Liphophilic Organic Cations Are Novel Environmental Dopaminergic

LIPOPHILIC ORGANIC CATIONS ARE NOVEL ENVIRONMENTAL DOPAMINERGIC TOXINS THAT MAY CONTRIBUTE TO THE ETIOLOGY OF PARKINSON’S DISEASE

A Dissertation by

Chamila Chathuranga Kadigamuwa

Master of Science, Wichita State University, 2015

Bachelor of Science, University of Kelaniya, 2007

Submitted to the College of Liberal Arts and Sciences and the faculty of the Graduate School of Wichita State University in partial fulfillment of the requirements for the degree of Doctor of Philosophy

December 2015

LIPOPHILIC ORGANIC CATIONS ARE NOVEL ENVIRONMENTAL DOPAMINERGIC TOXINS THAT MAY CONTRIBUTE TO THE ETIOLOGY OF PARKINSON’S DISEASE

The following faculty members have examined the final copy of this dissertation for form and content, and recommend that it be accepted in partial fulfillment of the requirement for the degree of Doctor of Philosophy with a major in Chemistry.

______Kandatege Wimalasena, Committee Chair

______David Eichhorn, Committee Member

______James G. Bann, Committee Member

______Moriah Beck, Committee Member

______George Bousfield, Committee Member

Accepted for the College of Liberal Arts and Sciences

______Ron Matson, Dean

Accepted for the Graduate School

______Kerry Wilks, Interim Dean

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DEDICATION

To my loving Parents & Wife

ACKNOWLEDGEMENTS

I would like to offer my sincerest gratitude to my supervisor Prof. K. Wimalasena, for his guidance, wisdom and support during the course of my research work. He encouraged me to develop my independent and critical thinking and research skills and greatly assisted me with scientific writing. Without his enthusiasm and persistent help this dissertation would not have been possible.

I wish to express my sincere appreciation to the members of dissertation committee,

Profs. David Eichhorn, James Bann, Moriah Beck and George Bousfield for serving in my committee and for their support and advice. I owe a debt of gratitude to Dr. Shyamali

Wimalasena for continuous support throughout my graduate studies and exhaustively proofreading all of the manuscripts and this dissertation.

My sincere thanks go to Dr. Yamuna Kollalpitiya, Dr. Viet Le, and Sumudu Mapa for their help and support in my research works. I would like to thank my family and especially my loving wife, Sumudu Mapa for her endless support and love during this period of endeavor and throughout my life.

Finally I wish to express my gratitude to the faculty and staff of the WSU Chemistry department and friends for assisting me in numerous ways.

ABSTRACT

Specific uptake through dopamine transporter (DAT) followed by the inhibition of the mitochondrial complex I has been accepted as the mechanism for specific dopaminergic toxicity of 1-methyl-phenylpyridium (MPP+), the most commonly used Parkinson’s disease toxin model.

However, MPP+ is taken up into many cell types through a number of other transporters suggesting that, in addition to the uptake, intrinsic vulnerability of dopaminergic cells may also contribute to their high sensitivity to MPP+. To test this possibility, a group of hydrophobic cyanines were employed in a comparative study based on their unique characteristics and structural similarity to MPP+. Here we show that cyanines freely accumulate in dopaminergic

(MN9D, SH-SY5Y) as well as in liver (HepG2) cell lines, but are specifically and highly toxic to dopaminergic cells (IC50s ~100 nM) demonstrating that1000 fold more toxic than MPP+. They cause mitochondrial membrane depolarization, mitochondrial complex I inhibition with potencies similar to the best known mitochondrial complex I inhibitor rotenone, and intracellular

ATP depletion without cell specificity. However, they increase reactive oxygen species (ROS) levels specifically in dopaminergic cells leading to the apoptotic cell death, parallel to well characterized PD models, MPP+ and rotenone. We propose that the specific sensitivity of dopaminergic cells towards all of these mitochondrial toxins (cyanine, MPP+ and rotenone) is most likely due to the expression of low levels of antioxidant enzymes together with the presence of high levels of oxidatively sensitive dopamine (DA) in these cells. The key similarities in structure and toxicity profile between MPP+ and these cyanines further suggest that the lipophilic, cationic cyanine dyes and other similar organic cations could be potent dopaminergic toxins, similar to MPP+. Based on the above findings, these cyanines may be stronger in vivo dopaminergic toxins than MPP+ and their in vivo toxicities must be evaluated.

TABLE OF CONTENTS

Chapter Page

1 INTRODUCTION ...... 1

2 BACKGROUND AND SIGNIFICANCE ...... 6

2.1 Neurological Disorders ...... 6 2.2 Neurodegenerative Diseases ...... 6 2.3 Parkinson's Disease ...... 7 2.4 Risk Factors of PD ...... 8 2.4.1 Genetics of PD ...... 8 2.4.2 Environmental Factors of PD ...... 9 2.4.2.1 Dopaminergic Toxin, 6-OHDA . . . . 10 2.4.2.2 MPTP and MPP+ Model Mechanisms of PD . . . 11 2.4.2.3 Pesticides ...... 15 2.4.2.3.1 Rotenone . . . . . 15 2.4.2.3.2 Paraquat . . . . . 17 2.4.2.3.3 Maneb ...... 18 2.5 Transporters and PD ...... 18 2.5.1 Dopamine Transporter (DAT)...... 18 2.5.2 Norepinephrine Transporter (NET) . . . . . 19 2.5.3 Organic Cation Transporter (OCT) . . . . . 20 2.5.4 Plasma Monoamine Transporter (PMAT) . . . . 20 2.5.5 Vesicular Monoamine Transporter (VMAT) . . . . 21 2.5.6 Calcium Channels ...... 21 2.5.6.1 Voltage-Gated Ca2+ Channels (VGCC) . . . 22 2.5.6.2 Ligand-Gated Ca2+ Channels (LGCC) . . . 23 2.5.6.3 Calcium Channel Inhibitors . . . . . 24 2.6 Mechanistic View of PD ...... 24 2.6.1 Mitochondrial Membrane Depolarization . . . . 24 2.6.2 Mitochondrial Complex I Inhibition . . . . . 25 2.6.3 Production of Reactive Oxygen Species . . . . 25

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TABLE OF CONTENTS (continued)

Chapter Page

2.6.3.1 Dopamine and Reactive Oxygen Species . . . . . 27 2.6.3.2 Anti-Oxidants and Reactive Oxygen Species. . . 30 2.6.3.3 Antioxidant Enzymes and Reactive Oxygen Species . . 31 2.6.3.3.1 Super Oxide Dismutase . . . . 32 2.6.3.3.2 Catalase (CAT) . . . . . 33 2.6.3.3.3 Glutathione Peroxidase (GPx) . . . 33 2.6.4 Apoptosis ...... 34 2.7 Dopaminergic Toxicity of Cyanine Dyes...... 36 2.8 Cell Culture ...... 39 2.8.1 PC12 Cells ...... 40 2.8.2 SH-SY5Y Cells ...... 40 2.8.3 MN9D Cells ...... 40 2.8.4 HepG2 Cells ...... 41

3 EXPERIMENTAL METHODS ...... 42

3.1 Materials ...... 42 3.2 Instrumentation ...... 43 3.3 Methods ...... 43 3.3.1 Cell Culture ...... 43 3.3.2 Differentiation of MN9D Cells . . . . . 44 3.3.3 Measurement of Cellular Uptake of Cyanine . . . . 44 3.3.4 Measurement of Cell Viability . . . . . 45 3.3.5 Mitochondrial Localization of Cyanines . . . . 46 3.3.6 Measurement of Mitochondrial Membrane Potential . . . 46 3.3.7 Rat Brain Mitochondria Extraction . . . . . 46 3.3.8 Mitochondrial Complex I inhibition . . . . . 47 3.3.9 Measurement of Reactive Oxygen Species . . . . 48 3.3.10 Mitochondrial ROS production by Cyanines . . . . 48

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TABLE OF CONTENTS (continued)

Chapter Page

3.3.11 Measurement of the Intracellular DA Content . . . 48 3.3.12 ATP Assay ...... 49 3.3.13 Estimation of Intracellular Superoxide Dismutase (SOD), Catalase and Glutathione Peroxidase (Gpx) Levels . . . 50 3.3.14 Chromatin Condensation Test for Apoptosis . . . . 50 3.3.15 Apoptotic DNA Fragmentation . . . . . 50 3.3.16 Protein Determination ...... 51 3.3.17 Data analyses ...... 51 3.3.12 Technical Statement ...... 52

4 RESULTS ...... 53

4.1 Cellular Toxicities of Cyanines, MPP+ and Rotenone. . . . 53 4.1.1 MPP+ and Rotenone are Specifically Toxic to Dopaminergic Cells . 54 4.1.2 Both 2,2'- and 4,4'- Cyanines are Highly Toxic to Dopaminergic Cells and Cell Specificity of Toxicities are Parallel to that of MPP+ and Rotenone ...... 56 4.1.3 Positively Charged Cyanines are Specifically Toxic to Dopaminergic Cells but Not the Negatively Charged Dye . . 59 4.2 Cellular Uptake of MPP+ and Cyanine . . . . . 60 4.2.1 MPP+ Uptake into All Three Cell Lines is Saturable at Low MPP+ Concentrations ...... 60 4.2.2 Both 2,2'- and 4,4'- Cyanines are Taken up into All Cells with Similar Specificity and Efficiency . . . . 62 4.2.3 2,2'-Cyanine Uptake into MN9D Cells is Non-Saturable and Time Dependent ...... 62 4.2.4 Both Positively and Negatively Charged Cyanine Dyes are Taken up into MN9D Cells . . . . . 63 4.2.5 The Effect of DAT and NET Inhibitors on 2,2’-Cyanine Uptake . 64 4.2.6 Effect of Extracellular Na+ and Ca2+ on 2,2’-Cyanine Uptake . 64 4.2.7 The Effect of Ca2+ Channel Blockers on 2,2’-Cyanine Uptake . 66

TABLE OF CONTENTS (continued)

Chapter Page

4.3 Effect of Ca2+ and Na+ Ions and Calcium Channels on Cyanine Toxicity . 66 4.3.1 Effect of Extracellular Ca2+ and Na+ on the MPP+ and 2,2’-Cyanine Toxicity in MN9D Cells . . . . 67 4.3.2 Effect of Intracellular Ca2+ on 2,2’-Cyanine Toxicity in MN9D Cells ...... 68 4.3.3 The Effect of Ca2+ Channel Blockers and Cation Channel Inhibitors on 2,2’-Cyanine Toxicity in MN9D Cells . . . 68 4.4 Intracellular Cyanines Dyes Localize in Mitochondria . . . 70 4.5 Cyanines, MPP+ and Rotenone Cause Mitochondrial Membrane Potential Depolarization ...... 72 4.6 Inhibition Studies of Mitochondrial Complex I with Cyanines, MPP+ and Rotenone ...... 75 4.7 Effect of Cyanine and Rotenone on ATP Levels in MN9D and HepG2 Cells ...... 77 4.8 Cyanine-, MPP+- and Rotenone-Induced ROS Production in Cells . . 78 4.8.1 2,2'-Cyanine and MPP+ Cause Concentration and Time Dependent Increases in Intracellular ROS Levels in MN9D and SH-SY5Y, but Not in HepG2 Cells ...... 81

4.8.2 Cyanine Induced ROS Production is Localized in the Mitochondrial Matrix ...... 81 4.8.3 The Antioxidant, Ascorbate (Asc) Pretreatment Inhibits the ROS Production by Cyanines, MPP+ and Rotenone Treatments in MN9D Cells ...... 83 4.8.4 Depletion of Intracellular Dopamine (DA) Reduces Cyanines, MPP+ or Rotenone Mediated ROS Production in MN9D Cells . 86

4.9 Asc Pretreatment Protect MN9D Cells from Cyanine, MPP+ and Rotenone Toxicities ...... 88

4.10 DA Depletion Protect MN9D Cells from Cyanines, MPP+ and Rotenone Toxicities ...... 90 x

TABLE OF CONTENTS (continued)

Chapter Page

4.11 Antioxidant Enzyme Levels are Low in MN9D Cells in Comparison to HepG2 Cells ...... 93 4.12 Cyanine, MPP+ and Rotenone Cause Apoptotic Cell Death in

MN9D Cells ...... 93

5 DISCUSSION ...... 97

6 CONCLUSIONS ...... 105

7 REFERENCES ...... 108

LIST OF TABLES

Table Page

1 Classification of the Major Types of Neurological Disorders . . 6

2 Ca2+ Chelators and Ca2+ Channel Inhibitors . . . . . 23

3 IC50 Values of Different Toxins in Different Cell Types . . . 56

4 Percentages of Rat Brain Mitochondrial Complex I Inhibition by Cyanines, MPP+ and Rotenone in NADH-Ubiquinone Oxidoreductase Activity Experiments ...... 76

5 Comparison of Relative Antioxidant Enzyme Activities of Rat Brain and Liver with Dopaminergic and Liver Cells . . . 93

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LIST OF FIGURES

Figure Page

1 Proposed Mechanism of MPTP Toxicity . . . . . 14

2 Morphological Images of MN9D, SH-SY5Y, HepG2 and PC12 Cells . 53

3 Comparative MN9D, SH-SY5Y and HepG2 cell toxicities of MPP+ and Rotenone ...... 55

4 Comparative MN9D, SH-SY5Y, PC12 and HepG2 Cell Toxicities of 2,2′- and 4,4′-Cyanines ...... 57

5 Comparative MN9D and HepG2 Cell Toxicities of DiOC2(3), DiSC3(3) and DiSC3(5) Cyanine Dyes and the Negatively Charged Dye, DiSBAC2(3) ...... 59

6 Characteristics of the Cellular Uptake of MPP+ and 2,2′-Cyanine . . 61

7 Characteristics of Cellular Uptake of Positively Charged Dyes, DiOC2(3), DiSC3(3) and DiSC3(5) Cyanine Dyes and a Negatively Charged Dye, DiSBAC2(3) by MN9D Cells ...... 63

8 Different Transporter and Ca2+ Channel Inhibitors and Effects of Extracellular Na+ and Ca2+ on 2,2’-Cyanine Uptake into MN9D Cells . 65

9 Effect of Different Buffers on 2,2’-Cyanine Toxicity of MN9D Cells . 67

10 The Effect of the Calcium Chelator, BAPTA and Calcium Channel Inhibitors on 2,2’-Cyanine Toxicity of MN9D Cells . . . . 69

11 Mitochondrial Accumulation of DiSC3(5) Cyanine and the Negatively Charged, DiSBAC2(3) ...... 71

12 The Effects of Cyanines, MPP+, Rotenone and the Negative Dye on Mitochondrial Membrane Potentials ...... 73

13 The Effects of Rotenone, Cyanines, MPP+ and the Negative Dye on Mitochondrial Membrane Potentials and Mitochondrial Complex I Inhibition ...... 76

14 Effects of DiSC3(5) Cyanine and Rotenone on Intracellular ATP Levels in MN9D and HepG2 Cells ...... 77

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LIST OF FIGURES (continued)

Figure Page

15 The Effects of Cyanines, the Negative Dye, MPP+ and Rotenone on the Intracellular ROS Levels ...... 79

16 Cyanine Induces ROS Production in the Mitochondrial Matrix . . 82

17 The Antioxidant, Ascorbate, Attenuates Cyanines, MPP+ and Rotenone Mediated ROS Production in MN9D Cells . . . . . 84

18 2,2′-Cyanine Decreases the Intracellular DA Levels and Prior DA Depletion Attenuates the Cyanines, MPP+ and Rotenone Mediated ROS Production in MN9D Cells ...... 87

19 The Effect of Antioxidant, Asc on MN9D Cell Toxicities of Cyanines, MPP+ and Rotenone ...... 89

20 The Effect of DA Depletion on Cyanines, MPP+ and Rotenone induced MN9D Cell Toxicity ...... 91

21 Cyanines, MPP+ and Rotenone Cause Apoptotic Dopaminergic MN9D Cell Death ...... 95

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LIST OF SCHEMES

Scheme Page

1 Formation of p-quinone by 6-OHDA . . . . . 11

2 Structure of MPPP and Pethidine ...... 12

3 Synthesis of MPPP and MPTP ...... 12

4 Conversion of MPTP to MPP+ ...... 14

5 Structure of Rotenone ...... 16

6 Structures of MPP+ and Paraquat ...... 17

7 Structure of Maneb ...... 18

8 Formation of Oxidants by Electron Transfer Reactions . . . 26

9 The Fenton Reaction (Equation 1) and Haber-Weiss Reaction (Equation 2) . 26

10 DA Oxidation Pathways ...... 29

11 Inhibition of Biosynthesis of DA by α-Methyl-L-Tyrosine . . . 30

12 Mechanism of Radical Scavenging Activity of Ascorbate . . . 31

13 Reaction Catalyzed by Superoxide Dismutase. . . . . 32

14 Involvements of Redox Metal Ions in Reactions Catalyzed SOD . . 32

15 Reaction Catalyzed by Catalase ...... 33

16 Mechanism of the Reaction Catalyzed by CAT . . . . 33

17 Reaction Catalyzed by Glutathione Peroxidase . . . . 33

18 Antioxidant Enzyme System ...... 34

19 Structures of MPP+, 2,2’-Cyanine and 4,4’-Cyanine . . . . 37

20 Structures of DiOC2(3), DiSC3(3) and DiSC3(5) Cyanines and Negatively charged, DiSBAC2(3) ...... 38

21 Types of Cyanines ...... 39

LIST OF ABBREVIATIONS

Abbreviation Systematic Name

2,2’-cyanine 1,1'-diethyl-2,2'-cyanine

2-APB 2-Aminoethoxydiphenylborane

4,4’-cyanine 1,1'-diethyl-4,4'-cyanine

6-OHDA 6-Hydroxydopamine

ΔΨm Mitochondrial Membrane Potential

μL Microliter

μM Micromolar

AAADC L-Aromatic Amino Acid Decarboxylase

AD Alzheimer’s disease

ALA Alpha Lipoic Acid

Asc Ascorbic Acid

ATP Adenosine Triphosphate bp Base pair

BAPTA-AM 1,2-Bis(2-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid tetrakis (acetoxymethyl ester)

BSA Bovine Serum Albumin

CAT Catalase

Caspase C-cysteine rich, asp- aspartic acid moiety containing, ase – proteases

CNS Central Nervous System

CoQ10 Coenzyme Q10

Da Dalton

DA Dopamine

DAPI 4, 6-Diamidino-2-phenylindole

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DAQ Dopamine o-quinone

DASQ Dopamine o-semiquinone

DAT Dopamine Transporter

DCFH-DA 2΄, 7΄-dichlorofluorescin diacetate

DiOC2(3) 3,3'-Diethyloxacarbocyanine

DiSBAC2(3) Bis-(1,3-Diethylthiobarbituric Acid)Trimethine Oxonol

DiSC3(3) 3,3′-Dipropylthiacarbocyanine

DiSC3(5) 3,3'-Dipropylthiadicarbocyanine

DMEM Dulbecco’s Modified Eagle’s Medium

DMF N,N-Dimethylformamide

DMSO Dimethyl Sulfoxide

DNA Deoxyribonucleic Acid

DOPA 3,4-dihydroxyphenylalanine

DOPAC 3,4-dihydroxyphenylacetic acid

EGTA Ethylene Glycol-bis(beta-aminoethyl ether)-N,N,N’,N’- tetracetic Acid

Em Emission

ER Endoplasmic Reticulum

ETC Electron Transport Chain

Ex Excitation

Fe-S iron-sulfur

FBS Fetal Bovine Serum

GPx Glutathione Peroxidase

GR Glutathione Reductase

GSH Glutathione

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GSSG Glutathione disulfide h Hour

H2O2 Hydrogen Peroxide

HEPES 2-[4-(2-hydroxyethyl) piperazin-1-yl]ethanesulfonic acid

HPLC High-Performance Liquid Chromatography

HPLC-EC HPLC with Electrochemical Detection

HPMP 4-hydroxyl-4-phenyl-N-methylpiperidine

HS Horse Serum

HVA High Voltage-Active

IC50 Half-Maximal Inhibitory Concentration

IP3 Inositol 1,4,5-Triphosphate

Km Michaelis Constant kDa Kilodalton

KRB Krebs-Ringer Buffer

L-DOPA 3,4-Dihydroxy-L-phenylalanine

LGCC Ligand Gated Calcium Channel

LRRK2 Leucine-Rich Repeat Kinase 2

LVA Low Voltage-Active

MAO Monoamine Oxidase

MAO-B Monoamine Oxidase B mg Milligram min Minutes mL Milliliter mm Millimeter mM Millimolar

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Mn-EBDC manganese ethylene-bis-dithiocarbamate

MP 1-methyl-4-piperidone

MPDP+ 1-methyl-4-phenyl-1, 2-dihydropyridinium

MPP+ 1-methyl-phenylpyridium

MPPP 1-Methyl-4-phenyl-4-propionoxypiperidine

MPTP 1-Methyl-4-phenyl-1,2,3,6-tetrahydropyridine

MTT 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) mV Millivolt

NADH Nicotinamide Adenine Dinucleotide (reduced form)

NCX Na+/Ca2+ Exchanger

ND Not Determined

NE Norepinephrine

NET Norepinephrine transporter nmole Nanomole nm Nanometer

NO• Nitric Oxide

• NO2 Nitrogen Dioxide

N2O3 Dinitrogen Trioxide

N2O4 Dinitrogen tetroxide

O2 Molecular Oxygen

•- O2 Superoxide

OCT Organic Cation Transporter

OH• Hydroxyl radical

ONOO- Peroxinitrite

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PARK Genes associated with Parkinson’s disease

Pmole Picomole

PBS Phosphate Buffered Saline

PD Parkinson’s Disease

PDF Parkinson’s Disease Foundation

PINK1 PTEN-Induced Putative Kinase 1

PLP Pyridoxal Phosphate

PMAT Plasma Monoamine Transporter

PQ Paraquat

PT Permeability Transition

RNS Reactive Nitrogen Species

ROI Region Of Interest

ROS Reactive Oxygen Species

SD Standard Deviation

SDS Sodium Dodecyl Sulfate

SLC Solute Carrier Gene

SNCA Synucein Alpha Gene

SOCE Store Operated Ca2+ Entry

SOD Superoxide Dismutase

TH Tyrosine Hydroxylase

THI Tyrosine Hydroxylase Inhibitor

TMRM Tetramethylrhodamine, methyl ester

TRPC Transient Receptor Potential Canonical

UV Ultraviolet

VGCC Voltage-gated Calcium Channel

VMAT Vesicular Monoamine Transporter

Vmax Maximum Velocity

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CHAPTER I

INTRODUCTION

Parkinson’s disease (PD) is the second most common neurodegenerative disease after

Alzheimer’s disease (AD), currently approximately 7-10 million people worldwide. According to the Parkinson’s Disease Foundation (PDF), in the United States alone, there are 50-60,000 new cases of PD diagnosed each year, adding to the one million people who are currently living with

PD (1). The medical cost of treatment and loss of productivity of PD patients were estimated to exceed $14 billion in 2010, and are expected to rapidly increase over the next few decades. The main pathological symptom of PD is the irreversible loss of dopaminergic neurons in the substantia nigra, a specific region in the midbrain causing movement disorders including shaking, muscular rigidity and slowness of movement (2-4). Since there is no cure for PD (5), the current research efforts are focused on the understanding of the etiology of PD in order to develop effective preventive and therapeutic strategies to decrease the financial and social burden of this debilitating disease.

The most common form of PD is sporadic (about 95%), while a minor population of PD

(about 5%) is familial in nature. During the past decade, 11 gene products associated with familial PD including α-synuclein (PARK4), parkin (PARK2), PINK1 (PARK6), DJ-1

(PARK7), and LRRK2 have been identified (6). The most commonly accepted view is that the exposure to environmental factors and toxins may be the cause of PD, especially in individuals with cumulative defects in pathways associated with handling of oxidative stress, mitochondrial function, calcium homeostasis, and the ubiquitin proteasome system. Although many specific examples of environmental factors associated with PD have been identified, the discovery of the specific dopaminergic toxicity of 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP)

1 provided the greatest stimulus for PD research (7-10). The discovery of MPTP not only stimulated studies relating its mechanism of action to pathogenesis in PD, but also provided the first viable model of the motor deficits of PD in primates. Extensive studies have shown that

MTPT itself is not toxic. MAO-B catalyzed conversion of MPTP to its oxidized product N- methyl-4-phenylpyridinium (MPP+), is essential for its neurotoxicity (Scheme 4). Exposure of the CNS to MPP+ itself has been shown to selectively destroy dopaminergic neurons in animal models, which induces the symptoms of PD, suggesting that MPP+ rather than MPTP is the active form of the toxin (11,12). Thus, the specific dopaminergic toxicity of MPP+ has been widely studied to model the etiology of Parkinson’s disease (7-9).

MPP+ is an aromatic molecule with a positively charged quaternary methyl ammonium group (pyridinium) with no other functional groups. The positive charge of the molecule is highly delocalized into its extended system resulting in significant reduction of the localized charge. However, the overall high hydrophilicity of MPP+ prevents it from freely crossing the cell membrane, therefore, requiring a mediated transport in to the cell. It is believed that MPP+ is specifically toxic to dopaminergic cells due to its specific accumulation in dopaminergic cells through the plasma membrane dopamine transporter (DAT) (13,14). The observation that MPP+ is a weak inhibitor of mitochondrial electron transport chain complex-I led to the widely accepted general conclusion that specific uptake into dopaminergic neurons through DAT followed by the inhibition of mitochondrial electron transport chain complex-I is responsible for the specific dopaminergic toxicity of MPP+ (15-17). Subsequently, a number of in vivo and in vitro studies have provided strong experimental support for these conclusions (18-20).

Numerous recent studies show that MPP+ is taken up not only into dopaminergic cells, but also into other neuronal and non-neuronal cells with varying efficiencies through a diverse

2 number of transporters including organic cation transporters (OCT) and non-specific plasma membrane amine transporters (PMAT), (21-25) etc. However, almost all in vivo as well as in vitro studies to date unequivocally show that MPP+ is specifically, highly toxic to dopaminergic cells in comparison to other cell types. These findings raise the intriguing question: “is the specific in vivo toxicity of MPP+ towards dopaminergic cells exclusively due to the specific uptake through DAT?” Since the MPP+ model is the current gold standard for non-familial PD research and pharmacological therapeutic screening, a proper understanding of the mechanism(s) of specific dopaminergic toxicity of MPP+ at the molecular level is of prime importance. Thus, to address this critical question and to gain further insight into the mechanism of specific MPP+ dopaminergic toxicity, we have chosen a group of hydrophobic cyanines for a comparative study based on their structural similarity to MPP+ and other unique properties (Schemes 19 & 20). For example, they are also inert aromatic quaternary ammonium salts with highly conjugated extended  systems similar to MPP+. However, they are significantly more hydrophobic in comparison to MPP+, due to the presence of non-polar steric bulk, suggesting that they may accumulate nonspecifically in any cell, through simple diffusion and not require a specific transporter. Therefore, they are good candidates to test whether the specific uptake is the primary determinant of the selective dopaminergic toxicity of MPP+ and also to gain further insight into the mechanism of MPP+ dopaminergic toxicity and similar toxins.

Here we show that 1,1'-diethyl-2,2'-cyanine (2,2'-cyanine), 1,1'-diethyl-4,4'-cyanine (4,4'- cyanine), 3,3'-diethyloxacarbocyanine iodide (DiOC2(3)), 3,3′-dipropylthiacarbocyanine iodide

(DiSC3(3)) and 3,3'-dipropylthiadicarbocyanine iodide (DiSC3(5)) non-specifically and freely accumulate in dopaminergic (MN9D, SH-SY5Y and PC12) and liver (HepG2) cells, but they are specifically and highly toxic to dopaminergic cells similar to MPP+. Remarkably, they are about

1000-fold more potent dopaminergic toxins in comparison to MPP+ under similar experimental conditions. We further show that cationic cyanines electrogenically accumulate in the negatively charged mitochondria of both dopaminergic MN9D and non-dopaminergic HepG2 cells at high concentrations causing mitochondrial membrane depolarization. They induce the over production of reactive oxygen species (ROS) specifically in dopaminergic cells, leading to apoptotic cell death, again parallel to MPP+. The ROS production by these cyanines was found to be due to their ability to inhibit the mitochondrial electron transport chain complex I, with a potency similar to that of the well characterized neurotoxin and complex I inhibitor, rotenone. Further studies have revealed that low levels of antioxidant enzymes expression, together with the presence of high levels of oxidatively sensitive DA in dopaminergic cells is the likely cause of excessive ROS production, specifically in dopaminergic cells, in comparison to HepG2 cells upon cyanine, MPP+ and rotenone treatments. These and other findings strongly suggest that the specific dopaminergic toxicity of these cyanines is primarily due to the inherent vulnerability of dopaminergic cells towards these types of mitochondrial toxins that lead to excessive production of ROS, and are largely independent of the specific cellular uptake. The key structural and toxicity profile similarities between MPP+ and these cyanines further suggest that the lipophilic cationic cyanine dyes and other similar organic cations could be potent in vivo dopaminergic toxins, similar to MPP+. This conclusion is further supported by the observation that a similar, negatively charged lipophilic dye, Bis-(1,3-Diethylthiobarbituric Acid) Trimethine Oxonol

[DiSBAC2(3)] shows no toxicity towards either dopaminergic MN9D or non-dopaminergic

HepG2 cells.

Finally, cyanines are a family of lipophilic cationic dyes that are commonly used in industry and scientific research. In spite of their wide usage, structural resemblance to MPP+

(Schemes 19 & 20) and the ability to freely accumulate in both cytosol and mitochondria of any cell type, their neurotoxicological properties have not been studied or reported. Based on the above findings, especially the mechanistic similarities of the toxicities of cyanines to that of well characterized PD models MPP+ and rotenone, we propose that cyanines are possible environmental neurotoxins that may contribute to the etiology of PD.

CHAPTER II

BACKGROUND AND SIGNIFICANCE

2.1 Neurological Disorders

Neurological diseases are associated with the disorders of the central and peripheral nervous system, which are composed of the brain, spinal cord, cranial nerves, peripheral nerves, nerve roots, autonomic nervous system, neuromuscular junction, and muscles. According to the

U.S National Library of Medicine, there are more than 600 neurological disorders. Classification of the major types of neurological disorders is listed in Table 1 below (26).

Table 1. Classification of the Major Types of Neurological Disorders

Type Examples Neurogenetic Diseases Huntington’s Disease and Muscular Dystrophy Developmental Disorders Cerebral Palsy and Spina Bifida Degenerative Diseases Alzheimer’s Disease and Parkinson’s Disease Metabolic Diseases Gaucher’s Disease Cerebrovascular Diseases Stroke Trauma Spinal Cord and Head Injury Cancer Brain Tumors Convulsive Disorders Epilepsy Infectious Diseases Meningitis

2.2 Neurodegenerative Diseases

The diseases accompanied by progressive loss of neuronal function or neuronal death are known as neurodegenerative diseases. The most common neurodegenerative diseases are

Alzheimer's disease (AD), Parkinson's disease (PD), Huntington’s disease and Amyotrophic

Lateral Sclerosis (ALS). These neurodegenerative diseases are irreversible, incurable, and result in progressive degeneration and/or death of neuronal cells (27).

2.3 Parkinson's Disease (PD)

PD is the second most common neurodegenerative disease after AD (1). PD is a debilitating neuromuscular motor disorder that is characterized by the following symptoms tremors, muscular rigidity, bradykinesia and altered speech (28). In addition, non-motor symptoms include: sleep abnormalities, fatigue, autonomic disturbances, mood disorders and cognitive dysfunction are also associated with PD. PD was originally described in 1817 by a

British physician, James Parkinson, in the classic report “An Essay on the Shaking Palsy”(29).

The basic pathophysiology of this disease includes degeneration of dopaminergic neurons and accumulation of cytoplasmic Lewy bodies in the pars compacta of the substantia nigra (30).

Numerous studies have shown that about 50-70 % loss of dopaminergic neurons leads to an overall reduction in dopamine (DA) in the striatum, ultimately resulting in the characteristic motor dysfunction observed in PD patients (2-4). The substantia nigra contains high levels of a dark pigment called neuromelanin, which is formed by extensive polymerization of DA (31). A physical manifestation of neurodegeneration in PD is the disappearance of this pigmentation.

Dopaminergic degeneration in PD is characterized by the accumulation of cytoplasmic Lewy bodies which are composed of the protein α-synuclein associated with other proteins such as ubiquitin and neurofilament protein (4,32,33). However the mechanisms responsible for the loss of dopaminergic cells in PD are still unclear. In addition, as mentioned above, there is no cure for PD and most treatments merely control symptoms. Similarly, there is no standard diagnosis that can facilitate accurate early diagnosis of PD (34,35). A combination of biological markers found in blood tests or brain imaging are currently used for the diagnosis of late stages of PD

(36). Since DA itself cannot cross the blood brain barrier (37), administration of the precursor of

DA, L-3,4-dihydroxyphenylalanine (L-DOPA), is used as a treatment for PD. L-DOPA crosses the blood brain barrier via an aromatic amino acid transporter and the blood brain barrier DOPA decarboxylase converts it to DA, leading to the increased striatal DA levels, which improve the

PD symptoms (38).

2.4 Risk Factors for PD

The most important risk factor for PD is age, with the average onset age approximately

50 to 60 years (39). Two other main etiological risk factors of PD are genetic and environmental factors (4,40). While about 10 % of the PD population is known to be associated with genetic factors, 90 % could be associated with environmental factors, including pesticides, herbicides, industrial toxins, and other environmental toxins (40). Many other minor risk factors, including use of well water, excess body weight, exposure to hydrocarbon solvents, farming or agricultural work, living in urban areas or industrialized areas with exposure to heavy metals, high dietary intake of iron, history of anemia and higher levels of education (41) have been also suggested.

2.4.1 Genetics of PD

Although PD is usually a sporadic disease, numerous genetic risk factors that are known to increase the risk to develop PD have been identified. There are 28 distinct chromosomal regions more or less convincingly connected to PD. However, only six of these specific regions contain genes with mutations that conclusively cause monogenic PD (6). Out of the six monogenic PD genes, mutations in SNCA (PARK1) and LRRK2 (PARK8) are responsible for autosomal dominant PD forms, and mutations in Parkin (PARK2), PINK1 (PARK6), DJ-

1 (PARK7), and ATP13A2 (PARK9) account for PD that displays an autosomal recessive mode

8 of inheritance (6). SNCA was the first gene associated with mutations reported to cause early- onset autosomal dominant PD before 50 years of age. However, the disease has a rapid progression and often presents with dementia, cognitive decline, and formation of Lewy bodies in the substantia nigra (42). Mutations in the LRRK2 gene cause the late-onset autosomal- dominant and sporadic PD and progress the disease slowly (43). LRRK2 linked PD usually shows the pathology of pure nigral degeneration without Lewy bodies (44). Parkin linked PD usually starts in the third or fourth decade of the patients’ life, and is usually slowly progressive.

Some of the Parkin mutations, however, are the most frequent cause of juvenile PD, with an age of onset less than 25 years (45). The clinical phenotypes of Parkin-, PINK1-, and DJ-1-linked PD are indistinguishable. All of these mutations show neuronal loss and gliosis, but do not show

Lewy body formation in the substantia nigra (46). ATP13A2 mutations have been found to cause a form of PD named Kufor-Rakeb syndrome. The disease is characterized by rapid progression accompanied by dementia (46).

2.4.2 Environmental Factors for PD

As mentioned above, about 90 % of cases are due to environmental factors. The first synthetic compound to induce PD symptoms which was subsequently used as a PD model is 6- hydroxydopamine (6-OHDA). However, 6-OHDA needed to be directly injected into the brain to cause PD symptoms (47). Therefore, the 6-OHDA is not an ideal model to test the environmental hypothesis for PD induction. The discovery of 1-methyl-4-phenyl-1,2,3,6- tetrahydropyridine (MPTP) as a Parkinson’s causing neurotoxin has provided the greatest stimulus, which has led to a new era of investigation into the etiology of PD. Intravenous administration of MPTP to squirrel monkeys has confirmed the observations in humans, including the specific destruction of substantia nigral dopaminergic neurons resulting in

Parkinsonian symptoms (48,49). These findings strongly support the proposal that environmental factors contribute to the etiology of PD. In light of this evidence, environmental influences such as exposure to pesticides, metals, organic solvents, carbon monoxide, carbon disulfide, plant- derived toxins as well as industrialization, rural environment, well water usage, bacterial and viral infections, have been proposed as contributing factors in the etiology of PD (50).

2.4.2.1 Dopaminergic Toxin, 6-OHDA

6-OHDA, a structural analog of dopamine (DA) and noradrenaline (NE), was first synthesized by Senoh and Witkop in 1959 (51). Similar to DA and NE, 6-OHDA is taken up into catecholaminergic cells through monoamine transporters such as the dopamine (DAT) and norepinephrine (NET) transporters (51,52). This compound causes a massive destruction of nigrostriatal dopaminergic neurons. Highly polar, 6-OHDA does not readily cross the blood- brain barrier, and thus intracerebral infusion is used to introduce it into the brain to observe the toxic effects (53). Uptake through DAT or NET into catecholaminergic cells followed by formation of reactive oxygen species (ROS) by the autoxidation of 6-OHDA is considered the main molecular mechanism underlying 6-OHDA neurotoxicity (54). 6-OHDA is highly susceptible to oxidation and produces high levels of toxic ROS, in comparison to DA, due to the presence of an additional ring hydroxyl group (55).

Autoxidation of 6-OHDA generates the para-quinone and hydrogen peroxide (H2O2) which are highly cytotoxic, and trigger the production of oxygen radicals (Scheme 1) (7,56).

Therefore 6-OHDA -induced cell death is proposed to be due to the production of extracellular toxic ROS and p-quinone (57). Even though 6-OHDA constitutes a valid and useful tool for the investigation of behavioral and cellular dysfunction associated with PD, because of its inability

10 to cross the blood brain barrier, 6-OHDA is not considered to be a good candidate to study the environmental toxin hypothesis for initiation of PD.

Scheme 1. Formation of p-quinone by 6-OHDA

2.4.2.2 MPTP and MPP+ Model of PD

In 1947 Dr. Albert Ziering first synthesized the synthetic opioid, desmethylprodine 1,3- dimethyl-4-phenyl-4-propionoxypiperidine (MPPP), but this drug was never completely developed or marketed. MPPP is an analog of pethidine or meperidine (Demerol) which is generally used to treat moderate to severe pain. Relative to pethidine, the ester group in MPPP is inverted (Scheme 2). The resulting molecule is slightly less potent and has a shorter duration of the effect than morphine. In the 1970s, Barry Kidston, a chemistry graduate student at the

University of Maryland, synthesized MPPP by using Dr. Ziering’s recipe, as a street drug, and injected it into himself (58). A few days after injecting himself with a sample from the synthesized MPPP drug, Kidston became frozen, unable to speak or walk. Later he died from a drug overdose and the autopsy showed that he had PD. Soon after, researchers analyzed the tainted drugs, and concluded that the synthesized MPPP was contaminated with an ester group eliminated byproduct, 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP).

Scheme 2. Structure of MPPP and Pethidine – Arrow shows the position of ester group

Scheme 3. Synthesis of MPPP and MPTP

MPPP is synthesized by the reaction of 1-methyl-4-piperidone (MP) with phenyl lithium followed by acylation of the intermediate product, 4-hydroxyl-4-phenyl-N-methylpiperidine

(HPMP) with propionic anhydride in the presence of sulfuric acid (Scheme 3). However, if this synthesis is not kept under tight control, then the ester group eliminated product, 1-methyl-4- phenyl-1,2,3,6-tetrahydropyridine (MPTP), may be produced. It was believed that Barry Kidston may have performed this synthesis at increased temperature and with shorter reaction times which may have led to the production of significant amounts of MPTP as a byproduct. Later in

1982, a group of young drug users developed a rapidly progressive Parkinsonian syndrome traced to intravenous use of MPPP, which was contaminated with MPTP. In 1983, Dr. J. William

Langston published the accidental discovery of the Parkinson-causing toxin MPTP (48), and later these findings were confirmed by experiments with monkeys (59). Currently, it is widely accepted that MPTP produces an irreversible and severe Parkinsonian syndrome including tremors, rigidity, bradykinesia and freezing in humans and nonhuman primates. These symptoms effectively respond to L-DOPA treatment (60,61). Therefore, MPTP provides a good model to study the mechanism of PD at the molecular level.

The mechanism implicated in dopaminergic toxicity of MPTP is depicted in Fig. 1.

MPTP itself is not toxic, but it’s lipophilicity allows it to readily penetrate the blood brain barrier and enter into the brain (48,62,63). There it accumulates into acidic organelles, mostly lysosomes, of glial cells (astrocytes). Astrocytes and serotonergic neurons contain monoamine oxidase B (MAO-B), which oxidizes MPTP to 1-methyl-4-phenylpyridinium (MPP+). First,

MPTP is oxidized by MAO-B to 1-methyl-4-phenyl-1,2-dihydropyridinium (MPDP+), then

MPDP+ is non-enzymatically disproportionated to produce MPP+ (Scheme 4) (11,12).

Fig. 1. Proposed Mechanism of MPTP Toxicity (20)

MAO-B

Scheme 4. Conversion of MPTP to MPP+

When MPP+ is extruded from glial cells via the organic cation transporter-3 (OCT3) to the extracellular space, it is taken up specifically into dopaminergic cells via the DAT (64).

Therefore, specific dopaminergic toxicity of MPP+ is proposed to be due to the uptake through the presynaptic dopamine transporter (DAT) into the dopamine nerve terminal followed by

14 inhibition of mitochondrial complex I in the electron transport chain (38). This disruption causes reduction in ATP synthesis and generation of free radicals and other reactive oxygen species (27) leading to the apoptotic cell death. MPP+ also competes with cytosolic DA for synaptic vesicle entry through the vesicular monoamine transporter-2 (VMAT-2) (65) and this leads to the increase of cytosolic DA, which undergoes enzymatic and non-enzymatic oxidation to generate hydrogen peroxide and highly reactive oxygen radicals (66). ROS damage mitochondria and other organelles as well as enhancing aggregation of α-synuclein (67) and increasing the demand on proteasomes and lysosomal degradative systems, which leads to further increase in the energy demand and ROS production that ultimately leads to dopaminergic neuron death.

2.4.2.3 Pesticides

Rural residence, well water drinking, and farming have been recognized risk factors for

PD (68), most likely due to high levels of pesticide exposure (69). Pesticides consist of multiple classes and subclasses of insecticides, herbicides, rodenticides, fungicides and fumigants; some of which are known to cause PD. Insecticides that are shown to be associated with PD include rotenone (70), pyrethroids (71), organophosphates (diazinon, chlorpyrifos) (72), and organochlorine (dieldrin, hexachlorohexane) (73,74). Herbicides associated with PD are paraquat and 2,4-dichlorophenoxyacetic acid (75). The dithiocarbamate fungicide, Maneb, is also suspected as a PD causing pesticide (76).

2.4.2.3.1 Rotenone

In 1902, a Japanese chemist, Nagai Nagayoshi, first isolated rotenone from the Derris elliptica plant, and the name rotenone was derived from the Japanese name of the plant

(“roten”). Later, rotenone (Scheme 5) was found as a naturally occurring substance in the roots

15 of many tropical plants (55). Rotenone is used as an effective insecticide, and as a nonselective piscicide. Indigenous peoples have used plants containing rotenone to catch fish, and it is still used as a piscicide to exterminate invasive fish species in lakes (77). The discovery that the dopaminergic toxicity of MPP+ is due to inhibition of mitochondrial complex I prompted the proposal that rotenone is also a PD causing toxin. Rotenone is a lipophilic compound that can freely cross the blood brain barrier without the need of a transporter and accumulates in the brain. In brain cells, rotenone could effectively inhibit the NADH:ubiquinone oxidoreductase in mitochondrial complex I of the electron transport chain (78,79) and reduce ATP production. It has been shown that rotenone binds to NADH:ubiquinone oxidoreductase and inhibits ubiquinone reduction (80) by inhibiting the transfer of electrons from iron-sulfur (Fe-S) centers to ubiquinone, leading to ROS production. Overproduction of ROS causes oxidative damage to

DNA and proteins. In addition, ROS can interact with intracellular nitric oxide (NO), especially superoxide and hydroxyl radicals, leading to peroxinitrite formation, resulting in cellular defects and further damage to DA neurons (81). Rotenone causes cytoplasmic accumulation of α- synuclein aggregates and selective degeneration of nigral dopaminergic neurons manifesting a

Parkinsonian syndrome including bradykinesia, rigidity, and tremors associated with PD (82,83).

Scheme 5. Structure of Rotenone

2.4.2.3.2 Paraquat

As shown in Scheme 6, Paraquat (PQ) (1,1′-dimethyl-4,4′-bipyridinium dichloride) is a quaternary nitrogen herbicide that structurally resembles MPP+ (25). Although it was first synthesized in 1882, PQ’s herbicidal properties were not recognized until 1955 (84).In 1962, PQ became commercially available and is one of the most widely used herbicides in the world (85).

Despite its high hydrophilicity and double positive charge, PQ is known to cross the blood brain barrier, probably via a neutral amino acid transporter (85,86), and is taken up into nigral dopaminergic terminals by DAT (87). However, several in vivo studies have demonstrated that the uptake of PQ is independent of DAT and does not interfere with DA uptake (88,89). After getting into the dopaminergic neurons, similar to MPP+ and rotenone, PQ also induces generation of large amounts of cytotoxic ROS (90). PQ induced Parkinsonian syndromes that recapitulate many features of PD have been shown in many animal models (91). PQ increases lipid peroxidation, decreases levels of antioxidants, impairs mitochondrial function as a weak complex

I inhibitor (86), increases aggregation of α-synuclein (92) and selectively kills nigral dopaminergic neurons.

Scheme 6. Structures of MPP+ and Paraquat

2.4.2.3.3 Maneb

Maneb (manganese ethylene-bis-dithiocarbamate) is a commonly used fungicide that belongs to the dithiocarbamate family. The major active ingredient of Maneb is manganese ethylene-bis-dithiocarbamate (Mn-EBDC) (Scheme 7). Epidemiological studies have shown that chronic exposure of humans to Maneb has been linked to the development of Parkinsonism (93).

Maneb causes selective dopaminergic neurodegeneration by disrupting mitochondrial function, increasing oxidative stress, and inhibiting proteasomal function (76).

Scheme 7. Structure of Maneb

2.5 Transporters and PD

Many transporters and channels in dopaminergic neurons that are directly or indirectly involved in PD have been identified. The most important transporters that are studied in detail related to PD are DAT, NET, OCT, VMAT and PMAT. In addition to these transporters, calcium and sodium channels are also being actively investigated in relation to PD, since calcium and sodium perturbations also play a role in dopaminergic cell death in PD.

2.5.1 Dopamine Transporter (DAT)

DAT is located on the plasma membrane of dopaminergic nerve terminals where neurotransmitters are released (94). It is an 80-kDa protein composed of 620 amino acid residues

18 with 12 transmembrane spanning regions and both carboxyl and amino termini residing intracellularly (95-97). DAT regulates the dopamine concentration in the synaptic cleft through reuptake of dopamine into presynaptic neurons (98). DAT activity is regulated by presynaptic receptors, protein kinases, and membrane trafficking (99). DAT is a Na+/Cl- dependent symporter that co-transports two Na+ ions and one Cl− ion with the dopamine substrate. When ionic gradients are altered and intracellular [Na+] or [Cl-] are increased, the DAT may function in the reverse mode (100). The ion concentration gradient generated by the plasma membrane Na+/K+ ATPase is the driving force for DAT-mediated dopamine reuptake. As mentioned above, 6-OHDA and MPP+ uptake are believed to be directly associated with DAT.

However DAT-dependent Paraquat uptake is controversial. The Km values determined for dopamine and MPP+ uptake through DAT are 1.2 ± 0.2 μM and 3.7 ± 0.3 μM in human neuroblastoma cells, respectively (101). Gene transfection and knockout studies as well as studies with a specific DAT inhibitor, GBR 12909, suggest that DAT plays a role in MPP+ uptake (102).

2.5.2 Norepinephrine Transporter (NET)

Norepinephrine transporter (NET), similar to DAT, contains 617 amino acid residues and 12 transmembrane segments. Both N- and C- termini and a large intracellular loop between the 3rd and 4th transmemebrane segments are located on the extracellular face (103). Similar to

DAT, NET also is a Na+/Cl--dependent symporter that is responsible for the reuptake of norepinephrine (NE). It is found that NET can also reuptake extracellular DA (104). The reuptake of these neurotransmitters is essential for regulating their concentrations in the synaptic cleft (104). The finding that NET may also be involved in the uptake of MPP+ suggests that

NET may also play a role in PD (105). Gene knockout models as well as the specific NET inhibitor, desipramine, are commonly used for studies involved in NET (106,107).

2.5.3 Organic Cation Transporter (OCT)

Similar to DAT and NET, OCT also possesses 12 transmembrane segments. Both N- and

C- termini are on the cytoplasmic face, also similar to DAT and NET, along with large intracellular and extracellular loops. OCT belongs to the solute carrier gene (SLC) superfamily, the largest superfamily of transporters, with 3 isoforms of SLC22A named OCT1, 2, and 3

(SLC22A1-3) which mediate facilitated transport of a variety of structurally diverse organic cations, including many drugs, toxins, and endogenous compounds (108,109). OCT-1 and OCT-

2 are mostly expressed in the kidney, liver, and intestine (110,111) while OCT-3 is expressed throughout the body including the heart, placenta, and brain (111-113). As described above,

MPP+ is extruded from glial cells via OCT-3, a Na+-independent bi-directional cation transporter, which is expressed throughout the brain (113). These results suggest that OCT may also play a role in environmental neurotoxin uptake.

2.5.4 Plasma Monoamine Transporter (PMAT)

Plasma Monoamine Transporter (PMAT) also belongs to the SLC superfamily and is encoded by the SLC29A4 gene (114). PMAT contains 530 amino acid residues containing 11 transmembrane segments. The C-terminus is located on the extracellular face and the N-terminus is on the cytoplasmic face along with a large intracellular loop (111,115). PMAT is a Na+- independent, reversible membrane potential-sensitive cation transporter which predominantly transports monoamine neurotransmitters such as serotonin, dopamine and norepinephrine as well as adenosine (116). In addition to monoamines, PMAT also transports various drugs, toxins, and

20 other compounds that carry a net positive charge at physiological pH, similar to OCT (22).

PMAT is known to be a low-affinity, high-capacity plasma membrane transporter for

+ catecholamines and MPP with Km values of 114 μM, 329 μM and 33 μM for serotonin, DA and

MPP+, respectively (115).

2.5.5 Vesicular Monoamine Transporter (VMAT)

Vesicular monoamine transporter (VMAT) is a 65-85 kDa synaptic vesicle membrane integrated transport protein, containing 12 transmembrane segments. Both N- and C- termini face the cytosolic inner face of the membrane (117,118). VMAT has a large hydrophilic loop containing multiple putative N-glycosylation sites facing the vesicular side of the membrane

(119). VMAT transports monoamine neurotransmitters such as dopamine, serotonin, norepinephrine, epinephrine, and histamine into synaptic vesicles. There are two isoforms of

VMAT named VMAT-1 and VMAT-2. VMAT-1 is predominantly expressed in the adrenal gland, whereas VMAT-2 is expressed in the central nervous system and within the chromaffin granules of the adrenal medulla (117,120,121). VMAT-2 is the most studied among the two, since catecholamines have a three-fold higher affinity for VMAT-2 than for VMAT-1. VMAT is relatively less specific than DAT and NET (66,118). Numerous studies have shown that MPP+ is a substrate for vesicular monoamine transporters, which sequester the toxin inside vesicles thereby protecting cells from its dopaminergic toxicity (122).

2.5.6 Calcium Channels

In addition to providing skeletal structure for bones and teeth, calcium acts as a second messenger and is involved in many cellular functions such as, contraction and relaxation of muscles and blood vessels, blood clotting, glycogen break down, oxidative metabolism as well as

21 conduction of nerve impulses. Under resting conditions, free cytosolic Ca2+ levels are maintained at very low concentrations, in the range of 200-500 nM (123,124), through the action of various plasma membrane transporters and specific endoplasmic reticulum (ER) and mitochondrial

Ca2+/ATPases (125). The major transporters known to regulate Ca2+ levels are voltage gated calcium channels (VGCC) and ligand gated calcium channels (LGCC) which are specifically important for neuronal function. In addition to VGCC and LGCC, there are other channels also involved in neuronal Ca2+ homeostasis such as, Na+/ Ca2+ exchanger (NCX) and transient receptor potential channels (TRPC).The perturbation of Ca2+ homeostasis for an extended period of time may disrupt cellular function and triggers exocytosis and apoptosis (126-128). Therefore, the physiological level of both extracellular and intracellular calcium ions may play a role in has been PD. In agreement with this proposal, the dopaminergic neurotoxin MPP+ s shown to perturb Ca2+ homeostasis in neurons (129).

2.5.6.1 Voltage-Gated Ca2+ Channels (VGCC)

Voltage-gated Ca2+channels (VGCC) are expressed on the membranes of excitable cells and play an important role in neuronal function (130). Under resting conditions, the membrane potential of a cell is in the range of -50 mV to -70 mV (131). This transmembrane potential is maintained by electrogenic Na+/K+ ATPase by pumping 3 Na+ ions out for every two K+ ions pumped into the cell. The resulting charge imbalance results in a net negative charge inside the cell membrane, relative to the outside. Therefore, under resting conditions the membrane is polarized. When this inner negative charge decreases, decreasing so called depolarization, it causes VGCCs to open and allow spontaneous Ca2+ influx (132) along the concentration gradient (under normal conditions intracellular Ca2+ concentration is ~100 nM and extracellular

Ca2+ concentration ~1.5 mM) (133). The increase in intracellular Ca2+ concentration leads to the

22 initiation of a number of Ca2+ sensitive pathways, including exocytotic release of neurotransmitters. VGCCs are classified into high voltage-active (HVA) and low voltage-active

(LVA), based on the voltage required to activate them. They are further classified into sub- groups of L, P/Q, N, R and T types, based on their pharmacological properties and amino acid sequences.

2.5.6.2 Ligand-Gated Ca2+ Channels (LGCC)

LGCC are typically composed of at least two different domains: a transmembrane domain which includes the ion pore, and an extracellular domain which includes the ligand binding location. They are activated through allosteric transition of the protein conformation upon binding of various ligands. Examples of LGCCs are inositol trisphosphate receptor (IP3 receptor), the ryoanodine receptor, the store operated Ca2+ entry channel (SOCE), etc.

Table 2. Ca2+ Chelators and Ca2+ Channel Inhibitors

Inhibitor Function Nifedipine L-type VGCC inhibitor (134) Nitrendipine L-type VGCC inhibitor (135) Mibefradil L-type VGCC inhibitor T-type VGCC inhibitor (136) Verapamil L-type VGCC inhibitor PMAT inhibitor (111,137) Benzamil T-type VGCC inhibitor NCX inhibitor TRPP3 inhibitor (138-140) Flufenamic Acid TRPC inhibitor (141,142) 2-Aminoethoxydiphenylborane IP3 antagonist (2-APB) SOCE inhibitor TRPC inhibitor (143-145) CGP 37157 Mitochondrial NCX inhibitor (146) Thapsigargin Inhibits Ca2+ ATPase of ER (147) BAPTA-AM Membrane permeable Ca2+chelator (148) EGTA Ca2+ Chelator (149)

2.5.6.3 Calcium Channel Inhibitors

In order to study the effects of various Ca2+ channels on uptake and toxicity of various toxins, commercially available, well-characterized pharmacological agents have been used. The inhibitory properties of these agents are listed in Table 2.

2.6 Mechanistic View of Parkinsonian Toxins

In spite of the differences in mechanisms of uptake, most commonly used PD models

MPP+, rotenone, paraquat, etc. appear to follow similar mechanisms of toxicity including mitochondrial membrane depolarization, mitochondrial complex I inhibition, excessive ROS production and finally, specific dopaminergic cell death by apoptosis.

2.6.1 Mitochondrial Membrane Depolarization

Mitochondrial membrane potential (ΔΨm) results from the difference in electrical potential across the inner membrane, generated by pH gradient and ion movement. The ΔΨm drives the synthesis of ATP in the electron transport chain (ETC). Several studies have shown that PD causing neurotoxins such as MPP+, PQ, rotenone, for example (150,151), depolarize the

ΔΨm and deplete intracellular ATP (152).

Depolarization of ΔΨm triggers the opening of the permeability transition [PT] pore (79) in the mitochondrial membrane (153). PT pore opening allows release of certain apoptogenic factors, including cytochrome C from the mitochondria into the cytosol, thereby triggering the apoptotic cell death pathway. In addition, Kroemer et al. (154) have shown that a reduction in

ΔΨm is accompanied by production of reactive oxygen species (ROS), contributing to apoptotic cell death.

2.6.2 Mitochondrial Complex I Inhibition

Mitochondrial complex I which is also known as NADH:ubiquinone oxidoreductase, is a crucial mitochondrial enzyme complex that oxidizes NADH with the concomitant reduction of ubiquinone (155). Schapira et al. (17) have found that mitochondrial complex I activity was significantly reduced in patients with PD. Based on this finding, they proposed that the compounds which inhibit mitochondrial complex I, such as MPP+, rotenone, paraquat, could be considered as environmental toxins that may cause PD.

Inhibition of mitochondrial complex I activity depresses rates of ATP synthesis and depolarization of mitochondrial membrane potential (156). In addition, complex I inhibition leads to excessive generation of ROS (157). The combination of reduced ATP synthesis, depolarization of the mitochondrial membrane, and increased ROS promote the opening of the

PT pore and apoptotic cell death (156).

2.6.3 Production of Reactive Oxygen Species

Overproduction of ROS is believed to be responsible for the toxicity of MPP+, pesticides and other environmental toxins that are implicated in neurodegenerative diseases, especially PD

(158). ROS are also implicated in many other diseases such as heart failure, atherosclerosis ischemia-reperfusion, endothelial dysfunction, hypertension (159), rheumatoid arthritis, and diabetes (159-162).

The production of intracellular reactive free radical species that exceed their deactivation through intracellular antioxidant defense mechanisms is defined as oxidative stress (163). Free radicals are unstable, very reactive molecules containing one or more unpaired electrons. The most common biological forms of free radicals are reactive oxygen species (ROS) and reactive

•- nitrogen species (RNS) (164,165). The most common ROS are superoxide (O2 ), hydrogen

• • peroxide (H2O2) and hydroxyl radical (OH ) and the most common RNS are nitric oxide (NO ),

- • peroxinitrite (ONOO ), nitrogen dioxide ( NO2), dinitrogen trioxide (N2O3), and dinitrogen tetroxide (N2O4).

There are many natural and unnatural cellular pathways that are responsible for the increased production of ROS including activities of enzymes such as NADH oxidases and cytochrome P450, as well as conditions such as inflammation, smoking, UV radiation, toxins, and aging (166). However, mitochondrial respiration is also responsible for the production of a large fraction of intracellular ROS. There are many mitochondrial toxins including those discussed above that could uncouple or inhibit the electron transport chain, resulting in the partial reduction

•- of O2 to produce reactive oxygen species such as superoxide (O2 ) and hydrogen peroxide

• (H2O2). H2O2 could be further broken down to highly reactive OH , under certain physiological conditions (Scheme 8) (167). For example, redox active metals like Fe2+and Cu2+ and organic

• compounds like quinones can catalyze the breakdown of H2O2 to produce OH through Fenton and Haber-Weiss reactions as shown in Scheme 9.

Scheme 8. Formation of Oxidants by Electron Transfer Reactions

Scheme 9. The Fenton reaction (Equation 1) and Haber-Weiss reaction (Equation 2)

Even though high concentrations of ROS and RNS are lethal to cells, at low or moderate concentrations, they are necessary for maturation of cellular structures and can act as weapons for the host defense system. For example, phagocytes release free radicals to destroy invading pathogenic microbes as part of the body’s defense mechanism against diseases. In addition,

ROS-mediated signaling pathways are required to maintain cellular homeostasis (168,169). On the other hand, the overabundance of these species causes oxidative stress leading to deleterious effects on DNA, proteins, lipids and other biological molecules (170-172). This can lead to apoptotic and/or even necrotic cell death. Reactions of ROS with the unsaturated fatty acids in the cell membrane leads to the formation of free radical chain reactions (173). These ROS induced lipid peroxidation chain reactions result in protein oxidation and loss or weakening of cell membrane structure and function (174). ROS induced protein oxidation also affect the activities of DNA repair enzymes and DNA polymerases, further impairing cellular integrity and function. These protein damages include modification of enzyme activities and damage to membrane transporters and receptors, which potentially, collectively lead to apoptotic cell death

(174-176).

2.6.3.1 Dopamine and Reactive Oxygen Species

DA oxidation has been proposed as one of the specific mechanisms for generation of

ROS in dopaminergic neurons (177). Several different pathways have been identified for the oxidation of DA; either enzymatically or spontaneously. Dissociation of protons from the two hydroxyl groups promotes the oxidation of DA to dopamine o-quinone (DAQ). This reaction could occur through either one or two steps. The one step reaction is catalyzed by the enzyme, tyrosinase (178) to convert DA to DAQ by a two-electron oxidation, with oxygen as an electron

−· acceptor that generates superoxide radical anions (O2 ) (Reaction 4) (Scheme 10). In the two-

27 step reaction, oxidation occurs through a one electron oxidation of DA to form a dopamine o- semiquinone radical (DASQ) (Reaction 1), which is subsequently oxidized to DAQ (Reaction 2)

−· there by producing two moles of O2 . In addition, two DASQ radicals can disproportionate to form one molecule of DAQ and one molecule of DA (Reaction 3) (Scheme 10). Dopamine o- quinone is not stable in the cytosol at physiological pH and it generates aminochrome through leukodopimnochrome. Next, aminochrome tautomerizes to 5,6-dihydroxyindole and then to 5,6- indolequinone, a precursor for the neuronal pigment neuromelanin (179), thereby generating one

−· more O2 (180) (Scheme 10). Accordingly, DA oxidation itself can form substantial amounts of

−· ROS (O2 ) which can specifically damage dopaminergic neurons.

−· In addition to formation of O2 by DA oxidation, the DA oxidation product, DAQ is a precursor for enzyme-assisted formation of tetrahydroisoquinolines like salsolinol, which is an endogenous neurotoxin known to cause increased ROS production and inhibition of the mitochondrial electron transport chain (181,182). In addition, it is found that DA oxidation products reduce α-synuclein levels (183). α-Synuclein is a negative regulator of DA biosynthesis due to its interaction with tyrosine hydroxylase. Thus reduction of α-synuclein levels prevents its inhibitory effect on DA synthesis, which leads to the production of more DA, possibly leading to more ROS production (184,185).

Scheme 10. DA Oxidation Pathways

The effect of DA on the cellular ROS production and neurotoxicity can be studied by using tyrosine hydroxylase inhibitors such as α-methyl-L-tyrosine, 3-iodotyrosine, and α-methyl-

DOPA (186). These inhibitors reduce intracellular DA levels by inhibiting tyrosine hydroxylase, which is the rate determining step in the catecholamine biosynthetic pathway (Scheme 11) (187).

In the current study, α-methyl-L-tyrosine was used to reduce the intracellular DA levels.

Scheme 11. Inhibition of Biosynthesis of DA by α-Methyl-L-Tyrosine. TH – Tyrosine Hydroxylase, THI – Tyrosine Hydroxylase Inhibitor, PLP – Pyridoxal Phosphate, AAADC – Aromatic Amino Acid Decarboxylase.

2.6.3.2 Anti-Oxidants and Reactive Oxygen Species

Treatment with antioxidants is a convenient and efficient way to protect cells from the noxious effects of ROS by eliminating surplus ROS by preventing their formation, scavenging, or promoting their decomposition (169). The antioxidant systems are categorized as endogenous and exogenous, as well as enzymatic and non-enzymatic (176). The endogenous antioxidants produced within the cell are the most powerful and efficient of the two. The five most important endogenous antioxidants are superoxide dismutase, catalase, glutathione, alpha lipoic acid

(ALA), and coenzyme Q10 (CoQ10). Exogenous antioxidants are obtained from our diet and dietary supplements, such as Vitamin E and Vitamin C (ascorbic acid), polyphenols and carotenoids (188).

In the current study, water soluble ascorbate (Asc) was used as an exogenous antioxidant to scavenge ROS produced in dopaminergic cells in response to various neurotoxin treatments.

The mechanism of ROS scavenging by Asc is initiated by the formation of the stable ascorbate free radical (Scheme 12), by donating an electron to the free radical in order to convert the free radical to a harmless ion or a molecule. The disproportionation of two ascorbate radicals produce one molecule each of dehydroascorbate and reduced Asc (189).

Scheme 12. Mechanism of Radical Scavenging Activity of Ascorbate

2.6.3.3 Antioxidant Enzymes and Reactive Oxygen Species

Antioxidant enzymes belong to the class of endogenous antioxidants. The major antioxidant enzymes directly involved in the neutralization of ROS and RNS are superoxide dismutase (38), catalase (CAT), glutathione peroxidase (GPx) and glutathione reductase (GR).

The report that antioxidant enzyme levels in the brain are low in comparison to the liver (190) prompted us to examine the relationship of antioxidant enzyme levels in catecholaminergic cells and the toxicities of various neurotoxins.

2.6.3.3.1 Superoxide Dismutase (38)

The first characterized antioxidant enzymes were SODs (191) whose active sites include metal ions, such as copper and zinc, or manganese, iron, or nickel. Three major types of

SODs are expressed in human cells: (1) copper-zinc SOD (CuZnSOD), which is present mainly in the cytoplasm; (2) mitochondrial manganese SOD (MnSOD) and (3) extracellular SOD

− (ECSOD). All SODs catalyze the dismutation (or partitioning) of superoxide (O2 ) radical into

H2O2 and molecular oxygen (Scheme 13).

Scheme 13. Reaction Catalyzed by Superoxide Dismutase

As shown in Scheme 14, redox metal ions in the active site of SOD are important for its

− catalysis of O2 dismutation.

Scheme 14. Involvements of Redox Metal Ions in Reactions Catalyzed SOD Where M = Cu (n=1) ; Mn (n=2) ; Fe (n=2) ; Ni (n=2).

2.6.3.3.2 Catalase (CAT)

Catalase is a tetrameric enzyme with a molecular weight of 240 kDa and a heme prosthetic group in the active site. CAT is responsible for detoxification of various phenols, alcohols and hydrogen peroxide. It is localized mainly in the peroxisomes, but small amounts are also present in the cytosolic fraction. CAT catalyzes the conversion of cytotoxic H2O2 into water and molecular oxygen (192) (Scheme 15).

Scheme 15. Reaction Catalyzed by Catalase

The complete reaction mechanism involving catalase is not known and Scheme 16 shows the hypothetical involvement of the heme group in the CAT mechanism which is believed to occur in two stages (193).

Scheme 16. Mechanism of the Reaction Catalyzed by CAT. Where E = Enzyme

2.6.3.3.3 Glutathione Peroxidase (GPx)

Selenocysteine is present at the catalytic site of GPx, and selenium availability regulates the GPx enzyme activity. GPx catalyzes the reduction of hydroperoxides, including lipid hydroperoxides, using glutathione (GSH) as co-substrate (176) (Scheme 17).

Scheme 17. Reaction Catalyzed by Glutathione Peroxidase

As shown above, GSH counteracts ROS by oxidation of reduced GSH to GSSG, reducing

GSH levels. Thus, loss of reduced GSH levels is a hallmark of excessive cellular ROS production. Glutathione reductase (GR) catalyzes the regeneration of reduced GSH from oxidized glutathione (GSSG) at the expense of reducing equivalents from NADPH (Scheme 18).

Together, SOD, CAT, GPx and GR constitute an effective antioxidant defense system to protect cells from oxidative damage by diverse ROS species (194) (Scheme 18).

Scheme 18. Antioxidant Enzyme System

2.6.4 Apoptosis

Apoptosis is the process of programmed cell death, which is accompanied by well- defined morphological changes such as cell shrinkage, chromatin condensation, nuclear fragmentation, chromosomal DNA fragmentation, and cellular blebbing. This process generates apoptotic bodies which are engulfed and destroyed by phagocytic cells. Unlike necrosis, where cell swelling, loss of cell membrane integrity and an uncontrolled release of cell death products into the extracellular space occurs, apoptosis does not include cell membrane rupture, thus

34 avoiding the release of degradative enzymes and other toxins into the extracellular space.

Apoptosis is an important process in multicellular organism development and homeostatic by maintaining fresh cell populations in tissues. Moreover, in immune reactions or when cells are damaged by disease or noxious agents, apoptosis occurs as a defense mechanism to eliminate damaged cells (195). Importantly, apoptosis can be induced by increased ROS (196) resulting from enzymatic reactions or toxic substances. The latter include neurotoxins such as MPP+, rotenone, paraquat. Neurotoxin stimulated apoptosis is considered to be one of the characteristic features of neurodegenerative diseases such as AD and PD (126,197).

Some factors like caspases, FasR, p53, Bax, etc., promote apoptosis, while members of the Bcl-2 family inhibit apoptosis. Caspases (C-cysteine rich, asp- aspartic acid moiety containing, ase – proteases) are crucial mediators of apoptosis. Among them caspase-3 is a frequently activated death protease, which catalyzes cleavage of many key cellular proteins in the apoptotic pathway (198,199) and is known to be implicated in PD (200,201).

One of the hallmarks of apoptosis is the cleavage of chromosomal DNA into multiples of

~200 bp oligonucleosomal fragments (202). This fragmentation is due to catalytic cleavage around histones of condensed chromosomal DNA into nucleosomal units (a human nucleosomal unit consists of ~180-200 bp), by an endonuclease (203,204). Therefore, observation of a repetitive electrophoretic banding pattern separated by ~180-200 bp of chromosomal DNA is an indication of apoptotic cell death. These repetitive banding patterns are commonly called apoptotic DNA laddering, which is widely used to identify apoptotic cell death.

In addition, 4,6-Diamidino-2-phenylindole (DAPI) experiments provide a rapid and convenient fluorescent assay for apoptosis. DAPI is a less membrane-permeable dye which can

35 bind to DNA and gives blue fluorescence (Ex/Em 358/461). With the process of apoptosis, the permeability for DAPI is increased and more dye can enter the cell and nucleus resulting in more dye binding to DNA, which produces greater blue fluorescence. Chromatin condensation in apoptosis also causes higher DAPI fluorescence as compared to normal cells. Thus DAPI staining is used in many studies, to reveal apoptosis (205).

2.7 Dopaminergic Toxicity of Cyanine Dyes

The discovery that MPTP/ MPP+ causes PD in both primates and other animal models, prompted a search for structural analogs of MPP+ that may contribute to the etiology of PD.

Accordingly, the current study was initiated with two simple cyanines 1,1'-diethyl-2,2'-cyanine

(2,2’-cyanine) and 1,1'-diethyl-4,4'-cyanine (4,4’-cyanine) which are structurally similar to

MPP+ (Scheme 19). In spite of their structural similarity, these cyanines are more hydrophobic than MPP+ due to bulky hydrophobic aromatic rings. Since MPP+ is charged and hydrophilic it cannot cross the blood brain barrier. On the other hand, cyanines not only possess structural similarity to MPP+, but also may accumulate freely in cells, due to their high hydrophobicity.

Therefore, they are good candidates to study the mechanisms of dopaminergic degeneration in

PD.

Scheme 19. Structures of MPP+, 2,2’-Cyanine and 4,4’-Cyanine

After finding that 2,2’-and 4,4’-cyanine are highly and specifically toxic to dopaminergic cells, we extended these studies to include some other fluorescent cyanines, 3,3'-

Diethyloxacarbocyanine (DiOC2(3)), 3,3′-Dipropylthiacarbocyanine (DiSC3(3)), 3,3'-

Dipropylthiadicarbocyanine (DiSC3(5)) (Scheme 20). These fluorescent cyanines have allowed us to conveniently determine their cellular localization in order to further our mechanistic studies. Our proposal that the positive charge in these toxins is vital for their toxicities has been addressed using the negatively charged lipophilic dye, Bis-(1,3-Diethylthiobarbituric Acid)

Trimethine Oxonol (DiSBAC2(3)) (Scheme 20).

Scheme 20. Structures of DiOC2(3), DiSC3(3) and DiSC3(5) Cyanines and Negatively charged, DiSBAC2(3)

Cyanine dyes are a family of synthetic dyes which consist of two nitrogen centers, one of which is positively charged and is linked to the other nitrogen through a conjugated carbon chain with an odd number of carbon atoms (206). There are three types of cyanines named streptocyanines or open chain cyanines, hemicyanines, and closed chain cyanines (Scheme 21)

(207). The cyanines used in this study are closed chain hetero aromatic and hetero atomic possessing sulfur or oxygen (Schemes 19 & 20).

Scheme 21. Types of Cyanines, I. streptocyanines or open chain cyanines, II. Hemicyanines and III. closed chain cyanines

Cyanines are commonly used in industry and scientific research. They are widely used in the production of photographic films, computer storage and in optical disks as video recording media (208). Since they carry an extended  electron system (Schemes 19 & 20), they are also used in solar cells (209), and in photochemical research (210). They are also frequently used as inhibitors of extra-neuronal noradrenaline (211), plasma membrane monoamine, and organic cation transporters (22,212), as precursors in the development of chemotherapeutic (213) and neuro-protective agents (214,215), and various cell imaging agents. However, the neurotoxicological properties of cyanines have not been reported.

2.8 Cell Culture

Cell culture models serve many purposes, such as studying passive and active transport of drugs, cell physiology, protein expression, and in vitro toxicity tests (216). Cell culture refers to one derived from dispersed cells obtained from an original tissue, from a primary tissue, from a cell line, or from a cell strain, by enzymatic, mechanical, or chemical disaggregation (217).

Mammalian cells are grown in suspensions or adherent cultures and maintained typically at 37˚C and 5% CO2 atmosphere. Culture conditions vary widely for each cell type. The growth medium, one of the culture conditions, can vary in pH, glucose concentration, growth factors, and other nutrients. Once cells are grown to a confluent layer, that is when all the available growth area is utilized and cells make close contact with one another, they need to be sub- cultured. The cell culture growth media is changed every 2-3 days until the cells again grow to confluency.

2.8.1 PC12 Cells

PC12 cells are derived from pheochromocytoma of the rat adrenal medulla (218), which was first characterized by Green and Tischler in 1976. This is a well characterized, well studied and widely used adrenergic neuronal cell model. PC12 cells synthesize and store catecholamine neurotransmitters, dopamine and norepinephrine, but not epinephrine (218). PC12 cells are also known to synthesize and store acetylcholine (219).

2.8.2 SH-SY5Y Cells

SH-SY5Y is a thrice cloned sub-clone of a human dopaminergic neuroblastoma cell line,

SK-N-SH which was established in 1970 from a bone marrow biopsy (220). The SK-N-SH parent cell line was first sub-cloned as SH-SY, was sub-cloned again as SH-SY5, then the third and final sub-cloning to produced the SH-SY5Y cell line (221,222).

2.8.3 MN9D Cells

MN9D is an immortalized cell line that is derived from the fusion of mouse embryonic ventral mesencephalic cells, rostral mesencephalic tegmentum (RMT) and neuroblastoma cell

40 line (N18TG2) (223). MN9D cells are widely used as a dopaminergic cell model, because they possess high levels of DA and DOPAC and express monoamine transporters such as DAT, NET, and VMAT-2 (224)

2.8.4 HepG2 Cells

HepG2 is a human liver carcinoma cell line derived from the liver tissue of a fifteen year old Caucasian American male with a well differentiated hepatocellular carcinoma. Since this cell line is derived from human liver and does not synthesize catecholamines, it has been extensively used as a control non-neuronal cell model.

CHAPTER III

EXPERIMENTAL METHODS

3.1 Materials

All reagents and supplies were purchased from Fisher Scientific (Pittsburg, PA, USA),

Sigma-Aldrich (Milwaukee, WI, USA) or Torcis (Bristol, United Kingdom) unless otherwise noted. Fetal bovine serum (FBS) was purchased from Valley Biomedical (Winchester, VA,

USA). F-12K medium and horse serum (225) were purchased from American Type Culture

Collection, ATCC (Manassas, VA, USA). Protein assay reagents were purchased from Bio-Rad

(Hercules, CA, USA). Catecholamine standards were purchased from ESA (Chelmsford, MA,

USA). Ultra-pure 10X TBE buffer and SYBR Safe DNA gel stain in 0.5X TBE were purchased from Invitrogen (Eugene, OR, USA). DNA marker was purchased from New England Biolabs

Inc. (Ipswich, MA, USA). The mouse hybridoma cell line MN9D (226) was graciously provided by Dr. Alfred Heller, University of Chicago. Human heptacellular liver carcinoma cell line (227) was obtained from Dr. Tom Wiese (Fort Hays University, Hays KS). Rat pheochromocytoma

PC12 and human neuroblastoma SH-SY5Y cell lines were purchased from ATCC (Manassas,

VA, USA). All solutions were prepared in Milli Q-deionized water (Millipore, Billerica, MA,

USA). Stock solutions of 3,3'-Diethyloxacarbocyanine Iodide (DiOC2(3)), 3,3′-

Dipropylthiacarbocyanine iodide (DiSC3(3)), 3,3'-Dipropylthiadicarbocyanine Iodide

(DiSC3(5)), Bis-(1,3-Diethylthiobarbituric Acid)Trimethine Oxonol (DiSBAC2(3)), Rotenone,

4′, 6-Diamidino-2-phenylindole, dilactate (DAPI), Tetramethylrhodamine, methyl ester

(TMRM), and 2΄, 7΄-dichlorofluorescin diacetate (DCFH-DA) were prepared in dimethyl sulfoxide (DMSO). In all experiments, the final dimethyl sulfoxide concentration was kept to a

42 minimum, usually < 0.05% v/v. KRB-HEPES contained 109.5 mM NaCl, 5.34 mM KCl, 0.77 mM NaH2PO4, 1.3 mM CaCl2, 0.81 mM MgSO4, 5.55 mM Dextrose, 25 mM HEPES, pH 7.4.

3.2 Instrumentation

UV-visible spectra were recorded on a Cary Bio 300 UV-visible spectrophotometer

(Varian, Inc). Fluorescence emission spectra were recorded on a JobinYvon-Spex Tau-3 spectrophotometer (ISA Instruments, Inc). Catecholamine levels were analyzed by using reversed-phase HPLC with electrochemical detection (HPLC-EC) on a C18 reversed-phase column (ESA, HR-80). HPLC-EC analyses were performed using ESA Model 582 solvent delivery module and Coulochem-II electrochemical detector with ESA 501 chromatographic software (ESA, Chelmsford, MA, USA). Mitochondrial membrane potentials were measured using a Nikon ECLIPSE-Ti-S microscope equipped with a Nikon S FLURO 40X lens. Images of

DAPI loaded apoptotic cells were captured using a Nikon ECLIPSE-Ti-S inverted fluorescence microscope equipped with a Nikon S FLURO 40X objective (Nikon Instrument Inc., Melville,

NY, USA). Mitochondrial dye localization and intracellular ROS levels in live cells were monitored using a Leica (228) confocal fluorescence microscope equipped with a 40X lens objective.

3.3 Methods

3.3.1 Cell Culture

MN9D, HepG2 and SH-SY5Y cells were cultured in 100 mm2 Falcon tissue culture plates in DMEM with high glucose (4500 mg/L) supplemented with 10% fetal bovine serum, 50

o g/mL streptomycin and 50 IU/mL penicillin at 37 C and 5% CO2. PC12 cells were cultured in

100 mm2 Biocoat tissue culture plates coated with rat tail collagen type I in F-12K medium supplemented with 2.5% FBS and 15% HS at 37˚C and 5% CO2 in humidified air. Cells were fed three times weekly, passaged by incubation in Trypsin-Versene, plated and then seeded into 12 well plates, 96 well plates or glass bottomed plates, depending on the nature of the experiment, and grown to 70-80% confluence for 2-3 days, unless otherwise stated.

3.3.2 Differentiation of MN9D cells

Differentiation of MN9D cells was stimulated with 1.0 mM sodium n-butyrate according to a published procedure (226). Briefly, cells were seeded at a density of 0.1 x 106 cells/well in

6-well plates and were treated with 1 mM n-butyrate for 3 days in DMEM media, at which time the media was replaced with fresh 1 mM n-butyrate containing media and incubated for an additional 3 days. The progress of differentiation was gauged by observing the expected morphological changes. All differentiated cells were used in appropriate experiments after the 6th day of differentiation.

3.3.3 Measurement of Cellular Uptake of Cyanine

Cells were grown in 12-well plates, washed with KRB-HEPES, and a desired concentration of cyanine in the same buffer was added, and incubated for 1 h at 37 °C. After the incubation, cells were washed, gently scraped, and collected in 1.0 mL of ice-cold KRB-HEPES.

Aliquots were removed for protein assay, the remainder was centrifuged for 3 min at 6,000 rpm, and the cell pellet was suspended in 0.1 mol/L perchloric acid for 2,2’- and 4,4’-cyanine (229) or

0.1 mol/L Tri- buffer (pH 7.5) containing 1% Triton X-100 for the other cyanine dyes. The cyanine content of cell extracts was determined by fluorescence using standard curves constructed with standard concentrations of the corresponding cyanine. All uptake readings were

44 normalized to the protein content of each sample and corrected for non-specific binding by subtracting the corresponding zero time readings.

3.3.4 Measurement of Cell Viability

Cell viability was determined by MTT [3-(4,5-dimethylthiazol-2-yl)-2,5- diphenyltetrazolium bromide)] assay (230,231). Briefly, cells were seeded in 96-well plates and allowed to grow until 70-80% confluence. Prior to the experiment, the medium was replaced with KRB-HEPES containing the desired concentration of cyanines or other reagents and incubated for 9 h to 12 h, depending on the experiment, at 37 °C. After the incubation, 10 μL of

5 mg/mL MTT solution was added to each well and was incubated for 2 h at 37 °C. The resulting formazan was solubilized by addition of 210 L detergent solution (50% DMF, 20% SDS) followed by incubation for 4 h at 37 °C and quantified based on the difference in the absorbance at 570 nm and 650 nm (231). Results are expressed as % viability of toxin treated cells with respect to control cells which were treated under the same conditions, except in the absence of the toxin.

In the experiments where tyrosine hydroxylase inhibitor (92) was used, cells were treated with the desired concentration of THI (L-α-methyltyrosine; 1-6 mM) at 37 °C in normal growth media for 24 h and then the medium was aspirated and toxin was added with the same concentration of THI in KRB-HEPES. Cells were incubated for 12 h at 37 °C and cell viabilities were determined as above. In experiments where ascorbate was used, cells were pre-incubated with the desired concentration of ascorbate for 1 h and then the toxin was added with the same concentration of ascorbate in KRB-HEPES. Cells were incubated for 12 h at 37 °C and the cell viabilities were determined as above.

3.3.5 Mitochondrial Localization of Cyanines

MN9D and HepG2 cells grown in glass bottomed plates were treated with 100 nM of

DiSC3(5) or DiSBAC2(3) in KRB-HEPES for 15 min at 37 °C and washed with KRB-HEPES.

Cells were then treated with 100 nM of Mitotracker Green or Red for 15 min at 37 °C and washed 3 times with KRB-HEPES. Cell images were captured at the corresponding wavelengths using a Leica confocal fluorescence microscope equipped with a 40X lens objective.

3.3.6 Measurement of Mitochondrial Membrane Potential

MN9D and HepG2 cells grown in glass bottomed plates were treated with 50 nM TMRM in KRB-HEPES for 45 min in the dark (232). After mounting on the microscope stage, ROIs were selected (20-30) and TMRM fluorescence (Ex/Em 543/573 nm) was measured in 5 sec intervals for 3 min using a Nikon ECLIPSE-Ti-S inverted fluorescence microscope. After 3 min,

2.5 M of toxin was added and fluorescence measurements were recorded for an additional 15 min. Parallel controls were carried out using an identical protocol except that the toxin was omitted from the incubation. The background fluorescence was subtracted from the ROI fluorescence of both tests and controls and averages of background were corrected. Control subtracted test data were used to estimate the mitochondrial membrane potential.

3.3.7 Rat Brain Mitochondria Extraction

Rat brain mitochondria were isolated according to the protocol of Iglesias-Gonzalez et al. (233).

Rat brains were kindly provided by Professor Li Yao at the Department of Biological Sciences,

Wichita State University. After sacrifice, the brain was quickly removed and immersed in an ice- cold isolation buffer (IB) containing 225 mM mannitol, 75 mM sucrose, 5 mM HEPES and 1 mg/mL fatty acid free BSA (pH 7.4, isotonized with KOH). Rat brain was weighed and cut in

46 small pieces for suitable manual homogenization in a Dounce-type glass homogenizer using IB.

The homogenates were distributed to polypropylene Falcon tubes and then centrifuged at 600 g for 10 min (JA-20 fixed angle rotor, set at maximum acceleration and low deceleration, Beckman

Instruments, Palo Alto, CA, USA). The supernatant was removed and the pellet was resuspended in 10 mL of IB and centrifuged to recover the fast-sedimented mitochondria. Then the obtained supernatants were pooled and centrifuged in 4 tubes at 10,000 g for 10 min. The pellets were resuspended in IB and centrifuged at 7,000 g for 10 min. The pellet obtained exhibited some layers and color with an upper white fluffy layer, containing myelin, synaptosomes, and lipids.

The upper white fluffy layer was discarded and the brown mitochondrial pellets were collected and resuspended in 10 mL of buffer and centrifuged in 2 tubes at 12,000 × g for 10 min. The resulting mitochondrial pellet was resuspended in a final volume of 150 μL assay buffer [25 mM potassium phosphate, pH 7.2, 5 mM MgCl2] and stocked in pre-cooled Eppendorf tubes. The mitochondrial solution was maintained in an ice-bath until the Mitochondrial Complex I assay was carried out. In order to maintain temperature control throughout the process, all isolation procedures were carried out at 4 °C.

3.3.8 Mitochondrial Complex I inhibition

Rat brain mitochondria were isolated as previously described. Mitochondrial membranes were opened by freeze-thawing mitochondria three times in hypotonic media [25 mM potassium phosphate, pH 7.2, 5 mM MgCl2] (234). NADH (0.13 mM), Antimycine A (2 μg/mL) and mitochondrial membrane extract (50 μg of protein) were added to the assay medium (potassium phosphate 25 mM, MgCl2, 5 mM, KCN 2 mM, bovine serum albumin 2.5 mg/mL) and incubated with or without toxin for 2 min at 37 oC. NADH:ubiquinone oxidoreductase was activated by adding ubiquinone1 (65 μM). Complex I specific activity was measured by following the

47 decrease in absorbance due to the oxidation of NADH at 340 nm, with 425 nm as the reference wavelength, for 10 min (235).

3.3.9 Measurement of Reactive Oxygen Species

Cells were grown in 12-well plates, rinsed with KRB-HEPES and incubated with 10 M

2΄, 7΄-dichlorofluorescin diacetate (DCFH-DA) for 60 min. Then, DCFH-DA loaded cells were washed with ice-cold buffer, and treated with the desired concentration of cyanines or other toxins for 1 h at 37 oC. Cyanines or other toxin-treated (or control) cells were washed, harvested, solubilized with 0.1 mol/L Tris buffer (pH 7.5) containing 1% Triton X-100. Cell debris was removed by centrifugation. The 2΄,7΄-dichlorofluorescein content in the supernatants was determined by fluorescence Ex/Em 504/526 nm (228). The data were normalized to the protein content of individual samples.

3.3.10 Mitochondrial ROS production by Cyanines

MN9D cells grown in glass bottomed plates were treated with 10 M DCFH-DA in

KRB-HEPES for 1 h. DCFH-DA-loaded cells were washed with buffer and treated with 1000 nM of DiSC3(5) or DiSBAC2(3) in KBR for 1 h at 37 °C and washed with KRB-HEPES and images were captured at the corresponding wavelengths using a Leica confocal fluorescence microscope equipped with a 40X lens objective (236).

3.3.11 Measurement of the Intracellular DA Content

Cells were grown in 12-well plates, washed with KRB-HEPES and treated with the desired concentrations of cyanine in KRB-HEPES for 1 h at 37 oC. The media was removed; cells were washed, gently scraped and suspended in KRB-HEPES. The cells were pelleted by low-speed centrifugation and lysed with 75 L of 0.1 mol/L HClO4. The dopamine in cell lysates 48 was separated and quantified by HPLC-EC as previously described (237,238) by using an ESA

Model 582 solvent delivery module and Coulochem-II electrochemical detector with ESA 501 chromatographic software (ESA, Chelmsford, MA, USA). Briefly, acidic cell extracts were separated by a C18 reversed phase column (ESA, HR-80) using a mobile phase composed of 75 mM NaH2PO4, 1.7 mM 1-octanesulfonic acid sodium salt, 25 μM Na2EDTA, 100 μL/L trimethylamine, pH 3.0 with 10 % CH3CN at a flow rate of 0.8 mL/min. The depletion of intracellular DA content in MN9D cells was accomplished by treating the cells with the tyrosine hydroxylase inhibitor L-α-methyltyrosine (92) (2-6 mM) for 24 h under normal growth conditions and carrying out the incubations with toxins in the presence of the same concentrations of THI.

3.3.12 ATP Assay

Cells grown in 12-well plates were treated with the desired concentration of cyanine or rotenone for 1 h in KRB-HEPES buffer. Following the incubation period, cells were collected in

1 mL KRB-HEPES buffer and a 50 μL sample was taken out for the protein assay. The extracted cell solution was centrifuged for 3 min at 3,500 g and the supernatant was aspirated and cell pellet was taken for the assay. Total final volume was adjusted to 1 mL with cell lysis buffer (0.1 mol/L Tris, 1 % Triton X-100, pH 7.5). ATP content was determined by using BioVision ATP colorimetric/fluorometric assay kit. Cellular ATP content was assayed by measuring the fluorescence excitation at 535 nm and emission at 587 nm using a Varian Cary Eclipse fluorescence spectrophotometer. The fluorescence readings were normalized to protein content of individual samples.

3.3.13 Estimation of Intracellular Superoxide Dismutase (38), Catalase and Glutathione

Peroxidase (Gpx) Levels

HepG2, MN9D, and SH-SY5Y cells were grown in 12-well plates, washed twice with ice-cold PBS, gently scraped, and collected in 1.0 mL cold PBS. Aliquots were removed for protein assay and the remainder was centrifuged for 3 min at 6,000 rpm, and the cell pellet was suspended in cold PBS. Next, the cell suspensions were lysed with lysis buffer (BioVision), centrifuged, and the supernatant was used to determine antioxidant enzyme activities using commercial assay kits [BioVision; SOD (K335-100); Catalase (K773-100) and Gpx (K762-

100)], according to the manufacturer’s guidelines. All data were normalized to protein content and the enzyme activities were expressed relative to the corresponding enzyme activities of

HepG2 cells (100%).

3.3.14 Chromatin Condensation Test for Apoptosis

MN9D cells grown in glass bottomed plates were treated with cyanines or other toxins in

KRB-HEPES for 12 h. Then, cells were treated with 300 nM 4', 6-diamidino-2- phenylindole, dilactate (DAPI) for 15 minutes in the dark (239), rinsed, reconstituted with KRB-

HEPES, and DAPI loaded cell plates were mounted on the microscope stage. Increase of DAPI fluorescence in the nuclei due to chromatin condensation was observed by a Nikon ECLIPSE Ti-

S inverted fluorescence microscope using a 40X objective with Ex/Em 358 nm/461 nm. The controls were treated similarly, except the toxins were omitted from the incubation medium.

3.3.15 Apoptotic DNA Fragmentation

MN9D cells grown in 12 well plates were treated with 500, 1000 or 2000 nM 2,2’- cyanine for 12 h. Total cellular DNA was extracted using the DNeasy DNA Isolation Kit

(QIAGEN, Valencia, CA, USA) according to the manufacturer's instructions, including treatment with proteinase K and RNase A. DNA samples were loaded onto 1.2% agarose gel and electrophoresed at 70 V for 2 to 3 h until the bromophenol blue band had migrated approximately two thirds of the way down the gel. Apoptotic DNA fragmentation bands were stained with SYBR safe DNA gel stain (Invitrogen, Eugene, OR, USA) visualized on a UV transilluminator, and photographed using a Kodak Gel Logic 100 system (Eastman Kodak,

Rochester, NY, USA) (240).

3.3.16 Protein Determination

Protein content of various cell preparations were determined by the method of Bradford

(241). Samples of cell suspensions (50 μL) in KRB-HEPES were incubated with 950 μL of

Bradford protein reagents for 10 min and absorbance at 595 nm was measured. The protein concentrations were determined using a standard curve constructed employing bovine serum albumin as a standard.

3.3.17 Data analyses

All data are averages of three or more determinations and presented as means ± SD. The error bars represent the SD of the data from the mean. In the experiments where uptake or inhibition kinetic parameters were determined, the means of the experimental data were directly fitted to the hyperbolic form of the Michaels-Menten equation. The ± values given in kinetic parameters were the fitting errors of the mean experimental data. All quantitative uptake, catecholamine levels, and ROS data were normalized to protein content of individual incubations to correct the results for the variations of cell densities between individual experiments.

3.3.18 Technical Statement

Due to the high toxicity and obvious health hazards of MPP+, rotenone and cyanines, extreme caution was used in their handling in accordance with published procedures (242).

CHAPTER IV

RESULTS

4.1 Cellular Toxicities of Cyanines, MPP+ and Rotenone

The relative sensitivity of dopaminergic MN9D (mouse mesencephalic neuronal cell line)

(243), SH-SY5Y (human neuroblastoma cell line) (244), PC12 (rat adrenal pheochromocytoma cell line) (245) and non-dopaminergic HepG2 (human liver cell line) (227) cells (Fig. 2) to MPP+, rotenone and cyanine toxicity were determined using standard MTT cell viability assays as detailed in Experimental Methods.

A B

Figure 2. Morphologies of MN9D, SH-SY5Y, HepG2 and PC12 Cells. (A), MN9D; (B), SH- SY5Y and (C), HepG2 were cultured in glass bottomed tissue culture plates in DMEM with high glucose (4500 mg/L) supplemented with 10% fetal bovine serum (FBS), 50 g/ml streptomycin and 50 IU/ml penicillin at 37 ˚C and 5% CO2. (D), PC12 cells were cultured in Biocoat tissue culture plates coated with rat tail collagen type I in F-12K medium supplemented with 2.5% FBS and 15% HS at 37 ˚C and 5% CO2 in humidified air. The image of PC12 was obtained from published literature (246). (E), MN9D cells were differentiated using n-butyrate as detailed in Experimental Methods. The images were captured using a Leica confocal microscope equipped with a 40x lens.

C D

Figure 2 (continued)

4.1.1 MPP+ and Rotenone are Specifically Toxic to Dopaminergic Cells

The most commonly used PD model toxin, MPP+ was highly toxic to mouse dopaminergic MN9D cells within the concentration range 0-300 M (IC50 ~100 M), however, no measurable toxicity was detected in non-dopaminergic, liver HepG2 cells under similar experimental conditions (Fig. 3A). Although, MPP+ was moderately toxic to human dopaminergic/adrenergic SH-SY5Y cells at higher concentrations (0-4 mM), they were still more sensitive to MPP+ than HepG2 cells cultured under similar conditions (Fig. 3B). Another well- known dopaminergic toxin (247) and Mitochondrial Complex I inhibitor (248), rotenone was also highly toxic to MN9D cells (IC50 ~25 nM), moderately toxic to SH-SY5Y cells (IC50 ~1.5

M) and not toxic to HepG2 cells under similar experimental conditions (Fig. 3C). However,

MN9D and SH-SY5Y toxicities of rotenone was about 1000 fold higher than that of MPP+

(Table 3).

120

100 80

60 HEPG2 40 MN9D % Cell ViabilityCell % 20 0 0 10 25 50 75 100 200 300 [MPP+] (μM)

120 100

60 SHSY-5Y

40 HEPG2

Viability% Cell 20

0 0 0.05 0.1 0.25 0.5 1 2 3 4 [MPP+ ] (mM)

Figure 3. Comparative MN9D, SH-SY5Y and HepG2 cell toxicities of MPP+ and Rotenone. Cells were grown in 96-well plates and were treated with MPP+ or rotenone in KRB-HEPES for 12 h at 37 °C. Cell viabilities were determined by the MTT assay as detailed in Experimental Methods. Results are expressed as % cell viability with respect to parallel untreated controls. Data presented as mean ± SD (n=6). (A), MN9D and HepG2; (B), SH-SY5Y and HepG2, cell toxicities of MPP+; (C), MN9D, SH-SY5Y and HepG2 cell toxicities of rotenone. 55

C 120

100 80 60 HEPG2

40 SH-SY5Y % Cell Viability% Cell 20 MN9D

0 0 5 10 15 20 25 30 50 75 100 [Rotenone] (nM)

Figure 3 (continued)

Table 3. IC50 Values for Different Toxins in Different Cell Types. None of the compounds were toxic to HepG2 cells under similar experimental conditions. ND - Not determined.

MN9D Differentiated SH-SY5Y PC12 Toxin MN9D MPP+ 100 μM ND 1 mM ND Rotenone 25 nM ND 1.5 μM ND 2,2’-cyanine 75 nM 100 nM 5 μM 2 μM 4,4’-cyanine 100 nM ND ND 3 μM DiOC2(3) 60 nM ND ND ND DiSC3(3) 60 nM ND ND ND DiSC3(5) 60 nM ND ND ND

4.1.2 Both 2,2'- and 4,4'- Cyanines are Highly Toxic to Dopaminergic Cells and Cell

Specificity of Toxicities are Parallel to that of MPP+ and Rotenone

Two simple cyanines, 2,2’- and 4,4’-cyanines were also specifically toxic to MN9D and

SH-SY5Y dopaminergic cells as well as adrenergic PC12 cells under similar experimental conditions (Figs. 4A & B). Remarkably, 2,2'-cyanine was about 1000 fold more toxic to MN9D cells in comparison to MPP+ (IC50s of 2,2’- and 4,4’-cyanines and MPP+ toxicities for MN9D cells were ~100 nM and ~100 M, respectively). However, rotenone was about 4 fold more toxic

56 to MN9D cells in comparison to that of 2,2’- and 4,4’-cyanines (IC50s of rotenone and 2,2’- and

4,4’-cyanines toxicities for MN9D cells were ~25 nM and ~100 nM respectively) (Table 3). Both

2,2’- and 4,4’-cyanines were toxic to adrenergic PC12 cells with IC50s of 2 M and 3 M, respectively. They were moderately toxic to SH-SY5Y (IC50 ~5 M) cells (Fig. 4C), but not significantly toxic to HepG2 cells under similar experimental conditions, parallel to the toxicity profile of MPP+. 2,2'- and 4,4’-cyanines, MPP+, and rotenone followed the same pattern of cell- specific toxicity, i.e. the order of toxicity was MN9D>SH-SY5Y>> HepG2 cells (Figs. 3 and 4A,

B & C) [note that PC12 data were not available for MPP+ and rotenone]. Furthermore, differentiated MN9D cell toxicity to 2,2'-cyanine was similar to that of undifferentiated cells

(Fig. 4D; IC50 of ~100 nM).

A 120

100

80 HEPG2

60 SHSY-5Y 40 PC12 % Cell Viability% Cell 20 MN9D

0 0 5 10 25 50 75 100 250 500 1000

[2,2'-cyanine] (nM) Figure 4. Comparative MN9D, SH-SY5Y, PC12 and HepG2 Cell Toxicities of 2,2′- and 4,4′- Cyanines. Cells were grown in 96-well plates and treated with 2,2′-cyanine or 4,4’-cyanine in KRB-HEPES for 12 h at 37 °C. Cell viabilities were determined by the MTT assay as detailed in Experimental Methods. Results are expressed as % cell viability with respect to parallel untreated controls. Data presented as mean ± SD (n=6). (A), MN9D, SH-SY5Y, PC12 and HepG2 cell toxicities of 2,2’-cyanine; (B), MN9D, SH-SY5Y, PC12 and HepG2 cell toxicities of 2,2’- cyanine; (C), MN9D and SH-SY5Y cell toxicities of high concentrations of 2,2’-cyanine; and (D), Differentiated MN9D, MN9D and HepG2 cell toxicities of 2,2′-cyanine.

B 120 100

80 HEPG2 60 SHSY-5Y 40

% Cell Viability% Cell PC12 20 MN9D 0

0 5 10 25 50 75 100 250 500 1000 C [4,4’ -cyanine (nM)

120 100

60 HEPG2 40

% Cell Vability % Cell SHSY-5Y 20 0 0 0.5 1 1.5 2 3 4 5 10 [2,2’-cyanine] (μM) D

120

100 HEPG2

80 MN9D

60 Diff. MN9D 40 % Cell Viability% Cell 20

0 0 5 10 25 50 75 100 250 500 1000 [2,2’ –cyanine] (nM)

Figure 4 (continued)

4.1.3 Positively Charged Cyanines are Specifically Toxic to Dopaminergic Cells but Not

the Negatively Charged Dye

All the positively charged cyanine dyes, 2,2’-cyanine, 4,4’-cyanine, DiOC2(3), DiSC3(3) and DiSC3(5) were about 1000-fold more toxic to MN9D than MPP+ with IC50s in the range of

75 nM – 100 nM and were not toxic to HepG2 cells (Figs. 3, 4 and 5). However, DiSBAC2(3) a negatively charged cyanine dye was not toxic to either MN9D or HepG2 cells (Figs. 5A & B).

120

100

80 DiOC2(3) 60 DiSC3(3) 40 DiSC3(5) % Cell Viability% Cell 20 0 DiSBAC2(3) 0 5 10 25 50 75 100 250 500 1000

[Dye] (nM) B 120

100 80 DiOC2(3) 60 DiSC3(3) 40

% Cell Viability% Cell DiSC3(5) 20 DiSBAC2(3) 0 0 5 10 25 50 75 100 250 500 1000 [Dye] (nM)

Figure 5. Comparative MN9D and HepG2 Cell Toxicities of DiOC2(3), DiSC3(3) and DiSC3(5) Cyanine Dyes and the Negatively charged dye, DiSBAC2(3). Cells were grown in 96-well plates and were treated with DiOC2(3), DiSC3(3), DiSC3(5) cyanine dyes or DiSCBAC2(3) in KRB for 12 h at 37 °C. Cell viabilities were determined by the MTT assay as detailed in Experimental

Methods. Results are expressed as % cell viability with respect to parallel untreated controls. Data presented as mean ± SD (n=6). (A), MN9D and (B), HepG2 cell toxicities of DiOC2(3), DiSC3(3) and DiSC3(5) cyanine dyes and the negatively charged dye, DiSCBAC2(3).

4.2 Cellular Uptake of MPP+ and Cyanine

Dopaminergic MN9D and SH-SY5Y and non-neuronal HepG2 cells (Fig. 2) were used for uptake experiments. The data for MPP+ uptake for different cell lines (Fig. 6A) were from Le and Wimalasena (249). The uptake of 2,2’-cyanine was studied in detail since it is more lipophilic than MPP+ and may not follow the same uptake characteristics. In addition, we assumed that the other cyanines may also follow the same uptake characteristics as 2,2’-cyanine.

4.2.1 MPP+ Uptake into All Three Cell Lines is Saturable at Low MPP+ Concentrations

The long range uptake kinetic data show that uptake of MPP+ into all three cell lines is complex. However, MPP+ uptake into all three cell types was saturable at low concentrations.

Increasing concentrations increased the rate of uptake further, indicating the presence of a non- specific and/or a low affinity, but efficient, high volume, secondary MPP+ uptake system in all these cells. The Vmax and Km parameters estimated for MN9D by fitting the data from the saturable concentration range (0-80 M; Fig. 6A1) to the Michaelis-Menten equation were 14.5

± 1.4 pmol/mg.min and 50 ± 9 M, respectively. SH-SY5Y cells showed apparent saturable kinetics in the concentration range of 0-200 M with Vmax and Km parameters of 30.5 ± 1.0 pmol/mg.min and 11.5 ± 1.5 M, respectively (Fig. 6A2). Similarly, the uptake of MPP+ into

HepG2 cells showed apparent saturation kinetics up to about 300 M with Vmax and Km parameters of 79 ± 9 pmol/mg.min and 105 ± 26 M, respectively (Fig. 6A3).

300 A 600 C 250 800 B 500 200 600 400 150 300 400 100 200 200 uptake (pmol/mg.30min) uptake uptake (pmol/mg.30 uptake min) 50 100 + + 0

0 MPP 0

MPP 0 20 40 60 80 0 50 100 150 200 MPP+ (pmol/mg.10 uptake mim) 0 50 100 150 200 250 300 + + [MPP ]M [MPP ] (M) MPP+ (M)

B1 0.9 B2 0.9 0.8 0.8

0.7 0.7 0.6 0.6 0.5 HEPG2 0.5 HEPG2 0.4 0.4 SHSY5Y SHSY5Y 0.3 0.3 mol/mg protein) mol/mg mol/mg protein) MN9D MN9D

(μ 0.2

(μ 0.2 0.1 0.1 0 0 0.25 0.5 1 0.25 0.5 1 [2,2’ –cyanine] (μM) [4,4’ –cyanine] (μM)

Figure 6. Characteristics of the Cellular Uptake of MPP+ and 2,2′-Cyanine. (A) MPP+ Uptake into MN9D Cells. The data were obtained from K. Wimalasena and V. Le from experiments of MPP+ uptake into MN9D, (A1); SH-SY5Y, (A2); HepG2, (A3) cells. (B1), 2,2′-cyanine and (B2),4,4′-cyanine uptake into MN9D, SH-SY5Y, and HepG2 cells. Cells were grown in DMEM to 70-80% confluence in 12 well plates treated with varying concentrations of 2,2′ or 4,4′- cyanine for 1 h at 37 °C and intracellular 2,2′ or 4,4′-cyanine levels were quantified by fluorescence (Ex/Em 320/420 nm) as detailed in Experimental Methods. (C) Concentration and time dependence of 2,2′-cyanine uptake into MN9D cells. MN9D cells were incubated with varying concentrations (or 1.0 M) of 2,2′-cyanine for 1 h (or a desired period of time) at 37 °C and intracellular 2,2′-cyanine levels were quantified as described above. All uptake data were normalized to respective protein concentrations and were corrected for nonspecific binding by subtracting the corresponding zero time readings. Data presented as mean ± SD (n=3). Concentration, (C1) and time dependence, (C2) of 2,2′- cyanine (1.0 M) uptake into MN9D cells.

C1 C2

1.0 1.0

0.8 0.8

0.6 0.6

0.4 0.4 mol/mg protein mol/mg protein.1h mol/mg   0.2 0.2

0.0 0.0 0.0 0.2 0.4 0.6 0.8 1.0 1.2 0 20 40 60 80 [2,2-cyanine]M Time (min)

Figure 6 (continued)

4.2.2 Both 2,2'- and 4,4'- Cyanines are Taken up into All Cells with Similar Specificity and

Efficiency

The uptake of 2,2'- and 4,4'-cyanines into MN9D, SH-SY5Y and HepG2 cells were determined by fluorescence spectroscopy (Ex/Em 320/420 nm) according to the protocol of Russ et al. (229). The uptake of 2,2'-cyanine and 4,4'-cyanine into all three cell lines during a 1 h incubation period was similar, and increased with increasing concentrations from 0-1 M (Figs.

6B1 & B2). Since both cyanine derivatives behaved similarly with respect to uptake and toxicity profiles, detailed characterization of the uptake and the other extended mechanistic studies were carried out with 2,2'-cyanine.

4.2.3 2,2'-Cyanine Uptake into MN9D Cells is Non-Saturable and Time Dependent

The concentration dependency of 2,2'-cyanine uptake into MN9D cells showed that the uptake was concentration dependent with no apparent saturation over the concentration range of

100-1000 nM during a 1 h incubation period (Fig. 6C1). The time dependence of the uptake of

2,2’-cyanine into MN9D at 1.0 M concentration showed an apparent saturation by about 1 h

(Fig. 6C2). An estimate of the intracellular 2,2'-cyanine concentration at saturation was 300-350

M (assuming a cellular volume of 3.0 L/mg of protein) suggesting that intracellular accumulation of 2,2'-cyanine was an apparently active, but non-mediated transport process.

4.2.4 Both Positively and Negatively Charged Cyanine Dyes are Taken up into MN9D

Cells

In spite of the difference in the net charges of 2,2’-cyanine, 4,4’-cyanine, DiOC2(3),

DiSC3(3) and DiSC3(5) (positively charged) and DiSBAC2(3) (negatively charged) all these dyes were taken up by MN9D cells in a concentration dependent manner during a 1 h incubation time period. Uptake increased with increasing concentration from 0-1 μM (Figs. 6B1, 6B2 and

7). However, the net uptake of the negatively charged DiSBAC2(3) was somewhat less than that of the positively charged dyes.

3.5

3 2.5 DiSBAC -ve 2 DiOC23 1.5 DiSC3 1 DiSC5 0.5 nmol dye / mg mg protein / of dye nmol 0 100 250 500 1000 [Dye] (nM) Figure 7. Characteristics of Cellular Uptake of Positively Charged Dyes, DiOC2(3), DiSC3(3) and DiSC3(5) Cyanine Dyes and a Negatively Charged Dye, DiSBAC2(3) by MN9D Cells. Cells were grown in DMEM to 70-80% confluence in 12 well plates treated with varying concentrations of DiOC2(3), DiSC3(3), DiSC3(5) and DiSBAC2(3) for 1 h at 37°C and intracellular DiOC2(3) (Ex/Em 482/497 nm), DiSC3(3) (Ex/Em 556/575 nm), DiSC3(5) (Ex/Em 651/675 nm) and DiSBAC2(3) (Ex/Em 530/560 nm) dye levels were quantified by fluorescence

63 as detailed in Experimental Methods. All uptake data were normalized to respective protein concentrations and were corrected for nonspecific binding by subtracting the corresponding zero time readings. Data presented as mean ± SD (n=3).

4.2.5 The Effect of DAT and NET Inhibitors on 2,2’-Cyanine Uptake

As mentioned above, DAT is believed to transport MPP+ into dopaminergic cells and there is some evidence that NET may also be involved in MPP+ uptake. Therefore, to determine whether DAT or NET are involved in 2,2’-cyanine uptake, a series of experiments measuring

2,2’-cyanine uptake by MN9D cells were carried out in the presence either specific DAT or NET inhibitors, GBR12909 (253) and desipramine (254), respectively. Some uptake experiments were also carried out in the presence of both DAT and NET inhibitors. As shown from the data in Fig.

8A, DAT and NET inhibitors had no significant effect on the uptake of 2,2’-cyanine into MN9D cells.

4.2.6 Effect of Extracellular Na+ and Ca2+ on 2,2’-Cyanine Uptake

It is believed that DAT-dependent MPP+ incorporation into dopaminergic cells (250), is dependent on extracellular Na+ (251). Therefore, to determine whether DAT is involved in cyanine uptake, the effect of extracellular Na+ on 2,2’-cyanine uptake into MN9D cells was investigated. In these experiments, Na+ in the incubation medium was replaced with equimolar concentrations of choline. In addition, previous studies from our lab have reported that extracellular Ca2+ affects MPP+ uptake into MN9D cells (252). Therefore, the effect of Ca2+ on the uptake of 2,2’-cyanine into MN9D cells was also investigated. In these experiments where complete removal of Ca2+ from the incubation medium was necessary, Ca2+ was omitted and 1.3 mM EGTA was included in the incubation medium. In all experiments, cells were incubated in a constant, 1 µM 2,2’-cyanine for 1 h and cellular uptake was measured by fluorescence (Ex/Em

320/420 nm) as detailed in Experimental Methods. As shown from the data in Fig. 8B, there was no effect of either extracellular Na+ or Ca2+ on 2,2’-cyanine uptake.

A 1.2

0.8

0.6

0.4

0.2 μmoles/mg protein. 1h protein. μmoles/mg 0

B 1

0.8

0.6

0.4

0.2 μmoles/mg protein. 1h protein. μmoles/mg

0 Regular No Ca2+ No Na+

Figure 8. Different Transporter and Ca2+ Channel Inhibitors and Effects of Extracellular Na+ and Ca2+ on 2,2’-Cyanine Uptake into MN9D Cells. MN9D cells were grown in DMEM to 70-80% confluence in 12 well plates as detailed in Experimental Methods. (A), Inhibitory Effect on 2,2’- cyanine uptake in MN9D cells. Cells were grown in 12-well plates under standard conditions as detailed in Experimental Methods. The growth media was removed and cells were washed with warm KRB-HEPES, pH 7.4 and incubated in 990 μL of KRB-HEPES, pH 7.4 containing a constant concentration of the desired inhibitor in KRB-HEPES, pH 7.4 for 10 min at 37 ˚C. Then, 10 μL 2,2’-cyanine was added and incubated for an additional 1 h at 37 ˚C. In all the experiments, the final inhibitor concentrations of the inhibitor and 2,2’-cyanine were fixed at 25

M and 1 M, respectively. Cells were washed several times, harvested and intracellular 2,2′- cyanine levels were quantified by fluorescence (Ex/Em 320/420 nm) as detailed in Experimental Methods. (B) In these experiments where Ca2+-free conditions were necessary Ca2+ is omitted from the incubation medium and 1.3 mM EGTA was included. Similarly, when Na+-free conditions were necessary Na+ in the incubation media were replaced with equimolar concentrations of choline. In all the experiments cells were incubated with 1 μM 2,2’-cyanine in the corresponding buffer for 1 h at 37 ˚C. Cells were washed, collected, and intracellular 2,2′- cyanine levels were quantified by fluorescence (Ex/Em 320/420 nm) as detailed in Experimental Methods. All the intracellular 2,2’-cyanine concentrations were normalized to respective protein concentrations as detailed in Experimental Methods. Data presented as mean ± S.D (n = 3).

4.2.7 The Effect of Ca2+ Channel Blockers on 2,2’-Cyanine Uptake

There is some literature evidence to suggest that Ca2+ channels may play a role in MPP+ uptake (255). Therefore, to examine whether or not 2,2’-cyanine uptake into dopaminergic cells is associated with calcium channels, the effects of Ca2+ channel inhibitors on the 2,2’-cyanine uptake into MN9D cells was investigated. The results of these experiments showed that L-type voltage gated calcium channel inhibitors, nitrendipine and nifedipine (256) had no effect on the

2,2’-cyanine uptake by MN9D cells, even at relatively high concentrations (Fig. 8A). Similarly, the dual T-type and L-type voltage gated calcium channel inhibitor, mibefradil (257), showed very little increase in 2,2’-cyanine uptake into MN9D cells, and the increase was not significant.

4.3 Effect of Ca2+ and Na+ Ions and Calcium Channels on Cyanine Toxicity

Ca2+ and Na+ are reportedly involved in many cases of dopaminergic neurodegeneration

(225,258). The toxicity of 2,2’-cyanine was measured in greater detail in dopaminergic MN9D cells in the presence of different buffers and different calcium channel inhibitors under the assumption that other cyanines follow the same trend.

4.3.1 Effect of Extracellular Ca2+ and Na+ on the MPP+ and 2,2’-Cyanine Toxicity in

MN9D Cells

Since the dopaminergic toxicity of MPP+ toward dopaminergic cells was shown to be dependent on extracellular Ca2+ and Na+ (252), the effect of extracellular Na+ and Ca2+ on 2,2’- cyanine toxicity was studied. In the Na+-free experiments, Na+ in the incubation buffer was replaced with equimolar concentrations of choline and in Ca2+-free experiments Ca2+ was omitted and 1.3 mM EGTA was included in the incubation medium. As shown from the data in

Fig. 9, removal of extracellular Ca2+ or Na+ had no significant effect on 2,2’-cyanine toxicity in

MN9D cells.

120

100

60 Regular No Ca2+ 40 % Cell Viability % Cell No Na+

0 Conrol 5 10 25 50 100 250 500 750

[2,2'-cyanine] (nM)

Figure 9. Effect of Different Buffers on 2,2’-Cyanine Toxicity of MN9D Cells. 2,2’-Cyanine Toxicity of MN9D in Regular, Ca Free and Na Free KRB buffers. Cells were grown in 96-well plates and were treated with 2,2’-cyanine in appropriate KRB-HEPES for 12 h at 37 °C. Cell viabilities were determined by the MTT assay as detailed in Experimental Methods. Results are expressed as % cell viability with respect to parallel untreated controls. Data presented as mean ± SD (n=6).

4.3.2 Effect of Intracellular Ca2+ on 2,2’-Cyanine Toxicity in MN9D Cells

To determine whether intracellular Ca2+ plays a role in 2,2’-cyanine toxicity, the effect of a membrane permeable Ca2+ chelator, 1,2-bis(2-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid tetrakis (acetoxymethyl ester) [BAPTA-AM] on 2,2’-cyanine toxicity was investigated. As shown in Fig. 10A, 50 μM BAPTA-AM showed no significant protection against 2,2’-cyanine toxicity in MN9D cells, suggesting that intracellular Ca2+ may not play a role in 2,2’-cyanine toxicity in MN9D cells.

4.3.3 The Effect of Ca2+ Channel Blockers and Cation Channel Inhibitors on 2,2’-Cyanine

Toxicity in MN9D cells.

To determine whether 2,2’-cyanine toxicity in dopaminergic cells is associated with voltage gated or other types of calcium or other cation channels, the effect of several Ca2+ channel blockers and cation channel inhibitors on 2,2’-cyanine toxicity in MN9D cells was investigated. These studies revealed that Ca2+ and cation channel inhibitors, nitrendipine, nifedipine, benzamil, mibefradil, verapamil, flufenamic, isadipine, 2-APB, CGP37517 and thapsigargin showed no protective effect on the MN9D cell toxicity of 2,2’-cyanine (Figs. 10B &

C). In fact, flufenamic acid and mibefradil appeared to increase the toxic effects.

A 120 Regular KRB 100 Regular KRB + BAPTA

40 % Cell Viability% Cell 20

0 0 50 100 500 1000

[2,2'- cyanine] (nM)

B 120 Control (W/O inh) 100 NITRENDIPINE

NIFENDIPINE 80 BENZAMIL 60 MIBEFRADIL 40 % Cell Viability% Cell VERAPAMIL 20 FLUFENAMIC

0 ISADIPINE Control 5 10 25 50 100

[2,2’-cyanine] (nM) Figure 10. The Effect of the Calcium Chelator, BAPTA and Calcium Channel Inhibitors on 2,2’- Cyanine Toxicity of MN9D Cells. (A), 2,2’-cyanine toxicity in MN9D with or without the calcium chelator, BAPTA in regular KRB-HEPES. Cells were grown in 96-well plates and were pre-incubated with 40 µL of 50 µM BAPTA-AM for 30 min followed by the addition of 10 µL

2,2’-cyanine where [2,2’-cyanine]final was 0-1000 nM in regular KRB-HEPES. (B), MN9D cells grown in 96 well plates were initially incubated with 25 µM inhibitor for 10 min followed by the addition of 0-100 nM 2,2’-cyanine in KRB-HEPES for 12 h at 37 °C. (C), MN9D cells grown in 96 well plates were initially incubated with 25 µM inhibitor or mixed inhibitors for 10 min followed by the addition of 0-500 nM 2,2’-cyanine in KRB-HEPES for 12 h at 37 °C KRB- HEPES for 12 h. Cell viabilities were determined by the MTT assay as detailed in Experimental Methods. Results are expressed as % cell viability with respect to parallel untreated controls. Data presented as mean ± SD (n=6).

C 120 Regular 2APB -25uM 100 CGP -50uM

FLU-50uM 80 THAP-25uM 60 2APB+FLU CGP+THAP 40 % Cell Viability% Cell

0 0 5 10 25 50 75 100 200 300 500 [2,2'-cyanine] (nM) Figure 10 (continued)

4.4 Intracellular Cyanine Dyes Localize in Mitochondria

Live cell imaging experiments with the mitochondrial labeling dye, Mitotracker, were performed to determine whether the positively charged cyanine dyes actively accumulated in the mitochondria of the cell due to the high negative potential of the inner mitochondrial membrane.

These experiments revealed that the positively charged cyanine dyes efficiently accumulated in the mitochondria (data only shown for DiSC3(5) dye) (Fig. 11A). On the other hand, the negatively charged cyanine dye, DiSBAC2(3) is distributed throughout the cell without detectable localization in the mitochondria (Fig. 11B).

A MN9D

DiSC3(5) - Cyanine Mitotracker Green Overlay HEPG2

DiSC3(5) - Cyanine Mitotracker Overlay

Green

Figure 11. Mitochondrial Accumulation of DiSC3(5) Cyanine and the Negatively Charged, DiSBAC2(3). MN9D and HepG2 cells grown in glass bottomed plates were initially incubated with 100 nM of DiSC3(5) or DiSBAC2(3) for 15 min at 37 °C, then the solution was removed, washed and incubated with 100 nM of Mitotracker Green or Mitotracker Red for 15 min at 37 °C. The solution was then removed, cells washed and images captured at corresponding wavelengths using a Leica confocal fluorescence microscope equipped with a 40X objective. Mitochondrial accumulation of dye was only observed for DiSC3(5) cyanine (A), but not with the negatively charged dye, DiSBAC2(3) (B) in both MN9D and HepG2 cells.

Figure 11 (continued)

4.5 Cyanines, MPP+ and Rotenone Cause Mitochondrial Membrane Potential

Depolarization

Since cyanines and MPP+ are positively charged hydrophobic compounds, they can accumulate electrogenically inside negatively charged mitochondria and depolarize the mitochondrial membrane. In addition, the mitochondrial complex I inhibitor, rotenone, is known to depolarize the mitochondrial membrane potential as a consequence of mitochondrial complex

I inhibition. Thus, the relative effects of cyanines, MPP+, and rotenone on the mitochondrial membrane potentials of live MN9D and HepG2 cells were determined using the fluorescent dye tetramethylrhodamine (TMRM) following well worked out literature procedures (232). The data

72 in Fig. 12A show that 2.5 M 2,2'-cyanine depolarized the mitochondrial membrane potentials of both MN9D and HepG2 cells to a similar extent i.e. 40-50 %. The images of TMRM fluorescence (Ex/Em 543/573 nm) before and after adding 2.5 M 2,2'-cyanine showed a remarkable decrease in fluorescence due to mitochondrial membrane potential depolarization in both MN9D and HepG2 cells (Fig. 12B). Parallel experiments with 500 M MPP+ with MN9D and HepG2 showed a similar but less pronounced mitochondrial depolarization (Fig. 12C). In addition, all of the positively charged cyanine dyes DiOC2(3), DiSC3(3) and DiSC3(5) at 2.5

M concentrations showed about 40 % membrane depolarization in MN9D cells, similar to

2,2’-cyanine. However, as expected, negatively charged DiSBAC2(3) did not depolarize the mitochondrial membrane potential to a measurable level. In a parallel study, 2.5 μM rotenone showed similar but less pronounced mitochondrial membrane depolarization, about 10-20 %

(Fig. 12D).

A 10

Time (min) 0 0 5 10 15 20 -10

-20 HEPG2

-30 MN9D -40

Potential Mem % Mito -50 -60

Figure 12. The Effects of Cyanines, MPP+, Rotenone and the Negative Dye on Mitochondrial Membrane Potentials. The effect of (A), 2,2′-cyanine and (C), MPP+ on the mitochondrial membrane potential of MN9D and HepG2 cells: Cells were treated with 2.5 M 2,2′-cyanine (or 500 M MPP+) and the mitochondrial membrane potentials were determined by using the fluorescent probe TMRM (Ex/Em 543/573 nm) as detailed in Experimental Methods. (B), Images of TMRM (Ex/Em 543/573 nm) fluorescence in MN9D (B1 & B2), and HepG2 (B3 & B4), cells before and after adding 1 μM 2,2’-cyanine. (C), The effect of DiOC2(3), DiSC3(3) and

DiSC3(5) cyanines, the negative dye, DiSBAC2(3) and rotenone on the mitochondrial membrane potentials of MN9D cells. Cells were treated with 2.5 M cyanines, negative dye or rotenone and mitochondrial membrane potentials were determined by using the fluorescent probe TMRM (Ex/Em 543/573 nm) as detailed in Experimental Methods.

B1 B2

B3 B4

C 10 Time /min 0 0 10 20 30 40 50 -10 -20

-30

(arbitorary units) (arbitorary -40 Decrease Pote. Memb. -50

Figure 12 (continued)

D 5 Time (min) 0 -5 0 2 4 6 8 10 DiOC23

-10 DiSC3 -15 DISC5 -20

(arb.units) (arb.units) DiSBAC -25 Rotenone -30

Mito. mem. potential decrese decrese potential mem. Mito. -35 -40

Figure 12 (continued)

4.6 Inhibition Studies of Mitochondrial Complex I with Cyanines, MPP+ and Rotenone

Inhibition studies of mitochondrial complex I were carried out with mitochondria isolated from rat brain according to the well-established NADH-ubiquinone oxidoreductase activity assay procedure (235). Complex I specific activity was measured by following the decrease in absorbance at 340 nm due to oxidation of NADH with rotenone, cyanine dyes, MPP+ and a negative charged dye (Fig. 13). The well characterized potent mitochondrial complex I inhibitor, rotenone, was found to inhibit 87.5 % of the complex I activity at 10 M concentration.

Interestingly and unexpectedly, cyanines were also found to be excellent inhibitors for mitochondrial complex I with potencies similar to rotenone itself. The data presented in Fig. 13

& Table 4 show that all the positively charged cyanines are potent inhibitors of complex I with

80-60% inhibition of complex I at 10 μM concentrations. However, the most commonly used PD model toxin, MPP+, showed a minor mitochondrial complex I inhibition potency (12.5%) even at

200 μM. As expected, negatively charged DiSBAC2(3) showed very little or no complex I inhibition at 10 M concentration.

0.1 Time (Sec)

0 -0.1 0 100 200 300 400 Control -0.2 22Cyanine - 10uM -0.3 44Cyanine-10uM -0.4 DiOC23-10uM DiSC3-10uM -0.5 DiSC5-10uM -0.6 DiSBAC-ve-10uM NADH Absorbance (340nm) AbsorbanceNADH -0.7 Rotenone-10uM -0.8 MPP+ - 200 uM

Figure 13. The Effects of Rotenone, Cyanines, MPP+ and the Negative Dye on Mitochondrial Membrane Potentials and Mitochondrial Complex I Inhibition. As described in Experimental Methods, isolated rat brain mitochondria were incubated with 10 μM cyanine dyes, the 10 μM negative dye, 10 μM rotenone or 200 μM MPP+ for 2 min and then 10 mM NADH was added and the decrease of absorbance at 340 nm was recorded for 10 min.

Table 4. Percentages of Rat Brain Mitochondrial Complex I Inhibition by Cyanines, MPP+ and Rotenone in NADH-Ubiquinone Oxidoreductase Activity Experiments. The data were derived from the traces in Fig. 13

Complex I Inhibition % Compound with respect to control Rotenone 87.5

DiOC2(3) 79.1

DiSC3(3) 83.3

DiSC3(5) 83.3

2,2’-cyanine 70.8

4,4’-cyanine 66.7

MPP+ 12.5

DiSBAC2(3) 0-2

4.7 Effect of Cyanine and Rotenone on ATP Levels in MN9D and HepG2 Cells

The above results showed that cyanine dyes actively accumulated in the mitochondria, depolarized the mitochondrial membrane potential and inhibited complex I. Therefore, it is expected that these toxins must also reduce mitochondrial ATP production. Accordingly, the effects of cyanines and rotenone on intracellular ATP levels of MN9D and HepG2 were determined using an ATP assay kit (Sigma) as detailed in Experimental Methods. As shown in

Fig. 14, cyanine (100 nM DiSC3(5)) treatment depleted intracellular ATP in both MN9D

( ~40%) and HepG2 (~20%) cells. As expected, rotenone-treated MN9D and HepG2 cells depleted ATP levels by 25 % and 50 % respectively.

120

100

60 MN9D % ATP HEPG2 40

Control DiSC3(5)-100 nMRotenone-25 nM

Figure 14. Effects of DiSC3(5) Cyanine and Rotenone on Intracellular ATP Levels in MN9D and HepG2 Cells. Cells were grown in DMEM to 70-80 % confluence in 12 well plates treated with 100 nM DiSC3(5) or 25 nM rotenone for 12 h at 37 °C and intracellular ATP levels were quantified as detailed in Experimental Methods. All the intracellular ATP levels were normalized to respective protein concentrations also as detailed in Experimental Methods. Data presented as mean ± S.D (n = 3).

4.8 Cyanine-, MPP+- and Rotenone-Induced ROS Production in Cells

In many neurodegenerative diseases, including PD, one of the major causes of neuronal cell death is proposed to be increased oxidative stress. In order to determine whether increased

ROS production is associated with the toxicities of the cyanines and other toxins, intracellular

ROS levels were measured in toxin-treated cells using a membrane permeable DCFH-DA fluorescence assay (Ex/Em 504/526 nm) as detailed in Experimental Methods.

4.8.1 2,2'-Cyanine and MPP+ Cause Concentration and Time Dependent Increases in

Intracellular ROS Levels in MN9D and SH-SY5Y, but Not in HepG2 Cells.

Treatment of MN9D cells with 2,2'-cyanine for 1 h increased the intracellular ROS levels by about two- to five-fold from baseline levels, depending on the concentration of 2,2'-cyanine

(250 to 1000 nM), (Fig. 15A). As shown in Fig. 15A, 2,2'-cyanine treatment also caused the increase of ROS in SH-SY5Y cells, but there was less pronounced in comparison to MN9D cells, under similar experimental conditions. Interestingly, in contrast to MN9D cells, baseline ROS levels were not measurably increased in HepG2 cells, even at the maximum concentration, 1 M

2,2'-cyanine, under similar experimental conditions (Fig. 15B). The time course studies of ROS production showed that the increased ROS levels in MN9D cells in response to 2,2'-cyanine treatment initiated after a lag period of about 20 min and steadily increased with time thereafter

(Fig. 15C). In addition, MPP+ also showed a similar but less pronounced concentration dependent increase in ROS, specifically in MN9D cells, but not in HepG2 cells (Fig. 15D).

Similar to 2,2'-cyanine, the other positively charged cyanines and rotenone also produced significant increase in intracellular ROS in MN9D cells. In contrast, the negatively charged

DiSBAC2(3) did not produce measurable changes in ROS in MN9D cells under similar experimental conditions (Fig. 15E).

600 A 500

400 HEPG2 300

% ROS SHS5Y 200 MN9D 100

0 0 0.25 0.5 1 [2,2' - cyanine] (μM)

Figure 15. The Effects of Cyanines, the Negative Dye, MPP+ and Rotenone on the Intracellular ROS Levels. Cells were grown in 12-well plates and treated with the desired concentration of compounds for 1 h at 37 °C in KRB-HEPES and intracellular ROS levels were quantified by the 2′,7′-dichlorofluoresceine diacetate method as detailed in Experimental Methods. All the intracellular ROS levels were normalized to respective protein concentrations as detailed in

Experimental Methods. Data presented as mean ± S.D (n = 3). (A), The effect of 2,2′-cyanine on intracellular ROS levels in MN9D, SH-SY5Y and HepG2 cells. (B), The trans, 2′,7′- dichlorofluoresceine diacetate (DCFH-DA) fluorescence corresponding to ROS and overlaid images were captured by a Leica confocal fluorescence microscope equipped with a 40X objective lens. MN9D and HepG2 cells after treating with 2,2’-cyanine (1.0 μM ) for 1 h. (C), time dependent increase of 2,2′-cyanine (1.0 M) mediated ROS production in MN9D cells. (D), The effect of MPP+ on intracellular ROS levels in MN9D and HepG2 cells. (E), The effect of DiOC2(3), DiSC3(3) and DiSC3(5) cyanines, the negative dye, DiSBAC2(3) and rotenone on ROS levels in MN9D cells.

C 180 160 140

120 100 80 60

% Increase in ROS in % Increase 40

20 0 5 10 20 30 40 50 60 Time (min)

D 180 160 140

120

100 80

% ROS HEPG2 60 MN9D 40

20 0 0 100 250 500 [MPP+ ] (μM)

Figure 15 (continued)

E 500 450 400

350

300 DiSBAC -ve 250 DiOC23 % ROS 200 DiSC3

150 DiSC5 100 Rotenone 50

0 0 250 500 1000 [ Dye] (nM) Figure 15 (continued)

4.8.2 Cyanine Induced ROS Production is Localized in the Mitochondrial Matrix

The above results show that cyanines inhibit the mitochondrial complex I in a manner similar to rotenone. Since rotenone is known to produce ROS in the mitochondrial matrix a series of experiments were carried out with DiSC3(5) to determine whether cyanine-mediated

ROS production also initiated in the mitochondria. As shown from the data in Fig 16, DCFH-DA fluorescence overlaped well with the DiSC3(5) fluorescence. Previous experiments with

Mitotracker showed DiSC3(5) localized in the mitochondria (Fig. 11A), thus the data in Fig. 16A confirm that cyanine also initiated ROS production in the matrix of the mitochondria, similar to rotenone. The enlarged single cell images of the overlays of DiSC3(5) and DCFH-DA fluorescence images clearly confirmed the production of ROS in the mitochondrial matrix (Fig

16B).

DCFH-DA DiSC3(5) Overlay

Figure 16. Cyanine Induces ROS Production in the Mitochondrial Matrix. (A), MN9D cells were grown in glass bottomed plates and treated with 50 μM DCFH-DA for 1 h at 37 °C in KRB-HEEPS, then the medium was removed and cells were further incubated with 1 M DiSC3(5) for 1 h at 37 °C in KRB-HEPES. Then incubation media was removed, cells were washed and DCFH-DA and DiSC3(5) fluorescence images were captured at appropriate wavelengths using a Leica confocal fluorescence microscope equipped with a 40X objective. (B) Enlarged single cell images of DCFH-DA, DiSC3(5), and overlay of the two images.

4.8.3 The Antioxidant, Ascorbate (Asc) Pretreatment Inhibits the ROS Production by

Cyanines, MPP+ and Rotenone Treatments in MN9D Cells

Asc is a potent antioxidant and is known to lower intracellular ROS levels under certain adverse cellular conditions (259). To test whether Asc could lower 2,2'-cyanine-mediated ROS production, MN9D cells were pretreated with increasing concentrations of Asc (0-10 mM) in

KRB-HEPES for 1 h at 37 °C and then exposed to 500 nM 2,2′-cyanine in the presence of the same concentration of Asc for an additional 1 h. Following the incubation, the intracellular ROS levels were determined by fluorescence. As shown in Fig. 17A, Asc treatment effectively reduced 2,2′-cyanine mediated ROS production in MN9D cells in a concentration dependent manner. Parallel experiments with DiOC2(3), DiSC3(3) and DiSC3(5) cyanines (500 nM),

MPP+ (500 M) and rotenone (500 nM) treated MN9D cells also showed that intracellular ROS production was significantly attenuated by Asc (Figs. 17B & C). Live cell fluorescence imaging experiments confirmed that ROS was produced intracellularly in response to 2,2′-cyanine (500 nM) treatment and Asc (2.0 mM) effectively modulated intracellular ROS production under the same incubation conditions (Fig. 17D).

A 120 100

80 60 % ROS % 40 20

0 0 0.25 0.5 1 2 5 10 [Ascobate] (mM) B 120

100

60 % ROS 40

20 0 0 0.5 1 2

[Ascorbate] (mM)

Figure 17. The Antioxidant, Ascorbate, Attenuates Cyanines, MPP+ and Rotenone Mediated ROS Production in MN9D Cells. MN9D cells were pre-incubated with the desired concentration of Asc in KRB-HEPES for 1 h at 37 °C. Then the cells were incubated with (A), 500 nM 2,2′- cyanine, (B), 500 M MPP+ or (C), 500 nM DiOC2(3), DiSC3(3), DiSC3(5) or rotenone in the presence of the same concentration of Asc for an additional 1 h in KRB-HEPES at 37 °C and the intracellular ROS levels were determined, as detailed above. Data presented as mean ± S.D (n=3). (D), Effect of Asc on 2,2′-cyanine mediated ROS production as visualized by live cell imaging. MN9D cells grown in glass bottomed plates were incubated without (first row) or with (second row) 2 mM Asc for 1h at 37 °C and then treated with 10 μM DCFH-DA for 1h. Then cells were washed and incubated with 500 nM 2,2′-cyanine in the absence (first row) or presence (second row) of 2 mM Asc. Trans images (Left), DCFH-DA fluorescence (Ex/Em 504/526) images (Middle) and the overlay of Trans and DCFH-DA fluorescence images.

C 120

100

DiOC23

60 DiSC3 % ROS 40 DiSC5 Rotenone 20

0 0 0.5 1 2 [Ascorbate] (mM) D

Without Asc

Trans DCFH-DA Overlay

With Asc

Trans DCFH-DA Overlay

Figure 17 (continued)

4.8.4 Depletion of Intracellular Dopamine (DA) Reduces Cyanines, MPP+ or Rotenone

Mediated ROS Production in MN9D Cells

Oxidatively sensitive DA is known to amplify ROS production in dopaminergic cells in the presence of various neurotoxins (177). To determine whether intracellular DA plays a similar role in the specific increase of ROS production in catecholaminergic cells in response to 2,2'- cyanine treatment, the effect of 2,2'-cyanine on intracellular DA levels of MN9D cells was examined (MN9D cells contain high levels of DA, but no other catecholamines are detectable under the standard growth conditions). As shown in Fig. 18A, treatment of MN9D cells with increasing concentrations of 2,2'-cyanine for 1 h decreased the intracellular DA in a concentration-dependent manner. In addition, treatment of MN9D cells, which were treated with the tyrosine hydroxylase inhibitor L-α-methyltyrosine (92) for 24 h to deplete intracellular DA, with 250 nM 2,2'-cyanine for 1 h showed a significant reduction in ROS production in comparison with cells grown without THI under normal conditions (Fig. 18B). The ROS levels decreased with increasing concentration of THI and a maximum 40% reduction of ROS was observed at 6 mM THI in treated cells as compared with untreated control cells (Fig. 18B). A similar trend was also observed with 500 M MPP+ under similar conditions (Fig. 18C).

Similarly, the data presented in Fig. 18D showed a similar reduction of DiOC2(3)-, DiSC3(3)-,

DiSC3(5)-cyanines and rotenone-induced ROS production in MN9D cells grown with THI under normal conditions. For example, 6 mM THI showed a maximum ROS depletion of about 35% in

DiOC2(3), DiSC3(3), DiSC3(5) cyanines and rotenone treated cells. Control experiments showed that treatment of MN9D cells with THI (2-6 mM) for 24 h under normal growth conditions, reduced intracellular DA levels by 80-95% depending on the concentration of THI

(data not shown). However, even 95% depletion of intracellular DA levels through tyrosine hydroxylase inhibition with THI only reduced the ROS production by about 40%.

A 120

100 80 60 40

% Dopamine 20 0 0 0.25 0.5 1

[2,2' -Cyanine] (μM) B 120

100

60 % ROS 40 20

0 0 1 3 6 [THI] (mM) Figure 18. 2,2′-Cyanine Decreases the Intracellular DA Levels and Prior DA Depletion Attenuates the Cyanines, MPP+ and Rotenone Mediated ROS Production in MN9D Cells. (A), 2, 2′-Cyanine decreases the intracellular DA levels in MN9D cells. Cells were treated with the desired concentration of 2,2′-cyanine for 1 h at 37 °C and intracellular DA levels were quantified by reversed-phase HPLC-EC as detailed in Experimental Methods. All the data were normalized to the respective protein concentration of each sample. Catecholamine levels are expressed as percentage of a control which was treated under identical conditions except that 2,2′-cyanine was omitted from the incubation medium. Data presented as mean ± S.D. (n=3). MN9D Cells were treated with 0-6 mM THI for 24 h to deplete intracellular DA and DA depleted cells were treated with (B), 250 nM 2, 2′-cyanine, (C), 500 M MPP+ or (D), 250 nM of each DiOC2(3), DiSC3(3), DiSC3(5) or rotenone for 1 h at 37 °C and intracellular ROS levels were quantified

87 by the DCFH-DA method as detailed in Experimental Methods. The data were normalized to respective protein concentrations of samples and presented as mean ± S.D (n=3).

120 C 100

80 60 % ROS 40

20 0

0 1 3 6 [THI] (mM) 120 D 100

80 DiOC23 60 DiSC3 % ROS 40 DiSC5 20 Rotenone 0 0 1 3 6 [THI] (mM)

Figure 18 (continued)

4.9 Asc Pretreatment Protect MN9D Cells from Cyanine, MPP+ and Rotenone

Toxicities

The effect of Asc pretreatment on cyanines, MPP+ and rotenone toxicity in MN9D cells was also determined. The data in Fig. 19A showed that 500 M Asc effectively protected the

MN9D cells from 2,2′-cyanine toxicity parallel to its effect on intracellular ROS levels (Fig.

17A). Similarly as shown from the data in Fig. 19B, C & D, the 500 M Asc treatment

88 significantly protected MN9D cells from DiOC2(3), DiSC3(3) and DiSC3(5) cyanines, MPP+ and rotenone toxicities as well.

A 120 No Ascorbic

100 With 500uM Ascorbic

40 Viability% Cell 20

0 0 5 10 25 50 75 100 250 500 1000 [2,2’ – Cyanine] (nM)

B 120

100

DiOC23 80 DiSC3 60 DiSC5

40 DiOC23+ASC

Viability % Cell DiSC3+ASC 20 DiSC5+ASC 0 0 5 10 25 50 75 100 250 500 1000 Concentration (nM)

Figure 19. The Effect of Antioxidant, Asc on MN9D Cell Toxicities of Cyanines, MPP+ and Rotenone. MN9D cells were pre-treated with 500 μM Asc for 1 h at 37 °C and then incubated with (A), 0-1000 nM 2,2′-cyanine or (B), 0-1000 nM of DiOC2(3), DiSC3(3) and DiSC3(5) (C) 0-300 μM MPP+ and (D), 0-100 nM rotenone in the presence of the same respective concentrations of Asc in KRB for 12 h at 37 °C, and cell viabilities were determined by the MTT assay. Data presented as mean ± SD (n=6).

C 120 No Inhibitor 100 500 uM ASC 80

40 % Cell Viability% Cell 20 0

0 10 25 200 300 + [MPP ] (μM)

D 120 + ASC 500uM - ASC 100

40 % Cell Viability% Cell 20

0 0 5 10 15 20 25 30 50 75 100 [Rotenone] (nM) Figure 19 (continued)

4.10 DA Depletion Protects MN9D Cells from Cyanines, MPP+ and Rotenone Toxicities

The tyrosine hydroxylase inhibitor, L-α-methyltyrosine (92), mediated DA depletion in which mitigated cyanine, MPP+ and rotenone toxicity in MN9D cells as shown in Fig. 20. The data clearly show that DA depletion reduced the 2,2′-cyanine toxicity effectively in a concentration dependent manner parallel to the effect on the ROS production (Figs. 20A & 18B).

In addition, the data clearly show that pretreatment of MN9D cells with 6 mM THI also reduces

90 the DiOC2(3), DiSC3(3) and DiSC3(5) cyanine toxicities effectively and parallel to the effect on the corresponding ROS productions (Figs. 20B & 18D). Similarly, those 6 mM THI treatments reduced the MPP+ and rotenone toxicities effectively, as well (Figs. 20C & D). Although the above data clearly show that the depletion of DA significantly reduces the toxicities of cyanines,

MPP+ and rotenone, the data also show that the drastic DA depletion only partially protected the cells from the toxic effect of these compounds.

A 120

100

Control 60 1mM THI 40 Viability % Cell 3mM THI 6mM THI 20

0 0 0.005 0.01 0.025 0.05 0.075 0.1 0.2 0.5 1 [2,2' -cyanine] (μM)

Figure 20. The Effect of DA Depletion on Cyanines, MPP+ and Rotenone induced MN9D Cell Toxicity. (A), MN9D cells were initially incubated with 0-6 mM tyrosine hydroxylase inhibitor, L-α-methyltyrosine (92) for 24 h. Then, DA depleted cells were incubated with 2,2′-cyanine or MPP+ in the presence of the same concentration of THI for 12 h at 37 °C, and cell viabilities determined by the MTT assay. The effects of 6 mM THI pre-incubation on the MN9D toxicities of (B) 0-1000 nM DiOC2(3), DiSC3(3) or DiSC3(5), (C) 0-300 μM MPP+ and (D), 0-100 nM rotenone. Data presented as mean ± SD (n=6).

B 120 DiOC23 100 DiSC3

80 DiSC5

60 DiOC23-THI DiSC3-THI 40 Viability% Cell DiSC5-THI 20

0 5 10 25 50 75 100 250 500 1000 Concentration (nM) C 120

100 No Inhibitor 80 6mM THI 60

40 % Cell Viability% Cell 20

0 0 10 25 50 200 300 [MPP+] (μM)

D 120 100 + THI 6mM

80 - THI

60 40

Viability% Cell 20

0 0 5 10 15 20 25 30 50 75 100 Rotenone (nM) Figure 20 (continued)

4.11 Antioxidant Enzyme Levels are Low in MN9D Cells in Comparison to HepG2 Cells

To determine whether the high vulnerability of dopaminergic cells towards 2,2′-cyanine toxicity as compared with liver HepG2 cells, is related to the relative expression of anti-oxidant enzymes, the relative activities of major anti-oxidant enzymes, superoxide dismutase (38), catalase and glutathione peroxidase in MN9D, SH-SY5Y, and HepG2 cells were determined using commercially available assay kits. As shown from the data in Table 5, all three enzyme levels were significantly lower in MN9D cells as compared with HepG2 cells. Similarly, SOD and catalase levels were significantly lower in SH-SY5Y cells, parallel to MN9D cells. However, glutathione peroxidase levels in SH-SY5Y cells were similar to those of HepG2 cells.

Table 5. Comparison of Relative Antioxidant Enzyme Activities of Rat Brain and Liver with Dopaminergic and Liver Cells. 1Data from (190) 2Determined as detailed in Experimental Methods. All three enzyme activities of HepG2 cells were adjusted to 100% and presented as mean ± SD (n=3). 3Data represent the activities of both Cu/Zn SOD and Mn SOD.

SOD Catalase Gpx Cu/Zn SOD Mn SOD Liver1 100 ± 7 100 ± 12 100 ± 9 100 ± 8 Brain1 37 ± 4 47 ± 7 3 ± 0 11 ± 3

HepG22 100 ± 73 100 ± 7 100 ± 10 Mn9D2 47 ± 23 16 ± 3 20 ± 2 2 3 SH-SH5Y 35 ± 4 26 ± 1 94 ± 13

4.12 Cyanine, MPP+ and Rotenone Cause Apoptotic Cell Death in MN9D Cells

To determine whether ROS induced apoptosis is the cause of cyanine, MPP+ and rotenone mediated dopaminergic cell death, apoptosis testes were carried out. As shown in the images in Fig. 21, 2,2′-cyanine mediated MN9D cell death was associated with cell shrinkage

(Figs. 21A1 & A2), chromatin condensation (Figs. 21A3 & A4), and DNA fragmentations

(laddering) (Fig. 21B) strongly suggesting the involvement of the apoptotic cell death pathway.

As shown from the data in Fig. 21C, DiOC2(3), DiSC3(3), DiSC3(5), MPP+ and rotenone mediated MN9D cell death was also associated with chromatin condensation as indicated by the increase in DAPI fluorescence in the nucleus, strongly supporting the involvement of the apoptotic cell death pathway. Moreover, control or negatively charged cyanine dye, DiSBAC2(3) treated cells did not show any increase in nuclear DAPI fluorescence, as expected.

A1 A2

A3 A4

Figure 21. Cyanines, MPP+ and Rotenone Cause Apoptotic Dopaminergic MN9D Cell Death. Cells were grown in glass bottomed plates and incubated with 1 μM 2,2′-cyanine in KRB- HEPES for 12 h at 37 °C and then treated with 300 nM DAPI for 15 min in the dark and apoptotic chromatin condensation was visualized by fluorescence microscopy (Ex/Em 358/461 nm). Control cells were treated in a similar manner except that 2,2′-cyanine was excluded from the incubation medium. (A1), Control MN9D cell morphology; (A2), 2,2-cyanine treated MN9D cell morphology; (A3) DAPI fluorescence of control cells without 2,2′-cyanine; (A4); DAPI fluorescence of 2, 2′-cyanine treated cells. (B), DAPI experiment was carried out with 1 μM cyanines or 100 μM MPP+ or rotenone or the negatively charged DiSBAC2(3) in KRB-HEPES for 12 h at 37 °C. The apoptotic chromatin condensation was visualized by fluorescence microscopy (Ex/Em 358/461 nm). Control cells were treated in a similar manner excluding dyes or rotenone from the incubation medium. (C), Apoptotic DNA fragmentation of MN9D cells treated with different concentrations of 2,2’-cyanine for 12 h at 37 °C.

B DiSBAC2(3) DiOC2(3) DiSC3(3)

+ DiSC3(5) MPP Rotenone

Figure 21 (continued)

CHAPTER V

DISCUSSION

In agreement with previous reports, the present study clearly demonstrated that the PD- causing neurotoxin, MPP+ is highly toxic to mouse dopaminergic MN9D cells (226), similar to mesencephalic dopaminergic primary cultures (260,261) and moderately toxic to human dopaminergic/adrenergic SH-SY5Y cells. Note that in addition to dopaminergic characteristics

(220), SH-SY5Y cells also have adrenergic characteristics, including the expression of both DAT and NET and the DM catalyzed conversion of DA to NE (262). They were relatively nontoxic to human liver HepG2 cells which don’t have DAT or NET (Figs. 3A & B). Both 2,2'- and 4,4'- cyanines were also specifically toxic to dopaminergic MN9D, PC12 and SH-SY5Y cells, but not to HepG2 liver cells (Figs. 4A, B & C). More importantly, both 2,2'- and 4,4'-cyanines followed a parallel cell specific toxicity pattern as MPP+, except that the former were remarkably ~1000 fold more potent toxins in comparison to MPP+ under similar experimental conditions (Figs. 3A,

4A & B). In addition, 2,2’-cyanine was also toxic to physiologically more relevant differentiated

MN9D cells, and the potency of toxicity was very similar to the undifferentiated MN9D cells

(Figs. 4D). These findings demonstrate that these cyanines are potent and specific dopaminergic toxins, similar to MPP+, and exhibit the same cell-specific pattern of toxicity; i.e. the order of toxicity is MN9D>SH-SY5Y>> HepG2 cells (Figs. 3, 4A, B & C).

MPP+ uptake into all cell types was saturable at low concentrations, demonstrating the presence of an efficient uptake system in these cells. Comparison of the corresponding Km parameters showed that while the uptake system of SH-SY5Y had the highest affinity for MPP+,

HepG2 had the lowest. However, the corresponding Vmax parameters show that HepG2 uptake

97 occured through a high volume transporter, in comparison to both SH-SY5Y and MN9D cells.

These findings showed that these cells took up MPP+ at varying efficiencies and the uptake characteristics was dependent primarily on the cell type, suggesting that multiple cell-specific transporters are present in these cells (Fig. 6A).

In contrast to MPP+, the uptake of cyanines into all cell types was not saturable and showed no cell specificity (Fig. 6B & C), indicating that the cellular accumulation exclusively occured through a non-mediated simple diffusion process, as predicted due to more hydrophobicity. It is believed that Na+ dependent dopamine transporter (DAT) (250) and norepinephrine transporter (NET) (263) are involved in MPP+ uptake. However, Na+ independence of the cyanine uptake together with the lack of inhibition with specific DAT and

NET inhibitors, GBR12909 and desipramine, respectively, confirmed that neither DAT nor NET play a role in cyanine uptake (Fig. 8A). Previous studies in our laboratory have also shown that

MPP+ uptake into the catecholaminergic cells is highly regulated by extracellular Ca2+ (252,255).

However, cyanine uptake experiments with Ca2+ free buffers showed that cyanine uptake did not depend on extracellular Ca2+ (Fig. 8B). In addition, uptake experiments with cyanine in the presence of L-type voltage gated calcium channel inhibitors, nitrendipine and nifedipine (256) and T-type and L-type voltage gated calcium channel inhibitor mibefradil (257), clearly showed that none of the voltage gated calcium channels play a role in cyanine uptake into MN9D cells

(Fig. 8A). Similarly, Ca2+ and cation channel inhibitors, nitrendipine, nifedipine, benzamil, mibefradil, verapamil, flufenamic, isadipine, 2-APB, CGP37517 and thapsigargin or intracellular

Ca2+ chelator BAPTA had no effect on cyanine uptake. As expected, parallel to the uptake characteristics, extracellular Ca2+ (Fig. 9) and Ca2+ chelator BAPTA (Fig. 10A) or Ca2+ channel inhibitors (Figs. 10B & C) had no effect on the MN9D toxicity of 2,2’-cyanine.

The time course of 2,2'-cyanine uptake into MN9D cells shows an apparent saturation with a trans-membrane concentration gradient of about 300 (assuming 3.0-4.0 L/mg cell volume), suggesting that intracellular accumulation must occur through a non-mediated, but active transport process. Clearly, such a large transmembrane concentration gradient could not be generated by plasma membrane potential-driven, simple diffusion which is equivalent to the free energy of a 6.5-13.7 concentration gradient (assuming -50 to -60 mV plasma membrane potential). Thus, non-specific intracellular accumulation of these cyanines in large quantities must be due to their compartmentalization into highly electrogenic organelles such as mitochondria.

Based on the above findings, we predicted that any lipophilic cationic dyes similar to

2,2’- and 4,4’-cyanines could be specifically and highly toxic to dopaminergic cells. Based on this prediction, we tested more complex cationic cyanine dyes DiOC2(3), DiSC3(3) and

DiSC3(5). As predicted, these were all found to be ~1000 fold greater toxic to dopaminergic

MN9D cells with toxicity than MPP+, similar to 2,2’- and 4,4’-cyanines. In addition, similar to

2,2’- and 4,4’-cyanines, other cationic cyanine dyes freely and effectively accumulate in all cells generating a high trans-membrane concentration gradient of about 300. Therefore, the characteristics of cellular accumulation and cell specific toxicity were similar in all the cyanine derivatives tested. One significant advantage to the use of DiOC2(3), DiSC3(3) or DiSC3(5) cyanines in further studies is that they are highly fluorescent (2,2’ and 4,4’-cyanines are not fluorescent) and therefore all ideal for cellular localization studies. Similarly, the negatively charged DiSBAC2(3) is also fluorescent and was used as a valuable negative control.

Accordingly, the experiments with mitochondrial labeling dye Mitotracker has shown that all positively charged cyanine dyes effectively accumulate in the negatively charged mitochondria

99 non-specifically in both MN9D and HepG2 cells. On the other hand, the negatively charged dye accumulated in all areas of the cell without significant specificity (Figs. 11A & B).

All cationic cyanine dyes tested (2,2'- and 4,4’-cyanines, DiOC2(3), DiSC3(3) and

DiSC3(5)) effectively depolarized the mitochondrial membrane potentials of both MN9D and

HepG2 cells (about 30-50%). However, the negatively charged DiSBAC2(3) did not measurably depolarize the mitochondrial membrane potentials of MN9D cells (Figs. 12A & D). These results further support that lipophilic, cationic cyanine dyes electrogenically accumulate at high concentration in the negatively charged mitochondrial matrix, independent of the nature of the cell. In agreement with previous reports, by Nakamura, et al., 2000 (150), MPP+ was also found to depolarize mitochondrial membrane potential in both MN9D and HepG2 cells, but the effectiveness appeared to be weaker in comparison to cyanines (Figs. 12A & C). In addition, cationic cyanines and MPP+ produced concentration-dependent, excessive ROS production in dopaminergic cells, but not in HepG2 cells. However, as expected, negatively charged

DiSBAC2(3) did not produce increased amounts of ROS in either MN9D or HepG2 cells. Since these cyanines accumulate in the mitochondria and depolarize the mitochondrial membrane with production of increased levels of ROS parallel to the well-known mitochondrial complex I inhibitor, rotenone, we hypothesized that cyanines may also behave as potent complex I inhibitors. Mitochondrial complex I NADH: ubiquinone oxidoreductase activity studies using isolated rat brain mitochondria showed significant inhibition of mitochondrial complex I activity with all tested cyanines (Fig. 13) consistent with that hypothesis. Surprisingly, DiOC2(3),

DiSC3(3) and DiSC3(5) were as effective as mitochondrial complex I inhibitors (83%) as the best mitochondrial complex I inhibitor, rotenone (87%) (Table 4). Both 2,2’ and 4,4’-cyanines also showed 70% and 66% mitochondrial complex I inhibition, respectively. However, MPP+

100 showed only a weak 12% inhibition and the negatively charged DiSBAC2(3) showed very little or no complex I inhibition. Rotenone is a well-studied complex I inhibitor and it is known to bind to a hydrophobic site in NADH: ubiquinone oxidoreductase and inhibit ubiquinone reduction activity of the complex (80). It is known that MPP+ also binds to the rotenone binding site to inhibit complex I (264). Therefore, we believe that cyanines may also bind to the NADH: ubiquinone reductase, most likely to the rotenone binding site, thereby inhibiting complex I.

However, further studies are certainly necessary to determine the exact mechanism for inhibition of complex I by these cyanines. The mitochondrial electron transport chain (ETC) (complexes I-

V) is the major site of ATP production via chemiosmotic phosphorylation in eukaryotes. Therefore, inhibition of mitochondrial complex I disrupt the ETC and reduce ATP production. As expected, both rotenone and cyanine treatment reduce ATP levels in the cells by inhibiting the mitochondrial complex I (Fig. 14).

Cyanines, MPP+ and rotenone cause concentration dependent, excessive ROS production specifically in dopaminergic cells (Figs 15), presumably as a consequence of the mitochondrial depolarization, complex I inhibition resulting in ATP depletion. In addition, it is observed that cyanine induced ROS production is initiated in the mitochondria (Fig 16). As expected, negatively charged DiSBAC2(3) does not depolarize mitochondrial membrane, inhibit complex I or increase the production of ROS. The proposal that excessive ROS production causes dopaminergic cell toxicity is strongly supported by the findings that the antioxidant Asc, reduced

ROS production in MN9D cells (Fig. 17) and effectively protected cells from cyanines, rotenone and MPP+ toxicity (Fig. 19). The observation that cyanines, MPP+ and rotenone accumulated in the mitochondria and depolarized the mitochondrial membranes of both MN9D and HepG2 cells to a similar extent, yet excessive ROS production and toxicity are limited to MN9D cells is

101 puzzling and must be due to some key physiological differences between these cells. One obvious difference between MN9D and HepG2 cells is that only MN9D cells contain high levels of DA, which is oxidatively sensitive and known to catalyze intracellular ROS production

(265,266). The depletion of intracellular DA prior to cyanines, rotenone or MPP+ treatment reduced ROS production (Fig. 18) provides strong support for the possibility that DA may exacerbate toxin-mediated ROS production in MN9D cells. This notion is also consistent with the observation that SH-SY5Y cells which normally contain comparatively low levels of DA and other catecholamines under normal growth conditions (262) produce moderate levels of ROS and show moderate vulnerability toward cyanines, MPP+ and rotenone, as compared with MN9D cells. DA depletion partially protected MN9D cells from cyanines, MPP+ and rotenone toxic effects (Fig. 20) in parallel with a decrease in ROS production providing further supports to the proposal that toxin mediated, DA amplified, increased ROS production could be the major cause of toxicity. However, even drastic DA depletion caused the reduction of ROS production and toxicities by only about 30-40%, and suggesting that there may be other important cellular differences contributing to the increased production of ROS, specifically in dopaminergic cells, in response to cyanine, MPP+ and rotenone treatments.

Based on the reported differences of antioxidant enzyme superoxide dismutase (38), catalase, and glutathione peroxidase (GPx) levels in animal brains and livers (190,267,268), we suspected that the extreme susceptibility of MN9D cells to cyanines, MPP+ and rotenone induced excessive ROS production could be due to expression of low levels of these enzymes in dopaminergic cells in comparison to liver (HepG2) cells. In agreement with this hypothesis,

MN9D cells, which are the most sensitive to these toxins, were found to contain substantially lower levels of all three enzymes as compared with HepG2 cells (Table 5), and parallel to those

102 reported for rat and mouse brains relative to their livers (190,267,269). In addition, while SH-

SY5Y cells expressed similar low levels of SOD and catalase, GPx levels in these cells were similar to or of HepG2 cells [note that as mentioned above, SH-SY5Ycells possess partial adrenergic characteristics (262) and adrenergic cells are known to express high levels of GPx

(270)]. Therefore, overall low expression of antioxidant enzymes and the presence of high concentrations of oxidatively labile DA may contribute to the excessive ROS production in response to cyanines, MPP+ and rotenone treatment causing the death of dopaminergic MN9D cells. The modest vulnerability of SH-SY5Y cells to cyanine toxicity could be due to relatively high levels of GPx and inherent low levels of DA present in these cells, in comparison to MN9D cells, under normal growth conditions. On the other hand, HepG2 cells possess the full capacity to cope with the excessive ROS production, most likely due to the presence of high levels of antioxidant enzymes and absence of oxidatively sensitive DA, resulting in strong protection from cyanines, MPP+ and rotenone toxicities.

Finally, we have shown that the increased levels of ROS in dopaminergic cells cause apoptotic dopaminergic cell death in response to cyanines, MPP+ and rotenone treatment. We have shown that 2,2’-cyanine treatment of MN9D cells causes cell shrinkage (from morphology)

(Figs. 21A1 & A2), chromatin condensation (from DAPI) (Figs. 21A3 & A4) and DNA fragmentation (from apoptotic DNA laddering) (Fig. 21C) which are the characteristic features of apoptotic cell death. Since chromatin condensation is a characteristic feature in apoptotic cell death, the DNA labeling dye, DAPI highly accumulates in the nucleus and gives bright fluorescence (239). As shown in Fig. 21B, bright DAPI fluorescence in cyanines, MPP+ and rotenone treated dopaminergic MN9D cells indicate that the cell death due to these toxins follows the apoptosis pathway.

103

Taken together, cyanines are a family of lipophilic cationic dyes that are commonly used in industry and scientific research. They are widely used in the production of photographic films, computer storage and video recording media (208), solar cells (209), and in photochemical research (210). They are also frequently used as inhibitors of extra-neuronal noradrenaline (211), plasma membrane monoamine, and organic cation transporters (22,212), and as precursors in the development of chemotherapeutic (213) neuro-protective agents (214,215). Fluorescent cyanines are widely used in organelles and protein labeling for cellular imaging research (271,272). In spite of their wide usage, structural resemblance to MPP+ (Schemes 19 & 20) and ability to freely accumulate in the cytosol and in the mitochondria of any cell type, their neurotoxicological properties have not been studied or reported. Based on the above findings, especially cell specific toxicity profiles and the mechanistic similarities in the toxic effects in well stablished PD models for MPP+ and rotenone, with cyanine, it is tempting to speculate that they could be stronger in vivo dopaminergic toxins that follow both toxicity characteristics of

MPP+ and rotenone, thus, the in vivo toxicities of these compounds must be investigated.

104

CHAPTER VI

CONCLUSIONS

Specific uptake through the dopamine transporter (DAT) followed by inhibition of mitochondrial complex I have been accepted as the cause of the specific dopaminergic toxicity of the most commonly used experimental PD model, MPP+. However, our findings show that MPP+ accumulates in both dopaminergic MN9D and SH-SY5Y and non-dopaminergic HepG2 cells through distinct mediated pathways with differing efficiencies, but is only toxic to dopaminergic cells. These results suggest that, in addition to efficient uptake, the intrinsic vulnerability of dopaminergic cells may also contribute to their high sensitivity to MPP+ and similar toxins. To test this possibility, a series of hydrophobic cyanine derivatives were employed in a comparative study based on their unique characteristics and structural similarity to MPP+. In contrast to

MPP+, cyanines accumulated in all these cells freely and non-specifically with similar high efficiency resulting in intracellular concentrations about 1000 times more than that of MPP+, under similar conditions. However, similar to MPP+, cyanines are also specifically and highly toxic only to dopaminergic cells.

Our findings show that positively charged cyanines and MPP+ accumulate non- specifically in the negatively charged mitochondrial matrix and depolarize the mitochondrial membrane potential in both MN9D and HepG2 cells, but are only toxic to dopaminergic MN9D cells. We also found that cyanines are a novel group of mitochondrial complex I inhibitors, as effective as the best known complex I inhibitor, rotenone; but MPP+ was found to be a relatively weak complex I inhibitor.

105

Cyanines, MPP+ and rotenone are taken up non-specifically into all cells, accumulate in the mitochondria, inhibit the mitochondrial complex I and depolarize the mitochondrial membrane potential, but cause excessive ROS production only in dopaminergic cells. In addition, cell imaging experiments have revealed that cyanine induced ROS production is initiated inside the mitochondria. Furthermore, cyanine, MPP+ and rotenone were found to cause dopaminergic cell death through the apoptosis pathway. The efficient protection of MN9D cells from cyanines, MPP+ and rotenone toxicity by the antioxidant, ascorbate, strongly suggests that the excessive ROS production specifically in dopaminergic cells is responsible for their observed toxicities. Experiments with the tyrosine hydroxylase inhibitor, -methyl-p-tyrosine which inhibits dopamine biosynthesis, showed that DA plays at least a partial role in the excessive production of ROS, specifically in dopaminergic MN9D cells. In addition, we found that dopaminergic MN9D and SH-SY5Y cells contain low levels of antioxidant enzymes in comparison to other cells which are responsible for the detoxification of ROS and protection of cells from oxidative damage. Taken together, low levels of antioxidant enzymes together with the presence of high levels of oxidatively sensitive DA in dopaminergic cells are likely to cause of excessive ROS production, specifically in dopaminergic cells in comparison to HepG2 cells upon cyanines, MPP+ and rotenone treatments. Thus, we propose that the dopaminergic cells are specifically and inherently vulnerable to mitochondrial toxins such as cyanines-, MPP+- and rotenone-induced oxidative stress in comparison to other cell types, regardless of the mechanism of cellular uptake. Based on the parallel toxicity profiles and mechanisms of cyanines, rotenone and MPP+, we strongly believe that specific dopaminergic toxicity of all these toxins must be due to a specific vulnerability of dopaminergic cells towards mitochondrial toxin mediated increased oxidative stress.

106

To determine whether the lipophilic cationic character and active accumulation in the mitochondria is vital for the specific dopaminergic toxicity of the above cyanines, a series of similar uptake and toxicity experiments were carried out with a lipophilic anionic dye. These experiments showed that the negatively charged dye also accumulated transporter independently and non-specifically in all cells with efficiency similar to cationic cyanines. However, in contrast to cationic cyanines, the anionic derivative did not show significant mitochondrial accumulation, inhibition of mitochondrial complex I, depolarization of mitochondrial membrane potential, excessive ROS production or apoptotic dopaminergic cell death. These findings strongly support the proposal that the lipophilic cation character and the effective mitochondrial accumulation is primarily responsible for the specific dopaminergic toxicity of these cyanines.

Finally, cyanines are a family of lipophilic cationic dyes that are commonly used in industry and scientific research. In spite of their wide usage and the structural resemblance to

MPP+ as well as their ability to freely accumulate in the cytosol and in the mitochondria of any cell type, their neurotoxicological properties have not been studied or reported to our knowledge.

Based on the above findings, especially, the cell specific-toxicity profiles and the mechanistic similarities of the toxin to MPP+, rotenone and cyanines, it is tempting to speculate that they could be stronger in vivo dopaminergic toxins with both characteristics of MPP+ and rotenone.

Thus, we propose that lipophilic cationic cyanines could be a novel group of environmental neurotoxins that contribute to the etiology of PD and their in vivo toxicities must be investigated.

107

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