ROLE of Ca and SODIUM-CALCIUM EXCHANGER

Home , Kowal (surname)

ROLE OF Ca2+ AND SODIUM-CALCIUM EXCHANGER (NCX) ON CELLULAR / MITOCHONDRIAL UPTAKE AND TOXICITY OF THE PARKINSONIAN TOXIN, MPP+

A Dissertation by

Mapa Mudiyanselage Sumudu Tharangani Mapa

Master of Science, Wichita State University, 2016

Master of Philosophy, University of Kelaniya, Sri Lanka, 2009

Bachelor of Science, University of Kelaniya, Sri Lanka, 2007

Submitted to the Department of Chemistry 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 2016

ROLE OF Ca2+ AND SODIUM-CALCIUM EXCHANGER (NCX) ON THE CELLULAR / MITOCHONDRIAL UPTAKE AND TOXICITY OF PARKINSONIAN TOXIN, MPP+

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

______Dennis Livesay, Dean

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DEDICATION

To my loving Parents & Husband

ACKNOWLEDGEMENTS

I would like to offer my sincerest gratitude to my supervisor Prof. K. Wimalasena, for his guidance and support throughout my research work in the graduate school. He encouraged me to develop my independent and critical thinking and research skills and greatly assisted me with scientific writing. I am grateful for the opportunity to work under his supervision and without his enthusiasm and persistent help this dissertation would not have been possible.

I am also very grateful 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. My deepest gratitude goes to Dr. Shyamali

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

My sincere thanks go to Dr. Yamuna Kollalpitiya, Dr. Viet Le, and Dr. Chamila

Kadigamuwa for their help and support in my research works. I would like to thank my dad, as well as my brothers for their encouragement during this period. Specially, I want to express my loving gratitude to my husband, Chamila for his endless support, patience and love during this time of endeavor and throughout my life.

Finally I would like to thank to the faculty and staff of the WSU Chemistry department and friends for assisting me in numerous ways.

ABSTRACT

Although the exact cause(s) of dopaminergic cell death in Parkinson's disease (PD) is not fully understood, the discovery that 1-methyl-4-phenylpyridinium (MPP+) selectively destroys dopaminergic neurons and causes PD symptoms in humans and other mammals has strengthened the environmental hypothesis of PD. The current model for the toxicity of MPP+ is centered on its specific uptake into dopaminergic cells through the dopamine transporter (DAT), electrogenic accumulation into the mitochondria, inhibition of the mitochondrial complex I, leading to ATP depletion, increased reactive oxygen species (ROS) production, and apoptotic cell death.

However, MPP+ is taken up into many cell types through a number of other transporters, challenging the major hypothesis of this model, that the specific dopaminergic toxicity of MPP+ is due to the specific uptake through DAT. In order to address this discrepancy, we have characterized a series of 4'-substituted MPP+ derivatives with varying hydrophilicities. The photophysical studies of these derivatives show that 4'I-MPP+ is fluorescent and can be used to localize the intracellular distribution of MPP+. Using these novel probes, we have shown that both cellular and mitochondrial uptake of MPP+ are modulated by the cell and mitochondrial membrane potentials and found that Ca2+ play a key role in these MPP+ uptakes and toxicities.

Furthermore, the effects of Ca2+ on MPP+ uptake and toxicity are mediated through mitochondrial and plasma membrane sodium calcium-exchangers (NCX) and the mitochondrial permeability transition pore (PTP). For the first time, we show that the specific mitochondrial

NCX (mNCX) inhibitor, CGP37157 inhibits mitochondrial uptake of MPP+ and protects dopaminergic MN9D cells from MPP+ toxicity. Thus, these findings could be further exploited for development of pharmacological agents to protect central nervous system (CNS) dopaminergic neurons from PD-causing environmental toxins.

TABLE OF CONTENTS

Chapter Page

1 INTRODUCTION ...... 1

2 BACKGROUND AND SIGNIFICANCE ...... 5

2.1 Neurologic Disorders ...... 5 2.2 Neurodegenerative Diseases ...... 5 2.3 Parkinson's Disease (PD)...... 6 2.3.1 Genetic Factors of PD...... 7 2.3.2 Environmental Factors of PD ...... 8 2.3.2.1 MPTP/MPP+ Model ...... 8 2.3.2.2 Rotenone ...... 11 2.4 MPP+ Uptake Mechanism(s) and Transporters . . . . 12 2.4.1 Dopamine Transporter (DAT)...... 12 2.4.2 Norepinephrine Transporter (NET) . . . . . 13 2.4.3 Organic Cation Transporter (OCT) . . . . . 14 2.4.4 Plasma Monoamine Transporter (PMAT) . . . . 14 2.4.5 Vesicular Monoamine Transporter (VMAT) . . . . 15 2.5 Ca2+ Homeostasis ...... 16 2.5.1 Intracellular Ca2+ Stores . . . . . 17 2.5.2 Voltage Gated Ca2+ Channels (VGCC) . . . 19 2.5.3 Ligand Gated Ca2+ Channels (LGCC) . . . 19 2.5.4 Na+/Ca2+ Exchanger (NCX) . . . . . 20 2.5.5 Mitochondrial Ca2+ Uniporter (MCU). . . . 22 2.5.6 Mitochondrial Permeability Transition Pore (PTP) . . 22 2.5.7 Transient Receptor Potential (TRP) Channels. . . 23 2.5.8 Calcium Channel Inhibitors . . . . . 23 2.6 Mechanism(s) of MPP+ Toxicity ...... 24 2.6.1 Mitochondrial Membrane Depolarization . . . . 25 2.6.2 Mitochondrial Complex I Inhibition . . . . . 25 2.6.3 Reactive Oxygen Species (ROS) Production . . . . 26 2.6.4 Apoptosis ...... 28 2.7 Cell Culture ...... 29 2.7.1 MN9D Cells ...... 30 2.7.2 SH-SY5Y Cells ...... 30 2.7.3 HEK293 Cells ...... 31 2.7.4 HepG2 Cells ...... 31

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TABLE OF CONTENTS (continued)

Chapter Page

3 EXPERIMENTAL METHODS ...... 32

3.1 Materials ...... 32 3.2 Instrumentation ...... 33 3.3 Methods ...... 34 3.3.1 Synthesis of 3'Hydroxy MPP+ Derivative and 4'Halogenated MPP+ Derivatives...... 34 3.3.2 Cell Culture ...... 35 3.3.3 Differentiation of MN9D Cells . . . . . 35 3.3.4 Measurement of Cell Viability . . . . . 35 3.3.5 Measurement of Cellular Uptake of MPP+ or MPP+ Derivatives . 36 3.3.6 MPP+ or MPP+ Derivative Uptake Into MN9D Mitochondria . 37 3.3.7 Measurement of the Mitochondrial Membrane Potential . . 38 3.3.8 Mitochondrial Localization of 4'I-MPP+ . . . . 38 3.3.9 Effect of FCCP and 85 mM K+ Ions on the Cellular Uptake of MPP+ and MPP+ Derivatives ...... 38 3.3.10 Measurement of the Mitochondrial Ca2+ Level . . . 39 3.3.11 NCX Transfection ...... 40 3.3.11.1 Determination of the NCX Expression Levels in Transfected Cells Using Western Blot . . . 41 3.3.12 Rat Brain Mitochondria Isolation . . . . . 42 3.3.13 Mitochondrial Complex I inhibition Studies . . . . 42 3.3.14 Measurement of Intracellular ATP Levels . . . . 43 3.3.15 Measurement of Reactive Oxygen Species (ROS) . . . 43 3.3.16 Chromatin Condensation Test for Apoptosis . . . . 44 3.3.17 Protein Determination ...... 44 3.3.18 Data Analyses ...... 45 3.3.19 Technical Statement ...... 45

4 RESULTS ...... 46

4.1 Photophysical Properties of MPP+ Derivatives and Intracellular Accumulation of 4'I-MPP+ ...... 46 4.2 Relative Dopaminergic Cell Toxicities of 4'-Substituted MPP+ Derivatives . 49 4.3 Characteristics of the Cellular Uptake of 4'-Substituted MPP+ Derivatives . 52 4.4 Mitochondrial Uptake of 4'-Substituted MPP+ Derivatives . . . 55 4.5 Intracellular 4'I-MPP+ Localized into the Mitochondria . . . 57 4.6 Membrane Potentials Drive Active Cellular and Mitochondrial Accumulation of MPP+ and its 4'-Substituted Derivatives . . . 58

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TABLE OF CONTENTS (continued)

Chapter Page

4.7 The Effect of Extracellular Ca2+ on Cellular Uptake of MPP+ . . 61 4.8 The Effect of Ca2+ on Mitochondrial Accumulation of MPP+ . . 63 4.8.1 The Effect of Extra-Mitochondrial (Cytosolic) Ca2+ and Na+ on Mitochondrial Accumulation of MPP+ . . . . 64 4.8.2 The Effect of Ca2+ Channel Blockers on Mitochondrial Accumulation of MPP+ ...... 65 4.8.2.1 The Effect of CGP37157 and KBR7943 on Mitochondrial Membrane Potential With 4'I-MPP+ Uptake . 67 4.8.2.2 CGP37157 Protect MN9D Cells From MPP+ and 4'I-MPP+ Toxicity ...... 69 4.9 Does mNCX Play a Role in Mitochondrial Accumulation and Toxicity of MPP+? ...... 69 4.9.1 NCX Transfection into HEK293 Cells Increases 4'I-MPP+ Uptake and Toxicity ...... 70 4.9.2 NCX Transfection into HEK293 Cells Depolarizes Mitochondrial Membrane Potential by 4'I-MPP+ Uptake . . . . 72 4.10 The Effect of Mitochondrial PTP Inhibitor on MPP+ Cellular Uptake . 72 4.11 The Effect of MPP+ on Mitochondrial Complex I . . . . 74 4.12 The Effect of MPP+ on ATP Depletion . . . . . 76 4.13 MPP+ and Its 4'-Substituted Derivatives Increase Intracellular ROS Production Specially in MN9D Cells . . . . . 77 4.14 MPP+ Cause Specific Apoptotic MN9D Cell Death . . . . 79

5 DISCUSSION ...... 81

6 CONCLUSIONS ...... 89

7 REFERENCES ...... 91

LIST OF TABLES

Table Page

1 Major Types of Neurological Disorders . . . . . 5

2 Ca2+ Channel Inhibitors and Functions . . . . . 24

3 Comparison of Relative Antioxidant Enzyme Levels in Various Cell Types ...... 28

4 Retention Times For MPP+ Derivatives . . . . . 46

5 UV-Vis Analysis of MPP+ Derivatives . . . . . 47

+ 6 IC50 Values of MPP Derivatives in MN9D Cells . . . . 52

7 FCCP and 85 mM K+ Percent Inhibition of MPP+ Derivatives in the Presence and Absence of Extracellular Ca2+ in MN9D Cells . . . 59

LIST OF FIGURES

Figure Page

1 Proposed Mechanisms of MPTP/MPP+ Uptake and Toxicity . . 11

2 Calcium Uptake and Extrusion Mechanisms in Mitochondria . . 18

3 Hypothetical Model of NCX ...... 21

4 UV-Vis Spectra of 4'I-MPP+ (5 M) in PBS, Methanol, and Acetonitrile . 47

5 Excitation and Emission Spectra of 4'I-MPP+ (5M) in PBS, Methanol, and Acetonitrile ...... 48

6 Light and Fluorescence Images of 4'I-MPP+ Treated Differentiated MN9D Cells ...... 48

7 MPP+ Derivatives are Specifically Toxic to Dopaminergic Cells Similar to MPP+ ...... 50

8 Uptake of MPP+ and 4'-Substituted MPP+ Derivatives into Cells . . 54

9 Mitochondrial Uptake of 4'-Substituted MPP+ Derivatives . . . 56

10 Intracellular Localization of Cytosolic 4'I-MPP+ . . . . 58

11 The Effect of FCCP and 85 mM K+ on MPP+ and 4'-Substituted MPP+ Derivative Uptake into MN9D and HepG2 Cells . . . . 60

12 The Effect of Extracellular Ca2+ on MPP+ and 4'I-MPP+ Uptake . . 62

13 The Effect of 4'I-MPP+ on the Mitochondrial Ca2+ Levels in Live MN9D Cells ...... 63

14 The Effect of Extra-mitochondrial Ca2+, Na+ or Li+ on MN9D Mitochondrial Uptake of 4'I-MPP+ ...... 64

15 The Effect of Ca2+ Channel Blockers on Mitochondrial Accumulation of MPP+ ...... 66

16 The Effect of CGP37157 and KBR7943 on the 4'I-MPP+-Mediated Mitochondrial Depolarization of HepG2 Cells . . . . 68

17 The Effect of CGP37157 on MN9D Toxicities of MPP+ and 4'I-MPP+ . 69

LIST OF FIGURES (continued)

Figure Page

18 NCX3 Transfection into HEK293 ...... 70

19 The Effect of NCX in 4'I-MPP+ Uptake and Toxicity . . . 71

20 Mitochondrial Membrane Potential Depolarization by 4'I-MPP+ in NCX/NCLX Transfected HEK293 Cells . . . . . 72

21 The Effect of Mitochondrial PTP Opening Inhibitor, Cyclosporine A on MPP+ and 4'I-MPP+ Uptake into MN9D Cells . . . . 73

22 Both MPP+ and 4'I-MPP+ Inhibit the Rat Brain Mitochondrial Complex I . 75

23 The Effect of MPP+ and 4'I-MPP+ on Intracellular ATP Levels of MN9D and HepG2 Cells ...... 76

24 The Effect of MPP+ and 4'-substituted MPP+ Derivatives on the Intracellular ROS Levels ...... 78

25 MPP+ or 4'-Substituted MPP+ Derivatives Cause Apoptotic MN9D Cell Death ...... 79

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LIST OF SCHEMES

Scheme Page

1 Synthesis of MPPP and MPTP ...... 9

2 Conversion of MPTP to MPP+ ...... 10

3 Structure of Rotenone ...... 12

4 The Electron Transport Chain Mediated Production of ROS . . . 27

5 Reaction Catalyzed by SOD (Equation 1) and Catalase (Equation 2). . 28

6 Structure of MPP+ and MPP+ Derivatives . . . . . 34

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LIST OF ABBREVIATIONS

Abbreviation Systematic Name

2APB 2-Aminoethoxydiphenylborane

6-OHDA 6-Hydroxydopamine oC Celsius

ΔF Fluorescence Change

ΔΨm Mitochondrial Membrane Potential

μg Microgram

μL Microliter

μm Micrometer

μM Micromolar

A Absorbance

AA Amino Acid

AD Alzheimer’s Disease

ALS Amyotrophic lateral sclerosis

ANT Adenosine Nucleotide Translocator

ATP Adenosine Triphosphate

BSA Bovine Serum Albumin

CBD Ca2+ binding domains

CICR “Ca2+”-Induced Ca2+ Release

CNS Central Nervous System

DA Dopamine

Da Dalton

DAPI 4, 6-Diamidino-2-phenylindole

DAT Dopamine Transporter

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DBS Deep Brain Stimulation

DCF 2΄, 7΄-dichlorofluorescin

DCFH-DA 2΄, 7΄-dichlorofluorescin diacetate

DMEM Dulbecco’s Modified Eagle’s Medium

DMF N,N-Dimethylformamide

DMSO Dimethyl Sulfoxide

DNA Deoxyribonucleic 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

FCCP Carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone

FBS Fetal Bovine Serum h Hour

H2O2 Hydrogen Peroxide

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

HPLC High-Performance Liquid Chromatography

HPLC-UV HPLC with Ultraviolet

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

HRP Horseradish Peroxidase

HVA High Voltage-Active

IC50 Half-Maximal Inhibitory Concentration

IP3 Inositol 1,4,5-Triphosphate

Km Michaelis Constant kb Kilobase kDa Kilodalton

KRB Krebs-Ringer Buffer

LB Lewy Bodies

L-DOPA L-3,4-Dihydroxyphenylalanine

LGCC Ligand Gated Calcium Channel

LVA Low Voltage-Active

MAO Monoamine Oxidase

MCU Mitochondrial Ca2+ uniporter mg Milligram min Minutes mL Milliliter mm Millimeter mM Millimolar mNCX Mitochondrial Na+/Ca2+ Exchanger

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

MS Mass Spectrometry

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

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NADH Nicotinamide Adenine Dinucleotide (reduced form)

NCLX Na+ or Li+ dependent Ca2+ exchanger

NCX Na+/Ca2+ Exchanger

NE Norepinephrine

NET Norepinephrine transporter ng Nanogram nM Nanomole nmole Nanomole nm Nanometer

NMR Nuclear Magnetic Resonance

NO• Nitric Oxide

• NO2 Nitrogen Dioxide

N2O3 Dinitrogen Trioxide

N2O4 Dinitrogen tetroxide

L-DOPA L-dihydroxyphenylalanine

•- O2 Superoxide

OCT Organic Cation Transporter

OH• Hydroxyl radical

ONOO- Peroxinitrite

PBS Phosphate Buffered Saline

PD Parkinson’s Disease

PDF Parkinson’s Disease Foundation

PMAT Plasma Monoamine Transporter

PTP Permeability Transition Pore

RNS Reactive Nitrogen Species

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ROI Region Of Interest

ROS Reactive Oxygen Species

SD Standard Deviation

SDS Sodium Dodecyl Sulfate sec Seconds

SOCE Store Operated Ca2+ Entry

SOD Superoxide Dismutase

SR Sarcoplasmic Reticulum

TBE Tris-Borate-EGTA

TBS Tris-Buffered Saline

TTBS Mixture of Tris-Buffered Saline and Tween 20%

TMRM Tetramethylrhodamine, methyl ester

TRP Transient Receptor Potential

UV Ultraviolet

UV-Vis Ultraviolet-Visible v Volume

VGCC Voltage-gated Calcium Channel

VMAT Vesicular Monoamine Transporter

Vmax Maximum Velocity

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CHAPTER I

INTRODUCTION

Parkinson’s, Alzheimer’s, Huntington’s and Amyotrophic lateral sclerosis (ALS) are neurodegenerative diseases which are characterized by progressive degeneration of the neurons in specific areas of the central or peripheral nervous systems. Among them, Parkinson's disease

(PD) is the second most common neurodegenerative disease and is caused by the loss of dopaminergic neurons in the substantia nigra, a specific region in the midbrain (1,2). Early stages of PD are associated with visible symptoms like shaking, muscular rigidity and slow movements.

This chronic and progressive disorder is commonly observed in the mid to late ages of life (2).

There are approximately 7-10 million people worldwide suffering from PD and about 60,000 new cases per year are diagnosed in the United States alone, according to the Parkinson's disease

Foundation (PDF) (3). The combined direct and indirect costs of PD treatment, social security payments due to loss of work and reduced productivity of patients is nearly $25 billion in the

United States, and is expected to increase significantly over the next few decades due to the anticipated increase in the aged population (3). The exact cause(s) of dopaminergic cell death in

PD is not fully understood, and no permanent treatment or cure for PD is currently available.

Therefore, understanding of the molecular causes of, and development of preventive and therapeutic strategies for PD is an immediate and urgent challenge for researchers in the field.

Increasing evidence suggests that PD is caused by genetic or environmental factors, or by a combination of both. While 5-10% of PD is known to be associated with genetic factors, the rest appears to be sporadic, and could be associated with environmental factors. In the genetic form of the disease, mutations have been identified in several genes, including parkin (PARK2),

PINK1 (PARK6), DJ-1 (PARK7) and LRRK2 (4). Similarly, a variety of environmental factors

1 and toxins such as permethrin, beta-hexachlorocyclohexane (β-HCH), paraquat, maneb, and rotenone have been shown to cause PD like symptoms in experimental models (5,6). The accidental discovery that exposure to the synthetic chemical, 1-methyl-4-phenyl-1,2,3,6- tetrahydropyridine (MPTP), causes PD symptoms in humans, other primates, and rodents has strengthened the hypothesis that environmental factors cause PD (7-9). Lipophilic MPTP, crosses the blood brain barrier and undergoes oxidation by monoamine oxidase-B (MAO-B) in glial cells to produce the fully aromatized terminal toxin, 1-methyl-4-phenylpyridinium (MPP+)

(10). Numerous in vivo as well as in vitro studies have shown that the toxic metabolite MPP+, rather than MPTP, selectively destroys dopaminergic neurons (10,11). Therefore, MPTP/MPP+ has been extensively used as a convenient model to study the mechanisms of specific dopaminergic cell death in PD (7-9). The current model for the dopaminergic toxicity of MPP+ is centered on several key steps including specific uptake into dopaminergic cells through the plasma membrane dopamine transporter (DAT), electrogenic accumulation of cytosolic MPP+ in the mitochondria, inhibition of the mitochondrial electron transport chain (ETC) complex I, which leads to intracellular ATP depletion, increased reactive oxygen species (ROS) production and apoptotic cell death (12-17).

Although many aspects of the above model are widely accepted, recent studies have challenged the proposal that the specific in vivo toxicity of MPP+ towards dopaminergic cells is due to the specific uptake through DAT (7,12), since MPP+ is taken up not only into dopaminergic cells, but also into other cells through diverse transporters, including organic cation transporters (OCT) and non-specific plasma membrane amine transporters (PMAT), etc.

(18-20). On the other hand, most in vivo as well as in vitro studies to date have unequivocally confirmed that MPP+ is specifically and highly toxic to dopaminergic cells in comparison to

2 other cell types. Clearly, some critical aspects of the mechanism of specific dopaminergic toxicity of MPP+ are still poorly understood. Since MPP+ is the most widely utilized model for the study of environmental contributions to the pathophysiology of PD (8), a better understanding of the mechanism of specific dopaminergic toxicity of MPP+ at the molecular level is of prime importance. Accordingly, the availability of molecular probes with structural and toxicological characteristics similar to that of MPP+, including specific dopaminergic toxicity, could provide additional information on the cellular distribution, mitochondrial accumulation and key cellular factors associated with these processes in live cells will certainly help in further advancement of the understanding of the mechanism of MPP+ dopaminergic toxicity at the molecular level (21,22).

We have synthesized and characterized a series of 4'-substituted MPP+ derivatives which were used to study details of cellular and mitochondrial uptake and toxicity of MPP+, and to identify various cellular constituents associated with these processes. We also characterized 4'I-

MPP+ as a fluorescent MPP+ mimic with unique photophysical properties, that could conveniently be used as a probe to map the cellular localization of MPP+ in live cells. In the present study, we have used those new probes with different cell models: dopaminergic human

SH-SY5Y, mouse MN9D, and non-dopaminergic HepG2 and HEK293 cells, to identify various proteins and factors responsible for the specific cellular and mitochondrial uptake and toxicity of

MPP+. Here we show that both cellular and mitochondrial uptake of MPP+ are modulated by the plasma and mitochondrial membrane potentials. In addition, we found that the extracellular, cytosolic, and mitochondrial Ca2+ play a key role in both cellular and mitochondrial uptake and toxicity of MPP+. Furthermore, the effects of Ca2+ on the MPP+ uptake and toxicities are mediated through the mitochondrial and plasma membrane sodium calcium-exchangers (NCX)

3 and mitochondrial permeability transition pore (PTP). For the first time, we found that the specific mitochondrial NCX inhibitor, CGP37157 (23,24), inhibits mitochondrial uptake of

MPP+ and protects dopaminergic MN9D cells from MPP+ toxicity. Thus, these findings could be exploited for the development of pharmacological agents to protect the dopaminergic system from PD-causing environmental toxins.

CHAPTER II

BACKGROUND AND SIGNIFICANCE

2.1 Neurologic Disorders

Neurological disorders or nervous system diseases are disorders that are associated with the brain, spinal cord or the peripheral nervous system. According to available information in the

U.S. National Library of Medicine, there are more than 600 neurological disorders, and the major types are listed below (25).

Table 1. Major Types of Neurological Disorders

Type Example Neurogenetic Diseases (Diseases caused Huntington’s Disease, Muscular Dystrophy by faulty genes) Developmental Disorders Cerebral Palsy, Spina Bifida Degenerative Diseases (Nerve cells get Alzheimer’s Disease, Parkinson’s Disease damaged or die) Metabolic Diseases Gaucher’s Disease Cerebrovascular Diseases (Diseases of the Stroke blood vessels that supply the brain) Trauma Spinal Cord, Brain Injury Convulsive Disorders (Seizure disorders) Epilepsy Infectious Diseases Meningitis

2.2 Neurodegenerative Diseases

Neurodegenerative diseases occur when nerve cells get damaged or die due to medical factors such as tumors, stroke, genetic factors or other environmental factors such as toxins.

Most neurodegenerative diseases are irreversible, progressive, and their causes are complex and not fully understood. The most common neurodegenerative diseases are Alzheimer's disease

(AD), Parkinson's disease (PD), Amyotrophic lateral sclerosis (ALS), Huntington's disease,

Friedreich's ataxiz, and Lewy body disease. These diseases are associated with common symptoms such as dementia, loss of memory, difficulty in motor control and movement, difficulty to talk or breathe, and malfunctions of the heart (26). These diseases can be life threatening depending on the type, and do not have a permanent cure; but treatments are available to suppress disease symptoms in most cases.

2.3 Parkinson’s Disease (PD)

PD is the second most common neurodegenerative disease after AD (15). This disease is more common in adults over the age of 50. PD symptoms initially show as rigidity of the arms, legs and trunk, rapid shaking of hands, arms and legs, slowness of movement, as well as impaired balance and coordination. The symptoms gradually get worse and finally patients may have trouble walking, talking, or doing simple tasks (27). They may also suffer from depression, sleep problems, or have trouble chewing or swallowing. PD is hard to diagnose in the early stages, because of the slow progression of the disease and lack of suitable diagnostic lab tests.

PD was first identified by James Parkinson and the symptoms were reported in “An

Essay on the Shaking Palsy” in 1817 (28). The degradation of dopaminergic neurons and accumulation of cytoplasmic Lewy bodies (LB) in the substantia nigra pars compacta area of the brain are the key pathophysiologcal features of PD (29). The loss of dopaminergic neurons leads to a decrease of dopamine (DA) in the striatum causing impairments in both voluntary (30) and involuntary (31) movements and in cognition (32). Cytoplasmic LB found in PD patients' brains are insoluble protein aggregates which consist of insoluble fibrils mainly composed of α- synuclein, a small 14 kDa protein, predominately expressed in neurons of the central nervous system (33), and small amounts of ubiquitin and β-crystallin (34,35). Although, researchers have

6 identified these two as the key pathophysiological features associated with PD, the exact cause of

PD is not fully understood. Physicians normally use medical and neurological history and a physical examination to diagnose PD (35). However, the diagnosis is complicated by the fact that most neurodegenerative diseases share common symptoms. Blood tests and brain imaging are currently used to identify late-stage PD (36). To suppress PD symptoms, the precursor of DA, L-

3,4-dihydroxyphenylalanine (L-DOPA) is commonly used, since DA itself is incapable of crossing the blood brain barrier to increase the amount of DA in the brain, which helps to control

PD symptoms (37,38). In severe cases, artificial deep brain stimulation (DBS) by implanted electrodes is used as a last resort for PD symptom treatment (39).

2.3.1 Genetic Factors of PD

A majority of PD patients do not directly inherit the disease, but it is known that there is a 4-9% higher chance of getting PD from first degree relatives compared to the general population. Approximately 15% of PD patients have a family history related to the disease (40).

In the inherited form of PD, genetic mutations were found in the LRRK2 (PARK8), Parkin

(PARK2), DJ-1 (PARK7), PINK1 (PARK6), ATP13A2 (PARK9) or SNCA (PARK1) genes (4).

LRRK2 gene mutations cause the late-onset of autosomal-dominant PD and show the pathology of nigral degeneration without LB (41,42). Parkin mutations are the most frequent genetic causes of PD and show pathology in the third or fourth decade of the patients' life (4). Parkin, DJ-1 and

PINK1 mutations are also tightly linked with PD and show the pathological evidence of neuronal loss and gliosis (43). However, it is not fully understood how the genetic factors cause PD and influence the risk of developing the disease.

2.3.2 Environmental Factors of PD

A broad range of environmental factors including pesticides, herbicides, fungicides, industrial toxins and other environmental toxins are known to contribute to the etiology of PD

(5,6). In addition, other minor factors such as being overweight, agricultural work, living in industrial or urban areas, exposure to heavy metals and hydrocarbon solvents, etc. are known risk factors for PD (44). Although, none of these factors are directly involved in developing the disease, acting together they may increase the risk of developing the disease. The oxidatively labile structural analog of DA, 6-hydroxydopamine (6-OHDA), was the first synthetic compound identified as a neurotoxin, which stimulate PD-like symptoms in animal models (45). Highly polar 6-OHDA is unable to cross the blood brain barrier and therefore, 6-OHDA has to be directly injected into the brain to observe its neurotoxic effects. Once inside the brain, it can specifically enter catecholaminergic cells through either the dopamine transporter (DAT) or norepinephrine transporter (NET) (8).

2.3.2.1 MPTP/MPP+ Model

1,3-Dimethyl-4-phenyl-4-propionoxypiperidine (MPPP), which is an analog of demerol, was first synthesized in 1947 by Dr. Albert Ziering as a synthetic opioid, but never released to the market as a developed drug. In the 1970s, Barry Kidston, who was a chemistry graduate student at the University of Maryland, synthesized MPPP as a street drug, using Dr. Ziering's protocol and injected it into himself (46). After a few days, Kidson exhibited symptoms similar to PD, and died later from a cocaine overdose. Autopsy confirmed that he was suffering from

PD. In 1982, six people in California were diagnosed with PD after having used MPPP. The

8 neurologist, J. William Langston, in collaboration with NIH, found that MPTP contamination in synthetic MPPP was the cause of PD (47).

In the synthesis of MPPP, 1-methyl-4-piperidone (MP), is treated with phenyl lithium to produce the intermediate product, 4-hydroxyl-4-phenyl-N-methylpiperidine (HPMP, Scheme 1).

In the second step, this product is converted to MPPP in the presence of propionic anhydride and sulfuric acid. This step is temperature- and acid-sensitive, therefore, it can produce the propionic acid eliminated product, MPTP at higher temperatures (48). This may have been the origin of the

MPTP contamination in Barry Kidson's MPPP preparation.

Scheme 1. Synthesis of MPPP and MPTP

Following the original discovery, MPTP was extensively used to model the molecular mechanisms associated with the specific degeneration of CNS dopaminergic neurons in PD.

Numerous studies showed that the high hydrophobicity of MPTP allows it to cross the blood brain barrier and enter into the brain cells (49). Within the brain, MPTP is converted to the fully aromatized product, MPP+, first by monoamine oxidase B (MAO-B) catalyzed oxidation to 1- methyl-4-phenyl-1,2-dihydropyridinium (MPDP+) followed by spontaneous disproportionation in glial cells (50). Further studies have unequivocally established that the MAO-B oxidized product, MPP+ is the dopaminergic toxin, not the parent compound, MPTP (Scheme 2) (10).

Scheme 2. Conversion of MPTP to MPP+

MPP+ produced in glial cells by MAO-B, is proposed to exit via the organic cation transporter-3 (OCT3) and that taken up specifically into dopaminergic cells via DAT (51). In dopaminergic cells, MPP+ is shown to accumulate in the mitochondria and inhibit mitochondrial

ETC complex I (52), leading to a decrease in ATP and an increase in ROS production, thereby causing apoptotic cell death (Figure 1) (13,37,50). In addition, MPP+ is proposed to compete with cytosolic DA for synaptic vesicle entry via the vascular monoamine transporter-2 (VMAT-

2), increasing cytosolic DA and leading to overproduction of ROS in the cytosol, further amplifying oxidative stress and cell death (Figure 1) (13,50,53).

Figure 1. Proposed Mechanisms of MPTP/MPP+ Uptake and Toxicity (50).

2.3.2.2 Rotenone

Rotenone is widely used as an insecticide, pesticide, or as a piscicide. Rotenone is known to effectively inhibit NADH:ubiquinone oxidoreductase of mitochondrial complex I, causing the inhibition of ATP and increase in ROS production (54). In addition, increased ROS could interact with intracellular nitric oxide (NO), superoxide and hydroxyl radicals to form peroxinitrite, further amplifying the oxidative stress (55). Overproduction of ROS can damage cellular DNA and protein, leading to cell death. In addition, rotenone is relatively lipophilic and freely passes through the blood brain barrier to enter the brain, similar to MPTP. In the brain, it accumulates in the mitochondria of cells and effectively inhibits mitochondrial complex I.

Rotenone is also known to cause dopaminergic neuron degeneration resulting in PD-like symptoms including rigidity, tremor and bradykinesia (56). It is also proposed to cause aggregation of α-synuclein in the cytoplasm, further amplifying PD symptoms (56).

Scheme 3. Structure of Rotenone.

2.4 MPP+ Uptake Mechanism(s) and Transporters

As mentioned above, the well-established mechanism involved in specific in vivo toxicity of MPP+ to dopaminergic neurons is due to its specific uptake through DAT (7,12). Scientists believed dopaminergic toxicity of MPP+ was due to this uptake mechanism, however, recent studies have challenged this proposed mechanism, since MPP+ is taken up not only by dopaminergic cells, but also by other cells via several transporters including NET, OCT and the non-specific plasma monoamine transporter, (PMAT), among others (18,19). In addition to these transporters, malfunction of calcium and sodium channels are also related to PD due to the perturbation of Ca2+ and Na+ concentrations in the cells (57-60).

2.4.1 Dopamine Transporter (DAT)

DAT is a Na+/Cl--dependent symporter that is specifically expressed on the plasma membrane of dopaminergic nerve terminals (61). It regulates the DA concentration in the synaptic cleft area by taking up the released DA into the presynaptic neurons. DA transport

12 through DAT is coupled with the cotransport of Na+ and Cl- with a stoichiometry of DA: Na+: Cl- of 1: 2:1 (62).

DAT is a transmembrane protein composing 620 amino acid residues (AA) consisting of

12 transmembrane helixes with both amino and carboxyl termini residing on the cytoplasmic face and a large extracellular loop between the third and fourth helixes (63,64). During DA transport through the DAT, Na+ initially binds to the extracellular domain, and then DA binds to stabilize the intermediate state of the DAT protein. In this state, disruption of hydrogen bonds between N82 and N353 and redistribution of salt bridges occurs and leads to a large conformational change in the protein resulting in the release of protein bound DA, Na+, and Cl- into the cytosol of the cell (65). The thermodynamic driving force for the active reuptake of DA is the ion concentration gradient generated by plasma membrane Na+/K+ ATPase. Previous reports suggest that the DAT transporter is directly involved in cellular uptake of the

+ + neurotoxins, 6-OHDA and MPP . The Km values for dopamine and MPP are 1.2 ± 0.2 μM and

3.7 ± 0.3 μM for DAT in human neuroblastoma cells, respectively (66). The best known inhibitor for DAT is the recreational drug, cocaine. However, GBR 12909 is extensively used as a specific and potent DAT inhibitor in pharmacological studies (67). DAT is sensitive to intracellular and extracellular Na+ and Cl- ions, and under certain conditions it functions in reverse mode.

2.4.2 Norepinephrine Transporter (NET)

Similar to DAT, NET is also a Na+/Cl--dependent symporter, which reuptakes extracellular norepinephrine (NE) and DA (68). NET is an important transporter present in presynaptic NE neurons to maintain the concentration of NE in the synaptic cleft area (68).

Similar to DAT, NET consists of 12 transmembrane domains composing 617 AA (69). NET

13 takes up 90% of released NE into the cell with 2 Na+ ions and 1 Cl- ion per a molecule of NE.

The ion gradients generated by Na+/K+ ATPase provides the thermodynamic driving force for effective re-uptake of NE. Desipramine is a commonly used specific inhibitor for NET (70).

2.4.3 Organic Cation Transporter (OCT)

OCT is an electrogenic Na+-independent bi-directional cation transporter which belongs to the large family of solute carriers with a structural similarity to DAT and NET. OCT plays a critical role in the uptake, distribution, and elimination of organic cations, including many drugs and toxins (71). In addition, it facilitates the excretion and distribution of endogenous organic cations such as choline, creatinine and cationic neurotransmitters (72). Since OCT transports positively charged substances, the energy for transport is derived from the concentration gradient of the substrate along with the membrane potential of the cell. A number of OCT sub-types including OCT1, OCT2, OCT3 and OCT6 are expressed in many different tissues (73,74). OCT1 and OCT2 are mostly expressed in the small intestine, liver and kidney (20,75,76), whereas

OCT3 is expressed throughout the body, including the brain and heart (20,72). OCT1, OCT2 and

OCT3 isoforms are known to transport both MPP+ and DA (72). Substrate binding site structures of OCT isoforms are responsible for the specificity of these proteins. The OCTs present in small intestine, liver and kidney operate together with cation/proton antiporters during cation uptake and efflux (76).

2.4.4 Plasma Monoamine Transporter (PMAT)

PMAT is an integral membrane protein composed of 530 AA, containing 11 transmembrane helixes with an extracellular face carboxyl terminus, cytoplasmic face amino terminus and a large intracellular loop between helixes VI-VII (77). Therefore, PMAT is not

14 homologous to other known monoamine transporters. This Na+-independent reversible cation transporter predominantly transports monoamine neurotransmitters like DA, NE and serotonin as well as adenosine from the corresponding synaptic areas into presynaptic neurons (77,78).

PMAT also transports drugs, toxins and other molecules with a net positive charge at physiological pH (20). Therefore, PMAT is abundantly expressed in the brain, heart, skeletal muscles and kidneys (79). PMAT is a low-affinity (high Km) high capacity (high Vmax)

+ + membrane transporter for catecholamines and cationic toxins such as MPP (Kms for MPP , DA and serotonin for PMAT are 33, 114 and 329 μM, respectively) (77).

2.4.5 Vesicular Monoamine Transporter (VMAT)

VMAT transports monoamine neurotransmitters through the monoaminergic neuron synaptic vesicles membranes. This ~70 kDa molecular weight integral membrane protein is composed of 12 transmembrane helices with both N- and C- termini facing the cytosolic side of the membrane (80,81). VMAT utilizes an intergranular proton gradient generated by V-ATPase in the membrane to import monoamine molecules from the cytosol into the vesicles against a concentration gradient (80). Two different isoforms of VMAT, VMAT-1 and VMAT-2, are expressed in different tissues. VMAT-1 is predominantly expressed in large, dense core vesicles of the peripheral nervous system and the medulla of the adrenal glands (82). VMAT-2 is prevalent in monoaminergic cells of the CNS, within the chromaffin granules of the adrenal medulla, β-cells of the pancreas and blood platelets (80,82,83). MPP+ is a good substrate for both

VMAT isoforms and it is a relatively less specific substrate for DAT or NET (13,81,84).

2.5 Ca2+ Homeostasis

All mammalian cells maintain relatively low levels of cytosolic Ca2+ (200-500 nM) under normal physiological conditions (85). Neuronal cells, which transmit information through electrical and chemical signals throughout the body, use Ca2+ ions as intracellular messengers

(86). They maintain a large Ca2+ gradient across the plasma membrane utilizing plasma membrane transporters (Ca2+-ATPase) and specific Ca2+ pumps present in Ca2+ storage sites such as the endoplasmic reticulum (ER) and mitochondria (87). ER and mitochondrial Ca2+ storage plays a critical role in the regulation of cytosolic Ca2+ levels. The major transporters responsible for Ca2+ ion movements through the plasma membrane are voltage-gated calcium channels

(VGCC) and ligand-gated calcium channels (LGCC) (87-89). In addition, plasma and mitochondrial membrane Na+/Ca2+ exchangers (NCX), mitochondrial Ca2+ uniporter (MCU), and transient receptor potential (TRP) channels are also involved in the maintenance of intracellular

Ca2+ levels (87,89).

Intracellular free Ca2+ concentration shows complex fluctuations during cell functions.

During trans-membrane signaling events, Ca2+ influx from the extracellular space is increased, while the Ca2+ efflux from cytosol and release of Ca2+ from intracellular Ca2+ stores are decreased, resulting in the elevation of intracellular Ca2+ concentrations to μM levels (88,89). On the other hand, intracellular Ca2+ concentrations are decreased via ATP-driven Ca2+ transport to the extracellular space and into the intracellular stores during cell relaxation. However, perturbation of Ca2+ homeostasis for a long period of time may disrupt cellular function and induce cell death (90,91). Although the exact mechanism of cell death due to the disruption of

Ca2+ homeostasis is not fully understood, some studies suggest that there is a strong correlation between perturbation of Ca2+ homeostasis and apoptotic cell death. Similarly, previous studies

16 suggest that intracellular Ca2+ may play a role in cellular (92,93) and mitochondrial accumulation of MPP+ (94).

2.5.1 Intracellular Ca2+ Stores

The ER and mitochondria serve as specific stores for intracellular Ca2+. Both organelles maintain their Ca2+ concentration at mM levels and facilitate the maintenance of cytosolic Ca2+ at μM levels (88,89). These organelles transport Ca2+ ions through specific Ca2+ transporters, some of which may also be present in the plasma membrane.

The ER, one of the largest intracellular organelles in the cell, mainly stores and releases

Ca2+ upon the activation of specific receptors. Two pathways mediate the release of ER Ca2+ into the cytosol; “storage-operated Ca2+ entry” (SOCE) (95,96) and “Ca2+-induced Ca2+ release”

(CICR) (97). While Ca2+ release through SOCE is stimulated by Ca2+ and inositol 1, 4, 5-

2+ 2+ triphosphate (IP3) together, Ca release through CICR is initiated by Ca and the ryanodine receptor.

The mitochondria have the ability to take-up Ca2+ ions from the cytosol through two specific high (nM) and low (M) affinity transporters. MCU involves rapid, massive Ca2+ entry into mitochondria (89). Other than MCU, NCX, Na+- or Li+- dependent Ca2+ exchanger,

(NCLX), Ca2+/H+ antiporter, etc, are present in the inner and outer membranes of mitochondria

(89,98)

Figure 2. Calcium Uptake and Extrusion Mechanisms in Mitochondria (98).

Mitochondria help to buffer the increase in intracellular Ca2+ during transmembrane signaling (98,99). Under these conditions, increased mitochondrial Ca2+ levels increase the cellular energy supply by activation of several tricarboxylic acid cycle or Krebs cycle enzymes, including pyruvate dehydrogenase, oxoglutarate dehydrogenase, among others. However, under extreme conditions, uncontrolled increase of intra-mitochondrial Ca2+ triggers mitochondrial permeability transition pore (PTP) opening, leading to mitochondrial swelling, cytochrome c release and eventually apoptotic cell death (91,100,101). Mitochondrial Ca2+ buffering mechanisms are closely associated with the spatial arrangement of mitochondria inside the cells.

For, example mitochondria are located in the vicinity of VGCC in the plasma membrane in excitable cells, to capture Ca2+ entering through these channels effectively, to prevent the increase in cytosolic Ca2+ (102).

2.5.2 Voltage Gated Ca2+ Channels (VGCC)

The influx of cations like Na+ or efflux of anions like Cl- depolarizes the plasma membrane of excitable cells resulting in the opening of the VGCC. The open VGCC facilitate spontaneous Ca2+ influx into the cell to initiate the transmembrane signaling cascades (102,103).

Therefore, VGCC are the primary mediators of depolarization-induced Ca2+ entry into neuronal cells, which serve as second messengers of electrical signaling initiators for many different cellular activities (103). High levels of VGCC are expressed not only in neuronal cells, but also in the membranes of many other excitable cells, such as in cardiac and smooth muscle cells, endocrine cells, etc. (104).

VGCC are structurally complex and are composed of several subunits, and are divided into two major classes: high voltage active (HVA) and low voltage active (LVA), based on the voltage required for their activation (104). They are further classified into L, N, P/Q, R and T- types based on their AA sequences and pharmacological properties. The L, N and P/Q-type channels are HVA channels, R-type is an intermediate voltage activated channel and T-type belongs to the LVA channels family. The L-type Ca2+ channels are found in muscle cells, neuronal cells and cardiac cells; N, P/Q and R-type Ca2+ channels are primarily present in neuronal cells, and T-type Ca2+ channels are present mainly in cardiac cells.

2.5.3 Ligand Gated Ca2+ Channels (LGCC)

LGCC are heteromeric proteins possessing at least two different domains: a transmembrane domain for the ion pore and an extracellular domain for ligand binding. As the name implies, LGCC are primarily regulated by allosteric mechanisms (105). Due to the presence of complex heteromeric structures, LGCC are divided into a wide range of subtypes

19 with diverse functional properties. For example, IP3 and the ryanodine receptors are present in the ER, sarcoplasmic reticulum (SR), while SOCE are present in the plasma membrane (95,96).

2.5.4 Na+/Ca2+ Exchanger (NCX)

The major Ca2+ ion exporter through the plasma membrane is NCX. This bidirectional transporter predominantly exports one Ca2+ ion for the uptake of three Na+ ions (106). This unique coupling of Ca2+ export against a large concentration gradient to the import of Na+ provides the thermodynamic driving force for the process. In the reverse mode, NCX transports three Na+ ions out against the electrochemical gradient across the membrane in exchange for the counter-transport of one Ca2+ ion into the cytosol (107). NCX is a voltage sensitive, electrogenic transporter with low affinity and high capacity for Ca2+ ions. The modes of NCX operation, forward or reverse depend on the relative cellular concentrations of Ca2+ and Na+ ions (106,108).

Therefore, NCX plays a critical role in the Ca2+ homeostasis of the cell, for proper physiological functioning.

Four closely related isoforms of NCX, NCX1, NCX2, NCX3 and NCX4 have been characterized. All these proteins show ~70% sequence homology with no striking functional differences (108). These are expressed in the plasma, ER, mitochondrial and nuclear membranes.

While NCX1 is widely expressed in many different cells, expression of NCX2 and NCX3 is limited to brain and skeletal muscle cells (109). A working model of NCX shows the presence of four domains, including two Ca2+ binding domains (CBD), a transmembrane and an intracellular domain (Figure 3) (106). Both CBDs are exposed to the cytosol of the cell and form a compact

β-sandwich using two antiparallel β-sheets with a large unstructured loop. The transmembrane domain is made out of 10 α-helices and embedded in the plasma membrane providing a path to

20 exchange Ca2+ and Na+ ions, through the membrane. The core of the transmembrane domain contains four ion binding sites (3 for Na+ and 1 for Ca2+) in a diamond shape (Figure 3). The intracellular domain connects the transmembrane domain with the CBDs.

Figure 3. Hypothetical Model of NCX (106).

The NCX structure and function are closely related to each other (106). When intracellular Ca2+ concentration increases, Ca2+ ions bind to the CBD1 and increase the rigidity and the electrostatic potential of the binding sites and induce a conformational change that triggers the transformation of NCX into an activated state, allowing it to translocate ions across the membrane. Both CBD1 and CBD2 undergo conformational changes upon binding of Ca2+ and the structures of Ca2+ bound forms are similar. However, these two domains show dramatic structural differences in the Ca2+ unbound (absence of Ca2+) forms, suggesting that CBDs regulate the activity of NCX.

2.5.5 Mitochondrial Ca2+ Uniporter (MCU)

The inner mitochondrial membrane protein, MCU, facilitates the entry of Ca2+ ions into the mitochondria from the cytosol. Ca2+ ion flow via the uniporter depends on the inner mitochondrial membrane potential and the relative concentrations of Ca2+ ions in the cytosol and mitochondrial matrices (110,111). The uniporter ion transportation allows to the electrochemical gradient maintenance across the membrane without direct involvement with ATP hydrolysis and other ion exchanges (110). The free Ca2+ concentration within the mitochondrial matrix is balanced by Ca2+ influx through the MCU and Ca2+ efflux through the NCX (112,113).

2.5.6 Mitochondrial Permeability Transition Pore (PTP)

The opening of mitochondrial PTP contributes to cellular injury by releasing cytochrome c into the cytosol to initiate the apoptotic pathway (100,101). Opening of PTP leads to a transition in the inner mitochondrial membrane permeability, leading to loss of mitochondrial solutes up to 1.5 kDa. Several factors can induce the opening of PTP. High mitochondrial matrix

Ca2+ levels, mitochondrial membrane potential depolarization, increased oxidative stress, and increased inorganic phosphate concentration by the depletion of ATP concentration, are some of the well known factors that can affect the opening of PTP (100,114). Open PTP allows Ca2+ to escape from the mitochondria, triggering increased ROS production. In addition, cyclosporin A,

Mg2+, ADP, adenosine nucleotide translocator (ANT) and 2-Aminoethoxydiphenylborane (2-

APB) are inhibitors of the opening of the PTP (100,114,115).

2.5.7 Transient Receptor Potential (TRP) Channels

TRP channels constitute a large and functionally versatile family of cation conducting channel proteins. They transport both monovalent and divalent cations, non-selectively through the plasma membranes of numerous excitable and non-excitable cells (116,117). So for, more than 50 TRP channels have been identified in different species and 28 of them are found in mammalian cells (117). Mammalian TRP channels are classified into 6 subfamilies: TRPC

(canonical), TRPV (vanilloid), TRPM (melastatin), TRPA (ankyrin), TRPML (mucolipin) and

TRPP (polycystin), according to the homology of the proteins (118). Except for a few, most TRP channels function as modulators of intracellular Ca2+ signaling, since they have a lower affinity for Ca2+ than for Na+. The non-specific Ca2+ uptake through these channels contributes to a change in the intracellular Ca2+ concentration and fine tunes membrane polarization (118). In some cases, TRP channels work together with VGCC, to further refine the efficiency of the respective transmembrane signaling pathway (118).

2.5.8 Calcium Channel Inhibitors

Commercially available, well characterized, pharmacological agents are extensively used to study the effect of various Ca2+ channels on the uptake and toxicity of various toxins such as

MPP+, rotenone, etc. The pharmacological properties and selectivity of these inhibitors are listed in Table 2.

Table 2. Ca2+ Channel Inhibitors and Functions.

Inhibitor Function Benzamil T-type VGCC inhibitor NCX inhibitor (119) TRPC3 inhibitor (120) Verapamil L-type VGCC inhibitor (121) PMAT inhibitor (20) Flufenamic Acid (FFA) TRPC3 and TRPC7 inhibitor (122) Mefenamic Acid (MFA) TRPC3 and TRPC7 inhibitor (123) Tolfenamic Acid (TFA) TRPC3 and TRPC7 inhibitor (123) Mibefradil T-type VGCC inhibitor L-type VGCC inhibitor (124) Nitrendipine L-type VGCC inhibitor (125) Nifedipine L-type VGCC inhibitor (126) Nickel T-type VGCC inhibitor (127) Gadolinium T-type VGCC inhibitor (127) CGP 37157 Specific mitochondrial NCX inhibitor (23,24) KBR7943 NCX inhibitor (128) 2-Aminoethoxydiphenylborane (2-APB) IP3 antagonist (129) SOCE inhibitor (130) TRP channel inhibitor (131) PTP inhibitor (115) Ruthenium red MCU inhibitor (110,132,133) EGTA Ca2+ chelator (134) BAPTA-AM Membrane permeable Ca2+chelator (135) Cyclosporin A PTP inhibitor (100,101)

2.6 Mechanism(s) of MPP+ Toxicity

The dopaminergic toxicity of MPP+ is associated with its active accumulation in the mitochondria. Numerous studies have demonstrated that MPP+ accumulates in the mitochondria isolated from the liver, brain and other cell types (21,94). Mitochondrial accumulation of MPP+ causes mitochondrial membrane depolarization, mitochondrial complex I inhibition (16,52), decreased ATP production (13), increased ROS production (13,136), culminating in specific dopaminergic cell death by apoptosis (137-139).

2.6.1 Mitochondrial Membrane Depolarization

The energy required for the synthesis of ATP is derived from the electrochemical gradient generated across the inner mitochondrial membrane by the electron transport chain

(ETC). The electrochemical gradient is composed of two components: mitochondrial membrane potential (∆Ψm, electrical gradient) and a proton gradient (chemical gradient). Under normal physiological conditions ∆Ψm is in the range of -150 mV (140). The magnitude of ∆Ψm may be altered significantly under the conditions where intracellular ionic concentrations are perturbed or ATP production is inhibited (141). Under extreme conditions, mitochondrial depolarization could trigger the opening of the PTP in the inner membrane of mitochondria. The opening of mitochondrial PTP allows the release of cytochrome c into the cytosol, which initiates the apoptotic cell death cascade (140).

Several commercially available fluorescent lipophilic cationic dyes, such as tetramethylrhodamine methyl (TMRM), tetramethylrhodamine ethyl (TMRE), 5,5',6,6'- tetrachloro-1,1',3,3'-tetraethylbenzimidazolylcarbocyanine iodide (JC-1) are commonly used to measure ∆Ψm (141). These dyes accumulate in the mitochondria electronically and the intra- mitochondrial dye concentration is proportional to the membrane potential. Therefore, the intensity of the fluorescence of the dye in the mitochondria measured by microscopy or flow cytometry can be used to estimate ∆Ψm.

2.6.2 Mitochondrial Complex I Inhibition

Complex I of ETC is located in the mitochondrial inner membrane. Complex I, which is referred to as NADH: ubiquinone oxidoreductase, oxidizes NADH, reduces ubiquinone, and transports protons across the inner membrane (142). Inhibition of complex I leads to the

25 reduction of ATP synthesis, overproduction of ROS and apoptotic cell death (136,143).

Numerous synthetic and natural products including rotenone, MPP+, the herbicide, paraquat, pesticides (benzimidazole, thiangazole, etc.) and some plant extracts (rotenoids, actenogenins, piericidins, etc.) are known to inhibit mitochondrial complex I (52,54,144,145). Epidemiological and biochemical studies with PD patients suggested that significantly reduced mitochondrial complex I activity is associated with PD pathogenesis (143). Therefore, exposure to mitochondrial complex I inhibitors may contribute to the etiology of PD.

2.6.3 Reactive Oxygen Species (ROS) Production

Two types of reactive free radicals are generated in living cells, i.e. reactive oxygen species (ROS; free radicals associated with oxygen) and reactive nitrogen species (RNS; free

•- radicals associated with nitrogen) (146,147). The most common ROS are superoxide (O2 ),

• • hydrogen peroxide (H2O2) and hydroxyl radical (OH ) and RNS are nitric oxide (NO ),

- • peroxinitrite (ONOO ), nitrogen dioxide ( NO2), dinitrogen trioxide (N2O3), and dinitrogen tetroxide (N2O4). These unstable, highly reactive species contain one or more unpaired electrons that cause chemical modifications of proteins, lipids, DNA and other biological molecules (148-

151). An imbalance between the production of intracellular free radical species and the capacity of cellular antioxidant defense mechanisms lead to oxidative stress. In humans, increased oxidative stress is closely associated with the development of neurodegenerative diseases

(152,153), cancer (154), cardiovascular diseases (155), sickle cell disease, depression, etc.

(151,156).

Many biological processes contribute to production of ROS under normal physiological conditions. The redox reactions of the mitochondrial ETC are major contributors to intracellular

ROS. In addition, activities of vital enzymes such as NADH oxidases, cytochrome P450, exposure to external hazardous conditions, including exposure to UV, heat and toxins, aging, smoking, and inflammation may also be responsible for the excessive ROS production (157,158). The mitochondrial toxins that interfere with the ETC could produce, partially reduced oxygen species

•- O2 leading to the production of other reactive oxygen species (Scheme 4) (159). As mentioned above, overproduction of ROS in the mitochondrial matrix could lead to mitochondrial dysfunction and apoptotic and/or necrotic cell death.

Scheme 4. The Electron Transport Chain Mediated Production of ROS.

Almost all cells have their own defense mechanisms and antioxidant systems to protect cellular components from ROS. The cellular antioxidant systems can be divided into two main categories: enzymatic and non-enzymatic or endogenous and exogenous (151). The most powerful and efficient endogenous enzymatic antioxidant systems are superoxide dismutase

(SOD), catalase, and glutathione peroxidase; and non-enzymatic endogenous systems are glutathione, alpha lipoic acid and coenzyme Q10. The exogenous antioxidants are ascorbic acid, vitamin E, carotenoids, etc., which are normally obtained from the diet or supplements. Copper- zinc SOD (Cu/Zn SOD) in the cytoplasm, mitochondrial manganese SOD (Mn SOD) and

•- extracellular SOD catalyze the conversion of O2 into H2O2 and O2 (Equation 1 in Scheme 5) and the widely distributed catalase, catalyzes the conversion of H2O2 to H2O and O2 (Equation 2 in

Scheme 5).

(1)

(2)

Scheme 5. Reactions Catalyzed by SOD (Equation 1) and Catalase (Equation 2).

Previous studies suggested that the antioxidant enzyme levels in the brain are low in comparison to the liver (160). In agreement with these reports, we previously reported that dopaminergic neuronal cell lines (MN9D and SH-SY5Y) contain less antioxidant enzymes relative to a liver (HepG2) cell line (Table 3) (161).

Table 3. Comparison of Relative Antioxidant Enzyme Levels in Various Cell Types

Cell Source SOD Catalase Cu/Zn SOD Mn SOD Liver 100 ± 7 100 ± 12 100 ± 9 Brain 37 ± 4 47 ± 7 3 ± 0 HepG2 100 ± 7 100 ± 7 MN9D 47 ± 2 16 ± 3 SH-SY5Y 35 ± 4 26 ± 1

2.6.4 Apoptosis

Apoptosis is a form of irreversible, energy-dependent programmed cell death that occurs in multicellular organisms. Mitochondrial membrane depolarization, complex I inhibition, ATP depletion and over-production of ROS are known to initiate the apoptotic cell death cascade.

Caspase-3 is a frequent mediator of apoptosis and is known to be implicated in PD (162-164).

Characteristically, apoptosis occurs in a single isolated cell without damaging the neighboring cells. In apoptosis mediated cell death, initially the cell shrinks and condenses, then the cytoskeleton collapses, the nuclear membrane disassembles and nuclear DNA fragmentation

28 occurs, leading to cell death (165,166). Therefore, apoptotic cell death is usually diagnosed by observing characteristic morphological changes such as blebbing, cell shrinkage, nuclear fragmentation, chromatin condensation, chromosomal DNA fragmentation and mRNA decay

(162). Apoptosis is highly regulated, tidy, and a controlled process that is important during development and aging.

Apoptosis is experimentally identified by observing the characteristic morphological and biochemical changes of the cells. The morphological changes can be directly observed by light or electron microscopy and DNA fragmentation is identified by TUNEL assay (167) or characterized by DNA electrophoresis (166). Chromatin condensation during apoptosis can be identified using the membrane permeable dye, 4,6-diamidino-2-phenylindole (DAPI), which binds to condensed DNA to give an intense blue fluorescence (Ex/Em 358/461 nm). DAPI dye enters preferentially into dying cells due to the mis-regulation of membrane permeability, resulting in stronger fluorescence in comparison to normal, healthy cells (161,167).

Measurement of intracellular caspase activity using colorimetric/fluorimetric assays or protein visualization by Western blots with specific antibodies, are commonly used in the biochemical assessment of apoptosis (167).

2.7 Cell Culture

Cell culture models are widely used in studies of cell physiology, in vitro toxicity tests, drug transport, in monitoring protein expression, and vaccine production (168). Cells can be removed from a primary tissue, a cell line or from a cell strain by enzymatic, mechanical, or chemical disaggregation (169). Cells obtained from a tissue or tumor and placed in culture are known as primary cultures. Most primary cell cultures have a limited lifespan, except when the

29 cells are derived from tumor or tumor tissues according by, primary cultures can be sub-cultured and passed down into a cell line. Culture conditions vary widely depending on the cell type.

Substrates with essential nutrients (glucose), growth factors, hormones and regulated physico- chemical environment (pH, temperature, gas concentration, and osmotic pressure) are maintained, as different culture conditions. Mammalian cells are usually cultured at 37 ºC in a sterile atmosphere of 5% CO2 and humidified air.

2.7.1 MN9D Cells

MN9D is an immortalized, dopaminergic cell line that is derived from the fusion of mouse embryonic ventral mesencephalic cells, rostral mesencephalictegmentum (RMT) with the

N18TG2 neuroblastoma cells (170). MN9D cells are extensively used as a model of dopamine neurons which resemble the rat embryonic dopaminergic primary cultures. They synthesize, store, and release catecholamines, DA and 3,4-dihydroxyphenylalanine, and express high levels of DAT, NET and VMAT-2 transporters (171). MN9D cells are also highly sensitive to MPP+ similar to rat embryonic dopaminergic primary cultures, suggesting that this cell line a good cell model to study the mechanistic aspects of the selective dopaminergic cell toxicity of MPP+ and related PD causing neurotoxins.

2.7.2 SH-SY5Y Cells

The human neuroblastoma cell line, SH-SY5Y, is a thrice cloned subline of SK-N-SK cells. The SK-N-SK parent cell line was isolated from a bone marrow biopsy, which was first subcloned as SH-SY, which was again subcloned as SH-SY5, and finally, subcloned as the SH-

SY5Y cell line (172). Cultures of SH-SY5Y cells produce viable adherent and floating cells with two distinct morphological phenotypes, tyrosine hydroxylase and dopamine β hydroxylase

30 containing neuroblast like catecholaminergic, and epithelial like counterpart cells (173,174).

Differentiated SH-SY5Y cells more resemble mature dopaminergic neurons, in comparison to the undifferentiated cells (175). Similar to MN9D cells, SH-SY5Y cells are also widely used as an in-vitro cell model in neuro-toxicological studies (176).

2.7.3 HEK293 Cells

The human embryonic kidney cell line, HEK293, is widely used as a convenient expression system in cell biology and biotechnology research due to its reliable growth and high efficiency of transfection with foreign DNA (177). HEK293 cells normally give high transfection efficiency with various transfection techniques, such as the calcium phosphate method, lipid based method, microinjection, etc. Commonly, HEK293 transient expression system is used for the structure-activity studies of various channel and transporter proteins, mechanisms of the cellular transport of various drugs and nutrients and the analysis of a nuclear export signal, etc. (178,179).

2.7.4 HepG2 Cells

HepG2 cell line is derived from a human liver carcinoma cells and is a good model to study the toxicities of various pharmacological agents and industrial chemicals. An adherent, epithelial like, HepG2 cells are easy to grow in large scale under laboratory conditions. Since

HepG2 are derived from liver tissue and does not express the characteristic catecholamines, they are often used as a non-neuronal control cell model in neuro-toxicological studies (180).

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. Glass bottomed plates were purchased from MatTek Corporation (Ashland, MA, USA).

Fetal bovine serum (FBS) was purchased from Valley Biomedical (Winchester, VA, USA).

Protein assay reagents, bovine serum albumin (BSA) standards, HRP-conjugated secondary antibodies and HRP color developing kits were purchased from Bio-Rad (Hercules, CA, USA).

Ultra-pure 10X TBE buffer, Lipofectamine 2000 reagent 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). MitoTracker Green FM and GeneJET plasmid miniprep kit was purchased from Life Technologies (Carlsbad, LA, USA).

Tetramethylrhodamine, methyl ester (TMRM) was purchased from AnaSpec (Fremont, CA,

USA). The mouse hybridoma cell line MN9D (170) was graciously provided by Dr. Alfred

Heller, University of Chicago, IL. Human heptacellular liver carcinoma cell line (HepG2) and human embryogenic kidney 293 cells (HEK293) (181) were a kind gift from Dr. Tom Wiese,

Fort Hays State University, KS. Human neuroblastoma SH-SY5Y cell lines were purchased from

ATCC (Manassas, VA, USA). Fresh rat brains for mitochondrial isolations were kindly provided by Dr. Li Yao, Wichita State University, KS. NCX1 and NCLX were obtained from Dr. Jonathan

Lytton, University of Calgary (Canada); NCX2, NCX3 and NCX3 antibody were obtained from

Dr. Michela Ottolia, Cedars-Sinai Heart Institute, LA. All the solutions were prepared in Milli Q- deionized water (Millipore, Billerica, MA, USA). Stock solutions of 1-methyl-4-

32 phenylpyridinium (MPP+) derivatives, Rotenone, Cyclosporine A, 4′, 6-Diamidino-2- phenylindole, dilactate (DAPI), 2΄, 7΄-dichlorofluorescin diacetate (DCFH-DA), and TMRM were prepared in 100% dimethyl sulfoxide (DMSO). In all experiments, the final DMSO concentration was kept to a minimum, usually < 0.05% v/v. Krebs-Ringer Buffer-HEPES (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. 109.5 mM NaCl, 5.34 mM KCl, 0.77 mM NaH2PO4, 1.8 mM CaCl2, 0.81 mM MgSO4, 44 mM NaHCO3 and 5.55 mM dextrose are

- consisted in DMEM-HCO3 . KCl buffer for mitochondrial suspensions contain 0.12 M KCl, 50.0

μM EGTA, 20.0 mM MOPS, pH 7.4.

3.2 Instrumentation

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

(Varian Inc, Palo Alto, CA). Fluorescence emission spectra were recorded on a JobinYvon-Spex

Tau-3 spectrophotometer (USA Instruments, Inc, Aurora, OH). MPP+ or MPP+ derivative uptakes were determine by C18-reversed-phase HPLC-UV on a Spectra Systems P4000 gradient pump coupled to an LDC Analytical SM 4000 UV detector using a C18 reversed-phase column

(Supelco Inc, Bellefonte, PA). HPLC elution buffer consisted of a 70:30 ratio of 20 mM

CH3COOH, 20 mM H3PO4, 30 mM TEA, 1.7 mM 1-Octanesulfonic acid sodium salt, pH 7.0:

+ CH3CN. Flow rate was 0.7 mL/min. Mitochondrial membrane potential, cellular 4'I-MPP uptake and intracellular Ca2+ levels were measured using a Nikon ECLIPSE-Ti microscope equipped with a Nikon S FLURO 40X objective. Images of mitochondrial dyes and DAPI loaded cells were captured using a Nikon ECLIPSE-Ti inverted fluorescence microscope equipped with a

Nikon S FLURO 40X objective (Nikon Instrument Inc., Melville, NY). Intracellular ROS levels

33 in live cells were monitored using a Leica (182) confocal fluorescence microscope equipped with a 40X objective (Leica Microsystems Inc, Buffalo Grove, IL).

3.3 Methods

3.3.1 Synthesis of 3'Hydroxy MPP+ Derivative and 4'Halogenated MPP+ Derivatives

All MPP+ derivatives used in this study were synthesized in our lab by Dr. Rohan Perera or other former graduate students or postdoctoral fellows (Scheme 6).

Scheme 6. Structures of MPP+ and MPP+ Derivatives

3'OH-MPP+ derivative was synthesized by O-demethylation of the -OMe pyridinium salt followed by N-methylation of pyridine nitrogen with CH3I (183). The 4'-halogenated (4'-F, Cl and Br) compounds were synthesized using the corresponding 4'-halogenated phenyl magnesium bromide and 1-methyl-4-piperidone as previously reported (183) The 4'-iodo derivative was synthesized using 4-iodo-phenylmagnisium bromide (prepared from 1-bromo-4-iodobenzene) and N-protected pyridine precursors using the same procedure as for other 4'-halogenated MPP+ derivatives.(183) Yield 42%; mp 305 °C (decomp.); 1H NMR (d6-DMSO) δ 4.3 (s, 3H), 7.8 (d,

2H), 8.0 (d, 2H), 8.5 (d, 2H), 9.0 (d, 2H); 13C NMR (d6-DMSO) δ 47.0, 100.0, 124.0, 130.0,

138.5, 146.0; MS (ESI) m/z 295.9804 (M+); Fluorescence Ex/Em 320/430 nm.

3.3.2 Cell Culture

MN9D, HepG2, HEK293 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 g/mL streptomycin and 50 IU/mL penicillin maintained at 37 oC in a humidified atmosphere with 7% CO2. Cells were cultured to about 70-80% confluence, seeded into12 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.

3.3.3 Differentiation of MN9D Cells

Differentiation of MN9D cells were carried out with 1.0 mM sodium n-butyrate according to the published procedure of Choi et al. (170). Briefly, MN9D cells were grown in

- DMEM-HCO3 containing 10% fetal bovine serum with 1.0 mM sodium butyrate at 37 °C in a humidified atmosphere with 7% CO2 for 3 days at which time the media was replaced with a fresh 1.0 mM sodium n-butyrate containing media and incubated for an additional 3 days. The progress of differentiation was judged by observing the expected morphological changes. All differentiated cells were used in appropriate experiments after the 6th day of differentiation.

3.3.4 Measurement of Cell Viability

Cell viabilities were determined by the MTT [3-(4,5-dimethylthiazol-2-yl)-2,5- diphenyltetrazolium bromide)] assay (184). Briefly, cells were seeded onto 96-well plates and allowed to grow to about 70-80 % confluence. Prior to the experiment, culture media was

- + + replaced with DMEM-HCO3 containing desired concentrations of MPP or a MPP derivatives and incubated for 16 h (unless otherwise stated) at 37 °C. After the incubation, 10 μL of 5 mg/mL MTT solution was added to each well and incubated for 2 h at 37 °C. The resulting formazan was solubilized by the addition of 210 L detergent solution [50% DMF/H2O (v/v),

20% SDS] followed by overnight incubation at 37 °C. The cell viabilities were determined by quantifying solubilized formazan by measuring the difference in the absorbance at 570 nm and

650 nm (185). Results were expressed as % viability of MPP+ or MPP+ derivatives 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 DMSO was used the final concentration was kept to a minimum, usually <0.5% v/v.

3.3.5 Measurement of Cellular Uptake of MPP+ or MPP+ Derivatives

Cells were seeded into 12-well plates at 0.25 x 106 cells/well and grown 2-3 days to about

70-80% confluency. The media was removed and a solution containing desired concentrations of MPP+ or MPP+ derivative in warm KRB-HEPES were added to each well and incubated at 37

°C for the desired period of time. After the incubation, incubation media was removed and cells were washed three times with ice-cold KRB-HEPES. Washed cells were scraped and suspended in 1.0 mL of cold KRB-HEPES and 50 L aliquots were withdrawn for protein determination.

The remaining cell suspensions were centrifuged at 5000 g at 4 °C for 5 min, the supernatants were discarded, and the cell pellets were treated with 75 μL of 0.1 M HClO4. The coagulated proteins were pelleted by centrifugation at 13,200 g at 4 oC for 8 min and the MPP+ or MPP+ derivative contents of the supernatants were determined by C18-reversed-phase HPLC-UV with the UV detection at 288 nm for 3'OH-MPP+, 295 nm for MPP+, 297 nm for 4'F-MPP+, 302 nm for 4'Cl-MPP+, 4'Br-MPP+ and 310 nm for 4'I-MPP+, respectively. All MPP+ and MPP+

36 derivative levels were normalized to the respective protein concentration of each sample and were corrected for non-specific membrane binding by subtracting the corresponding time zero time point readings for each MPP+ and MPP+ derivative concentration (normally less than 5% of the intra-cellular concentration).

To quantify 4'I-MPP+ uptake in to MN9D and HepG2 by fluorescence, cells were grown in glass bottomed plates for 2 days, the media was removed, washed, reconstituted with a 1 mL

KRB-HEPES, pH 7.4, and mounted on the state of Nikon ECLIPSE Ti-S inverted fluorescence microscope. After the region of interest (ROI)s were selected, cells were incubated for 2 min and

4'I-MPP+ was added to a final concentration of 100 M and the intracellular fluorescence at

Ex/Em 340/470-550 nm (although the excitation maxima of 4'I-MPP+ was 320 nm, an excitation wavelength of 340 nm was used to minimize the exposure of cells to UV) was recorded as a function of time for a period of 1 h. The averages of background corrected, control subtracted

ROI fluorescence intensities were used to estimate the cellular uptake of 4'I-MPP+.

3.3.6 Uptake of MPP+ and MPP+ Derivative Into MN9D Mitochondria

Intact mitochondria were isolated according to a published protocol (186) from MN9D cells which were grown in culture plates for two days. The isolated intact mitochondria were suspended in KCl buffer and 5 μL aliquots were withdrawn for protein determination. The desired amounts of mitochondrial suspensions were incubated with the desired concentrations of

MPP+ or MPP+ derivative for 45 min at 37 °C in KCl buffer. After the incubation, incubates were centrifuged at 7,000 g at 4 °C for 10 min, the supernatants were discarded, and the mitochondrial pellets were treated with 75 μL each of 0.1 M HClO4. The coagulated proteins were removed by centrifugation at 13,200 g for 8 min at 4 oC and MPP+ or MPP+ derivative

+ contents in HClO4 extracts were quantified by HPLC-UV as detailed above. All MPP (and

MPP+ derivative) levels were normalized to the respective protein concentration of each sample.

3.3.7 Measurement of the Mitochondrial Membrane Potential

Cells grown in glass bottomed plates were treated with 50 nM TMRM in KRB-HEPES for 45 min in the dark (187). After mounting on the stage of Nikon ECLIPSE Ti-S inverted fluorescence microscope, 15-20 ROIs with uniform fluorescence were selected and TMRM fluorescence (Ex/Em at 543/573 nm) was recorded at 5 sec time intervals for 3 min. Then, the appropriate concentrations of MPP+ or MPP+ derivative were added and TMRM fluorescence measurements were continued for the desired time at 15 sec time intervals. The background fluorescence was subtracted from the ROI fluorescence and averages of corrected data were used to estimate the magnitude of the mitochondrial membrane potential.

3.3.8 Mitochondrial Localization of 4'I-MPP+

Cells were grown in glass bottomed plates and incubated with 100 M 4'I- MPP+ and 200 nM MitoTracker Green FM (Life Technologies, Grand Island NY, USA) in KRB-HEPES at 37

°C for 30 min. Then, cells were washed, reconstituted with 1 mL KRB-HEPES, pH 7.4, and fluorescence images of intracellular 4'I- MPP+ (Ex/Em 340/470-550 nm) and MitoTracker Green

FM (Ex/Em 644/665 nm) were recorded using a Nikon ECLIPSE Ti-S inverted fluorescence microscope fitted with a 40X objective.

3.3.9 Effect of FCCP and K+ on the Cellular Uptake of MPP+ and MPP+ Derivatives

Cells grown in 12-well plates were washed and incubated with 5 μM carbonyl cyanide-4-

(trifluoromethoxy)phenylhydrazone (FCCP) in KRB-HEPES, no Ca2+-HEPES or EGTA-HEPES

38 for 30 min followed by 50 μM MPP+ or MPP+ derivative in the same buffer for 45 min at 37 °C.

To test the effect of K+, cells were incubated with 50 μM MPP+ or 4'I-MPP+ in the 85 mM K+-

HEPES buffer for 45 min at 37 °C. After the incubations, cells were washed with the same ice-

+ + cold HEPES buffer and treated with 0.1 M HClO4. The MPP or MPP derivative contents of

HClO4 extracts were quantified by C18 reversed phase HPLC-UV as detailed above. The controls were treated under the same conditions except that FCCP was omitted from the initial incubation. All uptake data were normalized to respective protein concentrations of each sample.

To visualize the effect of FCCP on the mitochondrial accumulation of 4'I-MPP+, cells were grown in glass bottomed plates and incubated with 5 M FCCP in KRB-HEPES for 30 min followed by 100 μM 4'I-MPP+ for 1 h at 37 °C. Then cells were washed and reconstituted with 1 mL of KRB-HEPES and the images of intracellular 4'I-MPP+ fluorescence (Ex/Em 340/470-550 nm) were recorded.

3.3.10 Measurement of the Mitochondrial Ca2+ Level

Cells grown in glass bottomed plates were treated with 5 μM Rhod-2-AM (Biotium,

Hayward, CA, USA) in KRB-HEPES, pH 7.4 for 1 h at room temperature in the dark. After incubation, cells were washed and reconstituted with KRB-HEPES and mounted on the stage of a Nikon ECLIPSE Ti-S inverted fluorescence microscope, 15-20 ROIs with uniform fluorescence were selected and Rhod-2-AM fluorescence (Ex/Em at 543/573 nm) was recorded at 15 sec time intervals for 2 min. Then, an appropriate amount of 100 μM 4'I-MPP+ was added and Rhod-2-AM fluorescence measurements were continued for an additional 58 min at 2 min time intervals. The background fluorescence was subtracted from the ROI fluorescence and averages of corrected data were used to estimate the mitochondrial Ca2+ levels.

3.3.11 NCX Transfection

NCX1, NCX2, NCX3 and NCLX plasmids obtained from Dr. Jonathan Lytton

(University of Calgary, Canada) and Dr. Michela Ottolia (Cedars-Sinai Heart Institute, LA,

USA) were amplified using competent E. Coli cells (kindly provided by Dr. Moriah Beck). In brief, 10 μL samples of cDNA were added to E. Coli and incubated for 20 min on ice. After incubation, samples were transferred to a 42 ºC water bath and incubated for 1 min followed by 5 min on ice. Then 1 mL Luria-Bertani (LB) medium was added to the samples and incubated at

37 ºC for 1 h. The transformed cells were selected by growing on the 100 μg/mL ampicillin antibiotic containing agar plates overnight at 37 ºC. Next day, single colonies were selected and grown in an antibiotic containing LB medium. Plasmid DNA was isolated using GeneJET plasmid miniprep kit (Life Technologies, Carlsbad, LA, USA) and quantified using 1% agarose gel electrophoresis with Quick-Load 1 kb DNA ladder (New England Biolabs Inc., Ipswich, MA,

USA). Extracted NCX plasmid DNA was transfected into HEK293 cells using Lipofectamine

2000, following the manufacturer's instructions. In brief, DNA was added to Opti-MEM media and incubated at room temperature for 30 min. Then diluted DNA was combined with diluted

Lipofectamine 2000 and further stabilized for 30 min at room temperature. The

DNA/Lipofectamine 2000 mixture was then added on to the HEK293 cells and incubated at 37

- ºC for 6 h. After the incubation, the mixture was replaced with fresh DMEM-HCO3 media for 24

- h followed by DMEM-HCO3 media containing the appropriate antibiotic (500 μg/mL ampicillin or neomycine) at 37 °C in a humidified atmosphere with 7% CO2 for 7 days. The selective media was changed every two days and all transfected cells were used in the appropriate experiments after the 7th day of transfection.

3.3.11.1 Determination of the NCX Expression Levels in Transfected Cells Using Western

Blot

NCX3 transfected HEK293 cells were grown in multi-well plates to 70-80% confluence under standard growth conditions. Cells were washed with cold phosphate-buffered saline (PBS), harvested, and centrifuged at 6,000 g at 4 °C for 5 min. The pellets were solubilized with 50 mM

Tris, 150 mM NaCl, 1% Triton X-100, and 2.5% protein inhibitor cocktail (Sigma), pH 7.5, for

1 h at 4 oC. After the solubilization, the samples were centrifuged for 10 min at 1200 g and the supernatants were used for the Western blotting experiments (see below). The protein contents of the cell preparations were determined by the method of Bradford.

Solubilized proteins (100 µg) were boiled in loading buffer (10X Bromophenol Blue loading dye with β-merchaptoethanol) with Cleland's reagent (DDT) for 5 min and subjected to

8.5% SDS gel electrophoresis under standard conditions. Protein bands were transferred onto 0.2

μm PVDF membranes (Bio-Rad, Hercules, CA) using standard protocols. After blocking the protein binding sites of PVDF membranes with 5% nonfat dried milk in Tris Buffered Saline

(TBS) containing 1% Tween 20, they were incubated overnight with rabbit polyclonal primary anti-NCX3 antibody (1: 1000; generously provided by Dr. Michela Ottolia, Cedars-Sinai Heart

Institute, LA, USA) with β-actin polyclonal antibody (1: 1000; Proteintech, Rosemont, IL, USA) in a primary antibody solution [20 mM Tris, 500 mM NaCl, 0.05% Tween-20, pH 7.5 (TTBS) with 10% nonfat dried milk]. Hybridized membranes were washed with TTBS, and incubated with the appropriate HRP-conjugated secondary antibodies (1:5000) in a hybridizing buffer

(TBS, 1% Tween-20, 1% nonfat dried milk) for 2 h. After washing the membranes, the protein bands were visualized using the HRP color development reagent kit following the manufacturer's instructions. The intensities of bands were quantified using a Gel Logic 100 imaging system.

3.3.12 Rat Brain Mitochondria Isolation

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

(188). Male Sprague Dawley rat brains (12 month old) were kindly provided by Professor Li Yao at Department of Biology, Wichita State University. After sacrificing, the brains were quickly removed and immersed in an ice-cold isolation buffer [225 mM mannitol, 75 mM sucrose, 5 mM

HEPES and 1 mg/mL fatty acid free BSA (pH 7.4, isotonized with KOH)]. The rat brains were weighed and cut into small pieces and homogenized using a Dounce-type glass homogenizer in the isolation buffer. The homogenates were distributed in polypropylene Falcon tubes and then centrifuged at 600 g for 10 min. The supernatants were removed and the pellets were suspended in 10 mL of isolation buffer and centrifuged to recover the fast-sedimenting mitochondria. The supernatants were pooled and centrifuged in four tubes at 10,000 g for 10 min. The pellets were suspended in the same buffer and centrifuged at 7,000 g for 10 min. The upper white fluffy layers containing myelin, synaptosomes and lipids were discarded and the brown mitochondrial pellets were collected and suspended in 10 mL of the same buffer and centrifuged at 12,000 g for

10 min. The resulting mitochondrial pellets were reconstituted with 150 μL of assay buffer [25 mM potassium phosphate (pH 7.2), 5 mM MgCl2] and stored in pre-cooled Eppendorf tubes.

3.3.13 Mitochondrial Complex I Inhibition Studies

Isolated intact mitochondria were lysed by freeze-thawing in hypotonic media containing

25 mM K3PO4 and 5 mM MgCl2, pH 7.2 (189). The complex I (i.e. NADH: ubiquinone oxidoreductase) activity of mitochondrial membrane fragments were determined according to the procedure of Birch-Machin & Turnbull (190). Briefly, mitochondrial membranes (50 μg protein) were incubated with 10 mM NADH and antimycine A (2 μg/mL) in a 1.0 mL assay solution

42 containing 25 mM K3PO4, 5 mM MgCl2, 2 mM KCN, 2.5 mg/mL bovine serum albumin, pH 7.2 for 2 min at 37 oC with or without rotenone, MPP+ or 4'I-MPP+. The complex I mediated ubiquinone dependent NADH oxidation reaction was initiated by adding ubiquinone1 (65 μM final concentration) and the initial rates of the reactions were monitored by following the decrease in absorbance at 340 nm with respect to 425 nm reference wavelength for 5 min.

3.3.14 Measurement of Intracellular ATP Levels

Cells grown in 12-well plates were treated with 100 μM MPP+ or 4'I-MPP+ in KRB-

HEPES for 6 h at 37 °C. After the treatments, cells were collected in 1 mL of KRB-HEPES and

50 μL samples were withdrawn for protein quantification. The remaining cell suspensions were centrifuged for 3 min at 3,500 g and the cell pellets were treated with 1 mL of lysis buffer (0.1 M

Tris, 1 % Triton X-100, pH 7.5) for 2 min and centrifuged at 3,500 g for 3 min to remove the cell debris. The ATP content of the supernatants were measured using a fluorometric ATP assay kit

(BioVision, Milpitas CA, USA) following the manufacturer's guidelines. All ATP levels were normalized to the protein content of each sample.

3.3.15 Measurement of Reactive Oxygen Species (ROS)

Cells grown in 12-well plates were incubated with 10 M DCFH-DA in KRB-HEPES for 1 h. DCFH-DA-loaded cells were washed with ice-cold KRB-HEPES and treated with 100

μM MPP+ or MPP+ derivatives in the same buffer for 1 h at 37 oC. After the incubation, cells were washed, harvested, solubilized with 0.1 M Tris buffer (pH 7.5) containing 1% Triton X-

100, and cell debris were removed by centrifugation at 13,200 g for 8 min at 4 oC. The content of ROS-oxidized DCFH product, 2΄,7΄-dichlorofluorescein, in the supernatants were quantified

43 by fluorescence [Ex/Em 504/525 nm](182). The data were normalized to the protein content of individual samples.

To visualize the intracellular ROS production by fluorescence microscopy, MN9D cells grown in glass bottomed plates were treated with 50 M DCFH-DA in KRB-HEPES for 1 h.

DCFH-DA loaded cells were washed with the same buffer and treated with 250 M 4'I-MPP+ in

KRB-HEPES for 4 h at 37 °C and washed with KRB-HEPES and images were captured by a

Leica TCS SP5II confocal fluorescence microscopy using appropriate instrumental settings

(Ex/Em 488/524 nm).

3.3.16 Chromatin Condensation Test for Apoptosis.

MN9D cells grown in glass bottomed plates were treated with 250 μM MPP+ or MPP+ derivative for 12 h and then with 300 nM DAPI in KRB-HEPES for 15 min in the dark (191).

Then cells were rinsed and PBS was added to cover the cell layer. Increase in the DAPI fluorescence (Ex/Em at 358/461 nm) of nuclei, due to chromatin condensation was observed using a Nikon ECLIPSE Ti-S inverted fluorescence microscope. The controls were treated similarly except that MPP+ or MPP+ derivatives were omitted from the incubation medium.

3.3.17 Protein Determination

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

(192) using BSA as the standard. Aliquots of (50 μL) of cell suspensions were incubated with

Bradford protein reagents (950 μL) for 10 min and absorbance at 595 nm was recorded. The absorbance readings were converted to the corresponding protein concentrations using a standard curve constructed employing BSA.

3.3.18 Data Analyses

All data are averages of three or more replicates of a single experiment and presented as mean ± SD (all the experiments were carried out multiple times and the representative data from a single experiment are shown). The error bars represent the standard deviations from the mean of the data. To correct for the minor variations in color development in the MTT assay of cell viability, all the absorbance values were converted to a percentages of controls, which were treated under the same conditions except that MPP+ and MPP+ derivatives were excluded from the incubations. All quantitative data i.e. uptake, ATP and ROS levels were normalized to the protein content of individual samples to correct the cell density variations. P values for pair-wise comparisons were calculated by two-tailed Student’s t test. For all the data a level of p<0.05 was considered statistically significant.

3.3.19 Technical Statement

Due to the high toxicity and obvious health hazards of MPP+ and MPP+ derivatives, extreme caution was used in their handling in accordance with published procedures (193).

CHAPTER IV

RESULTS

4.1 Photophysical Properties of MPP+ Derivatives and Intracellular Accumulation of 4'I-

MPP+

Relative hydrophilicities of MPP+ derivatives were estimated using the corresponding

C18-reversed-phase HPLC-UV retention times in three different solvent systems. As shown from the data in Table 4, the retention times increased in the order 3'OH<4'H~4'F<4'Cl<4'Br<4'I-

MPP+ in all three solvent systems.

Table 4. Retention Times For MPP+ Derivatives

+ MPP Derivative Buffer:CH3CN Buffer:CH3CN:MeOH Buffer:CH3CN:MeOH 70:30 (min) 60:30:10 (min) 65:25:10 (min) 3'OH-MPP+ 5.11 4.59 5.07 MPP+ 5.92 5.28 6.02 4'F-MPP+ 6.44 5.68 6.60 4'Cl-MPP+ 8.66 7.08 9.17 4'Br-MPP+ 9.67 7.74 10.30 4'I-MPP+ 11.76 8.88 12.56

The UV-Vis spectral analysis of MPP+ derivatives show that the characteristic absorption band of the parent MPP+ was red shifted significantly in 4'Cl, 4'Br, and 4'I- MPP+, while blue shifted in 3'OH-MPP+ (Table 5).

Table 5. UV-Vis Analysis of MPP+ Derivatives

MPP+ Derivative Maximum Absorption (nm)

3'OH-MPP+ ~288 MPP+ ~295 4'F-MPP+ ~297 4'Cl-MPP+ ~301 4'Br-MPP+ ~302 4'I-MPP+ ~313

The effect of the solvent polarity on the characteristics of the absorption spectra of 4'I-

MPP+ was further investigated. The results of these experiments show that the absorption band at 313 nm was not significantly affected by the change in the polarity of the solvent (Figure 4).

0.14

0.12 PBS MeOH 0.10 CH3CN 0.08

0.06

Absorbance 0.04

0.02

0.00 280 300 320 340 360 380 nm

Figure 4. UV-Vis Spectra of 4'I-MPP+ (5 M) in PBS, Methanol, and Acetonitrile

Fluorescence spectral analysis of 4'I-MPP+ in PBS show a moderately intense fluorescence band with the excitation and emission maxima at 320 and 430 mm, respectively

(Figure 5). In addition, the fluorescence of 4'I-MPP+ was moderately quenched by polar solvents, for example, fluorescence of 4'I-MPP+ in PBS and MeOH was quenched by about ~60% compared to acetonitrile (Figure 5).

Ex Em 40 PBS MeOH CH CN 30 3

Fluorescence Fluorescence (au)

0 300 400 500 600 nm

Figure 5. Excitation and Emission Spectra of 4'I-MPP+ (5M) in PBS, Methanol, and Acetonitrile

Furthermore, imaging of 4'I-MPP+ loaded differentiated MN9D cells clearly shows that the intracellular 4'I-MPP+ could easily be detected in live cells by observing the intrinsic fluorescence of 4'I-MPP+ (Figure 6). Fluorescence response of 4'I-MPP+ was observed inside the differentiated MN9D cells and the fluorescence intensity increased with time (Figure 6).

Figure 6. Light and Fluorescence Images of 4'I-MPP+ Treated Differentiated MN9D Cells. Differentiated MN9D cells were washed with KRB-HEPES, pH 7.4, reconstituted with the same buffer and the light image was recorded. Then 4'I-MPP+ in KRB-HEPES, pH 7.4 was added to a final concentration of 50 M and the fluorescence (Ex/Em 340/470-550 nm) was recorded at the time zero and after 2 h. (A) light image; the florescence image of the same cells (B) at zero time; (C) after 2 h.

4.2 Relative Dopaminergic Cell Toxicities of 4'-Substituted MPP+ Derivatives

The toxicity of 4' substituted MPP+ derivatives on dopaminergic MN9D, SH-SY5Y and non-dopaminergic liver HepG2 and kidney HEK293 cells were determined using the MTT cell viability assay. The data show that while MN9D cells are highly susceptible, SH-SY5Y cells are moderately susceptible to the toxicities of all the derivatives during 12 h incubation periods under similar experimental conditions. Similar to the cell specificity of the parent toxin MPP+,

HEK293 and HepG2 cells are relatively resistant to the toxicities of these derivatives under similar experimental conditions. As shown from the data in Figures 7A-D the toxicities of all these derivatives follow the same trend in cell specificity of toxicity and it is in the order

MN9D>SH-SY5Y>HEK293>HepG2 cells. In addition, 4'I-MPP+ is also toxic to differentiated

MN9D cells with somewhat less potency in comparison to the undifferentiated cells, but not significantly toxic to HepG2 cells under similar experimental conditions (Figure 7E), again parallel to the toxicity profile of MPP+ (161).

A MN9D

B SH-SY5Y

Figure 7. MPP+ Derivatives are Specifically Toxic to Dopaminergic Cells Similar to MPP+. Cells were grown in 96-well plates and treated with the desired concentrations of MPP+ or MPP+ - derivatives in DMEM-HCO3 for 12 h at 37 °C. Cell viabilities were determined by the MTT assay as detailed in section 3.3.4. Results are expressed as % viability with respect to parallel controls which were treated similarly except that the toxin was omitted from the incubation medium. (A) MN9D; (B) SH-SY5Y; (C) HepG2 and (D) HEK293. (E) Differentiated MN9D, SH-SY5Y cell toxicities of 4'I-MPP+ under similar experimental conditions. Data are presented as mean ± SD (n=6).

C HepG2

D HEK293

Figure 7. (continued)

The inspection of the relative MN9D cell toxicities of 4' substituted MPP+ derivatives show that the toxicities increased with increasing hydrophobicity of the 4' substituent, as

+ estimated by the corresponding IC50 parameters (Table 6). Consequently, 4'I-MPP was the most

+ toxic (IC50 ~50 μM) and 3'OH-MPP was the least toxic (IC50 >175 μM ) to dopaminergic MN9D cells under similar experimental conditions.

+ Table 6. IC50 Values of MPP Derivatives in MN9D Cells

+ MPP Derivative IC50 Value (μM)

3'OH-MPP+ > 175 4'F-MPP+ 175 MPP+ 93 4'Cl-MPP+ 100 4'Br-MPP+ 78 4'I-MPP+ 48

4.3 Characteristics of the Cellular Uptake of 4'-Substituted MPP+ Derivatives

The cellular uptake of 4'-substituted MPP+ derivatives into the different cell types were determined by C18-reversed-phase HPLC-UV as detailed in section 3.3.5. The results show that cellular uptake into all the cell types increases with increased hydrophobicity of the toxin (Figure

8A). The 4'I-MPP+ uptake into dopaminergic cells (MN9D and SH-SY5Y) was about twice as much as the 3'OH-MPP+ uptake. For example, 4'I-MPP+ and 3'OH-MPP+ uptake into MN9D cells were about 1.6 and 0.6 nmoles/mg of protein in 45 min, respectively (Figure 8A). In addition, while all 4'I-substituted MPP+ derivatives were taken up into all cell types with varying efficiencies, HepG2 cell uptake was the most efficient and HEK293 cell uptake was the least efficient, under similar experimental conditions for all the toxins (Figure 8A).

Time dependent uptake of MPP+ and 4'I-MPP+ into MN9D was relatively linear during the 60 min incubation period (at 100 M concentration). The uptake of 4'I-MPP+ was about two

52 times as fast as MPP+ (1.2 nmoles/mg versus 0.6 nmoles/mg of protein in 1h; Figure 8B), which is consistent with the higher toxicity of 4'I-MPP+ relative to MPP+. Both MPP+ and 4'I- MPP+ uptake into HepG2 cells were faster in comparison to MN9D cells and apparently saturable under similar conditions (Figure 8B). The MPP+ uptake reached to about 1.2 nmoles/mg in 60 min and 4'I-MPP+ reached up to 1.4 nmoles/mg of protein in 40 min again suggesting that 4'I-

MPP+ uptake is somewhat more faster than MPP+.

As mentioned above, intracellular 4'I-MPP+ could be easily detected by its intrinsic fluorescence in live cells (Figure 6). Thus, to determine whether the cellular uptake of 4'I-MPP+ could be measured by fluorescence, a parallel series of uptake experiments were carried out with live MN9D and HepG2 cells using epifluorescence microscopy. Data in Figure 8B show that 4'I-

MPP+ uptake time courses for both MN9D and HepG2 cells were similar to that of HPLC-UV time courses.

A * * *

* * * * * * * * * * * * * * * # * * * * *

0.8 1.4 250 + + + MN9D/MPP /UV 1.2 MN9D/4'I-MPP /UV MN9D/4'I-MPP /Flu 0.6 200 1.0 0.8 150 0.4 0.6 100 0.2 0.4 50

Uptake (nmoles/mg) Uptake Uptake (nmoles/mg) Uptake 0.2

Uptake (arbitory units) (arbitory Uptake 0.0 0.0 0 0 10 20 30 40 50 60 0 10 20 30 40 50 60 0 10 20 30 40 50 60 Time (min) Time (min) Time (min)

1.4 1.6 700 + + + 1.2 HepG2/MPP /UV 1.4 HepG2/4'I-MPP /UV 600 HepG2/4'I-MPP /Flu 1.0 1.2 500 1.0 0.8 400 0.8 0.6 300 0.6 200 0.4 0.4

Uptake (nmoles/mg) Uptake 100

Uptake (nmoles/mg) Uptake 0.2 0.2 units) (arbitory Uptake 0.0 0.0 0 0 10 20 30 40 50 60 0 10 20 30 40 0 10 20 30 40 50 60 Time (min) Time (min) Time (min)

Figure 8. Uptake of MPP+ and 4'-Substituted MPP+ Derivatives into Cells. Cells were grown in DMEM to 70-80% confluence in 12 well plates and the cellular uptakes were measured by C18 reversed phase HPLC-UV as detailed in section 3.3.5. For uptake measurements by 4'I-MPP+ fluorescence, cells were grown in glass bottomed plates and fluorescence was measured by a Nikon Eclipse Ti-S inverted fluorescence microscope. (A) Cells were incubated with 100 μM MPP+ or 4'-substituted MPP+ derivatives in KRB-HEPES, pH 7.4 for 45 min, at 37 °C. The intracellular toxin contents were determined as described in section 3.3.5. The data are presented as mean ± SD (n=3). *p<0.005 and #p<0.05 versus respective controls. (B) Time dependency of uptake was measured by HPLC-UV and 4'I-MPP+ fluorescence. MN9D and HepG2 cells were incubated with 100 μM MPP+ or 4'I-MPP+ in KRB-HEPES, pH 7.4 for a desired period of time, at 37 °C. The intracellular MPP+ or 4'I-MPP+ contents were determined as described in section 3.3.5. The data are presented as mean ± SD (n=3). To measure the cellular uptake of 4'I-MPP+ by fluorescence, MN9D cells were treated with 100 μM 4'I-MPP+ and the increase of intracellular fluorescence at Ex/Em at 340/470-550 nm was measured as a function of time.

4.4 Mitochondrial Uptake of 4'-Substituted MPP+ Derivatives

Numerous previous studies have demonstrated that MPP+ accumulates in mitochondria isolated from the liver, brain and other cell types (94). The permanent positive charge of MPP+ derivatives could facilitates them to electrogenical accumulation inside the negatively charged mitochondria to cause depolarize the mitochondrial membrane potential. To test this possibility, the mitochondrial uptake of 4'-substituted MPP+ derivatives were determined with isolated

MN9D cell mitochondria using HPLC-UV quantification. Results show that the mitochondrial uptake of these derivatives are dependent on their relative hydrophobicities (Figure 9A), following a pattern similar to that of cellular uptake. The most hydrophobic, 4'I-MPP+ (at 400

M concentration within 45 min), showed the highest mitochondrial uptake (1.0 nmoles/mg of protein) in comparison to other 4'-substituted MPP+ derivatives.

The mitochondrial uptake of MPP+ is also associated with the depolarization of mitochondrial membrane potential. To determine whether the 4'-substituted MPP+ derivatives behave the same way, their effect on the mitochondrial membrane potential was determined using the fluorescent mitochondrial dye, TMRM. As shown from the data in Figure 9B, while all 4'-substituted MPP+ derivatives depolarize the mitochondrial membrane potentials of MN9D cells with varying efficiencies, 4'I-MPP+ was found to be the most effective, which is again consistent with the relative mitochondrial uptake rates of these toxins. While 500 μM 3'OH-

MPP+ depolarizes the mitochondrial membrane potential by ~20% in about 45 min, the same concentration of 4'I-MPP+ depolarizes the potential by ~50% under similar conditions (Figure

9B). Similarly, while 500 μM MPP+ depolarizes the mitochondrial membrane by 20 - 50% in about 45 min, 100 μM 4'I-MPP+ takes only 20 min to exert a similar effect, under similar experimental conditions (Figure 9C). Mitochondrial membrane potential of HepG2 cells was also

55 depolarized by MPP+ and 4'I-MPP+, but with a higher efficiency, in comparison to MN9D cells

(Figure 9C). All 4'-substituted MPP+ derivatives except 4'I-MPP+ show ~10 min lag period in the initiation of the depolarization of the mitochondrial membrane potential at a concentration of 500

μM (Figure 9B). On the other hand, the mitochondrial depolarization with the most hydrophobic,

4'I-MPP+ is clearly visible within 6 min after the addition of the toxin.

A B 1.2 * 10 1.0 0 0.8 * * -10

0.6 -20 3OH MPP+ 0.4 -30 MPP+ n moles/mgn protein 4F MPP+ -40 0.2 * 4Cl MPP+ * fluorescence F-TMRM * 4Br MPP+ -50 0.0 4I MPP+ -60 MPP+ 4F MPP+ 4I MPP+ 3OH MPP+ 4Cl MPP+4Br MPP+ 0 10 20 30 40 Time (min) Figure 9. Mitochondrial Uptake of 4'-Substituted MPP+ Derivatives. (A) Uptake of MPP+ and 4'- Substituted MPP+ Derivatives into Isolated MN9D Cell Mitochondria. Mitochondria from MN9D cells were isolated according to the procedure described in section 3.3.6. Intact mitochondrial suspensions were incubated with 400 μM MPP+ or a 4'-substituted MPP+ derivative in KCl buffer for 45 min at 37 °C. Intracellular content of the toxin was determined by

C18 reversed phase HPLC-UV as detailed in section 3.3.6s. The data are presented as mean ± SD (n=3). *p<0.005 versus respective controls. (B) Mitochondrial Membrane Potential Depolarization by MPP+ and 4'-Substituted MPP+ Derivatives. MN9D cells were treated with 50 nM TMRM in KRB-HEPES at 37 °C in the dark for 45 min. The mitochondrial TMRM fluorescence was recorded for 3 min and then the media was reconstituted with the desired concentration of MPP+ or one of the 4'-substituted MPP+ derivatives and the recording was continued for the desired period of time. Controls were carried out using an identical protocol except that toxins were excluded from the incubation media. The averages of background corrected, control subtracted data were used to estimate the change in mitochondrial membrane potentials as a function of time. (C) Mitochondrial Membrane Potential Depolarization by 500 M MPP+ and 100 M 4'I-MPP+ in MN9D (blue) and HepG2 (purple) Cells. Experiments were carried out similar to (B).

10 10 0 0 -10 -10 -20 -20 -30

+ -30 -40 4IMPP + MPP -50 -40

Meb.potential deacrease (Arb.units) deacrease Meb.potential -60 -50 (Arb.units) deacrease Meb.potential 0 2 4 6 8 10 12 14 0 10 20 30 40 50 Time (min) Time (min)

Figure 9. (continued)

4.5 Intracellular 4'I-MPP+ Localized into the Mitochondria

The cytosolic distribution of intracellular fluorescent 4'I-MPP+ was examined in 4'I-

MPP+ loaded differentiated and undifferentiated MN9D and HepG2 cells after labeling the mitochondria with the commonly used mitochondrial marker MitoTracker Green FM. As shown from the data in Figure 10, the fluorescence of MitoTracker Green FM overlaps completely with that of 4'I-MPP+s in all cases without significant cell-type specificity. The fluorescence images further show that no significant fluorescence is associated with the cell nucleoli or the membrane.

Diff. MN9D

Undiff. MN9D

HepG2

Figure 10. Intracellular Localization of Cytosolic 4'I-MPP+. Cells were incubated with 100 M 4'I-MPP+ and 200 nM MitoTracker Green FM, washed and fluorescence (4'I-MPP+; Ex/Em 340/470-550 nm and MitoTracker Green FM; Ex/Em 644/665 nm) images were recorded.

4.6 Membrane Potentials Drive Active Cellular and Mitochondrial Accumulation of MPP+ and its 4'-Substituted Derivatives

The role of plasma membrane and/or mitochondrial membrane potentials on the overall cellular accumulation of MPP+ and its 4'-substituted derivatives was determined using FCCP

(acts as a depolarizing agent on the plasma membrane and mitochondrial membrane) and 85 mM K+ ions (depolarizes the plasma membrane). As shown in Figure 11A, 5 μM FCCP significantly decreased the cellular accumulation of both MPP+ and its derivatives into MN9D cells. Interestingly, the uptake inhibition by FCCP was significantly higher than that of 85 mM

K+ under similar conditions (Tables 7). In the absence of extracellular Ca2+, the effects of both

FCCP and 85 mM K+ were greater for all three toxins in comparison to that of in the presence of

Ca2+. For example, FCCP inhibited the MPP+ uptake by 26.6% in the presence of extracellular

Ca2+ and 66.8% in the absence Ca2+ with EGTA (Tables 7). The same concentration of FCCP had a stronger effect on the cellular accumulation of both MPP+ and 4'I-MPP+ in HepG2 cells in comparison to MN9D cells (Figure 11B). 4'I-MPP+ fluorescence imaging experiments further confirmed that cytosolic 4'I-MPP+ in FCCP treated differentiated MN9D and HepG2 cells is distributed largely throughout the cytosol and not specifically localized in the mitochondria of the cells, as opposed to FCCP untreated controls (Figure 11C).

Table 7. FCCP and 85 mM K+ Percent Inhibition of MPP+ Derivatives in the Presence and Absence of Extracellular Ca2+ in MN9D Cells

Compound % Inhibition by FCCP % Inhibition by 85 mM K+ + Ca2+ - Ca2+ + Ca2+ -Ca2+ 3'OH-MPP+ 30.5 48.9 16.0 38.8 MPP+ 26.6 66.8 13.3 38.4 4'I-MPP+ 33.7 50.7 28.5 54.2

A B 4.0 3.0 - K+ A -FCCP + K+ 2.5 +FCCP 3.0 2.0

2.0 1.5

1.0

nmoles/mg protein nmoles/mg 1.0 protein nmoles/mg 0.5

0.0 0.0

MPP+/Ca2+ MPP+/+Ca2+MPP+/-Ca2+MPP+/+EGTA MPP+/+EGTA4I MPP+/Ca2+ 4I MPP+/+Ca2+4I MPP+/-Ca2+4I MPP+/+EGTA 4I MPP+/+EGTA 3OH MPP+/+Ca2+3OH MPP+/-Ca2+3OH MPP+/+EGTA

Differentiated MN9D Differentiated MN9D HepG2 HepG2 + FCCP + FCCP

Figure 11. The Effect of FCCP and 85 mM K+ on MPP+ and 4'-Substituted MPP+ Derivative Uptake into MN9D and HepG2 Cells. (A and B) MN9D cells were incubated with 5 μM FCCP in KRB-HEPES, with KRB-HEPES containing no Ca2+ or KRB-HEPES containing no Ca2+ with EGTA for 30 min and then with 50 μM MPP+ or a derivative of 4'-substituted MPP+ for 45 min at 37 °C. In experiments where the effect of K+ was investigated, cells were incubated with 50 μM MPP+ or 4'-substituted MPP+ derivatives in 85 mM K+ containing KRB-HEPES for 45 min at 37 °C. After the incubation, intracellular MPP+ or 4'-substituted MPP+ levels were quantified by C18 reversed phase HPLC-UV as detailed in section 3.3.9. The data are presented as mean ± SD (n=3). (C) The Effect of FCCP on the Intracellular Distribution of 4'I-MPP+ in Differentiated MN9D and HepG2 Cells. Cells grown in glass bottomed plates were incubated with 5 μM FCCP for 30 min and then with 100 M 4'I-MPP+ for 1 h in KRB-HEPES at 37 °C. Cells were washed and 4'I-MPP+ fluorescence images (Ex/Em 340/470-550 nm) were recorded. Controls were carried out in an identical manner except that FCCP was omitted from the incubation medium.

4.7 The Effect of Extracellular Ca2+ on Cellular Uptake of MPP+

The effect of extracellular Ca2+ ions on the cellular uptakes of MPP+ and 4'I-MPP+ was investigated in Ca2+ media (KRB-HEPES). In these experiments, to assure the complete removal of Ca2+ from the incubation media, 1.3 mM EGTA was included in the Ca2+ free media.

Interestingly, the uptake of both toxins into dopaminergic MN9D and SH-SY5Y cells was increased by ~2 fold in the incubations of Ca2+ free KRB-HEPES. On the other hand, Ca2+ had no significant effect on the uptake of MPP+ and 4'I-MPP+ into non-dopaminergic HepG2 and

HEK293 cells, under similar incubation conditions (Figure 12A).

The Ca2+ sensitivity of the MN9D cell uptake of 4'I-MPP+ was further examined with known calcium and/or cation channel inhibitors. In these experiments, cells were pre-incubated with the desired inhibitor for 30 min, and then incubated with 50 M 4'I-MPP+ for 45 min and the intracellular toxin levels were quantified using HPLC-UV as detailed in the section 3.3.5. The results of these experiments revealed that while KBR7943, ruthenium red, 2APB, verapamil and

CGP37157 had no significant effect, GBR12909 [DAT inhibitor (194)], benzamil, and mibefradil [VGCC inhibitor (120,124,195)] showed a moderate inhibition (~20-25% inhibition) of 4'I-MPP+ uptake into MN9D cells under similar experimental conditions (Figure 12B).

2.5 2.0 2+ Ca2+-HEPES Ca - HEPES * EGTA- HEPES 2.0 EGTA -HEPES * * 1.6

1.5 1.2

1.0 0.8

* * uptake (nmoles/mg) + 0.5 # 0.4

MPP+ uptake (nmoles/mg) * * 4I' MPP 0.0 0.0

HepG2 MN9D HepG2 MN9D SH-SY5Y HEK293 SH-SY5Y

Figure 12. (A) The Effect of Extracellular Ca2+ on MPP+ and 4'I-MPP+ Uptake. Cells were grown in DMEM to 70-80% confluence in 12 well plates and the cellular uptakes were measured by C18 reversed phase HPLC-UV as detailed in section 3.3.5. Cells were then incubated with 50 μM MPP+ or 4'I-MPP+ in KRB-HEPES or EGTA-HEPES (1.3 mM EGTA; Ca2+ free condition), pH 7.4 for 45 min, at 37 °C. (B) Inhibition of 4'I-MPP+ Uptake into MN9D Cells by Various Pharmacological Agents. Cells were grown in DMEM to 70-80% confluence in 12 well plates and the cellular uptakes were measured by C18 reversed phase HPLC-UV as detailed in section 3.3.5. Cells were incubated with 990 μL of KRB-HEPES, pH 7.4 containing a 25 μM of the desired inhibitor (1 μM ruthenium red and 10 μM GBR12909) for 30 min, at 37 °C. Then 10 μL of 4'I-MPP+ (final concentration 50 μM) was added and the incubation was continued for an additional 45 min, at 37 °C. The data are presented as mean ± SD (n=3). *p<0.005 and #p<0.05 versus respective controls.

4.8 The Effect of Ca2+ on Mitochondrial Accumulation of MPP+

4'I-MPP+ was selected to examine the mitochondrial uptake since it had the highest mitochondrial uptake rate among the 4'-substituted MPP+ derivatives tested. Mitochondrial Ca2+ concentration in live MN9D cells was measured using the mitochondrial Ca2+ specific fluorescence dye Rhod-2 AM. As shown from the data in Figure 13, 100 M 4'I-MPP+ decreased the Rhod-2 AM fluorescence significantly within a specific time period in MN9D cells with respect to a control without 4'I-MPP+.

Figure 13. The Effect of 4'I-MPP+ on the Mitochondrial Ca2+ Levels in Live MN9D Cells. Cells were treated with 5 μM Rhod-2-AM in KRB-HEPES for 1 h at room temperature in the dark. After the incubation, Rhod-2-AM fluorescence was recorded for 2 min and then the media was reconstituted with 100 M 4'I-MPP+ (final concentration) and the recording was continued for additional 58 min. Controls were carried out using an identical protocol except that 4'I-MPP+ was excluded from the incubation media. The averages of background corrected, control subtracted data were used to estimate the change in mitochondrial Ca2+ levels as a function of time.

4.8.1 The Effect of Extra-Mitochondrial (Cytosolic) Ca2+ and Na+ on Mitochondrial

Accumulation of 4'I-MPP+

The effect of Ca2+ on the mitochondrial uptake of 4'I-MPP+ was investigated using the mitochondria isolated from MN9D cells. As shown from the data in Figure 14A, 500-1000 nM

Ca2+ in the incubation media inhibits the 4'I-MPP+ uptake into mitochondria by about 40-60%.

The effects of Na+ and Li+ on mitochondrial 4'I-MPP+ uptake, were also investigated by replacing equal concentrations of KCl with 10 mM NaCl or LiCl in the incubation buffer. While

Na+ inhibits the mitochondrial uptake by ~50%, Li+ had no inhibition effect on the uptake under similar experimental conditions (Figure 14B).

A B

* * *

Figure 14. The Effect of Extra-mitochondrial Ca2+, Na+ or Li+ on MN9D Mitochondrial Uptake of 4'I-MPP+. MN9D mitochondrial suspensions were incubated with 200 μM 4'I-MPP+ in KCl buffer containing (A) 0, 500 or 1000 nM Ca2+; (B) 10 mM NaCl or LiCl for 45 min at 37 °C. After the incubations, the intracellular 4'I-MPP+ contents were quantified by HPLC-UV as described in section 3.3.6. The data are represent as mean ± SD (n=3). *p<0.005 versus zero Ca2+/ Na+ controls.

4.8.2 The Effect of Ca2+ Channel Blockers on the Mitochondrial Accumulation of MPP+

To further characterize the role of Ca2+ on mitochondrial 4'I-MPP+ uptake, the effect of a series of pharmacological agents that are known to interfere with the mitochondrial Ca2+ homeostasis was examined using isolated intact MN9D mitochondria. Initial screening experiments showed that the specific mitochondrial Ca2+ uniporter inhibitor, ruthenium red

(133,196) or voltage gated Ca2+ channel inhibitors, verapamil (121) and benzamil (120,195) had no significant effect on mitochondrial uptake of 4'I-MPP+ (Figure 15A). But the mNCX inhibitors, CGP37157 [specific mNCX inhibitor (23,24)], KBR7943 (128), and 2APB significantly inhibited the uptake of 4'I-MPP+ into mitochondria (Figure 15B; 50 M inhibitor at

200 M 4'I-MPP+). The dose dependency studies with the most effective inhibitor, CGP37157, showed that with a pharmacologically relevant concentration of 10 M (23), 4'I-MPP+ uptake inhibition was ~40% and increase of the concentration to 25 M had only a minor effect (~45% inhibition) on 4'I-MPP+ uptake into mitochondria (Figure 15C).

To determine whether the mNCX inhibitors are also effective in inhibiting the mitochondrial uptake of 4'I-MPP+ in live cells, a series of live cell imaging experiments were carried out with CGP37157. The data presented in Figure 15D shows that while cytosolic 4'I-

MPP+ is primarily localized in the mitochondria (labeled with MitoTracker Green FM) of control

HepG2 cells, in cells pre-treated with CGP37157, the intracellular 4'I-MPP+ is dispersed throughout the cells.

* * *

Figure 15. The Effect of Ca2+ Channel Blockers on Mitochondrial Accumulation of MPP+. (A and B) Effects of Benzamil, Verapamil, FFA, Ruthenium red, CGP37157, 2APB, and KBR7943 on the MN9D Mitochondrial Uptake of 4'I-MPP+. MN9D mitochondrial suspensions were initially incubated with a 50 μM concentration (10 μM ruthenium red) of the desired agent in KCl buffer for 10 min at 37 °C and then incubated with 200 μM 4'I-MPP+ for 45 min at 37 °C. + After the incubations, the intracellular 4'I-MPP contents were quantified by C18 reversed phase HPLC-UV as described in section 3.3.6. The data are represent as mean ± SD (n=3). *p<0.005 versus controls. (C) Concentration Dependence of the Inhibition of MN9D Mitochondrial Uptake of 4'I-MPP+ by CGP37157. The experiments were carried out using a similar protocol as above with varying concentrations of CGP37157 (10-50 M). The data are represent as mean ± SD (n=3). *p<0.005 versus controls. (D) The Effect of CGP37157, on the Mitochondrial Accumulation of 4'I-MPP+ in HepG2 Cells. Cells were incubated with 50 μM CGP37157 for 30 min and then with 100 M 4'I-MPP+ and 200 nM MitoTracker Green FM in KRB-HEPES for 1 h at 37 °C. After the incubations, cells were washed and the fluorescence images at Ex/Em 340/470-550 nm (4'I-MPP+) and 644/665 nm (MitoTracker Green FM) were recorded. Controls were treated under similar conditions except that CGP37157 is excluded from the incubation medium.

* * *

Figure 15. (continued)

4.8.2.1 The Effect of CGP37157 and KBR7943 on Mitochondrial Membrane Potential With

4'I-MPP+ Uptake

Mitochondrial membrane depolarization was measured using the mitochondrial membrane potential sensitive fluorescent dye, TMRM as described previously, to further confirm the inhibitory effects of CGP37157 and KBR7943 on the mitochondrial accumulation of the toxin. CGP37157 and KBR7943 pre-treatments significantly reduced the typical mitochondrial membrane depolarization observed with 4'I-MPP+ in MN9D cells (Figure 16A). The experiments

67 with TMRM showed that CGP37157 slows down the 4'I-MPP+-induced mitochondrial membrane depolarization of HepG2 cells lasting at least 4 h (Figure 16B).

20 100 M 4IMPP 10 100 M 4IMPP+50 M KBR 100 M 4IMPP+50 MCGP 0

-10

-20

-30 4IMPP+ -40

Memb. Pot. Decrease (arb. units) (arb. Decrease Pot. Memb. -50 0 5 10 15 20 Time (min) B No 4I MPP + 100  M 4I MPP + 100  M 4I MPP + + CGP

T = 0 h

T = 4 h

Figure 16. (A) The Effect of CGP37157 and KBR7943 on the 4'I-MPP+-Mediated Mitochondrial Depolarization of HepG2 Cells. HepG2 cells were incubated with either 50 μM CGP37157 or KBR7943 for 30 min and then with 50 nM TMRM for 45 min in KRB-HEPES at 37 °C. After the incubation intracellular TMRM fluorescence (Ex/Em 543/573 nm) was measured for 2 min and then, the media was reconstituted with 4'I-MPP+ to a 100 M final concentration and the fluorescence measurements were continued for additional 20 min. Controls were treated identically except that CGP37157 and KBR7943 were excluded from the initial incubation media. The averages of background and baseline subtracted data were used to estimate the mitochondrial membrane potentials. (B) The Effect of CGP37157 on the 4'I-MPP+ Mediated Mitochondrial Membrane Depolarization under Longer Incubation Conditions. HepG2 cells were incubated with or without 10 μM CGP37157 followed by 50 nM TMRM as described above. After the incubation, the intracellular TMRM fluorescence was recorded. Then, the media was reconstituted with 4'I-MPP+ to a 100 M final concentration and the TMRM fluorescence images was recorded after 4 h incubation at 37 °C. Control was carried out using an identical protocol except that either CGP37157 or 4'I-MPP+ were omitted from the incubation media.

4.8.2.2 CGP37157 Protect MN9D Cells From MPP+ and 4'I-MPP+ Toxicity

As expected, CGP37157 was found to effectively protect MN9D cells in a concentration dependent manner from MPP+ and 4'I-MPP+ toxicity due to the toxin uptake inhibition (Figure

17).

* * * * * * * * * * * * * # # *

Figure 17. The Effect of CGP37157 on MN9D Toxicities of MPP+ and 4'I-MPP+. Cell were pre- incubated with 0-50 µM CGP37157 for 30 min followed by 80 µM MPP+ or 25 µM 4'I-MPP+ in KRB-HCO3 for 8 h. The relative cell viabilities were determiner using MTT assay as detailed in section 3.3.4. Data are presented as mean ± SD (n=6). *p<0.0001 and #p<0.005 versus no CGP37157 controls.

4.9 Does mNCX Play a Role in Mitochondrial Accumulation and Toxicity of MPP+?

The inhibition uptake into mitochondria by specific mNCX inhibitors suggests that mNCX may play a role in the mitochondrial uptake of 4'I-MPP+. This possibility was further investigated using the NCX isoforms transfected HEK293 cells. Wild type dopaminergic MN9D and SH-SY5Y cells express NCX3, while non dopaminergic HepG2 and HEK293 cells do not express significant levels of NCX3 (Figure 18A). Western blot analysis of the NCX3 transfected

HEK293 cells showed the expression of high levels of this protein in both plasma membrane and mitochondrial membrane (Figures 18B).

Figure 18. NCX3 Transfection into HEK293. NCX3 plasmid was transfected into wild type HEK293 cells as described in section 3.3.11. Mitochondria were extracted from NCX3 transfected HEK293 cells and wild type HEK293 cells. NCX3 expression levels (A) in cells and (B) in mitochondria were identified using Western blot analysis as explained in section 3.3.11.1. β-actin was used as a loading control.

4.9.1 NCX Transfection into HEK293 Cells Increases 4'I-MPP+ Uptake and Toxicity

The standard uptake experiments with NCX transfected and wild type HEK293 cells show that the NCX transfection increases the 4'I-MPP+ uptake relative to the wild type cells, significantly (Figure 19A). In parallel to the increase in uptake, NCX transfection also led to a significant increase in 4'I-MPP+ toxicity in comparison to the wild type cells (Figure 19B). The results further show that NCX2 and 3 transfected cells produced the highest and NCLX transfected cells showed modest effects on both uptake and toxicity.

A * *

* # * * * * * #

Figure 19. The Effect of NCX in 4'I-MPP+ Uptake and Toxicity. (A) 4'I-MPP+ Uptake into NCX/NCLX Transfected HEK293 Cells as Measured by HPLC-UV. HEK293 cells were transfected with NCX/NCLX plasmid and grown in 12 well plates. Cells were incubated with the desired concentration of 4'I-MPP+ in KRB-HEPES, pH 7.4 for 45 min, at 37 °C. The + intracellular 4'I-MPP contents were determined by C18 reversed phase HPLC-UV as described in section 3.3.5. The data are presented as mean ± SD (n=3). *p<0.005 and #p<0.008 versus non transfected controls. (B) 4'I-MPP+ is Specifically Toxic to NCX/NCLX Transfected HEK293 Cells. HEK293 cells were transfected with corresponding plasmids and grown in 96-well plates. + - Cells were treated with the desired concentrations of 4'I-MPP in DMEM-HCO3 or DMEM- - 2+ HCO3 with EGTA (to exclude Ca ) for 16 h at 37 °C. Cell viabilities were determined by the MTT assay as detailed in section 3.3.4. Results are expressed as % viability with respect to parallel controls which were treated similarly except that the toxin was omitted from the incubation medium. Data are presented as mean ± SD (n=6).

4.9.2 NCX Transfection into HEK293 Cells Depolarizes Mitochondrial Membrane

Potential by 4'I-MPP+ Uptake

Similarly, while 4'I-MPP+ does not cause a significant depolarization of mitochondrial membrane potential in wild type HEK293 cells, NCX transfected cells show a significant depolarization of the mitochondrial membrane potential under similar experimental conditions

(Figure 20).

10 5 0 -5 -10 -15 Non transfected -20 NCX1

F-TMRM-fluoresecence NCX2 -25 NCX3 NCLX -30 0 10 20 30 40 50 60 Time (min) Figure 20. Mitochondrial Membrane Potential Depolarization by 4'I-MPP+ in NCX/NCLX Transfected HEK293 Cells. Transfected cells were treated with 50 nM TMRM in KRB-HEPES at 37 °C in the dark for 45 min. The mitochondrial TMRM fluorescence was recorded for 2 min and then the media was reconstituted with 100 M 4'I-MPP+ and the recording was continued for 58 min. Controls were carried out using an identical protocol except that 4'I-MPP+ were excluded from the incubation media. The averages of background corrected, control subtracted data were used to estimate the change in mitochondrial membrane potentials as a function of time.

4.10 The Effect of Mitochondrial PTP Inhibitor on MPP+ Cellular Uptake

Previous studies suggest that the mitochondrial Ca2+ levels and mitochondrial membrane depolarization play a role in the opening of the mitochondrial PTP (100,101). Since the mitochondrial and cytosolic Ca2+ appear to play a role in MPP+ uptake and the fact that 4'I-MPP+ uptake leads to the depolarization of the mitochondrial membrane potential, the effect of the

72 well-known mitochondrial PTP opening inhibitor, cyclosporine A, on the 4'I-MPP+ uptake was investigated. In these experiments, cells were initially incubated with cyclosporine A, and then with MPP+ or 4'I-MPP+ in KRB-HEPES and intracellular toxin levels were quantified by HPLC-

UV as detailed in the section 3.3.5. The results show that cyclosporine A moderately inhibits the

MPP+ uptake into MN9D cells in a concentration dependent manner (Figure 21A). The uptake time course studies show that cyclosporine A has a significant effect on the MN9D cell uptake of 4'I-MPP+ in the presence, but no effect in the absence of Ca2+ in the incubation medium

(Figures 21B and C). As shown from the data in Figure 21B, cellular 4'I-MPP+ uptake increased in the presence of extracellular Ca2+ by cyclosporine A within 5 - 40 min with a maximum stimulation at ~ 10 min; the effect of 4'I-MPP+ largely disappeared at 60 min.

Figure 21. The Effect of Mitochondrial PTP Opening Inhibitor, Cyclosporine A on MPP+ and + + + 4'I-MPP Uptake into MN9D Cells. The MPP and 4'I-MPP uptakes were quantified by C18 reversed phase HPLC-UV as detailed in section 3.3.5. (A) Cells were pre-incubated with the desired concentration of cyclosporine A for 1 h and then with 100 μM MPP+ in KRB-HEPES, pH 7.4 for 1 h at 37 °C. (B and C) The Effect of Cyclosporine A on the Time Courses of 4'I- MPP+ Uptake into MN9D Cells in the (B) Presence and (C) Absence of Extracellular Ca2+. Cells were incubated in (B) KRB-HEPES, pH 7.4 and (C) EGTA-HEPES, pH 7.4 (with no Ca2+), containing a 5 μM cyclosporine A for 1 h at 37 °C. Then the incubation media was reconstituted to a final 50 μM concentration of 4'I-MPP+ and further incubated for the desired period of time, at 37 °C. Data are presented as mean ± SD (n=3).

Figure 21. (continued)

4.11 The Effect of MPP+ on Mitochondrial Complex I

Previous studies suggest that the toxicity of MPP+ and similar toxins [rotenone, cyanines, etc.(161)] are closely associated with their ability to accumulate in the mitochondria thus inhibiting the electron transport chain complex I (16,52). The relatively high mitochondrial uptake together with high MN9D toxicity of 4'I-MPP+ suggest that it could also be a complex I inhibitor. To test this possibility, the relative effects of MPP+ and 4'I-MPP+ on the NADH-

74 ubiquinone oxidoreductase activity of complex I were determined using isolated rat brain mitochondrial membranes employing the well-characterized ubiquinone dependent NADH- oxidation assay (190). The inhibition of complex I was determined by measuring the decrease in absorbance due to the ubiquinone dependent oxidation of NADH at 340 nm as a function of time. Under standard assay conditions, rotenone, a well-characterized specific and potent complex I inhibitor (54) inhibited about 90% of the NADH oxidation activity at 10 M confirming the suitability of the assay for specific complex I activity measurements (Figure

22).Under similar conditions, 4'I-MPP+ and MPP+ showed 46% and 21% inhibition of the complex I activity at 200 M concentrations, respectively (Figure 22).

0.0 A

-0.2

340

A -0.4 Control MPP (200 M) -0.6 4I MPP (200 M) Rotenone (10 M)

-0.8 0 50 100 150 200 250

Time (sec)

Figure 22. Both MPP+ and 4'I-MPP+ Inhibit the Rat Brain Mitochondrial Complex I. Isolated rat brain mitochondria were incubated with MPP+, 4'I-MPP+ or rotenone (a standard complex I inhibitor) in the standard assay solution for 2 min at room temperature. Then the reaction was initiated by adding 10 mM NADH and the rate of ubiquinone dependent oxidation of NADH was followed by measuring the decrease of absorbance at 340 nm with respect to the reference wave length of 425 nm for 5 min.

4.12 The Effect of MPP+ on ATP Depletion

The above results show that MPP+ or derivatives (4'I-MPP+) actively accumulate in the mitochondria, depolarize the mitochondrial membrane potential and inhibit the complex I.

Therefore, the exposure of cells to these toxins was expected to deplete the intracellular ATP levels. To test this possibility, intracellular ATP levels in MN9D and HepG2 cells after treatment of MPP+ or 4'I-MPP+ were determined using an ATP assay kit from Sigma as detailed in section

3.3.14. The treatment of MN9D or HepG2 cells with MPP+ or 4'I-MPP+ depletes intracellular

ATP levels in both cell types compared to controls, with 4'I-MPP+ showing a more pronounced decrease (Figure 23). As shown in Figure 23, 100 μM 4'I-MPP+ treatment decreased ATP levels by ~20% in both cell types.

* * * *

Figure 23. The Effect of MPP+ and 4'I-MPP+ on Intracellular ATP Levels of MN9D and HepG2 Cells. Intracellular ATP levels of MPP+ or 4'I-MPP+ (100 μM) treated MN9D and HepG2 cells (for 6 h at 37 °C) were quantified as detailed in section 3.3.14. The data are presented as mean ± SD (n = 3). *p< 0.0025 versus the controls with no toxin.

4.13 MPP+ and Its 4'-Substituted Derivatives Increase Intracellular ROS Production

Specifically in MN9D Cells

Numerous previous studies have shown that the dopaminergic toxicity of MPP+ is due to the apoptotic cell death caused by increased oxidative stress (137,161). To test whether the increased ROS production is also associated with the toxicity of 4'-substituted MPP+ derivatives, their effect on ROS levels was determined using the ROS sensitive fluorescent probe DCFH-DA

(197) as detailed in section 3.3.15. Treatment of MN9D cells with MPP+ and 4'-substituted MPP+ derivatives significantly increases the intracellular ROS levels within 1 h and the extent of ROS production is proportional to their cellular uptake rates (Figure 24A). Interestingly, MPP+ or its derivative treatments did not increase the baseline intracellular ROS levels in HepG2 cells in contrast to the dopaminergic cells under similar experimental conditions. As shown in Figure

24A, 4'Cl-MPP+, 4'Br-MPP+ and 4'I-MPP+ treatments caused a slight increase of ROS in

HEK293 cells under similar experimental conditions but was less pronounced in comparison to

MN9D cells. 4'I-MPP+-mediated intracellular ROS production in MN9D cells was also further confirmed by live cell fluorescent imaging experiments (Figure 24B). The potent antioxidants, ascorbic acid and cystine were found to inhibit the 4'I-MPP+ mediated increase in ROS in MN9D cells (Figure 24C).

* * *

* # #

B C

+ Control –Trans 4′I-MPP -Trans

* *

Control –DCF + 4′I-MPP -DCF

Figure 24. (A) The Effect of MPP+ and 4'-substituted MPP+ Derivatives on the Intracellular ROS Levels. Cells were treated with 10 μM DCFH-DA in KRB-HEPES for 1 h and then with 100 μM MPP+ or MPP+ derivatives for an additional 1 h at 37 °C. Intracellular ROS levels were measured by measuring the DCF fluorescence of MPP+ or 4'-substituted MPP+ derivatives treated cell extracts as detailed in section 3.3.15. The data are presented as mean ± SD (n=3). *p<0.005 and #p<0.05 compared with controls with no toxins. (B) 4'I-MPP+-Mediated ROS Production in MN9D Cells is Localized to the Mitochondria. MN9D cells were treated with 50 μM DCFH-DA for 1 h at 37 °C in KRB-HEPES. Then, cells were washed and incubated with 250 M 4'I-MPP+ for 4 h at 37 °C. The DCF fluorescence (Ex/Em 488/524 nm) images were recorded using a Leica confocal fluorescence microscope. Controls were treated under the same conditions except that 4'I-MPP+ was omitted from the incubation media (C) The Effect of Antioxidants on the 4'I-MPP+ Mediated ROS Production in MN9D Cells. MN9D cells were pre- incubated with 50 μM antioxidants in KRB-HEPES for 1 h at 37 °C. Then the cells were incubated with 100 μM 4'I-MPP+ for additional 1 h in KRB-HEPES at 37 °C and the intracellular ROS levels were determined as detailed in section 3.3.15. Data are presented as mean ± SD (n=3). *p<0.05 compared with zero antioxidant controls.

4.14 MPP+ Cause Specific Apoptotic MN9D Cell Death

Apoptotic cell death can be also diagnosed by observing the characteristic cell shrinkage and the increase of the fluorescence of the nuclear dye, DAPI, due to chromatin condensation. As shown from the data in Figure 25, MPP+- and 4'-substituted MPP+ derivative-mediated MN9D cell deaths were associated with the characteristic cell shrinkage (Figure 25A) and chromatin condensation (Figure 25B) as indicated by the increase of the nuclear DAPI fluorescence in toxin-treated cells in comparison to the controls.

Control 3'OH-MPP+ MPP+ 4'F-MPP+

4'Cl-MPP+ 4'Br-MPP+ 4'I-MPP+

Figure 25. MPP+ or 4'-Substituted MPP+ Derivatives Cause Apoptotic MN9D Cell Death. MN9D cells were grown in glass bottomed plates and incubated with 250 μM MPP+ or MPP+ derivatives in KRB-HEPES for 12 h at 37 °C and then treated with 300 nM DAPI for 15 min in the dark. Apoptotic chromatin condensation was visualized by observing the increase in nuclear DAPI fluorescence (Ex/Em 358/461 nm). Control cells were treated under similar conditions except that MPP+ and MPP+ derivatives were omitted from the incubation medium. (A) Trans images; (B) DAPI fluorescence images.

Control 3'OH-MPP+ MPP+ 4'F-MPP+

4'Cl-MPP+ 4'Br-MPP+ 4'I-MPP+

Figure 25. (continued)

CHAPTER V

DISCUSSION

MPP+ has been widely used for several decades as a convenient model to study the mechanisms of specific dopaminergic cell death in PD. In spite of extensive studies, the molecular details of the cellular uptake and the selective dopaminergic toxicity of MPP+ are still poorly understood and extensively debated. Therefore, the aim of this study was to develop and characterize convenient molecular probes to characterize the uptake and cellular toxicity in order to identify the cellular components involved in these processes in live cells and to advance the understanding of the mechanism of the specific dopaminergic cell toxicity of MPP+.

Accordingly, we have synthesized a series of 4'-halogen substituted MPP+ derivatives and characterized them with respect to their physical properties. These studies have led to the discovery that 4'I-MPP+, which is moderately fluorescent (Ex/Em 320/470-550 nm) (Figures 5 and 6) could be used to quantify the uptake and cellular distribution of MPP+ in live cells. In addition, since the hydrophilicities of these derivatives gradually increase with the increasing hydrophobicity of the 4'-substituent (Table 4), they could be used to determine the effect of increasing hydrophobicity on the rate of cellular uptake and toxicity. Based on the above justifications, we proceeded to characterize these derivatives with respect to their uptake and toxicity profiles against MPP+ to determine their suitability as MPP+ mimics for detailed mechanistic studies.

As expected, the standard cell viability assays showed that all 4'-substituted MPP+ derivatives are highly toxic to rat dopaminergic MN9D cells (170), moderately toxic to human dopaminergic SH-SY5Y cells (172), but relatively nontoxic to human liver HepG2 cells and

81 kidney HEK293 cells (Figures 7A-D). In addition, the dopaminergic cell toxicities of these derivatives increased with the increasing hydrophobicity of the molecule i.e. toxicity was in the order 4'I-MPP+> 4'Br-MPP+> 4'Cl-MPP+> 4'F-MPP+ ~ 4'OH-MPP+ (Table 6). In addition, the most effective toxic derivative, 4'I-MPP+, was also highly toxic to physiologically more relevant, differentiated MN9D cells parallel to MPP+ (Figure 7E) (161). These findings show that all the

MPP+ derivatives follow selective dopaminergic toxicity profiles similar to that of MPP+.

The traditional uptake measurements show that the rate of the cellular uptake of 4'- substituted derivatives into MN9D cells increased with the increasing hydrophobicity of the molecule. For example, the most hydrophobic derivative, 4'I-MPP+ showed the highest cellular uptake rate while the least hydrophobic derivative 3'OH-MPP+ had the lowest uptake rate. Thus, the rates of the uptakes appear to be parallel to their corresponding toxicities (Figure 8A).

Similar to MN9D cells, the non-neuronal HepG2 cells also take up these derivatives effectively.

In fact, the rates of uptake into HepG2 cells were significantly higher than that of MN9D cells, but followed the same trend as MN9D, i.e. uptake rates increased with increasing hydrophobicity. Furthermore, the uptake time courses determined by conventional HPLC-UV showed similar patterns for both MPP+ and 4'I-MPP+ in both cell types, suggesting that the mechanisms of uptake for the two derivatives could be similar (Figure 8B). More importantly, the time course of uptake constructed by measuring the intrinsic fluorescence of 4'I- MPP+ in live MN9D cells was similar to that obtained by conventional HPLC-UV (Figure 8B), suggesting that the intracellular 4'I-MPP+ in living cells could be conveniently quantified by fluorescence.

Lipophilic cations including MPP+ actively accumulate specifically in the mitochondria of many different cell types due to the large negative inner mitochondrial membrane potential

(161,198). In excellent agreement with these findings, all MPP+ derivatives were effectively

82 taken up into the MN9D cell mitochondria (Figure 9A) and the rates of uptake were found to be parallel to that of the cellular uptake. Specifically, similar to the cellular uptakes, the mitochondrial uptakes also increases with increase in the hydrophobicity of the molecule. In agreement with these findings, the most hydrophobic 4'-substituted derivative, 4'I-MPP+ was found to more strongly depolarize the mitochondrial membrane potential in comparison to the other less hydrophobic MPP+ derivatives (Figure 9B). These results strongly suggest that at least a portion of the cellular uptake must be due to the non-mediated simple diffusion through the cell membrane.

While the mitochondrial accumulation of MPP+ has not been directly demonstrated in live cells, fluorescent 4′-(dimethylamino)-MPP+ and related derivatives have been shown to accumulate in the mitochondria of various cells (199). However, the use of these derivatives to measure the cellular and mitochondrial uptake of MPP+ in live cells is hampered by the findings that they also concentrate into the nucleoli of the cell, specifically interact with and inhibit various cell membrane transporters, and non-specifically interact with the cell and sub–cellular organelle membranes in high levels, complicating the interpretation of the data (200,201). On the other hand, the unique photophysical properties of 4'I-MPP+ together with the similarity in toxicological and chemical properties to that of MPP+ suggest that 4'I-MPP+ could be a good probe to model the characteristics of the uptake of MPP+ into live cell mitochondria using epifluorescence. As shown from the data in Figure 10, the fluorescence of the mitochondrial specific dye MitoTracker Green FM overlaps completely with that of 4'I-MPP+ in all cases and no significant fluorescence is associated with the cell nucleoli or the membrane. These findings suggest that intracellular 4'I-MPP+ is primarily localized into the mitochondria of both cell types without significant cell-type specificity. Therefore, 4'I-MPP+ is an excellent probe to measure

83 the cellular and mitochondrial uptake of MPP+ and to model the intracellular distribution of

MPP+ in live cells.

The toxicities of MPP+ and similar toxins [rotenone, cyanines, etc. (161)] are closely associated with their ability to accumulate in the mitochondria, inhibit the electron transport chain complex I (16,52), decrease ATP production (13), and increase ROS production (13,136).

Parallel to the behavior of MPP+, 4'I-MPP+ was also found to be a complex I inhibitor with the inhibition potency higher than that of MPP+ (Figure 22). 4'I-MPP+ also decreased the intracellular ATP levels more efficiently than MPP+ (Figure 23). We and others have previously shown that dopaminergic cell toxicity of MPP+ is primarily due to the increased ROS-mediated apoptosis (137,138,161). Similar to MPP+, all the 4'-halogen substituted MPP+ derivatives were also capable of mitochondrial accumulation, membrane depolarization, complex I inhibition, and intracellular ATP depletion in both MN9D and HepG2 cells, but increased ROS production was observed only in dopaminergic MN9D cells (Figures 24A and B). This selective ROS production in MN9D cells in response to the MPP+ derivative treatments is in excellent agreement with our previous proposal that dopaminergic cells are inherently more prone to produce high levels of

ROS in response to mitochondrial toxins similar to MPP+, relative to other cell types (161,202).

The data further show that the MPP+ derivative mediated dopaminergic cell death is also due to apoptosis. These findings further confirm that the mechanisms of the selective dopaminergic cell toxicity of all MPP+ derivatives are similar to that proposed for MPP+. Previous studies have suggested that the high mitochondrial membrane potential contributes to the electrogenic accumulation of cations such as MPP+ in mitochondria (203,204). The present studies clearly show that the mitochondrial uptake is dependent on the relative hydrophobicity of the molecule, suggesting that their mitochondrial uptake is most likely due to a non-mediated simple diffusion

84 process. To further substantiate these proposals, plasma membrane and mitochondrial membrane potential contribution in toxin uptake were studied using mitochondrial and membrane depolarizing agents, FCCP and 85 mM K+ ions. These and parallel cell imaging studies (Figure

11A and C) clearly show that the mitochondrial and plasma membrane potentials play an important role in the cellular and mitochondrial accumulation of these toxins. Interestingly, the effect of FCCP on the 4'I-MPP+ uptake into HepG2 cells was more pronounced than that of

MN9D cells suggesting that there could be some fundamental differences between the uptake pathways of these into the two cell types (Figure 11B). The experiments carried out with membrane depolarizing agents in the presence and absence of Ca2+ show that the effects of membrane and mitochondrial potentials on the uptakes were more drastic in the absence of Ca2+

(Tables 7). Although the role of Ca2+ in the uptake of these toxins is not fully understood, these findings appear to suggest that cytosolic and/or mitochondrial Ca2+ may play a role in modulating the mitochondrial membrane potential and that mitochondrial and plasma membrane potentials play a role in cellular and mitochondrial uptake of these toxins.

All mammalian cells maintain low levels of cytosolic Ca2+ under physiological conditions and the endoplasmic reticulum and mitochondria play a critical role in the regulation of cytosolic Ca2+ levels, especially in excitable neuronal cells (99). Previous studies have indicated that intracellular Ca2+ may play a role in the cellular (92,93) and mitochondrial (94) accumulation of MPP+. For example, Frei and Richter have shown that Ca2+ depleted mitochondria take up more MPP+ than Ca2+ loaded mitochondria (94). In agreement with these proposals, we also have observed that the absence of extracellular Ca2+ significantly increases the MPP+ and 4'I-MPP+ uptake into dopaminergic cells (Figure 12A). Interestingly, the MPP+ and 4'I-MPP+ uptake into non-dopaminergic HepG2 and HEK293 cells were not significantly

85 affected by extracellular Ca2+ (Figure 12A). In addition, pharmacological agents that interfere with cellular Ca2+ homeostasis inhibit MPP+ and 4'I-MPP+ uptake (Figure 12B), suggesting that extra/intra cellular Ca2+ may play a role in the uptake of MPP+. This hypothesis was further evaluated by measuring the effect of 4'I-MPP+ on the mitochondrial Ca2+ levels using Rhod-2

AM and by determining the effect of extracellular Ca2+ on the mitochondrial uptake of 4'I-MPP+

(because of the relatively higher rate of 4'I-MPP+ uptake into the mitochondria (Figure 9A), we chose to use 4'I-MPP+ in these experiments). The results of these experiments show that the mitochondrial uptake of 4'I-MPP+ is coupled with the mitochondrial Ca2+ efflux (Figure 13). In addition, the extra-mitochondrial Ca2+ was found to inhibit the 4'I-MPP+ uptake into isolated

MN9D cell mitochondria at physiologically relevant concentrations of 500-1000 nM (Figure

14A). Furthermore, the 4'I-MPP+ uptake was effectively inhibited by the mNCX inhibitors

[CGP37157 (23,24) and KBR7943 (128); Figures 15B and C], but not by the specific mitochondrial Ca2+ uniporter inhibitor, ruthenium red (133,196) or voltage gated Ca2+ channel inhibitors, verapamil (121) and benzamil (120,195) (Figure 15A). These findings together with the effective inhibition of the 4'I-MPP+-mediated typical mitochondrial membrane depolarization

(Figures 16A and B) by CGP37157 strongly suggest that mNCX may play an indirect role in the uptake of 4'I-MPP+ into the MN9D cell mitochondria. This proposal is further strengthened by the finding that CGP37157 effectively protects MN9D cells from MPP+ and 4'I-MPP+ toxicities

(Figure 17).

To further investigate the roles of NCX transporter and Ca2+ on 4'I-MPP+ uptake, a series of experiments were carried out with NCX transfected HEK293 cells. While the wild type

HEK293 cells are known to express very low levels of, or no NCX (205,206), MN9D and SH-

SY5Y cells were found to express high levels of NCX3 (Figure 18A). In addition, the 4'I-MPP+

86 uptake into wild type HEK293 cells was significantly lower than that of MN9D cells (Figures 8A and 19A). NCX transfected HEK293 cells showed a significant increase in the uptake of 4'I-

MPP+ (Figures 19A), with strong mitochondrial membrane depolarization in comparison to wild type cells (Figure 20). In addition, NCX transfected cells were more sensitive to 4'I-MPP+ toxicity (Figure 19B) in comparison to the wild type, strongly suggesting that NCX plays a role in the uptake and toxicity of MPP+.

Early studies show that mitochondrial PTP opening induced by matrix Ca2+ overloads, oxidative stress and depolarized mitochondrial membrane potential lead to apoptotic cell death

(100,101). Since MPP+ toxicity is also associated with the intracellular Ca2+, high oxidative stress, and mitochondrial membrane depolarization; a series of experiments were carried out with a well characterized mitochondrial PTP opening inhibitor, cyclosporine A to determine whether mitochondrial PTP plays a role in the uptake and toxicity of MPP+. These studies have revealed that the cellular 4'I-MPP+ uptake is moderately inhibited by cyclosporine A in the presence and no effect in the absence of extracellular Ca2+ (Figures 21B and C). These findings appear to indicate that the effects of Ca2+ on the uptake and toxicity of MPP+ and its derivatives are mediated by both NCX and PTP. However, the molecular relationship between cytosolic/mitochondrial Ca2+/mNCX/PTP and the mitochondrial accumulation and toxicity of

MPP+ or 4'I-MPP+ is not fully understood at present and is currently under investigation.

Taken together, the above findings demonstrate that the plasma and mitochondrial membrane potentials play crucial roles in the cellular and mitochondrial accumulation of MPP+ and its 4'-halogen substituted derivatives. The mitochondrial accumulation of these toxins which is a non-mediated simple diffusion process is tightly coupled with the cellular uptake. The mitochondrial uptake of MPP+ is associated with the efflux of mitochondrial Ca2+. The depletion

87 of intracellular Ca2+ (i.e. removal of Ca2+ from the incubation media and addition of EGTA) increases the overall cellular uptake of MPP+ by 2-3 fold. The effects of Ca2+ on the uptake and toxicity of MPP+ are mediated by the plasma membrane NCX, mNCX and the mitochondrial

PTP. Our findings that mNCX and mitochondrial PTP play a role in the mitochondrial accumulation and toxicity of MPP+ are intriguing and open a new direction for mechanistic studies involving the toxicity of MPP+ and related toxins. More importantly, the discovery that specific mNCX inhibitors protect dopaminergic cells from the MPP+ toxicity could be further utilized for the development of pharmacological agents to protect the CNS dopaminergic system from PD causing environmental toxins.

CHAPTER VI

CONCLUSIONS

Numerous previous in vivo and in vitro studies have shown that MPP+ selectively destroys dopaminergic neurons. Therefore, MPP+ has been extensively used as a convenient model to study the mechanisms of specific dopaminergic cell death in PD. The current model for the dopaminergic neuronal toxicity of MPP+ is centered on several key steps including specific uptake into dopaminergic cells through DAT, electrogenic accumulation of cytosolic MPP+ in the mitochondria, inhibition of complex I of the mitochondrial ETC leading to intracellular ATP depletion, increased ROS production and apoptotic cell death. Although many aspects of this model have been widely tested and accepted, numerous recent studies show that MPP+ is taken up not only into dopaminergic cells, but also into other cells through diverse transporters including OCT and PMAT, etc. On the other hand, most in vivo as well as in vitro studies to date have unequivocally confirmed that MPP+ is substantially more toxic to dopaminergic cells in comparison to other cell types. This suggests that critical aspects of the mechanism of this selective toxicity to dopaminergic cells are still poorly understood. Similarly, although electrogenic accumulation of cytosolic MPP+ in the mitochondria, followed by the inhibition of complex I of the mitochondrial ETC has been proposed as the primary cause of the toxicity, the molecular details of the mechanism of the mitochondrial accumulation of MPP+ has not been fully explored or understood. Since MPP+ is the most widely used current model for the study of environmental contributions to the pathophysiology of PD, a better understanding of the mechanisms of cellular/mitochondrial accumulation and the selective dopaminergic cell toxicity of MPP+ at the molecular level is of prime importance.

In the present study, we have synthesized and characterized a series of 4'-halogen substituted-MPP+ derivatives to further examine the specific dopaminergic toxicity of MPP+. All

MPP+ derivatives show cellular and mitochondrial uptake and toxicity profiles similar to that of

MPP+. Therefore, they are good mimics to study the mechanism of the specific dopaminergic toxicity of MPP+. In addition, we found that 4'I-MPP+ is fluorescent and it could be used in live cell imaging. Using these probes we show that MPP+ uptake into the cells occurs through multiple pathways including the plasma membrane transporters such as DAT, PMAT, or OCT or through a simple diffusion process. The cellular accumulation of these toxins is coupled with the mitochondrial uptake and both cellular and mitochondrial uptakes are driven by the plasma and mitochondrial membrane potentials. We have previously shown that the extra-cellular Ca2+ inhibits cellular MPP+ uptake. We also found that mitochondrial MPP+ uptake is coupled to the efflux of mitochondrial Ca2+ through the mNCX. While the present studies confirmed these findings, they further show that the effect of extra-cellular Ca2+ on the MPP+ uptake is mediated through the mitochondrial and plasma membrane NCX and the mitochondrial PTP. Our finding that mNCX and PTP play a role in the mitochondrial accumulation and toxicity of MPP+ is intriguing and opens a new direction for mechanistic studies involving the toxicity of MPP+ and related toxins. More importantly, the discovery that specific mNCX inhibitors protect dopaminergic cells from the MPP+ toxicity could be further utilized for the development of pharmacological agents to protect the CNS dopaminergic system from PD causing environmental toxins. However, the molecular relationship between cytosolic/mitochondrial

Ca2+/NCX/PTP and the mitochondrial accumulation and toxicity of MPP+ is not fully understood at present and is currently under investigation.

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