic connections have also been shown
to be influenced by the presence of ST cells (Hemmendinger et al. 1981).
Our studies also revealed the role that GFAP-ir astrocytes may play in the co-cultures and
the SVM cultures. Growth of GFAP-ir astrocytes in the SVM was better in the first two
weeks of culture compared to that in the co-cultures. There was, however, no
277
significance difference in GFAP-ir cell numbers between the co-cultures and the SVM
cultures from the third week onwards. GFAP-ir cell numbers, however, never exceeded
the number of TH-ir cells in co-cultures but did so in SVM. These observations suggest
that astrocytes may play a supportive role in the development of DAergic neurons
particularly, in the absence of their trophic ST-target in the SVM.
Astrocytes represent the major non- neuronal cells in the brain, and these cells have been
implicated in the segregation, maintenance, and support of neurons. There is emerging
evidence that astrocytes are an important source of neuroactive substances such as growth
factors, eicosanoids, and neurosteroids, which may subsequently influence neuronal
development, survival, and neurosecretion (Kabbadj et al. 1993; Ojeda et al. 2000;
Araque et al. 2001; Azcoitia et al. 2001). Many studies have demonstrated the
therapeutic potential of astrocyte transplantation in neurodegenerative diseases (Smith et
al. 1987; Lundberg et al. 1996; Ridet et al. 2003; Ericson et al. 2005). It is also believed
that astrocytes may represent ideal vehicles for delivery of neurotrophic factors such as
GDNF into the CNS to provide protection in PD animal models (Ericson et al. 2005;
Yang et al. 2008).
Astrocytes have several unique properties that make them an ideal resource for cell or
gene therapy.
1) Astrocytes can provide growth factors or other molecules to support the growth and
functional maintainence of neurons (Ridet et al. 1997; Araque et al. 2001).
2) Astrocytes represent excellent carriers to help deliver therapeutic molecules into the
brain.
278
3) Astrocytes have the ability to survive for prolonged periods when transplanted into
their native environment (Andersson et al. 1993).
9.3 THE EFFECTS OF MPTP AND ROTENONE ON VM AND ST SLICE
CULTURES
9.3.1 MPTP and rotenone
The discovery of MPTP (Langston et al. 1983) paved the way for understanding the
molecular mechanisms of DAergic cell death in various animal and cell culture models
(Speciale 2002). MPTP is itself non-toxic, but conversion by MAO-B in astrocytes to its
active metabolite, MPP+, renders a much more toxic species which is selectively taken up
by DAergic neurons expressing DAT (Chiba et al. 1985; Javitch et al. 1985; Gainetdinov
et al. 1997). The mechanism of toxicity of MPP+ has been proposed to be mediated
through inhibition of mitochondrial complex I (Nicklas et al. 1985). The rotenone model
of PD (Betarbet et al. 2000) has reinforced the idea that complex I inhibition may actually
be a key factor involved in the death of DAergic neurons and the subsequent development
of Parkinsonism.
9.3.2 Summary of experimental findings
A dose- and time-dependent response was observed when the co-cultures were treated
with either MPTP or rotenone. 7 day old co-cultures treated with MPTP showed less of a
loss in numbers of their TH-ir cells, at lower doses, as compared to the higher doses
during the post-treatment period. 14 day old co-cultures, however, demonstrated a
279
greater vulnerability at both low and high doses, as shown by the significant reductions in
TH-ir cells at all analysis times during the recovery period. Interestingly, the higher
doses of rotenone (50 & 100 nM) were cytotoxic to most cells, whether they were
DAergic or not, whereas with the lower doses TH-ir cells were most affected. This could
be due to its highly lipophilic nature, thus, such that it freely crosses all cellular
membranes and accumulates in subcellular organelles such as the mitochondria (Talpade
et al. 2000; Schuler and Casida 2001a, b; Caboni et al. 2004). In contrast, MPTP, once
converted to MPP+, is specifically taken up by DAT in DAergic neurons, as mentioned
previously.
An MPTP concentration of 100 μM was required to dramatically reduce the number of
TH-ir neurons in the culture. Like rotenone, MPTP not only decreased the number of
TH-ir cells but also of all other cells in culture, indicating that high concentrations of
MPTP produced a general toxic effect. A similar result has been reported for MPP+ and
although most in vitro studies have used MPP+ in rat dissociated embryonic
mesencephalon and primary cell lines (Cohen and Mytilineou 1985; Mytilineou and
Cohen 1985; Mytilineou et al. 1985; Friedman and Mytilineou 1987; Mytilineou and
Cohen 1987; Schinelli et al. 1988; Danias et al. 1989; Otto and Unsicker 1993). Low
doses of 0.1 – 10 μM MPP+ produced a concentration-dependent decrease in the number
of surviving TH-ir neurons. Concentrations equal to, or higher than, 30 μM, however,
were toxic to all cell types (Sanchez-Ramos et al. 1986). Our data did not show any
significant difference in TH-ir cell loss between MPTP and MPP+ in the culture either at
low (5 & 10 μM) and high doses (50 & 100 μM).
280
The numbers of GFAP-ir astrocytes in culture plays an important role in the toxicity of
MPTP. MPTP is not toxic itself, as mentioned previously, its conversion to MPP+ has
been shown to be the main reason for DAergic cell death (Chiba et al. 1985; Javitch et al.
1985; Gainetdinov et al. 1997). MPTP conversion to MPP+ by MAO-B mainly occurs in
astrocytes (Markey et al. 1984; Takada et al. 1990; Di Monte et al. 1991) and not in
DAergic neurons (Westlund et al. 1985). MPP+ is then specifically taken up into
DAergic nerve terminals via plasma membrane-bound DA transporter (DAT) (Chiba et
al. 1985; Javitch et al. 1985). Genetically modified mice with a deficiency of DAT have
been shown to be resistant to MPTP toxicity (Gainetdinov et al. 1997; Takahashi et al.
1997). Thus, the amount of TH-ir cell death is likely to be dependent on the number of
astrocytes available to convert MPTP to its toxic by-product and on the presence of DAT.
Death of TH-ir and other cells at the higher treatment doses may be due to a number of
reasons for example, MPTP, once converted to MPP+, could enter DAergic neurons and
subsequently cause displacement of DA from secretory vesicles (Chang and Ramirez
1986. As a result, cytoplasmic oxidation occurs, due to the generation of oxygen free
radicals upon extracellular DA release {Sulzer, 2000 #1759; Speciale et al. 1998; Sulzer
et al. 2000; Inazu et al. 2001). Oxidation of DA produces free radicals and reactive
quinones (Tse et al. 1976; Graham 1978; Graham et al. 1978), which damage cell
components and therefore exert a toxic effect on neurons (Berman and Hastings 1999;
Olanow and Tatton 1999). It is generally accepted that MPP+ is one of the most potent
DA releasing agents (Markstein and Lahaye 1984; Pileblad et al. 1984; Chang and
281
Ramirez 1986; Rollema et al. 1986; Chiueh et al. 1992a; Chiueh et al. 1992b; Obata and
Chiueh 1992; Lotharius and O'Malley 2000; Obata et al. 2001; Uezono et al. 2001; Obata
2002; Chagkutip et al. 2005).
The common variability in TH-ir cell neurodegeneration was more obvious in the 7 day
old compared with the 14 day old co-cultures, since these cells are less likely to have
established a monolayer neuronal network. The toxic effects of both toxins may depend
on the numbers/density of the cells present. The VM slice contained different subsets of
DAergic neurons and other cells. Thus, the variation in TH-ir cell number after treatment
could be caused by the different susceptibility of the requisite subsets of DAergic neurons
present. For example, MPTP treatment in mice caused the loss of midbrain DAergic
cells; this was quantified as a 69% loss of nucleus A8 cells, a 75% loss of nucleus A9
cells and A10 subnuclei, and a 42% loss of ventral tegmental area cells (German et al.
1996). A single dose of MPTP induced a toxic effect that remained in the post treatment
period. This was demonstrated by using FJ-C staining, which is a marker for neuronal
degeneration.
Rotenone was more toxic than MPTP, as shown by the difference in dose concentrations
required to elicit toxicity. 100 nM rotenone induced a greater TH-ir cell loss than 100
μM MPTP. Even though the inhibition sites of both neurotoxins are thought to be the
same (Ramsay et al. 1991b; Ramsay et al. 1991a) rotenone is highly competitive whereas
MPP+ is a very weak complex I inhibitor (Tipton et al. 1993; Tipton and Singer 1993;
Schuler and Casida 2001a, b). Similar to MPTP, Rotenone has also been shown to
282
induce DA release from DAergic neurons (Zoccarato et al. 2005; Sai et al. 2008). Not
only does rotenone cause a decrease in TH and VMAT2 protein levels, it also increases
DAT protein level and MAO activity in a dose dependent manner (Sai et al. 2008). This
could contribute to increased ROS production and subsequent neuronal degeneration
(Radad et al. 2006; Sai et al. 2008). Both neurotoxins were able to induce casapse-3
activation particularly at the lower doses indicating cell death occurred by apoptosis. FJC
demonstrated that TH-ir cell exposed to MPTP or rotenone at low doses is able to induce
neuronal degeneration which continued in the following weeks of recovery.
9.4 THE NEUROTROPHIC ROLE OF GDNF ON TH-ir NEURONS IN SLICE
CULTURES
9.4.1 GDNF and DAergic neurons
Since the discovery that GDNF is a trophic factor for embryonic midbrain DAergic
neurons in culture (Lin et al. 1993) this compound has been widely studied as a survival
factor. This is the case for DAergic neurons in vivo after grafting (Stromberg et al. 1993;
Rosenblad et al. 1996) and in vitro (Meyer et al. 1999; Meyer et al. 2001). GDNF has
also been studied as a neuroprotectant and/or factor which can enhance regeneration of
injured neurons or augment phenotypic expression of surviving DAergic neurons in
models of PD, both in vitro (Krieglstein et al. 1995b; Krieglstein et al. 1995c; Hou et al.
1996; Eggert et al. 1999; Nicole et al. 2001) and in vivo (Choi-Lundberg et al. 1997;
Rosenblad et al. 2000b; Rosenblad et al. 2000a).
9.4.2 Summary of experimental findings
283
Co-cultures treated with increasing GDNF concentrations showed increases in both TH-ir
cell size and number of dendritic branches. However, the numbers of DAergic neurons
were not significantly influenced by GDNF in a dose-dependent manner, as compared to
controls. The neurotrophic influences of GDNF on TH-ir cell size and branching have
been reported in previous in vitro studies (Lin et al. 1993; Lin et al. 1994; Costantini and
Isacson 2000b; Jakobsen et al. 2005; Hirata and Kiuchi 2007) but not previously in
studies using postnatal SN tissue. Levels of the dopamine metabolites, DOPAC and
HVA, in the culture medium, as well as the amount of high-affinity [3H]-DA uptake were
also significantly increased by GDNF (Hou et al. 1996; Jakobsen et al. 2005). In vivo
studies have shown GDNF to increase the survival, growth and function of DAergic
neurons in particular in intrastriatal fetal nigral DAergic grafts (Lin et al. 1993;
Stromberg et al. 1993; Johansson et al. 1995; Rosenblad et al. 1996; Sinclair et al. 1996;
Apostolides et al. 1998).
Studies of GDNF neuroprotection mainly stem from PD toxin-induced animal models.
MPTP-treated non-human primates receiving intrastriatal and intranigral injections of
GDNF was observed to have more TH- and VMAT2-positive fibers in the areas of
delivery, as well as reductions in their clinical rating scores (Tomac et al. 1995; Gash et
al. 1996; Bjorklund et al. 2000; Kordower et al. 2000; Palfi et al. 2002; Oiwa et al. 2006;
Emborg et al. 2008). Furthermore, administration of GDNF not only improved MPTP-
induced disability and reversed DAergic cell loss in the SN but also diminished L-DOPA-
induced dyskinesia (Iravani et al. 2001). Gene transfer of GDNF to the ST of MPTP
284
monkeys also enhanced the survival and outgrowth of co-implanted fetal DAergic
neurons (Elsworth et al. 2008).
In the current study, GDNF treatment after toxin exposure did not provide any significant
neuroprotection against the higher doses of MPTP or rotenone but, at the lower doses
there was significant protection against both toxins, as indicated by the greater number of
TH-ir cells in GDNF-treated co-cultures compared to non-GDNF treated co-cultures.
These results are supported by various in vitro studies of DAergic neurons exposed to
both MPP+ and GDNF together. GDNF shown to has been shown to protect TH+ve
neurons against both MPP+-induced apoptosis and loss of neuronal processes, as well as
against formation of intracellular reactive oxygen species (ROS) (Krieglstein et al. 1995c;
Hou et al. 1996; Jakobsen et al. 2005; Zeng et al. 2006).
Similar to the current study, it was also observed that GDNF did not alter the total
number of TH+ve neurons, but increased (at high GDNF concentrations) DA cell soma
size and – most dramatically – the production of DA and its metabolites DOPAC and
HVA (Jakobsen et al. 2005). The restorative effect of GDNF on DA levels indeed, was
much stronger than its effect on preservation of TH+ve cell numbers. Additionally, in
vitro studies using MPP+ in primary mesencephalic cell cultures reported that GDNF
could not prevent the acute loss of TH+ve cells, but could prevent delayed cell death
and/or enhance recovery of injured neurons (Hou et al. 1996). Furthermore, pretreatment
with GDNF significantly protected these cultures against MPP+ toxicity (Chao and Lee
1999); {Jakobsen, 2005 #40}. This suggests that GDNF pre-treatment stimulates
285
expression of protective factors which prepares the cellular defence mechanisms against
possible toxic insults (eg. oxidative stress, apoptosis). This was not the case in the
current study, as pre-treatment with GDNF did not provide any significant
neuroprotection against MPTP and rotenone. However, there was a marked
morphological improvement of TH-ir cells treated with GDNF compared to co-cultures
that were not treated with GDNF. The low GDNF dose treatment (20 ng/mL) may be too
low to provide such a siginificant neuroprotection prior to MPTP and rotenone toxic
insult.
Fewer studies have compared the effect of GDNF against rotenone compared to those
against MPTP/MPP+. TH+ve cell-loss as induced by rotenone exposure was significantly
attenuated when mesencephalon cultures were co-incubated with GDNF (Yang et al.
2008). Blockade of GDNF with specific antibodies dramatically abolished the GDNF-
induced neuroprotective effect (Yang et al. 2008). Interestingly, rotenone and MPP+ have
been shown to down-regulate the GDNF receptor, Ret, but unlike MPP+, rotenone did not
affect the expression of other cellular components, including TH and actin, in cultured
cells (Hirata and Kiuchi 2007). This finding is important since there is known to be a
progressive and late-onset loss of DAergic neurons in mice where Ret has been
conditionally deleted (Kramer et al. 2007). These findings establish Ret as a critical
regulator of long-term maintenance for the nigrostriatal DAergic system.
The effects of other NTFs, including nerve growth factor (NGF) and brain derived
neurotrophic factor (BDNF), have been studied in cultures exposed to rotenone (Siegel
286
and Chauhan 2000; Hirata et al. 2006; Jiang et al. 2006). These NTFs and GDNF, as well
as their receptors have been localized by immunocytochemistry to DAergic neurons in
the SN (Nishio et al. 1998; Walker et al. 1998; Chauhan et al. 2001), suggesting both
autocrine and paracrine functions for these neurotrophins. All three NTFs have been
shown to significantly reduce rotenone toxicity to TH+ve neurons in midbrain neuronal
cultures (Jiang et al. 2006). The protective effect of NGF was completely abolished by
inhibiting both the microtubule-associated protein kinase (MEK) and by treating with the
microtubule-stabilizing drug taxol. In a MEK-dependent manner, NGF, BDNF and
GDNF also significantly attenuated rotenone-induced microtubule depolymerization and
ensuing accumulation of vesicles in the soma and elevation in protein carbonyls (Jiang et
al. 2006). NGF in particular, however, did not attenuate the rotenone-induced increase in
lactate levels indicating that NGF may not be involved in the inhibition of mitochondrial
respiration by rotenone (Hirata et al. 2006). However NGF was able to completely
prevent rotenone-induced DNA fragmentation and subsequent cell death. In the current
study GDNF treatment was able to reduce cytotoxicity levels induced by both rotenone
and MPTP measured by LDH assay. The intracellular signaling of NGF-induced cell
survival is largely dependent on Trk, a high affinity NGF receptor (Schramm et al. 2005;
Arevalo et al. 2006). Rotenone exposure of PC12 cells did not affect Trk expression, but
treatment with NGF caused down-regulation of Trk (Hirata et al. 2006), and
internalization of the activated receptor. Furthermore, a specific Trk inhibitor, K252a
(Ohmichi et al. 1992), completely attenuated the protective effect of NGF on rotenone-
induced DNA fragmentation (Koizumi et al. 1988). These results suggest that NGF
attenuates rotenone-induced apoptosis via the Trk signaling pathway. NGF, EGF and
287
FGF-b have also been shown to attenuate rotenone-mediated apoptosis, however IL-6 and
GDNF had little effect on rotenone-induced DNA fragmentation (Hirata et al. 2006).
GDNF may protect DAergic neurons through its activation of antioxidant enzyme
systems since both acute and chronic treatments with this factor caused an elevation
glutathione peroxidase (GPx) activity (Chao and Lee 1999). Acute GDNF treatment also
increased superoxide dismutase (SOD) and catalase (CAT) activities in rat striatum (Chao
and Lee 1999). In addition, chronic GDNF treatment decreased the DA level by
increasing DA turnover (Chao and Lee 1999). These results suggest that GDNF treatment
can activate antioxidant defences, resulting in the efficient removal of ROS. Another
antioxidant enzyme, manganese superoxide dismutase (MnSOD), a primary antioxidant
defence enzyme against superoxide radicals produced within mitochondria, was
overexpressed in transgenic mice (Klivenyi et al. 1998) and this overexpression
significantly attenuated MPTP toxicity (Przedborski et al. 1992). There have also been
conflicting reports in DAergic neuron cultures with overexpression of human SOD
(hSOD1) activity showing no significant resistance to MPP+ treatment when compared to
normal cultures (Sanchez-Ramos et al. 1997). This has been further supported by a more
sensitive measure of DA neuron integrity and function ([3H]-DA uptake) which failed to
demonstrate resistance of hSOD1 cultures to the MPP+ toxin (Sanchez-Ramos et al.
1997).
Apart from increases various antioxidant enzyme activities in midbrain DA neurons
(Chao and Lee 1999) GDNF also protects mesencephalic neurons against apoptosis by
288
up-regulation of bcl-2 and bcl-x through PI3 kinase activation (Sawada et al. 2000).
Further, MAPK/ERK pathway activation is necessary for GDNF to protect against
NMDA-induced neuronal death through reduction of Ca2+ influx (Nicole et al. 2001).
So far most GDNF studies have been undertaken in non-human primates (Gerhardt et al.
1999; Kordower et al. 2000; Iravani et al. 2001; Palfi et al. 2002; Elsworth et al. 2008)
and rodents (Cheng et al. 1998; Date et al. 1998; Date et al. 2001; McLeod et al. 2006;
Schober et al. 2007; Chen et al. 2008; Zhang et al. 2008) in conjunction with
MPTP/MPP+. In vitro studies mainly involved primary midbrain DAergic cells derived
from embryonic rats or mice (Krieglstein et al. 1995b; Krieglstein et al. 1995a; Hou et al.
1996; Jordan et al. 1997; Ling et al. 1998a; Ma et al. 2000; Jakobsen et al. 2005) or
human embryonic stem cells (Zeng et al. 2006) whereas the present study has been
carried out on postnatal rat slice co-cultures. Studies of the effects of GDNF on rotenone
exposure are limited (Hirata and Kiuchi 2007; Cho et al. 2008; Yang et al. 2008).
9.5 Summary
In this study we were able to maintain the growth of DAergic neurons derived from
postnatal rat pups using the organotypic slice cultures. The importance of the trophic
target-ST in the development and survival of DAergic neurons in culture was
demonstrated by poor growth in its absence. Both neurotoxins MPTP and rotenone,
induced DAergic cell degeneration in a dose-dependent manner and TH-ir cell
vulnerability which increased with culture time. Exposure of TH-ir cells to GDNF in co-
cultures increased cell size and induced significant cell branching and lengthening despite
289
no difference in cell numbers compared to controls. TH-ir cells in co-cultures treated
with MPTP and rotenone was protected by GDNF but only at the lower doses of the
neurotoxins.
9.6 Conclusion
Thus, this study has shown the importance of using organotypic slice cultures of VM and
ST, demonstrating TH-ir cell growth and development is regulated by its trophic target-
ST. By establishing this culture system we were able to study cell death of TH-ir cells,
in addition, not only are TH-ir cells vulnerable to MPTP and rotenone treatment, but they
can be protected by GDNF. These results represent important findings in the continued
investigation of the underlying basis against PD.
290
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APPENDICES
Table 1. Co-cultures of ST & VM (TH-ir cells)
Weeks 1 (n=9) 2 (n= 12) 3 (n=9) 4 (n=9) 5 (n=11) 6 (n=9) 7 (n=9)
AVG 8.45 8.609756 11.30556 12.32 11.30556 12.15909 10.89722
STDEV 1.217066 0.473862 0.634648 0.831865 1.716895 0.625227 1.116301
SEM 0.544289 0.136792 0.211549 0.372022 0.517663 0.188513 0.455728
Table 2. Single VM cultures (TH-ir cells)
Weeks 1 (n=10) 2 (n=9) 3 (n=11) 4 (n=9) 5 (n=9) 6 (n=9)
AVG 6 4.6875 6.545455 6.638889 6.888889 6.75
STDEV 0.751068 0.737804 0.926472 0.899294 0.784776 0.823446
SEM 0.237508 0.260853 0.279342 0.299765 0.261592 0.274482
387
Table 3. Co-cultures of ST & VM (GFAP-ir cells)
Weeks 1 (n=9) 2 (n=9) 3 (n=9) 4 (n=9) 5 (n=9) 6 (n=9)
AVG 4.9375 5.75 7.388889 7.305556 8.388889 7.944444
STDEV 0.726585 0.806226 1.076443 1.470072 1.12828 1.286067
SEM 0.209747 0.268742 0.358814 0.490024 0.376093 0.428689
Table 4. Single VM cultures (GFAP-ir cells)
Weeks 1 (n=9) 2 (n=9) 3 (n=9) 4 (n=9) 5 (n=9) 6 (n=9)
AVG 7 6.15625 7.045455 7.777778 7.972222 8.111111
STDEV 0.960769 0.766617 1.033273 1.221501 0.999603 1.115547
SEM 0.303822 0.27104 0.311544 0.407167 0.333201 0.371849
388
Table 5. 7 day co-cultures MPTP treated (TH-ir cells)
Control (n=12)
Vehicle (n=9)
0.5 uM (n=9)
1 uM (n=9) 5 uM (n=9) 10 uM (n=9) 50 uM (n=9) 100 uM (n=9)
AVG 11.22917 12.18519 9.925926 9.740741 8.740741 9.740741 8.166667 7.311111
1 week post STDEV 1.6789 1.144789 0.828619 0.712125 0.712125 1.059484 0.845154 0.792643 SEM 0.484657 0.381596 0.276206 0.237375 0.237375 1.059484 0.281718 0.264214
Control (n=10)
Vehicle (n=9)
0.5 uM (n=9)
1 uM (n=9) 5 uM (n=9) 10 uM (n=9) 50 uM (n=9) 100 uM (n=9)
AVG 13.25 14.11111 13.55556 11.81481 11.59259 10.25926 9.259259 6.977778
2 weeks post
STDEV 0.926809 0.918937 1.250641 1.001423 0.888355 1.129758 0.764229 0.690484
SEM 0.293083 0.306312 0.41688 0.333808 0.296118 1.129758 0.254743 0.230161
Control
(n=12) Vehicle (n=10)
0.5 uM (n=9)
1 uM (n=9) 5 uM (n=9) 10 uM (n=9) 50 uM (n=9) 100 uM (n=9)
AVG 13.89583 13.47222 12.11111 9.037037 9.407407 7.037037 6.638889 5.111111
3 weeks post STDEV 1.258834 1.081959 1.187542 1.453946 1.575272 0.854017 0.723198 0.831817 SEM 0.363394 0.360653 0.395847 0.484649 0.525091 0.284672 0.241066 0.277272
389
Table 6. 14 day co-cultures MPTP treated (TH-ir cells)
Control (n=9) Vehicle (n=9) 5 uM (n=9) 10 uM (n=9) 50 uM (n=9) 100 uM (n=9)
1 week post
AVG 11.51111 9.777778 7.088889 6.911111 4.911111 3.244444 STDEV 0.944415 0.926599 0.792643 0.848052 0.763432 0.679423 SEM 0.314805 0.308866 0.264214 0.282684 0.254477 0.226474
Control (n=9) Vehicle (n=9) 5 uM (n=9) 10 uM (n=9) 50 uM (n=9) 100 uM (n=9)
2 weeks post
AVG 12.53333 12.13333 6.177778 5.511111 5.311111 4.155556 STDEV 1.057441 1.07872 0.983706 0.968181 0.874441 0.928233 SEM 0.35248 0.359573 0.327902 0.322727 0.29148 0.309411
Control (n=9) Vehicle (n=9) 5 uM (n=9) 10 uM (n=9) 50 uM (n=9) 100 uM (n=9)
3 weeks post
AVG 10.57778 11.8 6.822222 5.288889 4.355556 2.355556 STDEV 0.865734 0.94388 0.833636 0.661342 0.773292 1.170772 SEM 0.288578 0.314627 0.277879 0.220447 0.257764 0.390257
390
Table 7. 7 day co-cultures Rotenone treated (TH-ir cells)
Control (n=12)
Vehicle (n=9)
1 nM (n=9) 5 nM (n=9) 10 nM (n=9) 50 nM (n=9) 100 nM(n=9)
1 week post
AVG 11.25 10.66667 10.91667 10.86111 9.25 7.388889 3.733333
STDEV 1.682197 1.603567 1.155731 0.899294 1.360147 1.177838 0.741085 SEM 0.485608 0.534522 0.385244 0.299765 0.453382 0.392613 0.26968
Control
(n=10) Vehicle (n=9)
1 nM (n=9) 5 nM (n=9) 10 nM (n=9) 50 nM (n=9) 100 nM(n=9)
2 weeks post
AVG 13.25 12.55556 8.194444 7.583333 6.638889 6.954545 1.472222
STDEV 0.926809 1.132493 1.037013 0.840918 1.099423 1.010516 1.133543 SEM 0.293083 0.377498 0.345671 0.280306 0.366474 0.304682 0.377848
Control (n=12)
Vehicle (n=10)
1 nM (n=9) 5 nM (n=9) 10 nM (n=9) 50 nM (n=9) 100 nM(n=9)
3 weeks post
AVG 14.11628 14.325 9.805556 7.861111 5.777778 8 6.861111
STDEV 1.383737 1.163273 1.348426 0.899294 0.831904 0.706099 0.960737 SEM 0.399451 0.367859 0.449475 0.299765 0.277301 0.235366 0.320246
391
Table 8. 14 day co-cultures Rotenone treated (TH-ir cells)
Control (n=9)
Vehicle (n=9)
5 nM (n=9) 10 nM (n=9) 50 nM (n=9) 100 nM (n=9)
1 week post
AVG 10.95556 11.08889 7.6 5.377778 4.933333 0.722222
STDEV 0.767391 0.874441 0.767391 0.7772 0.653661 0.80904 SEM 0.255797 0.29148 0.255797 0.259067 0.217887 0.247028
Control (n=9)
Vehicle (n=9)
5 nM (n=9) 10 nM (n=9) 50 nM (n=9) 100 nM (n=9)
2 weeks post
AVG 11.48889 11.08889 5.4 5.155556 1.111111 0.888889
STDEV 0.991377 0.949216 0.939052 0.767391 1.005038 0.982164 SEM 0.330459 0.316405 0.313017 0.255797 0.335013 0.327388
Control (n=9)
Vehicle (n=9)
5 nM (n=9) 10 nM (n=9) 50 nM (n=9) 100 nM (n=9)
3 weeks post
AVG 9.111111 9.533333 6.733333 4.222222 1.888889 1.288889
STDEV 0.9101 0.990867 0.80904 0.95081 1.210226 1.272475 SEM 0.303367 0.330289 0.26968 0.316937 0.403409 0.424158
392
Table 9. TH-ir cells in co-cultures treated with GDNF at varying doses
Naïve Control (n=5)
5 ng (n=5)
10 ng (n=5)
100 ng (n=5)
1 week AVG STDEV SEM
10.583330 0.7216879 0.322749
11.2500 1.682197 0.752301
12.159090 0.6252272 0.27961
13.250000 0.9268087 0.414481
2 week AVG STDEV SEM
11.305560 0.6346478 0.283823
11.0000 1.213954 0.542897
11.541670 1.141287 0.510399
12.555560 1.132493 0.506466
3 week AVG STDEV SEM
12.320000 0.8318654 0.372022
10.5625 1.740180 0.778232
11.600000 0.5163978 0.23094
12.320000 0.8318654 0.372022
393
Table 10. GDNF Dose-Dependent increase in TH-ir Cell branching
Control n=50 5 ng n=50 10 ng n=50 100 ng n=50
1 week exposure AVG
STDEV
SEM
2.714286
0.5
0.070711
4.367346939
0.698029296
0.09871625
4.510204
0.649437
0.091844
5.979592
0.749716
0.106026
2 week exposure AVG
STDEV
SEM
3.020408
0.594762
0.084112
3.897959184
0.467479855
0.066111635
4.16326531
0.65659915
0.09285714
5.632653061
0.487077918
0.06888322
3 week exposure AVG
STDEV
SEM
3.061224
0.555584
0.078571
3.673469
0.473804
0.067006
4.040816
0.454569
0.064286
5.102041
0.684498
0.096803
394
Table 11. GDNF Post-treatment of TH-ir cells in 14 day old co-cultures exposed to MPTP & Rotenone
MPTP Naïve Control
(n=9)
GDNF Control
(n=9)
5 uM
(n=9)
5 uM +
GDNF(n=9)
10 uM
(n=9)
10 uM +
GDNF(n=9)
50 uM
(n=9)
50 uM +
GDNF(n=9)
100 uM
(n=9)
100 uM+
GDNF(n=9)
AVG STDEV
SEM
10.4
1.069966
0.338353
10.2963
1.067521
0.35584
6.444444
0.800641
0.26688
7.666667
0.620174
0.206725
5.6
0.894427
0.282843
7.266667
0.691492
0.218669
3.133333
0.681445
0.215492
3.866667
0.730297
0.23094
1.966667
0.718395
0.227177
2.433333
0.85836
0.271437 Rotenone
5 nM
(n=9)
5 nM
+ GDNF(n=9)
10 nM
(n=9)
10 nM
+ GDNF(n=9)
50 nM
(n=9)
50 nM
+ GDNF(n=9)
100 nM
(n=9)
100 nM
+ GDNF(n=9)
AVG STDEV
SEM
6.259259
0.764229
0.254743
7.888889
0.57735
0.19245
5.066667
0.691492
0.218669
6.133333
0.730297
0.23094
4
0.830455
0.262613
4.266667
0.868345
0.274595
0.6
0.498273
0.157568
0.933333
0.639684
0.202286
395
Table 12. GDNF Pre-treatment of TH-ir cells for 14 days in co-cultures prior to exposure to MPTP & Rotenone
MPTP Naïve Control (n=9)
GDNF Control (n=9)
5 uM (n=5)
5 uM + GDNF (n=5)
10 uM (n=5)
10 uM + GDNF(n=5)
AVG STDEV
SEM
10.6
0.968468 0.306257
8.933333 0.784915 0.248212
6
0.755929 0.338062
7.133333 0.63994 0.28619
4.86666667 0.74322335 0.33237959
6.133333330.639940470.28619008
Rotenone
5 n M
5 nM +GDNF
10 nM
10 nM +GDNF
AVG
STDEV SEM
5.533333 0.915475 0.409413
7.266667 0.883715 0.395209
4.8
0.77459667 0.34641016
6.666666670.899735410.40237391
396
397
398
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