Raman Specroscopic Imaging Analysis of Signaling Proteins and protein Cofactors in Living Cells

RAMAN SPECTROSCOPIC IMAGING ANALYSIS OF SIGNALING PROTEINS AND PROTEIN COFACTORS IN LIVING CELLS

Achut P Silwal

A Dissertation

Submitted to the Graduate College of Bowling Green State University in partial fulfillment of

the requirements for the degree of

DOCTOR OF PHILOSOPHY

August 2018

Committee:

H Peter Lu, Adviser

Anita Simic Graduate Faculty Representative

John R Cable

Alexey T Zayak

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ABSTRACT

H Peter Lu, Advisor,

Proteins play a central role in biological processes. Broad classes of protein types are

involved in the innumerable functions of living systems, such as catalysis of the biochemical

reactions, transportation of the essential molecules, defense of the immune system, and the

transmission of messages from cells to cells. In our projects, we have combined the analytical

approaches including surface-enhanced Raman scattering (SERS), fluorescence microscopy,

electrochemistry, and computational methods to investigate the structures and functions of the

proteins and signaling molecules in the living cells. In our research, we have studied the

interactions and functions of dopamine transporters (DAT), dopamine receptors (DARs), and

several signaling molecules such as dopamine (DA), amphetamine (AMP), methamphetamine

(MAMP), and methylenedioxypyrovalerone (MDPV) in living cells. The interactions between

signaling molecules and DAT or DARs are crucial for the functioning of dopaminergic

pathways. In our project, we have probed interactions of signaling proteins including DAT and

DARs and investigated the changes that happen in signaling proteins, other interacting

compounds or intracellular contents of the second messenger such as cyclic adenosine

monophosphate (cAMP). The second messenger like cAMP is important in many biological

processes which are produced due to interactions of the signaling protein in living cells. Our

studies on DA-DAT or DA-DARs interactions mainly utilize the Raman spectroscopy to

characterize the selectivity and efficacy of psychopharmaceutic drugs. In addition, we have also

studied the redox states and mechanism of protein cofactors in different experimental conditions.

We have probed and characterized the redox states and mechanisms of FMN cofactors in

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biological and non-biological environments using electrochemistry, SERS measurements, and

computational measurements. The results obtained from our research could be useful in the

diagnosis of abnormalities or diseases originated from malfunctioning of proteins or protein-

ligand interactions.

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To my friends and family

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ACKNOWLEDGMENTS

Firstly, I would like to express my deepest gratitude to my advisor Prof. Dr. H. Peter Lu

for being a caring person and constant source of support, ideas, and inspiration throughout my

Ph.D. research.

Besides my advisor, I would like to thank my committee members: Dr. John R. Cable,

Dr. Alexey T. Zayak, and Dr. Anita Simic for their valuable time, insightful comments, and

encouragement.

I would also like to thank Dr. Jon E. Sprague, Director of the Ohio Attorney General’s

Center for the Future of Forensic Science at BGSU for his help and collaboration. I would also

like to thank Dr. Andrew Torreli for the stimulating discussion on protein biology.

My sincere thanks go to present and past members of my research group; especially Dr.

Yufan He and Dr. Takashige Fujiwara for their helpfulness and teaching. I would also like to

thank Dr. Bharat Dhital, Dr. Dibeyndu K. Sasmal, Dr. Rajeev Yadav, Dr. Nibedita Pal, Dr.

Vishal Govind Rao, Dr. Zijian Wang, Dr. Qing Guo, Dr. Maolin Lu, and Min Gu for all research

techniques and ideas they taught me.

I am grateful to my fellow friends Meiling Wu, Susovan Roy Chowdhury, Sunidhi

Jaiswal, and Lorena Alvarez for their friendship, supports, and stimulating discussions.

I am thankful to the Bowling Green State University (BGSU); Department of Chemistry

and Center for Photochemical Sciences; the Ohio Eminent Scholar endowment fund, and the

Ohio Attorney General’s center for the future of Forensic Science for funding.

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I am thankful to all my professors at BGSU for providing me the best education. I am

also thankful to staffs at the Center for Photochemical Sciences and the Department of Chemistry

particularly Nora R. Cassidy, Alita Frater, Hilda E. Miranda, Mary Toth, Charles Codding, and

Doug Martin for their kind help.

Last but not the least, I would like to thank my family: my parents and siblings for their

love and support. I am very thankful to my beloved wife, Rakshya Khatiwada, for being such a

wonderful person and supporting me all the time with love and understanding. I am thankful to

all who helped me unconditionally to make this work possible and enjoyable.

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TABLE OF CONTENTS

Page

CHAPTER 1. INTRODUCTION ......................................................................................... 1

1.1. Biological significances of proteins .................................................................... 1

1.2 Signaling proteins ................................................................................................ 2

1.3. Dopamine in the central nervous system (CNS) ................................................. 3

1.3.1 Mechanism of dopaminergic signaling ................................................. 5

1.3.2 Dopamine transporter ............................................................................ 7

1.3.3 Dysfunction of dopamine transporter and neurodegenerative diseases 11

1.3.4 Dopamine receptors .............................................................................. 11

1.4 Flavoproteins ........................................................................................................ 13

1.4.1 Flavin cofactors ..................................................................................... 15

1.4.2 Electron transfer process in Flavin coenzymes ..................................... 16

1.5 Raman spectroscopy ............................................................................................ 18

1.5.1 Surface-enhanced Raman spectroscopy (SERS) .................................. 19

1.5.2 Electromagnetic mechanism ................................................................. 20

1.5.3 Chemical enhancement mechanism ...................................................... 24

1.6 References ............................................................................................................ 25

CHAPTER 2. EXPERIMENTAL SECTION ........................................................................ 43

2.1 Theoretical background ....................................................................................... 43

2.1.1 Raman spectroscopy ............................................................................. 43

2.1.2 Resonance Raman spectroscopy ........................................................... 45

2.1.3 Surface-enhanced Raman spectroscopy ................................................ 46

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2.1.4 Two-photon excited (2PE) fluorescence microscopy ........................... 48

2.1.5 Transmission electron microscopy (TEM) ........................................... 50

2.1.6 Confocal microscopy ............................................................................ 53

2.2 Experimental set-ups ............................................................................................ 54

2.2.1 Raman measurements and optical imaging ........................................... 54

2.2.2 Electrochemical control and Raman measurement ............................... 55

2.2.3 Two-photon excitation (2PE) fluorescence imaging ............................ 57

2.2.4 Two-photon excited (2PE) fluorescence microscopy ........................... 58

2.3 Density functional theory calculations ................................................................. 59

2.4 Materials and sample preparation ........................................................................ 60

2.4.1 Synthesis of silver nanoparticles ........................................................... 60

2.4.2 Fabrication of silica shell on silver NP ................................................. 61

2.4.3 Preparation of Britton-Robinson (B-R) buffer ...................................... 62

2.4.4 HEK293 cell culture ............................................................................. 62

2.4.5 Preparation of hDAT inducible HEK293 cells ..................................... 62

2.4.6 Bacteria growth and Plasmid amplification .......................................... 63

2.4.7 Transfection of pcDNA3.1-hDAT in HEK293 cell .............................. 64

2.5 References ............................................................................................................ 65

CHAPTER 3. RAMAN SPECTROSCOPIC SIGNATURE MARKER OF DOPAMINE-

HUMAN DOPAMINE TRANSPORTER INTERACTION IN LIVING CELLS ................ 75

3.1 Introduction .......................................................................................................... 76

3.2 Experimental sections .......................................................................................... 83

3.2.1 Synthesis of silver nanoparticles and sample preparation .................... 83

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3.2.2 Surface-enhanced Raman measurements .............................................. 84

3.2.3 Two-dimensional SERS plot vs relative signal peak intensity ............. 84

3.2.4 Density functional theory calculations .................................................. 85

3.3 Results and discussion ......................................................................................... 85

3.4 Summary ............................................................................................................ 95

3.5 References ........................................................................................................... 96

CHAPTER 4. MODE-SELECTIVE RAMAN IMAGING OF DOPAMINE-HUMAN

DOPAMINE INTERACTION IN LIVE CELLS .................................................................. 109

4.1 Introduction .......................................................................................................... 110

4.2 Experimental sections .......................................................................................... 114

4.2.1 Synthesis of silica-coated silver nanoparticles ..................................... 114

4.2.2 HEK293 cells culture ............................................................................ 115

4.2.3 Preparation of hDAT inducible HEK293 cells ..................................... 115

4.2.4 Transfection of pcDNA3.1-hDAT in HEK293 cell .............................. 116

4.2.5 Two photons excited (2PE) fluorescence imaging ............................... 116

4.2.6. Surface-enhanced Raman measurements ............................................. 116

4.2.7 Mode-selective Raman measurement ................................................... 117



4.3.1 Conventional and mode-selective Raman measurement ...................... 118

4.3.2 Two-photon excited (2PE) fluorescence imaging approach ................. 124

4.3.3 Correlation between mode-selective Raman image and spectra ........... 128

4.4 Summary ............................................................................................................ 130

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4.5 References ............................................................................................................ 131

CHAPTER 5. RAMAN SPECTROSCOPIC ANALYSIS OF SIGNALING MOLECULES-

DOPAMINE RECEPTORS INTERACTIONS IN LIVING CELLS ................................... 139

5.1 Introduction .......................................................................................................... 140

5.2 Experimental section ............................................................................................ 145

5.2.1 Transfection of DRD1 and DRD2 DNA in live cells ........................... 145

5.2.2 Synthesis of silica-coated silver nanoparticles ..................................... 147



5.3.1 Internalization of silver nanoparticles ................................................... 148

5.3.2 Probing of intracellular cAMP in HEK293 cells .................................. 150

5.3.3 Probing of intracellular cAMP in HT22 cells ....................................... 153

5.3.4 Raman peaks assignment of cAMP ...................................................... 160

5.3.5 Raman peaks assignment of DRD1-HEK293 cells .............................. 165

5.4 Summary ............................................................................................................ 166

5.5 References ............................................................................................................ 167

CHAPTER 6. RAMAN SPECTROSCOPY PROBING OF REDOX STATES AND

MECHANISM OF FLAVIN COENZYME .......................................................................... 179

6.1 Introduction .......................................................................................................... 179

6.2 Experimental section ............................................................................................ 189

6.2.1 Synthesis of silver nanoparticles and sample preparation .................... 189


6.2.3 Surface-enhanced Raman measurements and electrochemical control 190

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6.2.4 2-D distribution SERS Plot vs conventional SERS spectrum .............. 191


6.4 Summary ............................................................................................................ 210

6.5 References ............................................................................................................ 211

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LIST OF FIGURES

Figure Page

1.1 Dopaminergic pathways in the central nervous system (CNS) of human ................. 4

1.2 Schematic representation of dopamine system in neuron cell ................................... 6

1.3 Schematic representation of dopamine system in neuron cells ................................. 8

1.4 D1 and D2 dopamine receptor ................................................................................... 12

1.5 Skeletal formula of Flavin mononucleotide and Flavin adenine dinucleotide .......... 16

1.6 The crystal structure (PDB 1 tll) of the neuronal nitric-oxide synthase (nNOS) ....... 17

1.7 Prolate spheroid to show the model proposed by Zeeman and Schatz ...................... 23

2.1 The schematic representation of Rayleigh, Stokes and anti-Stokes scattering .......... 44

2.2 The schematic representation of resonance Raman scattering .................................. 46

2.3 The schematic representation of the (A) electromagnetic enhancement in SERS

measurements ............................................................................................................ 47

2.4 The schematic representation of two-photon excitation fluorescence microscopy ... 49

2.5 Schematic representation of TEM in diffraction modes ............................................ 51

2.6 Schematic representation of TEM in imaging mode ................................................. 52

2.7 Confocal microscopy geometry including the light pathways ................................... 53

2.8 The schematic representation of the experimental set-up used in Raman measurement

and optical imaging .................................................................................................... 55

2.9 The schematic representation of the home-modified experimental setup for the

electrochemical redox process and Raman spectroscopy .......................................... 56

2.10 The percentage transmittance of dichroic mirror (chroma, ZT532rdc) and

bandpass filter (FES0500) .......................................................................................... 57

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2.11 Schematic representation of the optical setup for the two-photon excited (2PE)

fluorescence and Raman imaging .............................................................................. 58

2.12 The comparison of experimental Raman spectrum (dotted black) of dopamine with

DFT calculated Raman spectrum ............................................................................... 59

2.13 The plasmon resonance related to the bare silver nanoparticles (AgNPs) and silica-

coated silver nanoparticles (AgNP@SiO2) ................................................................ 61

3.1 Schematic representation of dopamine system in neuron cell showing hDAT, DA,

and DA receptor, (B) HEK293 cell, and (C) silica-coated silver nanoparticles ........ 77

3.2 DFT calculated and experimental Raman spectrum of DA and cell samples ............ 80

3.3 2D-distribution Raman spectrum of (A) DA, (B) HEK293, (C) hDAT-HEK293,

(D) DA-HEK293, (E) DA-hDAT-HEK293 cell ........................................................ 87

3.4 (A) Crystal structure of dDATmfc (PDB ID: 4xp1) .................................................... 89

3.5 Characteristic Raman peaks at 807 cm-1 and 1076 cm-1 of integrated Raman

spectrum of the DA-hDAT-HEK293 cell .................................................................. 91

3.6. The effect in vibrational mode at 807 cm-1, and 1057 cm-1 as bupropion interferes

DA-hDAT interaction after adding bupropion into DA-hDAT-HEK293. ................ 93

3.7 SERS spectra from Bupropion ................................................................................... 94

4.1 The two-photon excited (2PE) fluorescence images from baseline HEK293 cells ... 111

4.2 The Raman spectrum and mode-selective Raman spectrum from DA and baseline

HEK293 cells ............................................................................................................ 118

4.3 The Crystal structure of dDATmfc (PDB ID: 4XP1) .................................................. 120

4.4 Two-photon excited (2PE) fluorescence and mode-selective Raman images of

hDAT-HEK293 cells. ................................................................................................ 122

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4.5 Evidences for the two-photon excited (2PE) emission .............................................. 125

4.6 The consistence between conventional and mode-selective SERS spectra ............... 126

4.7 Summary of the comparative observations from mode-selective Raman

measurements on fifty hDAT expressed HEK293 cells after addition of DA. .......... 128

4.8 The crystal structure of dopamine transporter (PDB ID: 4XP1) showing the binding

site of dopamine ......................................................................................................... 128

4.9 The results from DFT calculation showing difference between unbound and bound

states of DA. ............................................................................................................ 130

5.1 (A) Homology model of D1 dopamine receptors; (B) Structure of the D2 dopamine

receptor bound to the atypical antipsychotic drug risperidone (PDB ID 6cm4.pdb) . 141

5.2 The DIC image of DRD1 over-expressed HEK293 cells in DA solution of 0.2 µM

concentration showing the internalization of AgNP@SiO2 in cells and mode-

selective Raman imaging ........................................................................................... 149

5.3 The Experimental and DFT calculated Raman spectrum of cAMP .......................... 150

5.4 The consistence of occurrence between cAMP-marker SERS image and peaks ...... 151

5.5 Integrated SERS spectra (left) and 2-dimensional distribution SERS spectra (right)

obtained from the DRD1 expressed HEK 293 cells .................................................. 154

5.6 The DFT calculated Raman spectrum of signaling compounds ................................ 155

5.7 The integrated experimental SERS spectrum obtained from DRD1 expressed

HEK293 cells in the absence (red) and presence (green) of signaling compounds ... 156

5.8 The integrated experimental SERS spectrum obtained from DRD2 expressed

HEK293 cells in the absence (red) and presence (green) of signaling compounds. .. 157

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5.9 The integrated experimental SERS spectrum obtained from DRD1-DRD2

co-expressed HEK293 cells in the absence (red) and presence (green) of

signaling compounds ........................................................................................... 158

5.10 The integrated experimental SERS spectrum obtained from baseline HEK293 cells

in the absence (red) and presence (green) of signaling compounds. ......................... 159

5.11 Integrated SERS spectra (left) obtained from the DRD1 expressed HT22 cells in

different experimental environments. ........................................................................ 160

5.12 The integrated SERS spectra and 2-d distribution SERS spectra due to interaction

of signaling compounds with different types of cell samples. ................................... 163

5.13 The experimental SERS spectrum of cAMP with major peaks ................................. 164

5.14 The experimental SERS spectrum obtained from cell samples in the absence of

signaling compounds ................................................................................................. 164

6.1 The DFT Raman spectra from different redox states of Flavin mononucleotide ...... 180

6.2 Zoom in view of integrated SERS spectra of Flavin mononucleotide in B-R buffer

at different pH environments ..................................................................................... 182

6.3 Schematic representation of the inner membrane of mitochondria showing the

involvement of FMN cofactor in electron transport process during ATP synthesis. . 183

6.4 The experimental SERS spectrum of FMN at the redox-sensitive region in pH 11,

pH 9, pH 7.6, pH 7.4, pH 7, pH 5, and pH 3. ............................................................ 184

6.5 (A) The UV-Vis absorption spectrum, and (B) emission spectrum of FMN (10 µM)

in B-R buffer solution. ............................................................................................... 187

6.6 The Cyclic voltammetry curve obtained from the mixture of FMN (50nM) and

AgNP@SiO2 in B-R buffer ........................................................................................ 196

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6.7 Two-dimensional (2-D) distribution Raman spectrum of FMN in acidic and

alkaline medium ......................................................................................................... 197

6.8 Two-dimensional (2-D) distribution Raman spectrum of FMN at pH 3 and 5 ......... 198

6.9 Two-dimensional (2-D) distribution Raman spectrum of FMN at pH 7 and 7.4 ...... 199

6.10 Two-dimensional (2-D) distribution Raman spectrum of FMN at pH 7.6 and 9 ...... 200

6.11 Two-dimensional (2-D) distribution Raman spectrum of FMN at pH 11 ................. 201

6.12 The calculated HOMO-LUMO energy state of different redox species of FMN ...... 204

6.13 The consistency of experimentally collected SERS peak position and DFT

calculated Raman peak position of different redox states of FMN ........................... 206

6.14 The experimental results from the study of FMN in nNOS of rat (Rattus norvegicus)

using combined approach of cyclic voltammetry and SERS technique .................... 207

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LIST OF TABLES

Table Page

3.1 Prominent Raman wavenumber of DA, HEK293 cell, and hDAT-HEK293 cells .... 82

3.2 Raman peak assignment of dopamine (DA) .............................................................. 90

3.3 Raman peak assignment of hDAT-HEK293 cell ....................................................... 92

5.1 The assignments of experimental SERS peaks obtained from pure cAMP ............... 162

5.2 The assignment of SERS peaks obtained from DRD1-HEK293 cells. ..................... 166

6.1 DFT calculated and experimental Raman wavenumber of the redox-sensitive mode

of different redox states of FMN with the energy of its HOMO and LUMO ............ 192

6.2 Redox potential of FMN (50nM) in different pH of Britton-Robinson buffer at

optimal scan rate 0.1 V/S ........................................................................................... 193

6.3 The Raman assignment of different redox states of FMN coenzymes ...................... 194

1

CHAPTER 1. INTRODUCTION

1.1. Biological significances of proteins

Proteins are biomolecules which contain one or more long chains of amino acids.

Proteins are differentiated base on their sequence of amino acids. The sequence of 20 standard

amino acid in proteins make the sequence of genes which serve as the genetic code in humans.

The residues in a protein are modified by the post-translational modification which alters the

properties and functions of proteins. The non-peptide groups attached to the proteins are called

prosthetic groups or cofactors. In mammalian cells the average half-life period of the proteins

is 2-3 days; the half-life period is the unit of proteins’ lifespan which varies from minutes to

several years. The aged or degraded proteins in living organisms are recycled by the

biochemical process which is called turnover process. Proteins are one of the most abundant

biomolecules in living cells. It is reported that smaller size bacterial cells like mycoplasma or

spirochetes contain 50,000 to 50×106 proteins molecules while larger and complex cells like

human cells contain 1-3×108 molecules of proteins.1 Even though different types of proteins

molecules are expressed in a single cell, a cell is not capable to express all gene coding

proteins. The number of proteins genome in the certain organism depends on its complexity.

The number of protein genome in a superior organism is higher than that of an inferior one.

For example, the number of proteins encoded by human genome is about 20,000 while the

average number of proteins encoded by bacterial genome is about 2358.2,3

Proteins play the central role in many biological processes like catalyzing the biochemical

reactions, transport of essential molecules, defend the immune system and transmit messages

from cells to cells.4-6 The immunoglobulin helps to combat pathogens, histone and cohesin

2

regulate gene expression and chromosome structure during cell division,7-9 actin and myosin

are involved in muscle contraction and movement,10-12 collagen and elastin provide support to

bodies,13 insulin controls blood sugar concentration by regulating the uptake of glucose into

cell, and hemoglobin transports the oxygen to the cells. Similarly, the enzyme protein helps the

metabolic process to occur at a precise rate in cells.14 Based on their functions, proteins in the

human body are classified into several groups including structural proteins, regulatory

proteins, transport proteins and signaling proteins. This dissertation is mainly dedicated to the

interaction of signaling proteins like dopamine receptors or transporters and the cofactor of

regulating proteins like flavoproteins in living cells.

1.2. Signaling proteins

Cells collect the signals from outside environments and process them to respond the

external changes. Special types of membrane proteins of cells, which are well recognized as

signaling proteins or receptors, are responsible to initiate the physiological responses due to

changes in environmental factors. The cells are not only the targets of outside signals, but also

the origin of the signals. As we mentioned before, the membrane proteins bind to the

signaling molecules and produce physiological responses. The nerve growth factors,

hormones, and neurotransmitters are commonly recognized signaling molecules in

multicellular organisms. Some signaling molecules such as neurotransmitters could have effect

for very short distances. For example, the dopamine and glycine neurotransmitter have effect

only for the few nanometers. However, some types of signaling molecules could have effect

for very long distance. For instance, the follicle stimulating hormones is produced by the

pituitary gland in the brain and can stimulate the follicle cells in the mammalian ovary.

The receptors are the signaling proteins which bind to signaling molecules like nerve

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growth factor, hormones, and different types of neurotransmitters to initiate the physiological

responses. Hundreds of thousand types of receptors are found in a single cell and the

populations of these receptors could vary depending on cell types. The receptors are

transmembrane proteins; the signaling molecules bind to the extracellular domain of the

receptors and start the sequences of the molecular switches and transmits the signal to the

internal signaling pathways. The interactions between receptors and signaling molecules are

important for the communication between cells. The receptors are also sensitive to the

environmental factors like light, temperature, and pressure outside of the cells. The receptors

are classified into three broad classes: (1) G protein-coupled receptors (GPCR), (2) Ion-

channel receptors (ICR) and (3) enzyme-linked receptors (ELR). In this dissertation, we

mainly are dedicated to probing the interaction of dopamine transporter or dopamine receptors

in living cells. Dopamine receptors are the members of G protein-coupled receptors (GPCR).

1.3. Dopamine in the central nervous system (CNS)

Dopamine (DA) stands for 3,4-dihydroxyphenethylamine which belongs to

catecholamine neurotransmitter. It controls many psychological and behavioral activities in the

mammalian central nervous system (CNS) through biochemical interactions with the

dopamine transporter (DAT) and dopamine receptors (DARs).15,16 The property of dopamine

as a neurotransmitter was discovered by Arvid Carlsson in 1957.17 Since then, DA

neurotransmission and signaling mechanisms have been established as a major field of the

research in neuropharmacology and

4

Figure 1.1. Dopaminergic pathways in the central nervous system (CNS) of human(top) and the

skeletal molecular structure of dopamine neurotransmitter (bottom).

psychiatry. Many antipsychotic drugs in modern neurology and psychiatry such as L-DOPA

and Ritalin target dopamine system in the brain. Dopaminergic neurons located in the ventral

tegmental area, substantia nigra pars compacta, and the arcuate nucleus of the hypothalamus

enzymatically convert tyrosine (Tyr) into L-DOPA and finally into DA.18-21 The sets of the

neurons in the brain which are involved in the synthesis and release of dopamine

neurotransmitters are called dopaminergic pathways. The nigrostriatal pathway, mesolimbic

pathway, mesocortical pathway, and tuberoinfundibular pathway are four commonly

recognized dopaminergic pathways in the human brain. All different dopaminergic pathways

are specific in their functions, for example, voluntary movement in a body is regulated by

nigrostriatal dopaminergic pathway. The depletion of this pathway is related to symptoms of

Parkinson’s diseases including tremor, rigidity, and postural imbalance. The nigrostriatal

pathway projects dopamine from the substantia nigra pars compacta (SNpc) of the midbrain to

5

the basal ganglia of dorsal striatum. The mesocortical and mesolimbic pathway are collectively

called mesocorticolimbic pathway which is responsible for the reward-related behaviors. The

mesolimbic pathway projects DA from the ventral tegmental area (VTA) to the ventral

striatum (nucleus accumbens, NAc) and mesocortical pathway extends from ventral tegmental

area (VTA) to the prefrontal cortex (PFC).22 The tuberoinfundibular pathway extends from

arcuate nucleus of the hypothalamus to the pituitary gland23-27 which controls the secretion of

specific hormones like prolactin from the anterior part of the pituitary gland.28,29 When

tuberoinfundibular pathways are blocked by some antipsychotic drugs, the prolactin level in

blood is increased which causes hyperprolactinemia. The hyperprolactinemia is related to

abnormal lactation, disruptions to the menstrual cycle in women, visual problems, headache

and sexual dysfunction. In addition to above-mentioned regions in the brain, DA is also

synthesized in the retina and olfactory bulb which helps in vision and odor processing.30,31

1.3.1. Mechanism of dopaminergic signaling

The DA neurotransmitter works in a sequence of several steps. When the action

potential is generated on neuron membrane from environmental stimulation, it causes the

opening of voltage-gated Ca2+ ions channel followed by the entry of Ca2+ ions into the neuron

cell, which induces the release of DA into the synaptic cleft. 32-34 Then, DA in the synaptic

cleft activates dopamine receptors (DARs) located in pre- or post-synaptic region to generate

the dopaminergic response.35-40 Dopamine receptors are primarily activated by dopamine and

produce dopaminergic signaling in the mammalian brain. Dopamine receptors have several

roles in human such as cognition, motivation, memory, motor control, and modulation of

neuroendocrine activities. Mainly, five kinds of human dopamine receptors: D1, D2, D3, D4,

and D5 are reported which are classified into D1 like (D1/D5) and D2 like (D2/D3/D4)

6

dopamine receptors based on sequence homology and functions. D1 like dopamine receptors

couple with Gs/olf subunit of heterotrimeric G protein to stimulate synthesis of cyclic

adenosine monophosphate (cAMP) while D2-like receptors couple with Gi/o/z subunit of G

protein and inhibit synthesis of cAMP. When D1 like dopamine receptors binds to signaling

molecules, the intracellular level of cAMP is increased which activates the enzyme protein

kinases A (PKA) and phosphorylates multiple protein substrates by donating phosphate

groups. Each step in the cascade further amplifies the initial signal, and the phosphorylation

reaction mediates both short and long-term response in the cell phosphodiesterase.

Phosphorylation allows for complicated control of protein functions. The cyclic adenosine

monophosphate (cAMP) is an important secondary messenger which has several roles

Figure 1.2. (A) Schematic representation of dopamine system in neuron cellshowing hDAT, DA,

and DA receptors;(B) dopamine receptors in “off” state before interaction to DA; (C) dopamine

receptors in “on” states after interaction to DA.

7

including regulation of downstream proteins such as kinases,41 ion channels,42-45and

transcription factors.46-48 Although a high degree of similarities are found between DARs of

the similar subfamily, each kind of dopamine receptors are distinct from each other. DARs are

expressed heterogeneously in the cells, therefore it is difficult to target them and uncover their

roles separately. The termination of G protein signaling and the initiation of G protein-

independent signaling is also regulated by DARs.49,50 51D1 like DARs are also known to

interact with NMDA and GABA receptors via their intercellular loops and C-termini.52-54 The

characterization of DARs has great importance to understand the synaptic and neural circuit

action of many signaling compounds including DA.

1.3.2. Dopamine transporter

High concentration of DA in the synaptic cleft causes over-stimulation of DARs

resulting several neurological and physiological disorders. The excessive neurotransmission

due to the accumulation of DA as well as other biogenic amino neurotransmitters like

serotonin, noradrenaline, and 𝛾-aminobutyric acid present in synaptic and perisynaptic space is

removed by the reuptake process mediated by dopamine transporters (DAT).17,55-57 DAT

belongs to the family of solute carrier 6 (SLC6) 58 which maintains homeostasis of DA

neurotransmitters by its Na+ and Cl- ions assisted reuptake processes. Therefore, the dopamine

transporter is also known as neurotransmitter sodium symporters (NSS). 59-63 Dysfunction of

NSS system is associated with several disorders like schizophrenia, depression64, attention

deficit hyperactivity disorder (ADHD)65, orthostatic intolerance66, epilepsy67, Parkinson’s

disease and infantile Parkinsonism dystonia.68 The agonists and antagonists of DARs has

influence in dopaminergic transmission by enhancing or blocking the actions of DA on

receptors.69-74

8

Figure 1.3. Schematic representation of dopamine system in neuron cells showing DAT, DA,

and DA receptors.The minimal construct drosophila dopamine transporter (dDATmfc; PDB ID:

4xp1) crystal structure and binding site of DA are shown in the red circle.

Inhibition of DA reuptake from synaptic cleft has several side effects, but it is applied as an

important pharmacological method for the treatment of depression.75,76 The targeted binding of

drugs to the human dopamine transporter (hDAT) has significant effect in the development

and function of the nervous system; thus, the study of DA-hDAT interaction has great

importance in the neurophysiology field.

Pharmacological properties of transporters have been a focus in the field of

neuroscience after Julius Axelrod first reported the involvement of the transporter in the

reuptake process of neurotransmitters.77 However, genes codes of transporter proteins were

first identified in the early 1990s.78,79 The identification of the SLC6 (solute carrier 6) gene

family which includes dopamine transporters (DAT) and serotonin transporter (SERT) were

discovered through homology screening in mammalian cDNA libraries. 80-83 The human

9

dopamine transporter (hDAT) encodes SLC6A3 gene which consists 15 exons and 14 introns.

It spans over 64 kbp and maps chromosome 5p15.3.84 The hydrophobicity analysis of the

many neurotransmitters including dopamine transporter revels that they contain cytoplasmic

amino and carboxyl termini, twelve transmembrane domains (TMDs), and one large

glycosylated loop between TMD3 and TMD4.82-86 The driving force for DAT-mediated

reuptake is the sodium concentration gradient across the plasma membrane generated by

Na+/K+ ATPase.87,88 The secondary structure of DAT has been elucidated by several

biochemical and mutagenesis studies. The binding of substrate or inhibitors to the residues of

transporter were studied using cysteine accessibility method (SCAM). Cysteine 342 residue is

supposed to be associated with cytoplasmic gating since it was more reactive to the thiol-

modifying agent during uptake process.89 The mutation of intracellular tyrosine to

alanine(Y335A) stops reuptake process of DA which further supports the role of intracellular

loop for the substrate translocation.90 The dopamine binding site in the dopamine transporter

has been analyzed by mutating aromatic and acidic amino acids in transmembrane domains.

The mutation of phenylalanine155 in TM3 domain to alanine and the replacement of residues

like serine and aspartate TM1 and TM7 domains significantly reduce the uptake of dopamine

which shows that these residues are essential for the substrate recognition.91,92 The sequence

homology and functional similarities of the high-resolution crystal structure of leucine

transporter (LeuT) to SLC6 transporters lead to the tertiary structure of DAT. 93 The crystal

structure of LeuT shows that transmembrane domains 1, 2, 3, 6 and 8 form the central

substrate binding pockets. The transport process of leucine is associated with three states of

transporter which are outward-facing, occluded and inward-facing states. Leucine and sodium

are bound in an occluded state of leuT which is free from water. However, this model has

10

some limitation to explain the mechanism of dopamine transporter due to certain

dissimilarities found between some parts of SLC6 and LeuT including substrate selectivity and

transport inhibition by addictive drugs. The solid information about the atomic level

interaction of substrate and hDAT was first reported by Gouaux and coworkers by X-ray-

crystallography based study of the dopamine transporter (dDATmfc). 94,95 They have explained

the molecular principle to distinguish the binding pattern of chemically distinct ligands to

dDATmfc near sodium and chloride ions. They showed that the open conformation of dDATmfc

is stabilized by the sodium and chloride ion coupled binding of compounds like nortriptyline,

dopamine, cocaine, and amphetamine. They have characterized central binding pocket of

dDATmfc into three subsites based on ligand specificity. Substrates including DA and its

analog are bound to DAT before full closure of the extracellular gate. In contrast, inhibitors

like cocaine are bound to the outward-facing DAT, acting like a wedge to block DAT in an

outward-open conformation. In addition to transport mechanism of DAT, it also has channel-

like activities. The voltage clamp studies in hDAT expressed Xenopus laevis oocytes show a

constitutive leak current which is different from the transport-related current in terms of ions

and voltage dependence.96,97 The transport current and constitutive leak current are associated

to the regulation of DA neuron excitability, it is reported that DA and amphetamine could

increase the firing rate of DA neurons. 98,99

In summary, dopamine transporter is a member of SLC6 gene which is considered as a

symporter since it clears out monoamine neurotransmitters from the synaptic cleft by the

sodium and chloride assisted reuptake process. It is the primary target for the psychostimulants

drugs like cocaine and amphetamines where cocaine is a competitive inhibitor that binds DAT

at its outward facing state, whereas amphetamines is a DAT substrate that could be transported

11

through DAT and subsequently lead to DA release. Therapeutic drugs like methylphenidate

and amphetamine/dextroamphetamine directly target DAT for treating certain types of

neuropsychiatric disorders.

1.3.3. Dysfunction of dopamine transporter and neurodegenerative diseases

The depletion of the dopamine transporter (DAT) in neurons causes hyperlocomotion

problems.100 The decrease of DAT reduces the rate of DA reuptake from the extracellular

space, which is associated to hyperlocomotion activity. Moreover, it is also reported that when

DAT is decreased in dopaminergic pathways the release of DA is also reduced. The analysis of

DA content in the dorsal striatum of knock out (KO) and wild-type (WT) mice shows that the

DA levels in KO mice are 95% smaller than WT mice.101 These experimental results show that

dopamine transporter has the crucial role to maintain DA homeostasis in the synaptic and

presynaptic regions. The malfunctioning of DAT is related to neurodegenerative diseases and

neuronal plasticity. The rate of DA uptake by DAT has a big effect on the amount of DA in the

cells. The depletion of DAT in cells causes cognitive deficits, motor abnormalities, and

hyperactivity which are the symptoms of the ADHD.102 The increased activity of DAT is

found to be related to several disorders including clinical depression.103 The ability of

dopamine agonist to stimulate the symptoms of bipolar disorder shows that dysfunction of

DAT is also associated to this disease.

1.3.4. Dopamine receptors

As we explained above, the signaling protein fall into three major classes: G-protein-

coupled receptors, ion channel receptors, and enzyme-linked receptors.104-109 Dopamine

receptors belongs to the types of G-protein coupled receptors (GPCR). GPCRs is the largest

12

Figure 1.4. (A) D1 dopamine receptor and (B) D2 dopamine receptor.

superfamily of membrane protein which performs many biological activities. The membrane

proteins which couple with heterotrimeric G protein are called G protein-coupled receptors.

They are activated by various types of signaling molecules such as hormones, pheromones,

odorants, and neurotransmitters. Dopamine receptors are primarily activated by dopamine and

produce dopaminergic signals in mammalia. Dopamine receptors have several roles in human

such as cognition, motivation, memory, motor control, and modulation of neuroendocrine

activities. Five kinds of human dopamine receptors (D1, D2, D3, D4, and D5) are reported

which are classified into D1 like (D1/D5) and D2 like (D2/D3/D4) dopamine receptors based

on sequence homology and functions. D1 like dopamine receptors couple with Gs/olf subunit

of heterotrimeric G protein to stimulate synthesis of cyclic adenosine monophosphate (cAMP)

while D2-like receptors couple with Gi/o/z subunit of G protein to inhibit synthesis of cAMP.

Although a high degree of similarities are found between DARs of the similar subfamily, they

are distinct from each other. When dopamine or other biogenic amino-neurotransmitters binds

to the dopamine receptors, it undergoes a conformational change, which introduces a series of

13

biochemical changes within cells. The activation of dopamine receptors by signaling

molecules trigger the changes in intracellular contents of second messengers such as adenosine

monophosphate (cAMP), diacylglycerol (DAG), and inositol 1,4,5-trisphosphate (IP3). The

interaction of D1 types of dopamine receptors with its agonist has the stimulatory effect on

adenylate cyclase activity and increases the intracellular cAMP level. On the other hand, the

interaction of D2 types dopamine receptors with its agonist has the inhibitory effect on

adenylate cyclase activity and reduces the intracellular cAMP level36,39,72,110-115. The D1 like

receptors includes D1 and D5 receptors which are 80% identical in terms of their

transmembrane (TM) domains. D2 like receptors includes D2, D3, and D4 receptors, where

D2 and D3 receptors are 75% identical, and D2 and D4 receptors are 53% identical.114 High

concentration of DA in the synaptic cleft causes over-stimulation of DARs resulting several

neurological and physiological disorders. The excessive neurotransmission due to the

accumulation of DA as well as other biogenic amino neurotransmitters like serotonin,

noradrenaline, and 𝛾-aminobutyric acid in synaptic and perisynaptic space is regulated by

dopamine transporter through Na+ and Cl- ions assisted reuptake process.17,55-57

1.4. Flavoproteins

The flavoproteins are proteins which commonly contains flavin coenzymes like flavin

adenine dinucleotide (FAD) and flavin mononucleotide (FMN) as the prosthetic group. The

requirement of prosthetic group for the activity of flavoprotein was first observed by Hugo

Theorell and coworkers in 1935.116 They separated apoprotein and a bright-yellow pigments

from a yeast protein (flavoproteins) and used these components separately for the oxidation of

NADH coenzyme where neither apoprotein nor a yellow pigment alone were able to catalyze

the oxidation of NADH coenzyme. When yellow pigments were combined back to apoproteins

14

then the oxidizing property was restored; this evidence shows that prosthetic group is required

for the functioning of flavoproteins. When yellow pigments of protein were replaced by

riboflavin, it was not functional which show that the yellow pigments were not riboflavin.

Later studies clearly show that prosthetic group present in flavoproteins were either FMN or

FAD and reported that FAD and FMN are deeply buried in the flavoproteins. 117-122 Some

flavoproteins contains covalently linked FAD or FMN which have stronger redox power. Most

of the flavoproteins are located in mitochondria because is of their redox power.123 The

spectroscopic property of flavin coenzyme is very useful to observe changes which occur

within the active site. The human genome encodes 90 flavoproteins, 76 of them require FAD,

14 require FMN and 5 require both FAD and FMN.123 The 90% of flavoproteins are

associated to redox reactions and 10% of them are associated to transferases, lyases,

isomerases, and ligases.124 Flavoproteins catalyze many redox reactions in biological systems

where FMN and FAD have key roles for their functions. Some studies report that FAD can

provide structural support for active sites and also provide the stabilization of intermediate

species during catalysis.125 Flavoproteins are involved in a broad range of biological processes

including bioluminescence, photosynthesis, DNA repair, apoptosis, and elimination of reactive

oxygen species (ROS).126-129 The ROS has been associated with the induction and

complications of diabetes mellitus, age-related eye disease, and neurodegenerative diseases

such as Parkinson's disease. Malfunctioning of flavoproteins is also related to oxidative stress

and the damage of extensive range of molecular species like lipids, proteins, and nucleic acids.

It is also associated to initiation and development of cancer, as well as the side-effects of

radiation and chemotherapy. Thus, flavoproteins and flavin coenzymes are extensively studied

and highly significant in protein science.

15

1.4.1. Flavin cofactors

The flavoprotein contains flavin moiety which could be FAD or FMN, are called flavin

cofactors. Flavoproteins was first discovered in 1879 in cow’s milk which was recognized as

lactochrome at that time. After a half-century of the discovery of the lactochrome protein, the

field of coenzyme research was introduced in the 1930s with the discovery of flavin cofactors

FAD and FMN. The flavin mononucleotide (riboflavin-5’-phosphate) is produced by the

catalytic activity of riboflavin kinase on riboflavin (vitamin B2). It functions as a prosthetic

group of various flavoenzymes including NADH dehydrogenase and blue-light

photoreceptors. FMN is the primary form of the riboflavin found in cells and tissue. The FAD

is composed of an adenosine monophosphate and flavin mononucleotide bridged together by

two phosphate groups (Figure 1.5). The FMN is formed by a C-N bond between an

isoalloxazine ring and ribityl chain, which is not a truly glycoside bond, therefore flavin

mononucleotide is not a truly mononucleotide. However, structure and chemical properties of

FMN is very close to nucleotide. FMN and FAD exist in four different form of redox states,

which are flavin-N(5)-oxide, quinone, semiquinone, and hydroquinone.130 Flavin coenzyme is

produced by bacteria and fungi but it cannot be synthesized by eukaryotes including human

and must be supplied from the dietary sources in the form of vitamin B2.

16

Figure 1.5. Skeletal formula of flavin mononucleotide (FMN) and flavin adenine dinucleotide

(FAD).

1.4.2. Electron transfer process in flavin coenzymes

Flavin coenzymes catalyze electron-transfer reactions in diverse ways on flavoproteins

which are associated with catalysis of one and two electrons transfer process.118,131 Sometimes,

they are involved in the catalysis of electron transfer reactions between two-electron donor and

one-electron acceptor.132,133 The electron transfer mechanism of flavin coenzymes changes as

their interactions with the flavoenzymes and the surrounding environment changes. The X-ray

crystallographic study shows that the electroactive site of flavin coenzymes like an FAD in

17

Figure 1.6. The crystal structure (PDB 1tll) of the neuronal nitric-oxide synthase (nNOS)of rat

(Rattus norvegicus), where the FMN domain is magenta, and the FAD domain is grey.

an aqueous environment is accessible to water molecules 134 and does not bind covalently to

enzymes. Under such conditions, flavin coenzymes undergo a two-electrons/two-proton (2e-

/2H+) reduction process and generate the fully reduced form of flavin coenzymes (FADH2) 132.

However, in a non-aqueous environment, coenzymes bind covalently to flavoenzyme and form

stable flavin semiquinone forms (FADH•). The difference is also reported in proton-coupled

electron-transfer (PCET) mechanism of flavin coenzymes in an aprotic organic solvent and the

aqueous solution.135,136The electrochemical studies show that the redox pathways of the FMN

molecules in aqueous solutions are pH dependent.137-149 The redox states of FMN has an

important role in the blue-light photoreceptor and electron transport process during ATP

synthesis. During ATP synthesis, FMN couples with the series of iron-sulfur (Fe/S) clusters in

the NADH dehydrogenase (complex I) to transport the electron from NADH to ubiquinone (Q)

and pump the protons from the mitochondrial matrix to its intermembrane space, this is an

important example where different redox states of FMN involve in a biological process. As

18

earlier studies show, FMN is involved in both one and two-electron transfer redox processes

taking part in reversible interconversion to generate its oxidized (FMN), semiquinone

(FMNH•) and reduced form (FMNH2). 118,131-133 Many studies have been carried out on flavin

coenzymes to understand biological properties and the importance of their redox states. Webb

and coworker applied the redox fluorometry technique based on the inherent fluorescence

from NAD(P)H and/or flavoprotein to monitor cellular energy metabolism as a function of

substrate availability.150 The kinetics of reduction of the flavocytochrome have been

investigated by using laser flash photolysis.151 Webster and coworker studied the

electrochemical reduction mechanisms of FMN in buffered aqueous solutions using variable-

scan-rate cyclic voltammetry, controlled-potential bulk electrolysis, UV−Vis spectroscopy,

and rotating-disk-electrode voltammetry.136 Similarly, Hazra and coworker used the steady-

state and time-resolved fluorescence quenching of flavin, circular dichroism and thermal

melting techniques to predict the structural reformation on aptamer due to flavin-aptamer

binding.152 Begum and coworker studied the translocation of Flavo enzyme in the nucleus of

Dictyostelium discoideum by utilizing immunofluorescence.153 Here, we found that surface-

enhanced Raman spectroscopy (SERS) is a powerful experimental approach for the fine

characterization of redox states of flavin coenzymes and redox reaction schemes in different

pH environments.

1.5. Raman spectroscopy

Raman spectroscopy utilizes inelastic scattering of light from molecules and provides their

characteristics vibrational fingerprint.154 The Raman spectroscopy has been used intensively after

the development of high powered continuous wave (CW) gas ion lasers in the late 1960’s.155-201

The Raman shift often expressed in cm-1 unit, represents energy differences between Eigenstates

19

of the molecule, and does not rely on the function of the wavelength of incident light.202 Raman

cross-section of many molecules are extremely small, which makes collection and analysis of

natural Raman spectra very difficult.203,204 Fluorescence of molecules further decreases

detectability of scattering signal. However, the progress in instrumentation including lasers,

spectrometers, imaging detectors, counting electronics, and digital spectral processing has

greatly reduced the weakness of Raman spectroscopy. These technical advancements in Raman

spectroscopy have greatly increase the significance of this technique to study wide varieties of

samples.205-250 When molecules are adsorbed on metallic substrates, Raman intensity is

increased by 106-1011 order, and this technique is called surface enhanced Raman spectroscopy

(SERS).251 Substrates used in SERS are usually nanoparticles252,253 or roughened surfaces of gold

or silver.

1.5.1. Surface-enhanced Raman spectroscopy (SERS)

The Surface-enhanced Raman Spectroscopy (SERS) mechanism was first observed by

Martin Fleischman and coworker in 1973 254 , which was obtained from pyridine monolayer

adsorbed on a silver electrode. Later in 1977, Jeanmaire 251 and Albrecht255 had found Raman

signal intensity from molecules adsorbed on a metal substrate is 105-106 times stronger than

normal Raman scattering of non-adsorbed molecules. The SERS spectrum has some difference

from ordinary Raman spectrum.256 First, the band intensity in SERS normally decreases with

the increase of vibrational frequency. For example, C–H stretches becomes relatively weaker

in SERS. In addition, combination bands and overtones are rare, selection rule becomes

flexible, normally forbidden Raman modes could appear, spectra become depolarized, and the

excitation profiles do not rely on the fourth power of laser frequency (𝜔#) as in non-resonant

Raman spectroscopy. Electromagnetic (EM), and chemical enhancement (CE) are two well-

20

explained mechanisms for signal enhancement in SERS.256-259

1.5.2. Electromagnetic mechanism

The electromagnetic mechanism has an important contribution to SERS. The incident

electromagnetic field acting on a probe molecule is increased due to the addition of a field

generated by the polarization of the metal particles. At the same time, the polarization of the

metal particles is induced by the Raman radiation produced from the probe molecules which

further amplify the surface enhancement. Many experimental evidences show that the surface

enhancement is mainly due to localized surface plasmon resonance (LSPR) which occurs due to

collective oscillation of the surface electrons localized on metal particles. The dependence of the

surface enhancement on the shape and surrounding medium of the particle supports LSPR

model. The electromagnetic mechanism can be demonstrated by a molecule adsorbed on a single

metal particle embedded in a homogenous medium252,260,261. The electrostatic calculations show

the significant enhancement of Raman cross-section when probe molecules are adsorbed on the

silver surface262-266. The theoretical calculation shows that electromagnetic enhancement

mechanism increases the Raman intensity up to 1011 factors of magnitude. However,

experimentally observed values are at the order of 106 which are105 times smaller than

theoretically predicted value. Two possible reasons such as radiation damping, and size effect are

often used to explain this deviation. The radiative damping accounts for the radiative loss of the

oscillating electric field when the radius of particles (r) exceeds the wavelength of excitation

laser (𝜆). 267

The smaller size of the metallic nanoparticle causes the loss of the conduction electrons

due to collisions with the particle surface. When the particle dimension is equal or smaller than

21

the mean free path of the electrons in the metal, there will be the additional loss of electrons due

to surface scattering. These effects can be expressed as a size-dependent imaginary part 𝜀&(𝜔) of

the electric function268. For small volume of nanoparticles, the enhancement is limited by the size

effect and for much larger particles, the enhancement fundamentally limited by radiation

damping. The electric field inside the metal nanoparticle (Ein) can be related to the applied

electromagnetic field E0 by

𝐸() = +

+- .(0).234

5 E0 ……………...…………………………………… (1.1)

Where A is the depolarization factor which is independent of size but dependent on

shape, ε(𝜔)= dielectric constants of the bulk metal, and ε7 = dielectric constant of surrounding

medium. This relation is only valid when the wavelength of the incident laser is much larger than

the radius of nano-particles (r≪ λ). The dielectric constant of all metals in the visible region can

be expressed as given below: 269,270

ε(𝜔) = 𝜀+(𝜔) + 𝑖𝜀+(𝜔) ……………...…………………………………… (1.2)

The value of the electric field inside the metallic nano-particle (Ein) is maximum when

the exciting photon energy satisfies the condition of ε (𝜔) = ε7(1-A-1) in equation (1.1). This

condition corresponds to the excitation of localized surface plasmons, and enhancement occurs

because of Ein ≫ E0.

Ein = ; < =;2; < -&;2

𝐸0 ……………...………………………………………. (1.3)

Ein is maximum when the real part of the denominator in equation (1.3) is zero and the

imaginary part in equation (1.2) is negligible. Both gold and silver nanoparticles satisfy these

22

two requirements in the region of the visible spectrum, hence they are a good source for SERS

substrate.

In addition, it is reported that the dipole moment located at the center of the particle 𝑃(<)

is induced by an external field which is given by 271

𝑃(<) = ; < =;2; < -&;2

𝑟3 E0 ……………...………………………………… (1.4)

Where r = radius of the sphere. The electric field is uniform inside the particle and decays

with the distance (d) from the surface according to the dipole decay law @@-A

B. Thus, a

molecule located at the surface acquires the largest field, but it is not compulsory for direct

connection with the surface.

Moreover, the Zeman and Schatz272 proposed a spheroid model for accurate calculation

of electromagnetic fields near the surface of small metal spheres. It can be explained under

following points: (1) solution of the static (Laplace) equations; (2) Adjustment of these results

for electrodynamics effects; and (3) addition of surface scattering effect in the dielectric

constants. The overall field enhancement factor 𝑅(<) is given by273

𝑅(<) = |EFGH,J

K |E2K

……………...……………………………………… (1.5)

Here, 2b and 2a can be taken as the major and minor axis of a prolate spheroid and

constant field E0 applied along the major axis. E is the resulting field just outside the spheroid.

23

Figure 1.7. Prolate spheroid to show the model proposed by Zeeman and Schatz.

The enhancement factor (𝑅(<)) is used to calculate single photon measurements; it is

more useful to calculate a Raman enhancement factor(𝑅(<,<L)) in SERS.

𝑅(<,<L) = 𝑅(<)𝑅(<L)……………...……………………………………. (1.6)

Furthermore, Gersten and Nitzan252 have identified three major electromagnetic

contributions to the Raman scattering enhancement ratio: (i) the image enhancement mechanism

that can be operative at close range; (ii) the lightning rod effect; and (iii) the resonance with the

surface plasmon associated with the eccentric surface particle feature. First two effects can

contribute SERS enhancement by increasing the electric field on the molecule and affecting the

polarization of the metal by the molecular dipole. When the equivalent point dipole located at the

center of particles replaces the polarized metal particle, this model totally underestimates the

maximum enhancement because the concentration of the electric field near the tip of a prolate

spheroid is neglected. The charge density in the sharper tip of the metal surface is higher which

contribute greater surface enhancement, this effect is called ‘lightening rod effect’274

.

24

1.5.3. Chemical enhancement mechanism

The chemical enhancement mechanism occurs due to charge transfer between metal and

analyte molecules. When analyte molecule is adsorbed on metal, its fermi level falls in the

symmetry of HOMO and LUMO of adsorbed molecules. When light interacts with adsorbed

analyte molecules it excites to the virtual energy states. The electron transfer from metal

substrate to the excited electronic level of the adsorbed molecules by the process of tunneling for

physically adsorbed molecules and by the hybridization for chemisorbed molecules. The electron

transfers back from the excited adsorbed molecule to the metal, leaving the adsorbed molecules

in the higher vibrational level in Eo (V = 1 of Eo); and (d) photon emission with energy ω by the

electron recombination with the hole below Fermi level (Ef). Charge transfer mechanism is short

range (0.1 - 0.5 nm) and depends on the factors like the site of adsorption, bonding geometry,

and energy level of the probe molecules. The charge-transfer processes contribute approximately

10-103 order to SERS.3 Raman polarizability and the polarizability of the molecule-metal

system,𝛼NO, due to CT excitation from metal to the molecule can be expressed in terms of the

density of states of the metal 𝜌Q(E) and the adsorbate 𝜌R(E). The intensity of the Raman

scattering can be expressed as252

𝐼 ω,ωU = VWX

# 𝛼YZ 𝜔 & 𝑣\ 𝑄 𝑣 &

……………...…………………. (1.7)

Where vg and ve are the vibrational wave functions of the ground and excited CT states,

respectively, and Q is related to the energy 𝜀R(Q) of the molecule supplemented by a

displacement Q associated with a molecular vibration whose frequency is𝜔 −𝜔Z.

25

Dividing equation (11) by the intensity for a free molecule, the enhancement factor

R(𝜔,𝜔Z) of the coupled molecule-metal system due to CT excitation from the metal to the

molecule takes the form:275

𝑅<,<L = 𝛼YZ 𝜔 −𝐸\^` 𝜔a −𝐸\^` &……………...…………………. (1.8)

𝐸\^` is the energy separation between the ground and the excited electronic state related

with a dipole transition on the free molecules.

Studies of SERS spectra of good electron acceptor or donor molecules at various excitation

energies and studies of the SERS spectra as functions of surface coverage should be useful to test

the CT contribution to SERS and to provide information on the structural and dynamic properties

of the adsorbed molecules. The Raman excitation profile of the background spectrum provide

useful facts concerning the CT mechanism.

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43

CHAPTER 2. EXPERIMENTAL SECTION

This chapter briefly explains the experimental techniques, instrumentations and the general

procedures used for the sample preparation during experiments.

2.1. Theoretical background

Here we explain about theoretical background of Raman spectroscopy, resonance Raman

spectroscopy, surface-enhanced Raman spectroscopy, two-photon excited (2PE) fluorescence

microscopy, transmission electron microscopy, and confocal microscopy.

2.1.1. Raman spectroscopy

The concept of Raman spectroscopy was invented by the Indian physicist sir C. V. Raman

who won the Nobel prize for his invention in 1930. Raman spectroscopy is a powerful technique

to probe the vibrational modes of the chemical bonds and symmetry of the molecules.1-6 Now,

different forms of Raman spectroscopy are successfully utilized to study the huge range of

molecules and their interactions in different physical conditions, which are evidence of its

importance.7-53 The technical advancement in Raman spectroscopy has greatly increased

significance of this technique to study wide varieties of samples.54-99

The principle of Raman spectroscopy is based on the inelastic scattering of light which

occurs due to energy transfer between the photons and molecules.31,51,100-103 When a

monochromatic light interacts with the molecule, molecule gets excited from the ground state to

the virtual states. When it relaxes back to the ground state, (1) the molecule goes back to the

exact same vibrational or rotational states causing no energy change between absorbed and the

emitted light, which is called elastic or Rayleigh scattering;104 or (2) the molecule returns to a

different vibrational or rotational state causing energy difference between the absorbed and

44

emitted photon, which is called inelastic or Raman scattering. If the final state of the molecule is

more energetic than its initial states, the scattering is called as stokes Raman. If the final state of

the molecule is less energetic than its initial state, the scattering is called anti-stokes Raman (Fig.

2.1).51

Figure 2.1. The schematic representation of Rayleigh, Stokes and anti-Stokes scattering.

The change of vibrational energy level is represented in wavenumbers (cm-1) and can be

calculated using following equation:

Δω = +bcdLef

− +bLgdH

×10k𝑐𝑚=+ ………………………………(2.1)

Where Δω is the Raman shift, and λlaser and λscat are the wavelength of the excitation laser

and scattered light respectively in nanometers units. This equation shows that the wavelength of

Rayleigh scattered light or excitation laser is the reference point to calculate the vibrational

wavenumber of the molecule. The Raman wavenumber of a scattered light is independent of the

laser frequency.

The vibrational modes of the molecule are Raman active when their polarizability is

changed. The molecules with the center of symmetry have Raman active vibrational mode; a

45

molecule without center of symmetry also could have some components such as in-phase

stretching or bending which contribute some Raman active modes. The intensity of Raman

scattering is expressed by the following equation.

𝐼Y ∝ 𝜔#𝐼7𝑁pqpr

&………………………………….…………..(2.2)

Where, 𝜔 is the wavenumber of the excitation light, I0 is the intensity of excitation laser,

N is the density of the scattering molecules, α is the polarizability, and Q is the vibrational

amplitude.

Intrinsically, Raman scattering is a less sensitive technique because Raman scattering

cross-section of most of the molecules is extremely smaller. In addition, the fluorescence coming

from the probe molecules or other sample components further decreases detectability of Raman

scattering signals. However, the technical advancement in instrumentation including lasers,

spectrometers, imaging detectors, counting electronics, and digital spectral processing has

greatly increased the effectiveness of Raman spectroscopy. In addition, the modified versions of

Raman spectroscopy such as Resonance Raman (RR), surface-enhanced Raman Scattering

(SERS), and tip-enhanced Raman Scattering (TERS) are used to magnify the Raman signal

intensity.

2.1.2. Resonance Raman spectroscopy

In RR spectroscopy the energy required for the electronic transition of molecules should

match the energy of the laser light (Figure 2.2). When the energy of excitation laser light

matches with the certain vibrational modes of the molecules, the Raman intensity related to that

certain vibrational mode is significantly increased.105-110 However, other vibrational modes

46

which are not resonant with excitation laser light do not show any enhancements. This property

of RR spectroscopy is beneficial for the mode selective Raman analysis of molecules. Despite its

many advantages, RR has some limitations, for example, the resonant excitation induces the high

fluorescence background which could interfere the Raman signal intensity. In addition, the

higher laser power required for the RR could induce the photodegradation of the probe

molecules.

Figure 2.2. The schematic representation of resonance Raman scattering.

2.1.3. Surface-enhanced Raman spectroscopy

Surface-enhanced Raman Spectroscopy (SERS) utilized the laser to excite the vibrational

transition of probe molecule located between metallic nanoparticles or adsorbed on a rough

metallic substrate. The enhancement of the signal depends on various factors such as size and

shape of nanoparticles, excitation wavelength, and location of adsorbed analyte molecules in

SERS substrate. In SERS, the Raman cross-section of the analyte is enhanced by 106 to 1011

orders of magnitude. Mainly two types of mechanisms are explained for the SERS enhancement:

47

(1) Electromagnetic mechanism (EM) and (2) Chemical enhancement or charge transfer

mechanism (CT).111 The contribution of EM and CT in Raman stokes component (𝜇YZ) are

summarized in equation 2.3.

𝜇YZ =∝YZ 𝐸+ 𝑟, 𝜔 exp[−i ω − 𝜔a 𝑡] ………………………………...…. (2.3)

Where, ∝YZ is the Raman Stokes polarizability and stands for CT mechanism, 𝐸+ 𝑟, 𝜔 is

the local field intensity and stands for EM, 𝜔 is the laser frequency, and 𝜔a is the frequency of

molecular vibration. The contribution of each effect depends on various factors such as nature of

probe molecules, the optical and chemical properties of the metallic surface, and experimental

parameters.

Figure 2.3.The schematic representation of the (A) electromagnetic enhancement in SERS

measurementswhere orange spheres represent the metallic nanoparticles and the green sphere is

the probe molecule and (B) chemical enhancement in SERS measurements.

The chemical enhancement factor arises due to adsorption of the molecule on the metallic

surface. When the adsorption occurs, the fermi level of the metal generally falls between the

highest occupied molecular orbital (HOMO) and lowest unoccupied molecular orbital (LUMO)

48

of the adsorbed molecule enabling the possibility of charge transfer between the metallic particle

and the molecule. The charge transfer works as a new channel for resonance Raman scattering

which increases the Raman intensity. The chemical enhancement factor is generally in the order

of 102.

2.1.4. Two-photon excited (2PE) fluorescence microscopy

Two-photon excited (2PE) fluorescence microscopy allows the imaging of submillimeter

thick biological samples like cells and tissues. In 2PE fluorescence microscopy, the wavelength

of emission light is shorter than the wavelength of excitation light. This technique typically uses

longer wavelength femtosecond laser for the excitation of molecules, and for each excitation,

two photons of excitation laser are utilized.112-116 The scattering of the excitation light from cell

or tissue with longer wavelength is smaller than that of shorter wavelength. The 2PE

fluorescence imaging is a crucial technique to reduce the background noise from the image,

increase the optical sectioning effect for the cell imaging, and suppression of the photobleaching

effect.113,117 The 2PE fluorescence imaging has been used for the study of different fields of

biology including physiology, neurobiology, embryology and tissue engineering.118-121

The concept of 2PE was first explained by Maria Goeppert-Mayer in 1931 which was

later observed experimentally by Wolfgang Kaiser in 1961.117,122,123 The 2PE microscopy was

invented by Watt W. Webb and coworker at Cornell University in 1990. 113,114 In this imaging

technique, the focused laser beam is scanned in a raster pattern to generate images which

increases the optical sectioning effect for the high-quality image. The optical sectioning effect is

higher in 2PE microscopes because the axial spread of the point spread function in this approach

is significantly lower than in single-photon excitation, which improves the resolution along the

49

Z-dimension. This technique utilizes two photons of comparably lower energy than the energy

required for the single photon excitation process to excite the molecule in one quantum event.

The photon energy required for the 2PE fluorescence approach is approximately half of the

energy required for the single photon excitation. This technique typically requires a high flux of

excitation photons because simultaneous absorption of two photons is less probable.

Figure 2.4. The schematic representation of two-photon excitation (2PE) fluorescence

microscopy.

The single photon fluorescence imaging of cells requires UV light for the excitation of

intracellular contents, which is toxic to the cell when exposed longer. In addition, the optical

sectioning effect of UV light is comparatively poorer. In contrast, the 2PE fluorescence imaging

utilizes visible or infrared light which is non-toxic for the longer exposure and gives improved

optical sectioning effect. The emission intensity of 2PE fluorescence is quadratically

proportional to the excitation intensity, therefore the excitation light for 2PE is tightly focused in

50

the small volume. The fluorescence from the sample is then collected by a high-sensitivity

detector, such as a photomultiplier tube (PMT) or avalanche photodiode (APD). We used

frequency doubled femtosecond 532 nm pulsed laser (Chameleon Discovery, coherent, ~100 fs

fwhm) for the excitation of samples. The excitation laser is shone on samples through 532 nm

dichroic mirror (Chroma, ZT532rdc) and oil immersion objective of the inverted microscope.

The diffraction limited (300 nm) epifluorescent light beam returns to the 532-nm dichroic mirror

and collected by single photon avalanche photodiode (APD) (PerkinElmer SPCMAQR-14). The

bandpass filter (FES0500) is positioned in front of APD to prevent entry of any excitation laser

and single photon excited (1PE) fluorescence.

2.1.5. Transmission electron microscope (TEM)

The transmission electron microscope (TEM) was invented by Max Knoll and Ernst

Ruska in 1931.124 Transmission electron microscopy (TEM) is the imaging technique which

generates an image of the specimen (less than 100 nm thick) or a suspension on a grid by the

transmission of electrons beam.125-132 The image of the extremely small specimen is magnified

and focused onto an imaging device, such as a fluorescent screen, photographic film, or a charge-

coupled device (CCD). TEM is a robust analytical tool which is used for the analysis of chemical

identity, crystal orientation, electronic structure, and sample induced electron phase of the

specimen. TEM utilizes smaller de Broglie wavelength of electrons, therefore it provides highly

resolved image than light microscopes.133 For lower magnification, the contrast in TEM image is

obtained because the electron absorption properties of the different material are different. For the

higher magnification, the image intensities from the complex wave interactions are modulated

and analyzed to obtain contrast.

51

The TEM utilizes the information of electron waves from the sample to form an image.

The electron wave distribution is correctly positioned onto the viewing system by projector

lenses. The intensity of TEM image is proportional to the time-averaged amplitude of the

electron wavefunctions 𝜓.134

𝐼 𝑥 = }~43~2

𝜓𝜓∗~4~2

𝑑𝑡 …………………………………. (2.4)

Figure 2.5. Schematic representation of TEM in diffraction modes

Equation (2.4) tells that the information about the sample and the electron beam itself could be

obtained by modifying the electron wave existing in the sample. The image intensity depends on

52

the amplitude of the electron beam as well as on the phase of the electrons. However, the phase

effect is normally ignored for the lower magnifications.

The TEM has two basic operation modes: diffraction mode and imaging mode. In both

modes, the specimen is illuminated with the parallel beam of electrons through the system of

condenser lenses and condenser aperture. After illumination of specimens, the scattered and

unscattered electrons exit from specimens; unscattered electrons form bright central beam on the

diffraction pattern while scattered electrons change their trajectories.

Figure 2.6. Schematic representation of TEM in imaging mode

In imaging mode, the objective aperture is inserted in a back focal plane of the objective

lens (Figure 2.6). When the objective aperture is used to select the central beam and block the

53

rest of the signal, it gives the bright field image. When the objective aperture is used to select the

signal from the diffracted beam, the dark field image is formed. The intermediate and projector

lenses are used to magnify and project the image on a detector, and the image of the sample is

obtained. In the diffraction mode, selected area aperture is inserted to determine the specimen

area (Figure 2.5) and display the signal intensity of the image. The diffraction pattern is obtained

on a screen by changing the strength of the intermediate lens. The diffraction mode in TEM

technique is considered a powerful tool to determine cell reconstruction and crystal orientation.

2.1.6. Confocal microscopy

Confocal microscopy or confocal laser scanning microscopy (CLSM) is the optical

imaging technique for the increasing optical resolution which requires the use of a spatial pinhole

Figure 2.7. Confocal microscopy geometry including the light pathways.

to block out-of-focus light. In this technique, only the light coming from focal plane or very

close to the focal plane is detected which increase the optical resolution but decreases the signal

54

intensity.135-143 This technique requires the longer exposure time and sensitive detector like

photomultiplier tube (PMT) or avalanche photodiode (APD). This technique is useful for

noninvasive optical sectioning of thick living specimens with the improvement of lateral

resolution. The fluorescent dyes are often used for the selective imaging of the part of the

biological samples. Only one point of the sample is illuminated at a time in confocal microscopy,

therefore scanning over a regular raster in the specimen is required for the 2D or 3D scanning.

The scanning method has a short response time and scanning rate can be varied; the slower rate

of raster scanning increases the signal-to-noise ratio, contrast, and resolution of the image.

2.2. Experimental set-ups

2.2.1. Raman measurements and optical imaging

Raman measurements and optical imaging are recorded by an Axiovert 135 inverted

scanning confocal microscope, equipped with a 100 x 1.3 NA oil immersion objective (Zeiss

FLUAR) (Figure 2.7). The CW laser (488 nm, Melles-Griot 35-IMA-040-120) is directed by a

single-mode optical fiber (Thorlabs) into the experiment-box. The output of the optical fiber is

collimated by an objective and directed into the microscope. A dichroic mirror (ZT488rdc

Chroma) is used to reflect the excitation light into the microscope objective. Before focusing the

scattered light into a monochromator (Triax 550, Jobin Yvon), a bandpass filter (HHQ495LP) is

positioned before the entrance slit to further eliminate the Rayleigh scattering. The Raman

spectra are recorded by an LN-CCD (Princeton Instruments) cooled at about -100°C with a

resolution of ~2 cm-1 with 600 g/mm grating and ~1 cm-1 with 1200 g/mm grating. The setup is

carefully calibrated by using mercury lamp and cyclohexane (mode at 801.3 cm-1) before the

Raman measurements. The laser light of ~10-15 µW power is used to pump the sample for SERS

55

measurements. For optical imagining, APD (Perkin-Elmer, SPCM-AQR14) is engaged with the

microscope.

Figure 2.8. The schematic representation of the experimental set-up used in Raman

measurement and optical imaging.

2.2.2. Electrochemical control and Raman measurement

The Spectro electrochemistry measurements were performed with CH Instruments 600C

electrochemical analyzer coupled with a home-made 3-electrode electrochemical cell (Figure

2.9) coupled with confocal Raman microscopy where a platinum wire, silver chloride, and ITO

worked as the counter electrode, reference electrode, and the working electrode respectively. We

used the combined approach of electrochemical control and Raman measurement to probe the

redox states and mechanism of flavin coenzymes (Chapter 6). The ~40 µl solution of FMN

56

(prepared in B-R buffer and 0.4 M KCl) and silver nanoparticles are incubated overnight on the

ITO surface and dried, then a solution of 0.1 M KCl was put on the top of the ITO as a

supporting electrolyte in the home-made electroscopic cell (Figure 2.9B). We have tested

different scan rates and identified that the scan rate at 0.1 V/s is optimal to obtain CV signals at

an adequate signal-to-noise ratio. The cyclic voltammetry provides the driving force which may

induce certain transitions and produces many redox states of FMN, these redox states would not

have occurred physiologically in the provided pH range.

Figure 2.9. (A) The schematic representation of the home-modified experimental setup for the

electrochemical redox process and Raman spectroscopy(not to scale). The optical requirement

for SERS are met by the combination of 488 nm CW argon ion laser (12±2 µW power) for the

excitation of the sample, confocal microscope (Axiovert 135) equipped with a XY piezo-

controlled scanning stage (Zeiss FLAUR, 100×; 1.3 NA), dichroic mirror (Chroma, Zt488 drc),

long pass filter (HHQ495 LP), and monochromator (Triax 550, Jobin Yvon). (B) The schematic

representation of the home-made electrochemical cell coupled with confocal Raman microscopy

57

where platinum (Pt) wire is the counter electrode, silver chloride (AgCl) is the reference

electrode, and ITO is the working electrode.

2.2.3. Two-photon excitation (2PE) fluorescence imaging

Frequency doubled femtosecond 532 nm pulsed laser (Chameleon Discovery, coherent,

~100 fs fwhm) is used for the excitation of samples. The excitation laser is shone on samples

through 532 nm dichroic mirror (Chroma, ZT532rdc) and oil immersed objective of the inverted

microscope's. The diffraction limited (300 nm) epifluorescent light beam returns to the 532-nm

dichroic mirror and collected by single photon avalanche photodiode (APD) (PerkinElmer

SPCMAQR-14). The bandpass filter (FES0500) is positioned in front of APD to prevent entry of

any excitation laser and single photon excited (1PE) fluorescence. The percentage transmittance

of the dichroic mirror (chroma, ZT532rdc) at the region of 400-500 nm is ≥ 55%, which is good

enough to achieve the purpose of our experiment. The bandpass filter (FES0500) is used in front

of detector to block the light longer than 500 nm including excitation laser of 532 nm. This

experimental set up ensures the collection of fluorescence emission from two-photon excitation

(2PE) of DA using 532 nm excitation laser.

Figure 2.10.The percentage transmittance of (A) dichroic mirror (chroma, ZT532rdc) and (B)

bandpass filter (FES0500)collected using Varian UV-Vis spectrophotometer (S/N EL07013173).

58

2.2.4. Mode-selective Raman measurement

The mode-selective Raman spectra are collected using experimental set up of SERS

measurements with required modification (Figure 2.11). In mode selective Raman measurement,

we replaced the long pass filter (HHQ495LP) of SERS measurement with the digital mini-chrom

monochromator ((DMC1-03) and selected desired wavenumber of the scattered light. However,

mode-selective Raman images are collected using experimental set up of 2PE fluorescent

imaging. Here, we replace the 532 nm fs pulsed laser by 488 nm CW argon ion laser and use the

bandpass filter (FES0500). In addition, we also replaced 532 nm dichroic mirror (Chroma, Zt532

drc) with 488 nm dichroic mirror (Chroma, Zt488 drc).

Figure 2.11. Schematic representation of the optical setup for the two-photon excited (2PE)

fluorescence and Raman imaging(not to scale). For two-photon excited (2PE) fluorescence

imaging, frequency doubled 532 nm fs pulsed laser (Chameleon Discovery, coherent, ~ 100 fs

fwhm) is used. For the Raman experiment, 488 nm CW argon ion laser is used. M=Mirrors and

MC=monochromator (digital mini-Chrom monochromator (DMC1-03) are foldable.

59

2.3. Density functional theory calculations

Geometry optimization and Raman wavenumber calculations were performed using the

density functional theory (DFT) method on a B3LYP level with a basis set of 6-31G (d) and

Gaussian 09 package to observe Raman wavenumber of probe molecules. Literature shows that

the scaling factors of 0.9804 could be more applicable for the DFT calculated vibrational

Figure 2.12. The comparison of experimental Raman spectrum (dotted black) of dopamine with

DFT calculated Raman spectrumon a B3LYP level with basis set of 6-31G(d), and Gaussian 09

package. The calculated wavenumber is scaled by a factor of 0.9614.

wavenumber at the lower wavenumber region (<1000 cm-1), and 0.9625 for the wavenumber at

higher wavenumber region (>1000 cm-1).144,145 Based on our control experiment for the

calibration of the scaling factor (Figure 2.12), and a comprehensive evaluation reported146, we

have scaled the DFT calculated Raman frequencies by a factor of 0.9614, which is suitable to our

60

region of interest (higher wavenumber region). The molecular orbitals were calculated with the

same basis set and visualized with Avogadro software (Avogadro: an open-source molecular

builder and visualization tool. Version1.XX.). All calculations were carried out on a vector

processor (Ohio Supercomputer Center, Columbus, Ohio).

2.4. Materials and sample preparation

All chemicals were purchased from Sigma-Aldrich and were used as received unless

otherwise stated.

2.4.1. Synthesis of silver nanoparticles

Silver nanoparticles (AgNPs) were synthesized by a standard sodium citrate reduction

method.147 Briefly, A three-necked 100 mL round bottom flask was washed with freshly

prepared Aqua Regia and dried. Then the solution of 9 mg of silver nitrate and 50 ml of

deionized water was boiled in this flask equipped with a water condenser. The solution was

brought to boiling under vigorous stirring in the oil bath. Once the boiling started, 1 mL of 0.039

M aqueous sodium citrate solution was added to the solution and it was refluxed for 90 minutes.

The absorption spectrum of particles was checked time by time after 45 minutes of boiling until

the wavelength of absorption maximum is found at 420 nm. The characterization of the size of

silver nanoparticle using transmission electron microscopy shows that the diameter of silver

nanoparticles is found between 50-70 nm.

61

Figure 2.13. The plasmon resonance related to the bare silver nanoparticles (AgNPs) and silica-

coated silver nanoparticles (AgNP@SiO2)collected using Varian UV-Vis spectrophotometer

(EL07013173). The images of nanoparticles obtained using transmission electron microscopy

(TEM) are given in inset.

2.4.2. Fabrication of silica shell on silver NP

The ultrathin layer of silica was generated on the surface of silver nanoparticles using a

standard protocol.148 In brief, when the synthesis of silver nanoparticles is completed,147 we

added 11.7 µl of 98% (3-aminopropyl)triethoxysilane to the refluxed solution and continued

stirring for other 15 min. Then, 2 ml solution of active sodium silicate (0.54 % weight) was

added to the solution and stirred for 1 hour at 90°C to generate ultrathin silica shells over

AgNPs. The synthesized nanoparticles were characterized by transmission electron microscopy

(TEM) and Varian UV-Vis spectrophotometer (EL07013173). According to our TEM

62

measurement, the size of AgNP@SiO2 was found 58 ± 5 nm with SiO2 shell thickness 5 ± 1 nm.

The UV-Vis spectroscopy shows that the maxima (λmax) of the plasmon resonance band

position related to silver nanoparticles is found at 420 nm which was shifted to 407 nm after

generating an ultra-thin layer of silica on its surface.

2.4.3. Preparation of Britton-Robinson (B-R) buffer

The Britton-Robinson (B-R) buffer was prepared by adding equal volumes of an

equimolar concentration of acetic acid, phosphoric acid, and boric acid (0.04 M). Then the pH

was adjusted by careful addition of NaOH solution (0.2 M).

2.4.4. HEK293 cell culture

The vial of frozen HEK293 cells is quickly thawed in a warm water bath (37°C) and

decontaminated by spraying 70% ethanol. Then cells are transferred into a T-75 flask with 15 ml

of complete medium. The complete medium contains Dulbecco's Modified Eagle's Medium

(DMEM, Sigma-Aldrich D5796), 10% fetal bovine serum (Sigma-Aldrich, F2442), and 1%

penicillin-streptomycin (ATCC, 30-2300). Then T-75 flask containing cells is stored in the

incubator at 37°C temperature with 5% supply of CO2 atmosphere. After 24 hours, all the

complete medium from the T-75 flask is aspirated out and replaced by the fresh complete

medium. When a confluence of the cells reached to 70-80% on the surface of the T-75 flask,

cells are taken out from the surface of T-75 flask using trypsin EDTA and subcultured on a 25-

mm circular cover glass in 35 mm petri dish.

2.4.5. Preparation of hDAT inducible HEK293 cells.

After 24 hours of subculture, we followed the standard protocol of Invitrogen for the

preparation of hDAT inducible HEK293 cells. In brief, all the complete medium from the T-75

63

flask is aspirated out and replaced by fresh complete medium containing 1×10-4 gm/ml zeocin

(invivoGen, ant-zn-1), and 1×10-3 gm/ml blasticidin (Millipore/Calbiochem, 203350). When the

cells reached 70-80% confluence on the surface of the T-75 flask, hygromycin B (Sigma-

Aldrich, H3274) is added in its final concentration 25 µg/µl, which kills all the cells within 2

days except hygromycin resistance cells, which are isolated, and cultured in complete media.

When cells are attached (approx. 24 hrs) on the surface of the T-75 flask, doxycycline (Tocris

Bioscience, 4090) is added to make final concentration 0.5 µg/µl and waited for ≥ 48 hrs to

obtain hDAT inducible HEK293 cells.

2.4.6. Bacteria growth and Plasmid amplification

Plasmid DNA encoding pcDNA3.1-hDAT (32810) was purchased from Addgene and

amplified with a standard method. The amplified plasmid was treated with hDAT inducible

HEK293 cells for the expression of the human dopamine transporter (hDAT). Here the extraction

of pcDNA3.1-hDAT from bacterial cells and transfection of the plasmid in HEK293 has been

explained briefly. Firstly, we received pcDNA3.1-hDAT plasmid of human dopamine receptor as

the bacteria in agar stab, which were streaked on the surface of LB agar plate using sterile wire

loop. All LB plates were kept inside the small incubator for 16 h at a temperature of 37°C.

Bacteria were grown on an LB plate as a colony with a copy of pcDNA3.1-hDAT plasmid. From

there, one of the separate bacterial colony was taken out and added into a falcon round-bottom

tubes containing 6 mL of LB solution with 100 µg/ml ampicillin. Then these falcon tubes were

placed in a rotating incubator for 24 h at 37°C to grow bacteria having pcDNA3.1-hDAT

plasmid. Bacteria from a falcon tube in LB solution were extracted in a 2 ml centrifuge tube by

centrifuging process. For that, the liquid part was thrown after each centrifuge and solid residue

finally deposited on the bottom of centrifuge tube. The solid residues collected in a centrifuge

64

tube was dissolved by 250 ml of resuspension reagents (Thermo Scientific, Gene JET Plasmid

Miniprep kit, cat. no. K0502). Then, 250 ml of lysis solution and 350 ml of neutralization

solution were added to separate the cell membranes and plasmids. The resulting solution was

centrifuged for 15 min at 1200 rpm, and the supernatant solution was taken out from the

centrifuge tube and filtered through the miniprep gene jet spin column to remove filtrate. The

content adsorbed on the filter was washed two times with 500 ml wash buffer and centrifuged to

remove any remaining alcohol present in the plasmid. Finally, the filter was washed twice with

25 µL of elution buffer to collect total 50 µl plasmid solution. The similar procedure is applied to

amplify the gene of D1 dopamine receptors (DRD1) and D2 dopamine receptors (DRD2)

plasmids which were also obtained in bacterial agar stab from Addgene.

2.4.7. Transfection of pcDNA3.1-hDAT in HEK293 cell

For the efficient gene transfection, we followed a protocol of lipid-mediated transfection

with Lipofectamine LTX and Plus reagent (Invitrogen, 11668-019). After subculture of hDAT

inducible HEK293, we normally waited for 1-2 days to allow them to attach on the cover glass

and cover 50 % of its area. This was the condition when cells were ready for the hDAT

expression. At one day before transfection, the growth medium having antibiotics (Penicillin

streptomycin) was replaced with the growth medium that had not antibiotics. For the

transfection, 5 µL of pcDNA3.1-hDAT plasmids was first added to 250 µl Opti-MEM solution.

Then 5 µl of Plus reagent was added to this resulting mixture. The 50 µl solution from this

diluted plasmid was then mixed with the previously prepared solution of 50 µl Opti-MEM and 4

µl of lipofectamine LTX (Invitrogen). This complex DNA reagent was incubated for 20 min at

room temperature. Finally, 50 µl solution of complex DNA reagent was added to the Petri dish

containing cells and waited for 2-3 days for the expression of hDAT in HEK293 cells. Before the

65

experiment, cells were washed with PBS solution to remove culture medium and used fresh PBS

solution to avoid drying of cells. The similar procedure is applied to express DRD1 or DRD2

genes in HEK293 or HT22 cells.

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75

CHAPTER 3. RAMAN SPECTROSCOPIC SIGNATURE MARKER OF DOPAMINE-

HUMAN DOPAMINE TRANSPORTER INTERACTION IN LIVING CELLS

Dopamine (DA) controls many psychological and behavioral activities in the central nervous

system (CNS) through interactions with the human dopamine transporter (hDAT) and dopamine

receptors. The roles of DA in the function of the CNS are affected by the targeted binding of

drugs to hDAT; thus, hDAT plays a critical role in neurophysiology and neuropathophysiology.

An effective experimental method is necessary to study the DA-hDAT interaction and effects of a

drugs like psychostimulants and anti-depressants which can influence DA-hDAT interactions. In

searching for obtaining and identifying the Raman spectral signatures for DA-hDAT interactions,

we have used surface-enhanced Raman scattering (SERS) spectroscopy to record SERS

spectrum from DA, Human Embryonic Kidney 293 cells (HEK293), hDAT-HEK293, DA-

HEK293, and DA-hDAT-HEK293. We have demonstrated a specific 2D-distribution SERS

spectral analytical approach to analyze DA-hDAT interaction. Our study shows that the Raman

modes at 807, 839, 1076, 1090, 1538, and 1665 cm-1 are related to DA-hDAT interaction, where

Raman shift at 807 and 1076 cm-1 are the signature marker for the bound state of DA to probe

DA-hDAT interaction. On the basis of density functional theory (DFT) calculation, Raman shift

of bound state of DA at 807 cm-1 is related to combination of bending modes α(C3-O10-H21),

α(C2-O11-H22), α(C7-C8-H18), α(C6-C4-H13), α(C7-C8-H19), α(C7-C8-N9), and Raman shift at 1076

cm-1 is related to combination of bending modes α(H19-N9-C8), γ(N9-H19), γ(C8-H19), γ(N9-H20),

γ(C8-H18), and α(C7-C8-H18). Our findings demonstrate that protein-ligand interactions can be

confirmed by probing the change in Raman shift of ligand molecules, which could be crucial to

76

understanding molecular interactions between neurotransmitters and their receptors or

transporters. The results have been published in American Chemical Society Journals.

3.1. Introduction

Dopamine (DA) is the catecholamine neurotransmitter, which controls many

psychological and behavioral activities in the central nervous system (CNS) of Mammalia

through biochemical interactions with the dopamine transporter (DAT) and dopamine receptors

(DARs).1,2 Dopaminergic neurons located in the ventral tegmental area, substantia nigra pars

compacta, and the arcuate nucleus of the hypothalamus enzymatically convert tyrosine (Tyr) into

L-DOPA and finally into DA.3-6 The DA neurotransmitter works in a sequence of several steps.

First, the action potential is generated on neuron membrane from environmental stimulation. It

causes the opening of voltage-gated Ca2+ ions channel followed by the entry of Ca2+ ions into the

neuronal cell, which induces the release of DA into the synaptic cleft. 7-9 Then, DA in the

synaptic cleft activates G protein-coupled DARs located in pre- or post-synaptic region to

generate the dopaminergic response.10-15 High concentration of DA in the synaptic cleft between

neuron cells causes over-stimulation of DARs resulting several neurological and physiological

disorders. The excessive neurotransmission due to accumulation of DA as well as other biogenic

amino neurotransmitters like serotonin, noradrenaline, and 𝛾-aminobutyric acid present in

synaptic and perisynaptic space is removed by the reuptake process; DA is preferentially taken

up into the dopaminergic nerve terminal by DAT.16-19 DAT belongs to the family of solute

carrier 6 (SLC6), 20 which maintains homeostasis of biogenic amino neurotransmitters by Na+

and Cl- ion assisted reuptake process and thus it is also known as neurotransmitter sodium

symporters (NSS). 21-25 Dysfunction of NSS system is linked with several disorders like

schizophrenia, depression26, attention deficit hyperactivity disorder (ADHD)27, orthostatic

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intolerance28, epilepsy29, Parkinson’s disease and infantile Parkinsonism dystonia.30 The agonists

and antagonists of DARs has influence in dopaminergic transmission by enhancing or blocking

the actions of DA on receptors.31-36 Inhibition of DA reuptake from synaptic cleft has several

side effect, but it is an important pharmacological method for the treatment of depression.37,38

The targeted binding of drugs to the human dopamine transporter (hDAT) has effect in the

development and function of the nervous system; thus, the study of DA-hDAT interaction has

importance in neurophysiology.

Figure 3.1. (A) Schematic representation of dopamine system in neuron cell showing hDAT,

DA, and DA receptor, (B) HEK293 cell, and (C) silica-coated silver nanoparticles.

Different types of experimental techniques and computational modeling have been

applied to understand the structure and function of membrane proteins and their interactions with

ligands 19,39-57 Imaging of hDAT has been conventionally studied by fluorescence microscopy,

positron emission tomography (PET), and single photon-electron tomography (SPET). Booij and

coworkers used [123I]-CIT single photon emission computed tomography (SPECT) to

78

demonstrate the loss of striatal dopamine transporter content in Parkinson's disease in human.45

The trafficking of hDAT has been studied by using Photoswitchable fluorescence microscopy

and fluorescence resonance energy transfer (FRET) microscopy. The real-time trafficking of

hDAT in a response of the compound like DA and amphetamine has been studied by total

internal reflection fluorescence (TIRF) microscopy.39-41 Lukyanov and coworkers reported the

position and movement of hDAT protein in living cells using photoswitchable cyan fluorescence

protein39, similarly, Sorkin and coworkers reported hDAT interaction and oligomerization during

trafficking in living cell40 utilizing FRET microscopy. However, solid information about the

atomic level interaction between DA and hDAT was first time reported by Gouaux and

coworkers after their X-ray-crystal based study of the dopamine transporter (dDATmfc) 54,55 They

have explained the molecular principle to distinguish the binding pattern of chemically distinct

ligands to DAT in the vicinity of sodium and chloride ions. In recent years, computational

molecular dynamic (MD) simulation has been applied to study atomic-level interactions between

hDAT and DA.19,46-53 Bahar and coworker have reported molecular mechanism of DA transport

by hDAT using homology modeling and full-atomic microsecond simulation.19 De Felice and

coworker used voltage clamp method to understand the stimulatory, and inhibitory action of DA

and other drugs in neurophysiological system.58

All the conventional techniques applied to study biological sample have intrinsic

advantages and limitations. For instance, fluorescence microscopy is one of the powerful tools

for biological imaging; however, photo-bleaching effect, phototoxicity of shorter wavelength

light to living cells, and toxicity due to byproduct formed in situ photochemical reaction are its

limitation.59-65 In addition, the biological property of the sample could be changed after

fluorescence labeling of samples. Xu and coworkers found the change in efflux function of

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different transporters when they were tagged with a fluorescent protein.66,67Although the PET

and SPET techniques are strong techniques to study protein trafficking and their interaction in

biological samples, the radioactive waste produced in this technique is harmful to in vivo studies.

Thus, it is highly desirable and necessary to develop and demonstrate other powerful technique

to study biological samples. Here, we found those qualities in SERS technique to study DA-

hDAT interaction in a live HEK-293 cell.

Raman spectroscopy was first used to study bacteriorhodopsin as a biological sample;68

salmon's sperm cell as living cells,69 and eosinophilic granulocytes cells as human living cells.70

Now, it has been established as a powerful analytical technique to study the broad range of

biological samples and living systems. Initially, Raman study of membrane proteins was very

difficult due to their low concentration at the cell membrane, extremely small Raman scattering

cross-section and very weak Raman scattering intensity compared to background noise.

However, the sensitivity of Raman spectroscopy has been improved tremendously by using

different approaches The technical advancement in Raman spectroscopy has greatly increases

significance of this techniques to study wide varieties of samples.68,71-111 Different forms of

Raman spectroscopy are extensively used to study the wide range of molecules and their

interactions in different physical conditions, which are evidence of its importance.112-158 Surface

Enhanced Raman Scattering (SERS) is one of the powerful technique to improve Raman

sensitivity, which is done by exploiting electromagnetic field enhancement and chemical

enhancement mechanism using metallic substrate especially nanoparticles or rough surface of a

noble metals.136,159-161 Van Duyne and coworkers have quantitatively reported in vivo

transcutaneous glucose sensing by using surface-enhanced spatially offset Raman spectroscopy

(SESORS).162 Henry and coworkers applied dark field microscopy and SERS technique

80

combining nanotags with a microfluidic device for the continuous detection of biomarkers in

blood.163 Recently, Haynes and coworkers have reported the sensing of ricin B chain in human

Figure 3.2. DFT calculated and experimental Raman spectrum of DA and cell samples.DFT

Calculated Raman spectrum of (A) unbound DA, and (C) DA in dDATmfc are calculated by

Gaussian 09 with B3LYP/6-31G(d) basis functions, scaling factor 0.9614. Experimental SERS

spectrum from (B) DA, (D) HEK293 cell, (E) hDAT-HEK293 cell, (F) DA-HEK293 cell and

(G)DA-hDAT-HEK293 cell are collected using silica coated silver nanoparticle as SERS

substrate, with excitation frequency 488.16 nm.

81

blood applying aptamer-conjugated silver film-over-nanosphere substrate, and SERS

technique.164 Moskovits and coworkers developed and synthesized SERS-based biotags (SBTs)

with highly reproducible optical properties and correctly identified the cancerous cells using this

SBTs on the mixture of cancerous and noncancerous prostate cells utilizing deconvolution

strategies.165,166 Nie and coworkers have reported biocompatible and nontoxic pegylated gold

nanoparticles and SERS technique for in vivo targeting and detection of tumor.167 Recently, they

have developed highly sensitive SERS-based assays which can detect 1 stem cell among

106 cells.168 The cytotoxicity on the living cell due to bare metal nanoparticles used as SERS

substrate is one of the detriments of SERS technique;169 which can be solved by using an

ultrathin layer of silica on SERS substrates. We have used a thin layer of silica (4-6 nm) over

silver nanoparticles to prevent cytotoxicity without reducing electromagnetic field enhancement.

Moreover, the metal nanostructures used in SERS technique quenches fluorescence background

giving higher signal-to-noise ratio, reduces photo-bleaching, and water background effect and

making SERS as a strong analytical tool for varieties of biological study including protein

structure, its interaction to other protein or ligand molecules.170-177 Coronado and co-workers

have studied the influence of protein kinase D1 on identification, localization, and quantification

of neuronal cell membrane receptor using SERS and other plasmonic probes.178 Ben-Amotz and

co-worker reported the ultra-filtration Raman difference (UFRD) method to obtain

thermodynamics and structural information after label-free detection and quantitation of protein-

ligand binding.177 Some research groups have already reported the interactions between

neurotransmitters and their binding protein using Raman spectroscopy.178,179 Peticolas and co-

worker applied the time-dependent UV Raman spectroscopy to observe the conformational

change of the acetylcholine and its analog during due to binding with protein. The probing of the

82

binding was based on changes in frequency and excited state electronic structure that brings

variations in the relative intensity of the Raman bands.179 In this chapter, the SERS technique is

applied to probe progress of DA-hDAT interaction by analyzing the change in Raman mode of

DA molecules.

Table 3.1. Prominent Raman wavenumber of DA, HEK293 cell, and hDAT-HEK293 cells

The SERS spectra from DA, HEK293, hDAT-HEK293, DA-HEK293, and DA-hDAT-

HEK293 has been collected and converted into 2D-distribution Raman spectrum to analyze DA-

hDAT interaction in the live HEK293 cell. Analysis showed that new Raman modes present at

807, 839, 1076, 1090, 1538, and 1665 cm-1 in DA-hDAT-HEK293 are markers of DA-hDAT

interaction. Similarly, the analysis of DFT calculated Raman spectrum of bound and unbound

states of DA shows that Raman modes at 807, 894, 971, 1076, and 1403 cm-1 are signature to the

bound state of DA (Figure 3.2A and 3.2C). Our rigorous analysis shows that only two Raman

83

modes present at 807 cm-1 and 1076 cm-1 are signature marker Raman shifts of bound states of

DA molecules, which are capable to identify and probe DA-hDAT interactions. Other modes

present at 894, 971, and 1403 cm-1 are not used to characterize the DA-hDAT interaction since

they are overlapped on Raman shift of hDAT or other cellular proteins.

3.2. Experimental sections

3.2.1. Synthesis of silver nanoparticles and sample preparation

Silver nitrate (AgNO3), sodium citrate, and sodium silicate were purchased from Sigma Aldrich

and were used as received. Silver nanoparticles (AgNPs) is synthesized by a standard sodium

citrate reduction method,180 followed by the addition of active sodium silicate to generate ultra-

thin silica shells over AgNPs.181 The synthesized nanoparticles are characterized by

Transmission electron microscopy (TEM), and UV-Vis spectroscopy. The size of AgNP@SiO2

is found between 50-65 nm with SiO2 shell thickness 4-6 nm. The absorption maximum of

AgNP@SiO2 is blue shifted in UV-vis spectrum with respect to bare AgNPs.

Prof. Louis J De Felice (Virginia Commonwealth University, Richmond, VA) had

generously gifted the hDAT inducible HEK293 cells. The vial of frozen cells was thawed

quickly in a 37oC water bath and then decontaminated by spraying with 70% ethanol and

transferred into a T-75 flask containing 15 ml of culture medium containing Dulbecco's Modified

Eagle's Medium (DMEM, Sigma-Aldrich D5796) supplemented with 10% fetal bovine serum

(Sigma-Aldrich, F2442) and 1% penicillin-streptomycin (ATCC, 30-2300). The cells were then

maintained in an incubator at a temperature of 37oC with 5% CO2 atmosphere. After 24 hours,

the medium is aspirated off and replaced with fresh, complete medium containing 1×10-4 gm/ml

zeocin (invivoGen, ant-zn-1), and 1×10-3 gm/ml blasticidin (Millipore/Calbiochem, 203350).

When the cells reached 70-80% confluence on the surface of T-75 flask, hygromycin B (Sigma-

84

Aldrich, H3274) of final concentration 25 µg/µl was added in to it to select hygromycin

resistance cells, which are isolated, and subculture in complete media on a 25 mm circular cover

glass in 35 mm petri dish. When cells are attached (approx. 24 hrs) on the cover glass,

doxycycline (Tocris Bioscience, 4090) was added at a concentration of 0.5 µg/µl. In 2 to 3 days

hDAT expression was completed in the HEK293 cells using the protocol as explained in in

chapter 2. Before SERS experiment, samples of HEK293 cells, and hDAT-HEK293 cells were

washed with PBS solution to remove culture medium, and wet with 250 µl fresh PBS solution to

prevent drying of the cell. DA-hDAT interaction is established adding 50 µl of 50nM DA

solution into cell sample at 15 minutes prior to SERS experiment.

3.2.2. Surface-enhanced Raman measurements

All SERS spectra were collected using home-modified confocal Raman microscope182

(Figure 2.8 in chapter 2) using 488-nm continuous-wave (CW) argon ion laser of approximately

10-15 µw power with an integration time of 30 seconds. Mercury lamp and cyclohexane were

used to calibrate the setup before Raman measurements with spectral resolution 2 cm-1, the range

of SERS spectrum was set to 700–1700 cm-1 and experimental parameters are set identical for

each record to avoid parameter's effect. Non-reproducibility of Raman spectrum (i.e. fluctuation

of peaks position and intensity) is removed by generating 2D-distribution Raman spectrum.

3.2.3. Two-dimensional SERS plot vs relative signal peak intensity

The temporal SERS fluctuation observed in individual spectrum makes it complicated to

determine the exact value of wavenumber for a Raman mode. The two-dimensional SERS plot of

spectral mode frequency for a sample is obtained by the combination of clearly visible Raman

peaks of 50-60 normalized SERS spectra. In this picture, Raman shift is represented by colorful

spot instead of peak, where color scale shows the recurrence of the Raman mode. The

85

broadening of the spot along x-axes shows the trend of wavenumber fluctuation for Raman

modes over numbers of SERS spectra. The mean position of the spot is taken as the average

Raman frequency. The fluctuation in relative intensity of a Raman mode could be noticed from

broadening or splitting of the spot along y-axes. The colorful spot is promising to select

prominent Raman modes and helps to get rid of signal fluctuation problem. Besides this

advantage, 2D-distribution Raman spectrum is generated using clearly visible Raman peaks from

normalized SERS spectra, hence it is always free from noise and unusual peaks. High degree of

consistency is found between individual SERS spectrum and corresponding 2D-distribution

Raman spectrum of a sample neglecting some minor discrepancies. Hence, the 2D-Raman

spectrum is more powerful than relative signal-peak intensity SERS spectrum.

3.2.4. Density functional theory calculations

Geometry optimization and Raman frequency calculations were performed using density

functional theory (DFT) method on the B3LYP level with a basis set of 6-31G (d) and Gaussian

09 package to see the difference between free DA and DA bound to hDAT. According to a

comprehensive evaluation of Scott and Radom183, the obtained frequencies were scaled by a

factor of 0.9614. Molecular orbitals were calculated with the same basis set and visualized with

Avogadro software (Avogadro: an open-source molecular builder and visualization tool). All

calculations were carried out on a vector processor (Ohio Supercomputer Center, Columbus,

Ohio).

3.3. Results and discussion

All the SERS spectra for DA, HEK293 cell, DA in HEK293 cell (DA-HEK293), hDAT

expressed HEK293 cell (hDAT-HEK293) and DA with hDAT-HEK293 as represented in Figure

3.2 were recorded using home-modified Raman microscope (Figure 2.8 in chapter 2) to

86

determine the binding interactions between DA and hDAT in a live cell. The analysis of SERS

spectra of DA, HEK293 cell, DA-HEK293, hDAT-HEK293, and DA-hDAT-HEK293 are

remarkably useful to probe DA-hDAT interactions. The expression of the hDAT protein in

HEK293 cell, the interaction of DA to HEK293 cell and human dopamine transporter (hDAT)

could be noticed by the spectral change in the SERS spectrum of DA, HEK293 cell, and hDAT-

HEK293 cell (Figure 3.2B-G). The analysis is easier and precise when 2D-distribution Raman

spectrum is generated (Figure 3.3A-E). At first, the wavenumber of prominent spots in 2D-

distribution Raman spectrum is determined based on intensity and their recurrence (Table 3.1).

The remarkable spectral difference is found in HEK293 cell after expression of hDAT protein

showing five extra peaks located at 1012, 1110, 1311, 1463, and 1578 cm-1 with the comparison

of the HEK293 cell. These new peaks might be contributed by hDAT since the new contributor

is only the hDAT protein. New Raman modes appeared in a hDAT-HEK293 cell could be

assigned on the basis of literature and our DFT calculation (table 3.3).184 The tentative

assignments for the Raman mode of hDAT protein are as follows. Raman mode at 1012 cm-1

could be related to (1010 cm-1) deformation frequency of C-O-H bond of alcoholic hydroxyl

group present in amino acid residues. The possible contributors of Raman mode at 1110 cm-1

could be summarized as follows: (1113-1115 cm-1) C-asymmetric bending, Cα-Cβ stretch, Nt-Cα,

Cβ-Cα-Ct stretch (Ala), and (1108-1113 cm-1) NtH3+ asymmetric rocking of (His). Similarly,

Raman mode of hDAT at 1311 cm-1 is given as (Near 1300 cm-1) amide III band, 40% C-N

stretch, 30% N-H bending, and 30% skeleton stretches, (1308-1310 cm-1) ring stretch of

phenylalanine, (1304-1308 cm-1) Hα-Cα-Ct, COO- symmetric stretching, Cβ-Cα-Ct (Ala), and

(1308-1310 cm-1) νring (Phe). Raman mode at 1463 cm-1 could be originated from synergic

contribution of (1464 cm-1) C-asym rocking (Ala), (1464-1470 cm-1) Cδ-bending (Lys), (1463-

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1464 cm-1) Cβ asymmetric rocking (Ala), and (1463-1465 cm-1) Cγ1 asym rocking, Cδ asym

bending (Ile). The Raman mode at 1578 cm-1 could be related to (1566 cm-1) C=N stretch, and

(1582 cm-1) structural stretching of Phe and Trp.

Figure 3.3. 2D-distribution Raman spectrum of (A) DA, (B) HEK293, (C) hDAT-HEK293, (D)

DA-HEK293, (E) DA-hDAT-HEK293 cellgenerated from 40-50 high quality normalized SERS

spectra of corresponding samples where color scale represents the occurrence of Raman modes,

higher occurrence showing prominent Raman modes. Red circled Raman modes in 3C and 3E

represents characteristic Raman modes of hDAT, and DA-hDAT interaction respectively.

88

Beside these new Raman shifts, original Raman shifts of the HEK293 cell are still present

in a hDAT-HEK293 cell (Figure 3.2E). When DA is added into hDAT-HEK293 cells and waited

for 5-10 minutes, new Raman modes for DA-hDAT interaction are observed. Characteristic

Raman modes located at 807, 839, 1076, 1090, 1538 and 1665 cm-1 are a landmark to DA-hDAT

interaction which is represented by red circles in Figure 3.3E. These new Raman modes are

nowhere present in Raman spectrum of DA, hDAT-HEK293, HEK293 cell, and DA-HEK293

themselves, and only appeared in the DA-hDAT-HEK293 system. The new Raman modes

observed in the DA-hDAT-HEK293 system could be contributed by different sources. The new

bond formed between DA and protein residues of hDAT could be one of the chief contributors;

crystal structure of hDAT analog supports those types of the bond formation during DA

bindings. The change of electronic density in the bonds of DA molecule due to DA-hDAT

interaction could be next contributors; the change in electronic density of DA is supported by

DFT calculation (Figure 3.4B). The change of electronic density within the structure of protein

residue of hDAT could be another contributor of new signature Raman modes. However, the

contribution from all factor are not equally relevant for analysis. For instance, the dynamics of

DA reuptake by hDAT is reported to be 3.3 µM/S-4.0 µM/S which means bond formation and

breaking phenomenon are faster process184,185, hence analysis of bond formation between DA

and protein residue is not preferred to study DA-hDAT interaction. Similarly, it is harder to

study DA-hDAT interaction by probing the change in Raman modes of many protein residues

available in hDAT.

89

Figure 3.4. (A) Crystal structure of dDATmfc (PDB ID: 4xp1)showing the binding site of DA;

(B) Calculated HOMO orbitals for unbound and bound states of DA, visualized with Avogadro

(Avogadro: an open-source molecular builder and visualization tool.

Version1.XX.http://Avogadro.openmolecules.net).

90

Table 3.2. Raman peak assignment of dopamine (DA).

Ultimately, the change in Raman mode of DA due to redistribution of electronic density

is preferred to probe DA-hDAT interaction. This approach is even justified since DA is the

center of DA-hDAT interaction, which is discussed thoroughly as following. The crystal

structure of dDATmfc (PDB ID: 4xp1) mimics the structure of hDAT (Figure 3.4A), which shows

the location of DA molecule within central binding site surrounded by the transmembrane

helices (TMs). The amine group of DA interacts with the carboxylate group of Asp46 within a

distance of 3A˚; while its catechol group is enclosed in a cavity formed by Ala117, Val120,

Asp121, Tyr124, Ser422 and Phe325 and it interacts with the carboxylate group of Asp121

through hydrogen bonding. The meta-hydroxyl group of DA molecules interacts with the side

chain of Asp121 at 2.7A˚ where it is oriented towards Ser422 in TM8 at 3.8A˚. The para-

91

hydroxyl group interacts with both the carbonyl oxygen of Ala117 and the carboxylate of

Asp121 at distances of 2.8 and 3.1A˚ respectively. The coordinates of DA molecule located in

the binding site of dDATmfc crystal has been used to calculate HOMO orbital and DFT Raman

spectrum of bound states of DA. HOMO orbital calculation (Figure 3.4B) of bound and unbound

states of DA shows a variation of electron density in bound state of DA molecule in O11-H22, O10-

H21, C3-C2=C1, N9-H19, and C7-C8 bonds with respect to its unbound state. Raman frequency

calculations of unbound states of DA is performed with geometry optimization by density

functional theory (DFT) method on the B3LYP level with a basis set of 6-31G (d) using

Gaussian 09 package. However, the DFT Raman calculation of bound state of DA is carried out

with the same basis set without geometry optimization, to conserve the coordinates of atoms, as

they are present in the binding cavity of the crystal structure.

Figure 3.5. Characteristic Raman peaks at 807 cm-1 and 1076 cm-1 of integrated Raman

spectrum of the DA-hDAT-HEK293 cell (averaged from 32 spectra).Characteristic Raman peaks

of bound states of DA molecules are associated to DA-hDAT interaction. Force vectors (green

92

arrow) in the structure of DA show Raman vibrational mode at 807 cm-1 and 1076 cm-1 in bound

states of DA molecules.

Table 3.3. Raman peak assignment of hDAT-HEK293 cell.

In a biological system, DA molecules are present in soluble form, hence experimental

SERS spectrum of DA molecules are required to collect both from solution and dried states to

93

test the solvent effect. We found SERS spectra of DA in solution state are consistent to that from

dried states; the only exception is that there is broadening effect in some Raman peaks in a

solution state of DA. The Raman mode assignment of unbound DA molecule based on our DFT

calculation and published literature,87 are shown in table 3.2. The Raman peaks of unbound DA

observed in our experiment are consistent with theoretical calculations (Figure 3.2A and 3.2B).

However, the experimental peaks for bound state of DA are not consistent (Figure 3.2G) with

Figure 3.6. The effect in vibrational mode at 807 cm-1, and 1057 cm-1 as bupropion interferes

DA-hDAT interaction after adding bupropion into DA-hDAT-HEK293.SERS spectrum of (A)

DA-hDAT-HEK293, (B) Bupropion-hDAT-HEK293, and (C) Bupropion-DA-hDAT-HEK293;

SERS spectra of Bupropion-DA-hDAT-HEK293 are collected after 10 min of bupropion

addition.

94

Figure 3.7. SERS spectra from Bupropion.The bupropion solution (50 mM) was incubated

overnight in AgNP@SiO2 colloids before collection of SERS spectrum; the integration time for

SERS collection was 10 second and power of excitation laser was 10-15µW.

DFT calculated Raman peaks (Figure 3.2C). It might be because experimental SERS spectrum of

bound DA is recorded from DA-hDAT-HEK293 sample, and it could be mixed up with Raman

peak of hDAT, HEK293, and unbound DA. On the other hand, DFT Raman spectrum of bound

DA is obtained by using coordinates of DA molecule located in the binding cavity of dDATmfc

95

crystal and hence should be free from any spectral disturbance. Analysis of DFT Raman

spectrum of unbound and bound states of DA is convenient to determine characteristic Raman

mode due to DA-hDAT interaction (Figure 3.2A and 3.2C). It shows that out of five new Raman

modes of bound states of DA, only two Raman mode which are located at 807, and 1076 cm-1

stand out as a landmark of DA binding to hDAT because other new Raman modes are

indistinguishable from Raman modes of unbound state of DA, HEK293 cell, or hDAT protein.

The visualization of DFT calculated vibrational mode of bound states of DA molecule

shows that Raman mode at 807 cm-1 is related to combined contribution of following bending

processes: α(C3-O10-H21), α(C2-O11-H22), α(C7-C8-H18), α(C6-C4-H13), α(C7-C8-H19), α(C7-C8-

N9). Similarly, another Raman mode at 1076 cm-1 is found to be related to a combination of

bending processes: α(H19-N9-C8), γ(N9-H19), γ(C8-H19), γ(N9-H20), γ(C8-H18), and α(C7-C8-H18).

We found these types of study is useful to show the progress of DA-hDAT interaction by

probing appearance and disappearance of the intensity of these two peaks. If any drugs, for

instance, psychostimulant, anti-depressant or potent are applied to replace DA molecules from its

binding sites, the relative intensity of these characteristic peaks present at 807 cm-1 and 1076 cm-

1 in SERS spectrum could be disappeared or decreased due to the interference of DA-hDAT

interaction. This statement is supported by the result of our control experiment (Figure 3.6 and

3.7).

3.4. Summary

We have recorded SERS spectrum of DA, HEK293, hDAT-HEK293 cell, DA-HEK293,

and DA-hDAT HEK293 cell to analyze DA-hDAT interaction. Overexpression of hDAT protein

in HEK293 cell is remarked by new Raman shift in the hDAT-HEK293 cell. Similarly, the

progress of DA-hDAT interaction is noted by the appearance of new Raman mode after addition

96

of DA into the hDAT-HEK293 cell. Our emphasis in this study is finding of marker Raman shift

of bound state of DA molecule to examine DA-hDAT interaction. The careful analysis of

experimental and DFT Raman spectrum of DA in bound and unbound states shows that Raman

shift of bound states of DA at 807 cm-1 and 1076 cm-1 are benchmarks for DA-hDAT interaction.

On the basis of our experimental and theoretical study, it is found that the characteristic Raman

shifts of bound states of DA observed at 807 cm-1 corresponds to the bending modes of 𝛼(C3-

O10-H21), 𝛼(C2-O11-H22), 𝛼(C7-C8-H18), 𝛼(C6-C4-H13), 𝛼(C7-C8-H19), and 𝛼(C7-C8-N9)), while

Raman shift at 1076 cm-1 corresponds to bending modes of 𝛼(H19-N9-C8), 𝛾(N9-H19), 𝛾(C8-

H19),𝛾(N9-H20), 𝛾(C8-H18), and 𝛼(C7-C8-H18). Our study concludes that protein-ligand

interaction could be confirmed by probing changes in Raman shift of ligand molecules, which

could be crucial for understanding molecular interactions of neurotransmitters with their

corresponding receptor or transporters.

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109

CHAPTER 4. MODE-SELECTIVE RAMAN IMAGING OF DOPAMINE-HUMAN

DOPAMINE INTERACTION IN LIVE CELLS

Dopamine (DA) is the catecholamine neurotransmitter, interacting with both dopamine

receptors (DARs) and dopamine transporters (DAT) to generate dopaminergic signals and

maintain its homeostasis in synaptic and perisynaptic space. Roles of the dopamine system in the

central nervous system are associated with the targeted binding of drugs to the DAT. Thus, the

interaction of DA, or its analog with DARs, or DAT has been studied extensively to uncover the

mechanism of dopamine system and dopaminergic signaling process. However, there is still a

lack of risk-free, label-free, and minimally invasive imaging approach to probe the interaction

between DA and DAT or DARs. Here, we have probed the location and interaction of dopamine

(DA) and human dopamine transporter (hDAT) by a spatial selection of signature Raman mode

using AgNP@SiO2-nanoparticle based surface-enhanced Raman scattering (SERS) approach.

We utilized signature Raman frequency 1287 cm-1 of DA to probe its location, and 807 and 1076

cm-1 of a bound state of DA to visualize the Raman image of DA-hDAT interaction in living

cells. We also utilized two-photon excitation (2PE) fluorescence imaging approach for the quick

probing of DA in live HEK293 cells. Our approach of mode-selective Raman imaging is

successful to generate background free image of DA, hDAT and DA-hDAT interactions in living

cells. This approach could be extended to explore the signaling proteins, neurotransmitters, and

several types of proteins-ligands interactions. The probing of signaling proteins,

neurotransmitters and other targeting drugs in living cells is crucial for the diagnosis and cure of

several neurodegenerative diseases. Our experimental approaches could be a convenient and

powerful technique for probing these types of interactions.

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4.1. Introduction

Dopamine (DA) is the biogenic amino neurotransmitter that plays a significant role in the

behavioral and psychological activities of human beings.1-6 DA neurotransmitter diffuses into the

synaptic cleft by the sequence of activities. It starts with the generation of the action potential

from the environmental stimulation which opens the voltage-gated Ca2+ ion channel and allows

Ca2+ flux to enter neuron cells. The elevation of Ca2+ ions inside cells stimulates the

degeneration of synaptic membrane and diffusion of DA into the synaptic cleft.7,8 The diffused

DA molecules interact with dopamine receptors (DARs) located at the postsynaptic region of

neighboring dendrites and generates dopaminergic signals.9-12 The prolonged interactions of DA

with DARs cause hyperstimulation of DARs which is detrimental for the well-functioning of the

dopaminergic system. The prolonged DA-DARs interactions and hyperstimulation of DARs are

avoided by the Na+ and Cl- assisted reuptake process of DA which is regulated by the dopamine

transporter (DAT).13-16 They are located at presynaptic region (axon terminal) of the neuronal

junction, which interacts with DA to pull them back to the cytosol of the neuron. The interactions

of DA with DAT and DARs are ongoing processes in synaptic cleft which determine the

effective functioning of dopaminergic systems. The imbalance in these activities is one of the

reasons for psychiatric and neurological disorders such as depression, schizophrenia, Parkinson’s

disease, and Alzheimer’s disease. The probing of DA, DARs, DAT and their interactions in the

nervous system is the prerequisite for the mapping of neural circuits and revealing the

pathophysiology of many neurodegenerative diseases and gaining new visions to develop

therapeutic treatments.17-20

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Figure 4.1. The two-photon excited (2PE) fluorescence images from baseline HEK293 cells.The

2PE image from (A) cleaned empty cover glass (B) spin coated 30 µl solution of 100 nM DA.

(C) HEK293 cells before addition of DA. (D) HEK293 cells after addition of DA. The

fluorescence signal intensity of HEK 293 cells is increased by 41 ± 6 % after addition of DA. (E)

and (F) are mode-selective Raman images of HEK293 cells corresponding to 1414 cm-1 and

1145 cm-1 respectively. For 2PE fluorescence imaging approach, frequency doubled femtosecond

532 nm pulsed laser (Chameleon Discovery, coherent, ~ 100 fs fwhm) of average power 15±2

µW is used.

The different types of experimental approaches such as fluorescence microscopy, positron

emission microscopy (PET), single photon-electron tomography (SPET), X-ray

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crystallography and computational molecular dynamics (MD) simulation have been applied to

explore the facts on structures and functions of several signaling proteins.21-41 Booij and

coworker used single photon emission computed tomography (SPECT) to observe the

relationship between Parkinson’s diseases and loss of human dopamine transporters (hDAT) in

the striatal region.27 The photo switchable fluorescence microscopy and fluorescence resonance

energy transfer (FRET) microscopy has been used to study the modes of hDAT trafficking.

Similarly, the total internal reflection fluorescence (TIRF) microscopy has been used to study the

real-time trafficking of hDAT in the response to the signaling molecules like DA and

amphetamine.22,23,42 Despite the fact that the fluorescence imaging has excellent results in several

circumstances, the labeling molecules used in this technique are not useful to probe DA, DAT,

DARs and their interactions since the sizes of labeling molecules are usually larger than DA

molecules. The label-free approaches are highly desirable to probe the interactions of DA and

other signaling molecules with membrane proteins like DAT or DARs. In the search of label-free

imaging methods to probe neurotransmitters in living cells, Webb and coworker first time used

the multiphoton fluorescence imaging approach to probe the serotonin neurotransmitters in rat

basophilic leukemia cells.43 Similarly, Maiti and coworkers utilized two photons fluorescence

imaging of dopamine in living cells and tissues for the quantitative mapping of DA

concentration.44 The aforementioned approaches including two photons excitation methods are

typically used for the mapping of signaling proteins and quantitative measurement of the

neurotransmitters content in cells or tissues. However, these techniques are not convenient for

probing the atomic level interaction between neurotransmitters and signaling proteins. In this

context, Gouaux and coworkers are the first who utilized X-ray-crystallography based studies to

resolve the atomic level interaction of DA with drosophila dopamine transporter (dDATmfc).36,37

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Computational modeling and X-ray crystallographic studies provide the important concept about

the interaction of DA with dopamine transporter, but there is still a need of minimal invasive and

effective technique to study DA-hDAT interaction in living system. We found that surface-

enhanced Raman spectroscopy is important approach to fulfill this purpose.

Surface-enhanced Raman spectroscopy (SERS) is already established as a powerful

experimental approach for the characterization of dopamine-human dopamine transporter (DA-

hDAT) interactions in living cells.45 Other forms of Raman spectroscopy are also successfully

utilized to study the huge range of molecules and their interactions in different physical

conditions.46-92 However, this has been established as more powerful analytical approach for the

label-free characterization and analysis of biomolecules like neurotransmitters, enzymes,

membrane protein, and highly organized systems such as membrane preparation and

photosynthetic bacteria.45,93-99 In this chapter, we discuss the combined approach of SERS

technique and two photons excited (2PE) fluorescence imaging to probe the DA-hDAT

interaction in living cells. In SERS technique, roughen metallic surface or nanoparticles of gold

or silver are used as a substrate which increases the Raman cross-section of the probe molecules

by electromagnetic (EM) and charge transfer (CT) mechanism.100 Although, SERS has important

applications in many fields, metal nanoparticles used in SERS has some disadvantage, for

example, proteins or enzymes are denatured due to direct interaction with Ag+ ions or AgNPs

during SERS experiments. The denaturation of proteins or enzymes is reported due to bond

formation between Ag+ ions or AgNPs with sulfhydryl groups of proteins.101,102 Therefore,

prevention for the direct contact between metallic SERS substrates and biomolecules is highly

required and recommended to avoid the denaturation problem of protein or enzyme. To achieve

this purpose, we have fabricated ultra-thin layer (4-6 nm) of silica (SiO2) on the surface of silver

114

nanoparticles (AgNPs), which effectively prevents the possible damage of biomolecules by Ag+

ions or AgNPs without reducing the electromagnetic field enhancement of Raman cross-

section.53,103 The silica-coated silver nanoparticles (AgNP@SiO2) are also used to obtain

fluorescence free and high-quality Raman spectra. It is reported that metal nanostructures have a

higher fluorescence quenching efficiency in contrast to the quenching efficiency of protein or

potassium iodide.104-106 In our SERS experiment, the metal nanostructures help to quench the

fluorescence background produced from the intracellular content of HEK293 cells. In the search

of background free, and label-free imaging approach to probe the atomic level interaction

between dopamine (DA) and human dopamine transporter (hDAT) in living human cells, here,

we have developed the mode-selective Raman imaging technique. This experimental approach is

found useful for the precise mapping of DA-hDAT interactions in living cells and could be

important for the detection of many protein-ligand interactions in native biological systems.

4.2. Experimental sections

4.2.1. Synthesis of silica-coated silver nanoparticles

Silver nanoparticles (AgNPs) are synthesized by a standard sodium citrate reduction

method,107 followed by the addition of active sodium silicate to generate ultra-thin silica shells

over AgNPs.108 Silver nitrate (AgNO3), sodium citrate, and sodium silicate required for the

synthesis of nanoparticles were purchased from Sigma Aldrich, and used without further

purification. Then, the synthesized nanoparticles were characterized by transmission electron

microscopy (TEM) and Varian UV-Vis spectrophotometer (EL07013173). According to our

TEM measurement, the average sizes of silica-coated silver nanoparticles are found 58 ± 8 nm

with SiO2 shell thickness 5 ± 1 nm. The UV-Vis measurement shows that the maxima (λmax) of

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the plasmon resonance band position related to silver nanoparticles is found at 420 nm which

was shifted to 407 nm after generating the ultra-thin layer of silica on its surface.45

4.2.2. HEK293 cells culture

The vial of frozen HEK293 cells are thawed quickly in a warm water bath (37oC) and

decontaminated by spraying 70% ethanol. Then cells are transferred into a T-75 flask with 15 ml

of complete medium. The complete medium contains Dulbecco's Modified Eagle's Medium

(DMEM, Sigma-Aldrich D5796), 10% fetal bovine serum (Sigma-Aldrich, F2442), and 1%

penicillin-streptomycin (ATCC, 30-2300). Then T-75 flask containing cells is stored in the

incubator at 37oC temperature with 5% supply of CO2 atmosphere. After 24 hours, all the

complete medium from T-75 flask is aspirated out and replaced by fresh complete medium.

When a confluence of the cells reached to 70-80% on the surface of the T-75 flask, cells are

taken out from the surface of T-75 flask using trypsin EDTA and sub-cultured on a 25-mm

circular cover glass in 35 mm petri dish.

4.2.3. Preparation of hDAT inducible HEK293 cells

After 24 hours of subculture, we followed the standard protocol of Invitrogen for the

preparation of hDAT inducible HEK293 cells. In brief, all the complete medium from the T-75

flask is aspirated out and replaced by fresh complete medium containing 1×10-4 gm/ml zeocin

(invivoGen, ant-zn-1), and 1×10-3 gm/ml blasticidin (Millipore/Calbiochem, 203350). When the

cells reached 70-80% confluence on the surface of the T-75 flask, hygromycin B (Sigma-

Aldrich, H3274) is added in its final concentration 25 µg/µl, which kills all the cells within 2

days except hygromycin resistance cells, which are isolated, and cultured in complete medium.

When cells are attached (approx. 24 hrs) on the surface of the T-75 flask, doxycycline (Tocris

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Bioscience, 4090) is added in its final concentration 0.5 µg/µl and waited for ≥ 48 hrs to obtain

hDAT inducible HEK293 cells.

4.2.4. Transfection of pcDNA3.1-hDAT in HEK293 cell

Plasmid DNA encoding pcDNA3.1-hDAT (32810) was purchased from Addgene and

amplified with a standard method. The amplified plasmid was treated with hDAT inducible

HEK293 cells for the expression of the human dopamine transporter (hDAT). The extraction of

pcDNA3.1-hDAT from bacterial cells and transfection of the plasmid in HEK293 has been

explained briefly in chapter 2.

4.2.5. Two photons excited (2PE) fluorescence imaging

Frequency doubled femtosecond 532 nm pulsed laser (Chameleon Discovery, coherent,

~100 fs fwhm) is used for the excitation of samples. The excitation laser is shone on samples

through 532 nm dichroic mirror (Chroma, ZT532rdc) and oil immersed objective of the inverted

microscope's. The diffraction limited (300 nm) epifluorescent light beam returns to the 532-nm

dichroic mirror and collected by single photon avalanche photodiode (APD) (PerkinElmer

SPCMAQR-14). The bandpass filter (FES0500) is positioned in front of APD to prevent entry of

any excitation laser and single photon excited (1PE) fluorescence. The detail of 2PE fluorescence

imaging is explained briefly in chapter 2.

4.2.6. Surface-enhanced Raman measurements

SERS spectra were collected using home-modified confocal Raman microscope109

(Figure 2.11 in chapter 2) using 488-nm continuous-wave (CW) argon ion laser of approximately

13±2 µw power. We used 30 second integration time for the recording of a SERS spectrum.

Mercury lamp and cyclohexane were used to calibrate the setup before Raman measurements

117

with spectral resolution 2 cm-1. For the analysis of conventional SERS spectrum, we selected the

range of 700–1700 cm-1. We maintained the same experimental parameters throughout the

experiment to avoid effect from parameters. For the collection of conventional SERS spectra,

long pass filter (HHQ495LP) was positioned in front of entrance slit of the monochromator

(Triax 550, Jobin Yvon). The Raman spectra were collected by an LN2-CCD (Princeton

instruments) which was cooled at about -100°C.

4.2.7. Mode-selective Raman measurement

The mode-selective Raman spectra are collected using experimental set up of SERS

measurements with some required modification (Figure 2.11 in chapter 2). In this experiment, we

replaced the long pass filter (HHQ495LP) of SERS measurement with the digital mini-chrom

monochromator ((DMC1-03). On the other hand, mode-selective Raman images are collected

using experimental set up of 2PE fluorescent imaging with its modification (Figure 2.11 in

chapter 2) For mode-selective Raman imaging, we replaced 532 nm fs pulsed laser by 488 nm

CW laser and used mini-Chrom monochromator instead of the bandpass filter (FES0500).


Geometry optimization and Raman frequency calculations were performed using density

functional theory (DFT) method on the B3LYP level with a basis set of 6-31G (d) and Gaussian

09 package to find the difference between free state and bound states of DA to hDAT. According

to a comprehensive evaluation of Scott and Radom110, the obtained frequencies were scaled by a

factor of 0.9614.

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4.3.1. Conventional and mode-selective Raman measurement

Our earlier work shows that Raman frequencies at 807 cm-1 and 1076 cm-1 are the

signatures of the bound states of DA molecules in human dopamine transporter (hDAT).45 Here,

we have reproduced and validated our earlier findings. Briefly, Raman spectral analysis of DA,

HEK293 cells, hDAT-HEK293 cells, DA-HEK293 cells, and DA-hDAT-HEK293 cells showed

that five new signature peaks at 807, 839, 1076, 1090, 1538, and 1665 cm-1 are related to the

interaction between DA and hDAT.45 These new five peaks could be contributed from several

changes that happen during DA-hDAT interactions.

Figure 4.2. The Raman spectrum and mode-selective Raman spectrum from DA and baseline

HEK293 cells.The Raman spectrum from (A) empty cover glass, and (B) DA coated cover glass,

appearance of Raman peak 1145 cm-1, and 1414 cm-1 respectively which shows the increase of

fluorescence signal in DA coated cover glass is contributed by DA only; (C) 1287 cm-1 mode-

selective Raman spectrum from DA coated cover-glass; (D) and (E) are Raman spectrum

119

corresponding to HEK293 cells before and after addition of DA respectively. mode-selective

Raman spectrum related to 1414 cm-1 and 1145 cm-1 are shown in figure (F) and (G)

respectively. The dotted red line and black line shows the consistency in mode-selective Raman

spectrum of HEK293 cells, and full range Raman spectrum of HEK293 cells before and after the

addition of DA. The dotted blue line shows the consistency appearance of Raman peak 1287 cm-

1 in full range Raman spectrum of DA, DA added HEK293 cells, and mode-selective Raman

spectrum of DA For Raman measurement 488 nm CW argon ion laser of average power 15±2

µW is used.

The X-ray crystallographic studies show the bond formation by hydroxy and primary

amine group of DA with protein residues of dopamine transporter (Figure 4.3).36,37 The direct

measurement of the bonds formation in living cell is difficult since it is faster dynamic processes;

particularly, the direct probing of bond formation using Raman experimental approach is very

difficult.111,112 The electron densities on protein residues, and DA are changed due to DA-hDAT

interaction, the change of electron density is associated to the Raman intensity which can be used

to probe the DA-hDAT interaction. The change in electron density of DA is more convenient and

effective to analyze the DA-hDAT interaction rather than the analysis of other proteins residues,

since DA has comparatively smaller and simpler structure than other protein residues.

Our DFT calculation shows that the electron density distribution of DA in its bound and

unbound states are different (Figure 4.9A). According to our DFT calculation, it is also found

that HOMO of bound states of DA is stable than it unbound states by 0.466 eV energy (Figure

4.9A). This stabilization energy interprets that the DA-hDAT interaction is an energetically

feasible process. In addition, DFT calculation shows that Raman wavenumber of a certain bonds

of DA is changed according to the change of its electrons density. The combined approach of

120

Raman experiments and theoretical calculations show that the two Raman peaks at 807 cm-1 and

1076 cm-1 are signature peaks for the bound states of DA, which could be utilized to probe the

progress of DA-hDAT interactions in living cells. The coordinates of DA for DFT calculation

was obtained from the crystal structure of dDATmfc (Figure 4.3). In this crystal structure, the

amine group of dopamine interacts with the carboxylate group of Asp46 at 3 A°. The catechol

group of dopamine is trapped in a cavity formed by Ala117, Val120, Asp121, Tyr124, Ser422,

and Phe325 and which interacts with the carboxylate group of ASP121 by the formation of

hydrogen bond. The meta-hydroxyl group of dopamine interacts with the side chain of Asp121 at

2.7 A° orienting itself towards Ser422 in TM8 at 3.8 A°. The para-hydroxyl group interacts with

the carbonyl oxygen of Ala117 and the carboxylate of Asp121 at distances of 2.8 and 3.1 A°

respectively.

Figure 4.3. The Crystal structure of dDATmfc (PDB ID: 4XP1)showing the binding site of

dopamine. The crystal structure of dDATmfc mimics structure of human dopamine transporter

(hDAT) where DA is located at the binding site surrounded by transmembrane helices (TMs)

121

The Raman frequency of bound states of DA is calculated without geometry optimization using

density functional theory (DFT) method on a B3LYP level with a basis set of 6-31 G(d), and

Gaussian 09 package. The geometry optimization is skipped for the bound states of DA to

conserve its coordinates. The DFT Raman calculation of unbound state of DA is carried out in

the same basis set with the geometry optimization. The analysis of DFT calculated Raman

spectra of bound and unbound states of DA (Figure 4.9 B) show that two Raman modes at 807

and 1076 cm-1 are landmarks for the bound states of DA. The analysis of DFT Raman and

experimental SERS spectra show a convenient way to probe the progress of ligand-protein

interaction.

In addition to DFT Raman calculation and conventional SERS study, here we have also

utilized the mode-selective Raman imaging approach which can visualize the high-quality and

background-free Raman images of DA-hDAT interactions in live HEK293 cells. The optical

requirements for our mode-selective Raman imaging approach are met by the combination of

488 nm continuous wave (CW) argon ion laser, inverted confocal microscope (Axiovert 135)

equipped with a XY piezo-controlled scanning stage (physic instrumente), oil immersion

objective (Zeiss FLUAR, 100×; 1.3 NA), digital mini-Chrom monochromator (DMC1-03), and

single photon avalanche photodiode (APD) (PerkinElmer SPCMAQR-14). The schematic

representation of experimental set up is shown in Figure 2.8 (Chapter 2). The photons from a

Raman frequency are selected using digital mini-Chrom monochromator and focused on APD to

obtain mode-selective Raman images. In addition to the mode-selective Raman images, we have

collected the mode-selective Raman spectra focusing same photons to spectrophotometer (Triax

550, Jobin Yvon). The consistency between the occurrence of mode-selective Raman image and

mode-selective SERS spectra is required to authenticate any mode-selective image. The

122

occurrence of mode-selective Raman images and spectra in our experiments are highly correlated

(Figure 4.7). For mode-selective Raman measurement, we allowed the photons corresponding to

Raman wavenumber 807 cm-1 and 1076 cm-1 (~508 nm and ~515 nm) to pass through digital

mini-Chrom monochromator. The mode-selective Raman images related to frequencies 1076 cm-

1 and 807 cm-1 are shown in figure 4.4E and 4.4F, respectively. The corresponding Raman peaks

of these frequencies are shown in figure 4.6E and 4.6F respectively.

Figure 4.4. Two-photon excited (2PE) fluorescence and mode-selective Raman images of

hDAT-HEK293 cells.2PE fluorescence image of (A) hDAT-HEK293 cells before addition of

DA and (B) after addition of DA. The fluorescence signal intensity of hDAT-HEK 293 cells is

increased by 42 ± 5 % after addition of DA. The mode-selective Raman images of hDAT-

HEK293 cells (C) before DA addition which are corresponding to 1463 cm-1 and are signature

for the hDAT protein; (D) after DA addition which are corresponding to 1287 cm-1 and are

signature for DA; (E) and (F) after DA addition which are corresponding to 1076 cm-1 and 807

cm-1 respectively and are signature for DA-hDAT interactions.

123

We tested the accuracy of our experimental results with many controlled experiments.

For that, we collected the mode-selective Raman images from HEK293 cells, DA-HEK293 cells,

and hDAT-HEK293 cells. The mode-selective Raman image of HEK293 cells was collected

using photons of Raman frequencies 1414 cm-1 (Figure 4.1E) and 1145 cm-1 (Figure 4.1F). The

mode-selective Raman image was confirmed as we were also able to collect mode-selective

Raman spectra corresponding to 1414 cm-1 (Figure 4.2F) and 1145 cm-1 (Figure 4.2G) from same

spot. Furthermore, we obtained the mode-selective Raman image (Figure 4.4C) and spectrum

(Figure 4.6C) from hDAT-HEK293 cells at the Raman frequency 1463 cm-1. According to our

experimental analysis, this Raman frequency is the signature for the hDAT protein. On the basis

of literatures, this Raman mode would be originated from the individual or combined

contribution of C-asym rocking of Ala, Cδ-bending of Lys, and Cγ1-asym rocking and Cδ-asym

bending of Ile.112 The assignments of other important Raman peaks from DA and cells are

provided elsewhere in this dissertation. In separate experiment, we have obtained the mode-

selective Raman image of dopamine (Figure 4.4 D) in HEK293 and hDAT-HEK293 cells

utilizing its signature Raman frequency 1287 cm-1. According to our DFT calculation and

literature, the signature Raman mode 1287 cm-1 is related to in-plane –CH2 bending vibration

arising from DA cationic form. We also obtained the mode-selective Raman spectra of this

frequency (Figure 4.6F). The concentration of dopamine, we used in our experiment is ~4µM,

which is a way smaller than the concentration of neuronal dopamine in the synaptic vesicles

(~100s of mM).113 The experimental results demonstrate that our approach is capable for the

direct imaging of dopamine present in brain cells.

124

4.3.2. Two-photon excited (2PE) fluorescence imaging approach

Before the mode-selective SERS experiments, we used the two-photon excited (2PE)

fluorescence imaging approach to visualize the quantitative changes of the fluorescence signal

from DA in HEK293 cells and hDAT-HEK293 cells. For that, we analyzed the 2PE fluorescence

images of HEK293 cells and hDAT-HEK293 cells at the absence and presence of DA. The red

spots in figure 4.1D and figure 4.4D represent the increase of fluorescence signal intensity in

HEK293 cells and hDAT-HEK293 cells respectively after addition of DA. We observe the

signature SERS peaks for DA, hDAT, membrane proteins, and their interaction in a SERS

spectrum originated from the bright red spot. The bright red spots obtained from 2PE

experiments, basically indicate the presence of DA in that area. However, this bright region does

not necessarily represent the interaction of DA with hDAT or other membrane protein. The

benefit of 2PE fluorescence is that: it works as a guide to find DA-rich region in cells where we

could expect a higher probability for DA-hDAT interactions. The interaction of DA, hDAT and

other membrane protein are probed and confirmed by the combined approach of mode-selective

Raman measurements and 2PE fluorescence imaging.

125

Figure 4.5. Evidences for the two-photon excited (2PE) emission.2PE emission spectra of 1mM

DA solution using frequency doubled 532 nm pulsed femtosecond laser (chameleon Discovery,

coherent, ~ 100 fs fwhm) (B) The emission spectra of same DA sample using 488 nm

continuous-wave (CW) laser (C) The excitation-emission spectra of 4 µM DA solution. (D) The

logarithm of fluorescence intensity as a function of logarithm of excitation laser power from 30

µl of 1mM air dried dopamine solution on cleaned cover glass (slope 1.91 ± 0.07). The

fluorescence counts from DA sample are recorded using single photon avalanche photodiode

(PerkinElmer SPCMAQR-14).

As we mentioned before, 2PE fluorescence imaging approach is important to find the

potential region of DA-hDAT interaction in HEK293 cells. The requirements for 2PE

fluorescence image is achieved by the combination of frequency doubled femtosecond 532 nm

pulsed laser (Chameleon Discovery, coherent, ~100 fs fwhm), inverted confocal microscope

(Axiovert 135) equipped with a piezo-controlled scanning stage (physic instrumente), oil

immersion objective (Zeiss FLUAR, 100×; 1.3 NA), and APD (PerkinElmer SPCMAQR-14).

The bandpass filter (FES0500) is positioned before the entrance slit of APD to collect the

fluorescence emission originated from two-photon excitations. The percentage transmittance of

126

the dichroic mirror (chroma, ZT532rdc) and bandpass filter (FES0500) show that they are

suitable optics to meet the criteria of 2PE (Experimental section, Figure 2.9).

Figure 4.6. The consistence between conventional and mode-selective SERS spectra.(A) and (B)

are the Raman spectrum from hDAT-HEK293 cells before and after addition of DA. (C) The

mode-selective Raman spectrum corresponding to Raman frequency 1463 cm-1 which are

signatures for the hDAT protein. (E) and (F) are mode-selective Raman spectrum corresponding

to 1076 cm-1 and 807 cm-1 respectively which are signature for DA-hDAT interactions. The

dotted lines show the consistency between signature peaks observed in mode-selective Raman

spectrum and conventional Raman spectrum.

The 2PE fluorescence images of HEK293 cells (Figure 4.1C) are originated from their

intracellular contents, the characteristic SERS spectrum of HEK293 cells is shown in figure

4.2D. When HEK293 cells are treated with 4 µM DA solution for ≥ 30 minutes, the 2PE

127

fluorescence is enhanced as the function of dopamine concentration. According to our

quantitative analysis, it is found that the fluorescence signal intensity of HEK293 is increased by

41 ± 6 % after addition of DA (Figure 4.1D). Figure 4.2C represents the mode-selective Raman

spectra from HEK293 cells after addition of dopamine. HEK293 cells were washed three times

with warm PBS buffer before 2PE fluorescence imaging experiments. Washing of cells is

important to remove the effect of extracellular DA fluorescence. We have also measured the

function of DA concentration in hDAT expressed HEK293 (hDAT-HEK293) cells. Here, we

analyze the 2PE fluorescence image of hDAT-HEK293 cells in the absence (Figure 4.4A) and

presence (Figure 4.4B) of DA. To test the sensitivity of the dopamine in hDAT-HEK293 cells,

the 4µM concentration of DA solution is treated with cells sample and waited for >30 minutes

for the incubation. Incubation time allows DA to enter hDAT-HEK293 cells and bind with

hDAT protein. The fluorescence signal intensity from hDAT-HEK293 cells is increased by 42 ±

5 % after addition of DA (Figure 4.2B). The 2PE fluorescence imaging of hDAT-HEK293 cells

are collected before (Figure 4.4A) and after (Figure 4.4B) addition of DA to probe the location of

DA and possible region for the DA-hDAT. The correlation between the appearance of

characteristic SERS spectrum of DA and increase of 2PE fluorescence signal intensity from cells

shows that 2PE fluorescence is contributed from DA. The mode-selective Raman image (Figure

4.4D) and SERS spectrum (Figure 4.2D) corresponding to Raman frequency 1287 cm-1 are

appeared after DA treatments which is signature for the presence of DA compound.

128

4.3.3. Correlation between mode-selective Raman image and spectra

Figure 4.7. Summary of the comparative observations from mode-selective Raman

measurements on fifty hDAT expressed HEK293 cells after addition of DA.The black and green

bar represents the appearance of mode-selective Raman images and mode-selective Raman

spectra respectively.

Figure 4.8. The crystal structure of dopamine transporter (PDB ID: 4XP1) showing the binding

site of dopamine(left), and the mode selective Raman image and spectra of DA-hDAT

interaction (right).

We performed the mode-selective Raman measurements in fifty dopamine treated hDAT-

HEK293 cells. For these experiments, cells were cultured in seven separate Petri dishes. From

each Petri dishes, 6-8 dopamine-rich hDAT-HEK293 cells were selected for the 2PE

fluorescence imaging and mode-selective Raman measurements. The dopamine-rich cells were

129

confirmed by 2PE fluorescence imaging. In our experiment, we observed Raman frequency 1287

cm-1 from all 50 dopamine-rich cells. However, the Raman frequency 1463 cm-1 were seen only

from 43 cells (Figure 4.7). These results indicate that hDAT protein was not expressed by a

significant density in all 50 HEK293 cells but only in 43 cells. The Raman frequencies: 1076 cm-

1 and 807 cm-1 were observed only from 38 dopamine-rich cells. These results show that

signature Raman frequencies related to DA-hDAT interactions were not found for all hDAT

expressed dopamine-rich cells. We found that 5 out of the 38 cells where both hDAT and DA

were present; but, failed to produce DA-hDAT signature peak. As we mentioned before, the

Raman wavenumber 1287 cm-1 and 1463 cm-1 are signatures for DA and hDAT protein

respectively, and 807 cm-1 and 1076 cm-1 are signatures for DA-hDAT interactions. These

experimental results related to occurrence of mode-selective Raman image and spectra indicate

that DA-hDAT interaction is not possible all the time even though DA and hDAT are found

together.

The results from the mode-selective Raman image (black bar) have good agreements with

the occurrence of mode-selective Raman spectra (green bar) which are summarized in figure 4.7.

However, we found one case where a mode-selective Raman image for 1076 cm-1 was obtained

but we did not find the mode-selective Raman spectra. This discrepancy could open another

scope of study, but this is not focus of this study. In our control experiment, we performed 2PE

fluorescence imaging experiment and Raman measurement on blank and dopamine coated cover

glass where fluorescence signal from blank cover glass (Figure 4.1A) was negligible with

comparison of dopamine coated cover glass (Figure 4.1B). Moreover, the Raman spectrum from

a blank cover glass is just like a background line (Figure 4.2A) while characteristic SERS

spectrum of dopamine was observed from the dopamine coated cover glass (Figure 4.2B). In

130

summary, our experimental results showed that dopamine, hDAT proteins, and their interactions

can be effectively and sensitively studied in living cells using combined approach of mode-

selective Raman imaging and 2PE fluorescence imaging techniques.

Figure 4.9. The results from DFT calculation showing difference between unbound and bound

states of DA.(A) The DFT calculated HOMO-LUMO energy level of unbound and bound states

of dopamine. The calculation shows HOMO orbital of bound states of DA molecule is more

stable than that of unbound states of DA by 0.466 eV. (B) The DFT calculated Raman spectra of

unbound and bound states of dopamine. The HOMO-LUMO energy level and Raman calculation

of bound states of DA is performed without geometry optimization by using density functional

theory (DFT) method on B3LYP level with a basis set of 6-31 G(d) using Gaussian 09 package.

4.4. Summary

We have recorded SERS spectra of dopamine, HEK293 cells, and hDAT expressed

HEK293 cells in the absence or presence of dopamine. The analysis of experimental and DFT

Raman spectra shows that Raman frequency of 807 cm-1 and 1076 cm-1 are benchmark for the

131

DA-hDAT interactions. As a matter of fact, these Raman frequencies are generated from the

bound states of DA. Utilizing these signature mode, we generated the mode-selective Raman

images and visualized the interaction between DA and hDAT proteins in living HEK293 cells.

Apart from Raman measurements, we utilized two-photon excited (2PE) fluorescence imaging

approach to probe the dopamine molecules in living cells which guides the region of the DA-

hDAT interactions in living cells. This is one of a simpler and effective technique to probe the

location of DA-hDAT interactions in living cells, which could be extended to probe the location

of other protein-ligand interactions. The combined approach of Surface-enhanced Raman

scattering and two-photon excited (2PE) fluorescence imaging could be developed as the

powerful tool to visualize the several signaling molecules, signaling proteins, and their

interactions in living systems.

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CHAPTER 5. RAMAN SPECTROSCOPIC ANALYSIS OF SIGNALING MOLECULES-

DOPAMINE RECEPTORS INTERACTIONS IN LIVING CELLS

Here we have reported a method to probe interactions of D1 and D2 types of dopamine

receptors with signaling compounds in living cells using surface-enhanced Raman spectroscopy

(SERS). In our experimental approach, four different signaling compounds including dopamine

(DA), amphetamine (AMP), methamphetamine (MAMP), and methylenedioxypyrovalerone

(MDPV) interact with D1 and D2 types of dopamine receptors expressed in HEK293 or HT22

cells, which is associated with the change of intracellular cAMP level. When agonist signaling

molecules interact with DRD1 expressed HEK293 or HT22 cells, cAMP levels increase in cells.

We probed the intracellular cAMP adsorbed on internalized silica-coated silver nanoparticles

(AgNP@SiO2); which have double roles in this experiment: the first role is the enhancement of

Raman cross-section of the samples, and the second is the adsorption of intracellular cAMP

molecules. The characteristic Raman peaks of cAMP are observed in the SERS spectrum of

DRD1 over-expressed cells when they interact with signaling compounds. Our experimental

approach is successful to collect SERS spectra from the intracellular cAMP with short exposure

time and low input power of incident laser without significant cell damage. Our experimental

results and DFT calculation show that 780 cm-1 and 1503 cm-1 are signature Raman peaks to

probe the cAMP formation in living cells. The SERS peak at 780 cm-1 is associated with C-O, C-

C, and C-N stretching; symmetric and asymmetric bending of two O-H bonds of cAMP while the

SERS peak at 1503 cm-1 is contributed by O9-H3 bending mode.

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5.1. Introduction

The G protein-coupled receptor (GPCRs) is the largest superfamily of membrane proteins

which comprises more than 800 protein-coding human genes.1 It controls and regulates many

activities including vision, test, smell, thinking, and behaviors. They are activated by the huge

range of stimuli such as hormones, pheromones, light, peptides, odorants, and neurotransmitters.2

Dopamine receptors (DARs) are heptahelical receptors and belongs to a type of GPCRs which

are primarily activated by dopamine and produce dopaminergic signals in mammalian brains

(Figure 1.1, chapter 1). Mainly, five types of human dopamine receptors are reported in the

literature, which are D1, D2, D3, D4, and D5 dopamine receptors. These five types of dopamine

receptors are classified into two classes: (1) the D1-like dopamine receptors which comprise D1

and D5 dopamine receptors and (2) the D2-like dopamine receptors which comprise D2, D3, and

D4 dopamine receptors. This classification is based on sequence homology and functions of

dopamine receptors. Although a high degree of similarities is found between DARs of similar

classes, each DAR is distinct from other types of DARs in terms of its encoded genes. For

example, both D1 and D5 dopamine receptors belong to D1-like dopamine receptors are encoded

by different genes: the D1 dopamine receptor is encoded by DRD1 and D5 dopamine receptor is

encoded by DRD5 genes.3-6 DARs are expressed heterogeneously in cells which makes it

difficult to target specifically to one type of dopamine receptors in vivo. Nevertheless, it is

important to characterize DARs to make understanding of the synaptic and neural circuit action

produced by DA and other signaling compounds.

DARs have several roles in the central nervous system (CNS) of a human such as

cognition, motivation, memory, motor control, and modulation of neuroendocrine activities.

141

Figure 5.1. (A) homology model of D1 dopamine receptors; (B) Structure of the D2 dopamine

receptor bound to the atypical antipsychotic drug risperidone (PDB ID 6cm4.pdb).

The termination of the G protein signaling and the initiation of the G protein-independent

signaling are also regulated by DARs.7-9 The D1-like DARs are also known to interact with

NMDA and GABA receptors via their intercellular loops and C-termini.10-12 D1-like dopamine

receptors couple with the Gs/olf subunit of the heterotrimeric G protein which stimulates

synthesis of cyclic adenosine monophosphate (3',5'-cyclic adenosine monophosphate, cAMP) in

cells, 13 while D2-like receptors couple with the Gi/o/z subunit of the G protein and inhibit the

synthesis of cAMP.14 The cAMP is an important secondary messenger which has several roles

including the regulation of downstream proteins,15 ion channels,16-19and transcription factors.20-22

The cAMP is synthesized from the adenosine triphosphate (ATP) by adenylate cyclase (AC)

which could be converted back into AMP by the catalytic action of phosphodiesterase.23-25 The

Mg2+ ion has an important role in these enzymatic conversions. The specific K+ currents are

decreased by Mg2+ ions in cells, which increases the influx of Ca2+ ions into the presynaptic

142

region and enhances the release of neurotransmitters.26 The cAMP has regulatory roles in

several biochemical processes such as metabolism of glycogens and lipids.27 One of the

important functions of the cAMP is that it activates the protein kinase A (PKA) which induces

the phosphorylation of substrate proteins. PKA usually exists in inactive tetrameric forms

containing two catalytic and two regulatory units where the catalytic center of PKA are blocked

by the regulatory units. When the cAMP binds to the N-terminal of PKA, it dissociates and

separates into regulatory and catalytic subunits; the catalytic subunit catalyzes the transfer of the

phosphate (PO43-) group from ATP to serine (Ser) or threonine (Thr) of the substrate proteins.

The phosphorylated proteins are associated with the regulation of ion channels and transcription

factor of DNA. Beside the PKA dependent functions, the cAMP also has other roles such as

activation of the calcium channel for the releasing of growth hormones.28-30

The biochemical functions of dopamine receptors only start when they interact with

signaling molecules or any other stimuli. Therefore, the probing of interactions between

signaling molecules like DA, AMP, MAMP, and MDPV with dopamine receptors have high

medical impacts. Both D1 and D2 dopamine receptors have a crucial role as targets for

antipsychotic drugs. Most of the clinically approved drugs show poor selectivity between

different types of dopamine receptors. The application of highly specific drugs is extremely

important to solve the neuropsychiatric and endocrine disorders since the poor selectivity of a

drug has severe side effects. Many experimental and computational approaches including ligand

binding assay (LBA),31-36 positron emission tomography (PET),37 single photon emission

computed tomography (SPECT),38 and high-throughput screening (HTS)39 are used to probe

interactions between ligands and different membrane proteins.40-60 These techniques are used for

143

the pharmacological and chemical characterization of signaling molecules (drugs) and their

selectivity to a specific dopamine receptor. The fluorescence microscopy is one of the important

techniques for the imaging of biological samples; but this technique is suffered from some

limitations including the photo-bleaching effect, photo-toxicity in living cells due to the shorter

wavelength of light, and cytotoxicity due to byproduct formation in situ photochemical

reactions.61-66 Some literature has also reported the changes of the biological property of the

sample after fluorescence labeling.67,68 Similarly, the radioligand binding assay (RBA)69

produces toxic radioactive wastes which are extremely harmful to in vivo studies. Although

above-mentioned techniques are important to screen the binding specificity of signaling

molecules like drugs and neurotransmitters they are suffered from some serious limitations. For

example, analytical results obtained from above-mentioned techniques are based on formation of

a ligand-receptor complex rather than a product formation scheme. Therefore, the result from

earlier experimental technique has a complexity to explain precisely about effective interactions

between signaling molecules and proteins. Therefore, it is highly desirable and necessary to

develop and examine other product-based technique to probe the ligand-protein interactions.

Here, we found those qualities in surface-enhanced spectroscopy (SERS) techniques to study the

interaction between dopamine receptors and signaling molecules such as DA, AMP, MAMP, and

MDPV in live cells.

The Raman spectroscopy was introduced to study the biological sample from the analysis

of bacteriorhodopsin which was later extended to study the salmon sperm and other living

cells.70-72 Now, different forms of Raman spectroscopy are already established as a powerful

analytical approach for the analysis and label-free characterization of diverse biological

144

molecules including heme protein, membrane protein, protein coenzymes, porphyrins, and highly

organized systems such as photosynthetic reaction centers and membrane preparation bacteria.73-

80 The initial attempt to study biological samples using Raman spectroscopy were unsuccessful

due to their low Raman cross-section; however, the Raman cross-section of biological samples

has been improved by hundreds of thousand folds using different approaches such as surface-

enhanced Raman spectroscopy (SERS), resonance Raman (RR) spectroscopy, and tip-enhanced

Raman spectroscopy.81-127 The tremendous enhancement of Raman cross-section in SERS

technique is achieved by exploiting electromagnetic field enhancement and chemical

enhancement mechanism using metallic substrate especially rough surface or nanoparticles of

noble metals.128 The metal nanoparticle used in Raman experiments also has another advantage

like quenching of fluorescence background.129-131 The combined approach of SERS and other

techniques including fluorescence microscopy, electrophysiology are widely used to investigate

the huge range of biological samples.132-138 Nevertheless, the SERS technique has a wide range

of useful application, the SERS substrate denatures proteins and enzymes during experiment

which is due to direct interaction of the Ag+ ion or Ag metal with the sulfhydryl group of protein

or enzymes.139,140 The direct interaction of metal nanoparticle with cells also causes cytotoxicity

problems.141,142The prevention of direct interactions between metallic surface and biological

molecules are highly desirable and recommended to protect biological samples during SERS

experiment which was achieved in our experiment by generating ultrathin layer (4-6 nm) of silica

over silver nanoparticles; it is known that these nanoparticles are capable to prevent the

denaturation of proteins and cytotoxicity in cells without reducing the electromagnetic field

enhancement.88,143

145

In this report, SERS spectra from DRD1 and DRD2 expressed cells in the presence and

absence of signaling molecules including DA, AMP, MAMP, and MDPV are collected and

analyzed to probe the effective interaction between signaling molecules and DARs. When

signaling molecules were added to DRD1 expressed cells, the characteristic Raman peaks of

cAMP were observed in the SERS spectrum of cells which help us to probe the effective

interaction between DRD1 and signaling molecules in a product-based scheme. Our

experimental results and DFT calculations show that 780 cm-1 and 1503 cm-1 are signature

Raman peaks to probe the cAMP formation in living cells. The visualization of the calculated

molecular vibration shows that SERS peak at 780 cm-1 is associated with C-O, C-C, and C-N

stretching; and symmetric and asymmetric bending of two O-H bonds of cAMP, while the SERS

peak at 1503 cm-1 is contributed by O9-H3 bending mode.

5.2. Experimental section

5.2.1. Transfection of DRD1 and DRD2 DNA in live cells

HEK293 and HT22 cells were cultured in 75 cm2 flasks in the complete medium which

was obtained by mixing DMEM (Sigma-Aldrich D5796) with 10% fetal bovine serum (Sigma-

Aldrich, F2442) and 1% penicillin-streptomycin (ATCC, 30-2300) at 37°C in 5% CO2

atmosphere.144 When cells reach ∼75% confluence, they are ready for the subculture. For the

SERS measurement, cells are sub-cultured on a 25 mm circular cover glass in 35 mm Petri dish.

When cells are reached 50% confluence on the cover glass, they are transfected with the plasmid

for protein expression. We used tango-DRD1 (66268) and GFP-DRD2 (24099) encoded plasmid

DNA respectively for the expression of the D1 and D2 dopamine receptors. We purchased these

plasmids from Addgene as the bacteria in agar stab. Plasmids were amplified according to the

146

standard protocol. Briefly, the bacteria from the agar stab were separately streaked on the surface

of LB agar plate using sterile wire loop. Each LB plate was kept inside the small incubator for 16

h at 37°C. Bacteria were grown on an LB plate as colonies with a copy of plasmids. Then, a

single bacterial colony was separated and transferred into a falcon round-bottom tubes containing

6 mL of LB solution and 100 µg/ml ampicillin. This falcon tube is placed in a rotating incubator

for 24 h at 37°C to grow bacteria. Then, bacteria were extracted from LB solution in 2 ml

centrifuge tube in the form of the solid residue by centrifuge process. This residue was dissolved

with the resuspension reagent (Thermo Scientific, Gene JET Plasmid Miniprep kit, cat. no.

K0502) and added with the lysis and neutralization solution to separate cell membranes and

plasmids from bacteria. The resulting solution is centrifuged for 15 min at 1200 rpm, and

supernatant solution is filtered through the miniprep or gene jet spin column to remove filtrate.

The content adsorbed on a filter is washed with the wash-buffer to remove any adsorbed alcohol.

Finally, the filter is washed twice with 25 µl of elution buffer to collect a total 50 µl of 1 mg/ml

plasmid DNA solution. For efficient gene transfer by transfection, we followed a protocol of

lipid-mediated transfection with Lipofectamine LTX and Plus reagent (Invitrogen, 11668-019).

The day before transfection, regular growth medium is replaced with growth medium without

antibiotics (Penicillin streptomycin). The preparation of DNA reagent is required to add into cell

samples which is explained briefly as following. The 50µl of Opti-mem reduced serum medium

is taken in the 2ml centrifuge tube and added with 4µl of Lipofectamine LTX solution; however,

the amount of Lipofectamine LTX could be varied. The mixture is vortexed and labeled as (1).

The 250 µl of Opti mem medium and 5 µl of plasmid DNA are taken in 2ml centrifuge tube and

added with 5µl plus reagent which is labeled as (2). Then, 50µl of solution (2) is mixed with

147

solution (1) and kept this solution at room temperature for 5 min for the incubation. After

incubation, we took 50 µl of this solution and added into cells. The successful transfection of

DRD1 and DRD2 plasmid in cells is confirmed by tango and green fluorescence respectively.

5.2.2. Synthesis of silica-coated silver nanoparticles

Silver nanoparticles (AgNPs) are synthesized by a standard sodium citrate reduction

method,145 followed by the addition of active sodium silicate to generate ultra-thin silica shells

over AgNPs.146 Silver nitrate (AgNO3), sodium citrate, and sodium silicate required for the

synthesis of nanoparticles were purchased from Sigma Aldrich and used without further

purification. The AgNPs and AgNP@SiO2 were characterized by transmission electron

microscopy (TEM) and Varian UV-Vis spectrophotometer (EL07013173). The size of

AgNP@SiO2 particles is found 80 ± 10 nm with SiO2 shell thickness 5 ± 1 nm. The UV-Vis

measurement shows the maxima (λmax) of the plasmon resonance band position of AgNPs colloid

at 420 nm, which was shifted to 407 nm for [email protected]


The geometry optimization and Raman frequency calculation of signaling compounds and

cAMP were performed using the density functional theory (DFT) on B3LYP/6-31G(d) and

B3PW91/LANL2DZ levels/basis set and Gaussian 09 package. The obtained frequencies from

B3LYP/6-31G(d) calculations were scaled by a factor of 0.9614147 while the frequency obtained

from B3PW91/LANL2DZ are used without supplying any scaling factor (Figure 5.2). All

calculations were carried out on a vector processor (Ohio Supercomputer Center, Columbus,

Ohio).

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5.3.1. Internalization of silver nanoparticles

We studied the interaction of signaling compounds with different types of dopamine

receptors using the combined approach of SERS measurement, wide-field microscopy, and DFT

calculations. For that, we added finely dispersed AgNP@SiO2 on cells which enter through the

semipermeable membrane of cell to cytoplasm. For the fine dispersion of nanoparticles, the

AgNP@SiO2 colloid was vortexed for 15 minutes before adding to cell samples. The differential

interference contrast (DIC) imaging was used to examine the internalization of AgNP@SiO2 in

cells (Figure 5.3); the internalization of nanoparticles was examined by changing the focusing

mode of the microscope. We assumed that the internalized nanoparticle is in a similar plane as

outsider nanoparticles on cover glass. The outsider nanoparticles on the upper surface of the cell

membrane are observed darker during the up-focusing mode of the microscope objective (figure

5.3A). In this mode, both outsider nanoparticles on the cover glass and internalized nanoparticles

are fainter. However, these outsider nanoparticles on the cover glass and internalized

nanoparticles become darker when the objective of the microscope was brought to down-

focusing mode (Figure 5.3C). Since our nanoparticles are not specifically modified to target

cytoplasm, we waited 18 h for the internalization of nanoparticles. Some internalized

nanoparticles were also able to form cluster inside cells which is important for the enhancement

of Raman intensity. The time for the internalization of the nanoparticle can be decreased

significantly by attaching cytoplasm targeting lipid on metal nanoparticles.148

149

Figure 5.2. The DIC image of DRD1 over-expressed HEK293 cells in DA solution of 0.2 µM

concentration showing the internalization of AgNP@SiO2 in cells and mode-selective Raman

imaging.The internalized AgNP@SiO2 in a cell is confirmed by changing focusing modes of

microscope: (A) up-focus, (B) Focus, (C) down-focus, and (D) further down-focus conditions.

Based on DIC image, we assumed that internalized nanoparticles are in a similar plane as

outsider nanoparticles on cover glass. The Raman image related to 780 cm-1 due to the

interaction of DRD1 with DA is collected using 488 nm CW excitation laser of 10-15 µW power

and a bandpass filter of 508.5±2 nm by CMOS industrial CCD camera (RS-500C). Images in

figure (E) is obtained using both white light and CW laser light while figure (F) is obtained using

CW laser only.

150

5.3.2. Probing of intracellular cAMP in HEK293 cells

Figure 5.3. The Experimental and DFT calculated Raman spectrum of cAMP.The experimental

SERS spectra are collected using home-modified confocal microscope (Figure 2.9 in Chapter 2).

The Raman frequency obtained from DFT calculation using B3LYP/6-31G(d) is corrected with

scaling factor 0.9614, but Raman frequency obtained from B3PW91/LANL2DZ (black) is not

corrected with supplying any scaling factors.

151

Figure 5.4. The consistence of occurrence between cAMP-marker SERS image and peaks.The

SERS image and peaks are associated to 780 cm-1 observed from DRD1, DRD2 and DRD1-

DRD2 expressed HEK 293 cells after adding dopamine.

After internalization of AgNP@SiO2 in four types of cells: baseline HEK293 cells, DRD1

expressed HEK293 cells, DRD2 expressed HEK293 cells, and DRD1 and DRD2 co-expressed

HEK293 cells, the SERS spectrum from these cells were collected in two different conditions:

(1) with the addition of signaling compounds such as DA, AMP, MAMP, and MDPV, and (2)

without the addition of signaling compounds. For condition (1), the signaling compound was

added 2 h later the adding AgNP@SiO2. When DRD1 expressed cells interact with signaling

molecules, the intracellular content of cAMP was increased which was tested by the observation

of cAMP signature peaks at 780 and 1503 cm-1. We used five Petri dishes of DA-DRD1 cells to

test the cAMP-related SERS peaks, and found that 4-5 cells out of twenty randomly selected

152

cells from a petri dish, showed signature peaks of cAMP. Altogether, 22 cells out of 100 cells

from five Petri dishes showed cAMP-related peaks. In similar experiments, none of 100 DA-

DRD2 cells were able to give cAMP-related peaks. In addition, we also collected mode-selective

SERS image of 780 cm-1 using 508.5±2 nm bandpass filter and CMOS industrial CCD camera

(RS-500C). In this experiment, out of 20 randomly selected DA-DRD1 cells from a Petri dish, 5-

8 cells showed the 780 cm-1 related Raman image, and out of 100 DA-DRD1 cells from five Petri

dishes, 37 cells showed the Raman image related to 780 cm-1. For similar experiments in 100

DA-DRD2 cells, 7 cells out of 100 showed the Raman image related to 780 cm-1. The minor

discrepancies are found between occurrence of Raman peaks and Raman image which is due to

wide range of bandpass filter. The spectral range of bandpass filter (508.5±2 nm) is particularly

higher to give off the mode-selective Raman image related to 780 cm-1, that is why mode-

selective Raman image could be contributed by another wavenumber too. Similarly, 2 DRD1-

DRD2 co-expressed cells out of 100 showed cAMP-related SERS peaks while 9 of them showed

780 cm-1 related mode-selective Raman image. The statistics of cAMP related peaks and images

are summarized in figure 5.4. The imperceptible amount of cAMP in DRD1-DRD2 co-expressed

cells is due to the competitive action of DRD1 and DRD2 receptors for the synthesis and

depletion of cAMP. We assumed that the competitive action for synthesis and depletion of cAMP

is favored by the non-specific interaction of four signaling molecules with DRD1 and DRD2

receptors respectively. Intracellular cAMP molecules formed during interactions of DRD1 with

signaling compounds are trapped by internalized AgNP@SiO2 and probed using SERS where the

formation of cAMP was confirmed by the appearance of its signature SERS peaks at 780 and

153

1503 cm-1. The Raman assignment of these signature peaks of cAMP is provided elsewhere in

this chapter.

We consider SERS peaks at 780 cm-1 and 1503 cm-1 are signature peaks of cAMP

because these peaks were only given by two samples; first: DRD1 expressed HEK293 cells in

the presence of signaling compounds (Figure 5.5A-D and 5.6A), and second: pure cAMP

compounds (Figure 5.5E). Our controlled experiments show that these signature peaks are not

observed in the SERS spectrum of the pure form of all four signaling compounds (Figure 5.5F-I,

and 5.6) and all types of cells in the absence of signaling compounds (Figure 5.5J, 5.7-5.10, and

5.11E, F). Moreover, these SERS peaks are also absent for all signaling compounds in all cells

except in DRD1 expressed cells (Figure 5.12). Our experimental results show that interaction

between signaling molecules with DRD1 receptors is required for the synthesis of cAMP in cells.

5.3.3. Probing of intracellular cAMP in HT22 cells

In addition, we have extended the same experimental approach to test the interaction

between DRD1 and signaling molecules in mouse hippocampal neuronal cell (HT22 cell). In this

experiment, we over-expressed D1 dopamine receptor in HT22 cells by following the similar

standard procedure as explained in the experimental section. In this experiment, we found that

interactions between DRD1 expressed HT22 cells with signaling molecules increases the

intracellular content of cAMP. The synthesis of cAMP in HT22 cells could be determined by

observing cAMP related signature peaks (Figure 5.11). In this figure, we could observe the

Raman peak at 780 cm-1 and 1503 cm-1 in the SERS spectra from DRD1-HT22 cells in the

presence of signaling compounds (Figure 5.11A-D) while these peaks are absent in baseline

HT22 cells or DRD1-HT22 cells in the absence of signaling molecules (Figure 5.11E, F).

154

Figure 5.5. Integrated SERS spectra (left) and 2-dimensional distribution SERS spectra (right)

obtained from the DRD1 expressed HEK 293 cells in different experimental environments. All

SERS spectra were collected by home-modified Raman microscope (Figure 2.9 in chapter 2)

using 488 CW argon ion laser (10-15 µW) with integration time 30 seconds.The Raman peaks at

780 cm-1 and 1503 cm-1 are observed in the SERS spectra from DRD1 expressed cells in (A)

AMP, (B) MAMP, (C) MDPV, and (D) DA. These two peaks are also observed in (E) pure

cAMP, but absent in (F) AMP, (G) MAMP, (H) MDPV, (I) DA and (J) DRD1 expressed

HEK293 cells.

155

Figure 5.6. The DFT calculated Raman spectrum of signaling compoundsincluding DA (pink),

cAMP (green), ATP (purple), MDPV (red), MAMP (black) and AMP (blue). The geometry

optimization and frequency calculation of compounds are obtained by using B3LYP level and 6-

6-31G(d) basis set.

156

Figure 5.7. The integrated experimental SERS spectrum obtained from DRD1 expressed

HEK293 cells in the absence (red) and presence (green) of signaling compounds.Signature peaks

of cAMP (blue) located at 780 cm-1 and 1503 cm-1 are also observed in the SERS spectrum

obtained from the DRD1 expressed HEK293 cell in the presence of signaling compounds.

157

Figure 5.8. The integrated experimental SERS spectrum obtained from DRD2 expressed

HEK293 cells in the absence (red) and presence (green) of signaling compounds.Signature peaks

of cAMP (blue) located at 780 cm-1 and 1503 cm-1 are not observed in the SERS spectrum

obtained from the DRD2 expressed HEK293 cell in the presence of signaling compounds.

158

Figure 5.9. The integrated experimental SERS spectrum obtained from DRD1-DRD2 co-

expressed HEK293 cells in the absence (red) and presence (green) of signaling

compounds.Signature peaks of cAMP (blue) located at 780 cm-1 and 1503 cm-1 are not observed

in the SERS spectrum obtained from the DRDDRD2 co-expressed HEK293 cell in the presence

of signaling compounds.

159

Figure 5.10. The integrated experimental SERS spectrum obtained from baseline HEK293 cells

in the absence (red) and presence (green) of signaling compounds.Signature peaks of cAMP

(blue) located at 780 cm-1 and 1503 cm-1 are not observed in the SERS spectrum obtained from

the baseline HEK293 cells in the presence of signaling compounds.

160

Figure 5.11. Integrated SERS spectra (left) obtained from the DRD1 expressed HT22 cells in

different experimental environments.The Raman peaks at 780 cm-1 and 1503 cm-1 are observed

in the SERS spectra from DRD1 expressed cells in (A) AMP, (B) MAMP, (C) MDPV, and (D)

DA These two peaks are absent in (E) DRD1 expressed HT22 cells and (F) baseline HT22 cells.

5.3.4. Raman peaks assignment of cAMP

The Cyclic adenosine monophosphate (cAMP) is an important second messenger, which

is utilized as a marker in this study to probe effective interactions between D1 dopamine receptor

(DRD1) and signaling molecules such as DA, AMP, MAMP, and MDPV. We have

collected SERS spectra from pure cAMP and compared with the SERS spectrum obtained from

intracellular cAMP content (Figure 5.5A-J); based on our experimental results and DFT

calculations (Figure 5.2), here we have assigned the Raman peaks of cAMP compounds in the

161

range of 700 to 1800 cm-1. Major SERS peaks obtained from experimental SERS spectrum of

cAMP and its geometry optimized molecular structure are provided in supporting information

(Figure 5.13). Raman peaks of cAMP in the lower frequency region (<1000 cm-1) are related to

780 cm-1: stretching of C-O, C-C, C-N bonds, and sym. and asym. bending of O-H bond; 824

cm-1: normal mode and bending of rings (III) and (IV); 858 cm-1: C1-H28 bending, and ring (III)

and (IV) breathing mode; 946 cm-1: sym. stretching and asym. bending of PO32-, stretching of

O10-H13, C8-C19, and C19-C18 bonds, and C-H twisting in the ring (I). Raman peaks of cAMP in

the higher frequency region are related to 1013 cm-1: normal mode, NH2 rocking, and sym.

stretching of PO32-; 1060 cm-1: ring (II) and (III) stretching, and NH2 rocking; 1076 cm-1: C12-O9

stretching, and CH2 rocking; 1096 cm-1 and 1115 cm-1: Ring stretching; 1145 cm-1: C-C

stretching, and CH2 rocking; 1177 cm-1: C-C stretching, O9-H3 bending, and Ring (II), (III) and

(IV) breathing mode; 1228 cm-1: H28-C1-H21 bending, and NH2 rocking; 1270 cm-1: C-H

bending, and C-N stretching; 1323 cm-1 and 1388 cm-1: C-H bending, C=N stretching, and C-N

stretching; 1407 cm-1: ring (III) and (IV) stretching, and C-H bending; 1503 cm-1: O9-H3

bending; 1556 cm-1: CH2 scissoring, ring (III) and (IV) stretching, and NH2 scissoring; 1602 cm-

1: ring (III) and (IV) deformation, and N-H bending. The assignments of experimental SERS

peaks obtained from pure cAMP compound are summarized in table 5.1.

162

Table 5.1. The assignments of experimental SERS peaks obtained from pure cAMP.

163

Figure 5.12. The integrated SERS spectra and 2-d distribution SERS spectra due to interaction

of signaling compounds with different types of cell samples.Integrated SERS spectra (left) and 2-

dimensional distribution SERS spectra (right) obtained from (A) DRD1 expressed HEK293 cells

(B) DRD2 expressed HEK293 cells (C) baseline HEK 293 cells (D) and DRD1 and DRD2 co-

expressed HEK293 cells in amphetamine (red line), methamphetamine (blue line),

Methylenedioxypyrovalerone (black line), and dopamine (green). The Raman peaks at 780 and

1503 cm-1 are only observed in DRD1 expressed cells which were added with signaling

compounds.

164

Figure 5.13. The experimental SERS spectrum of cAMP with major peaks.The energy

optimized molecular structure of cAMP obtained from DFT calculation using B3LYP level and

6-31G(d) basis set is shown in figure inset.

Figure 5.14. The experimental SERS spectrum obtained from cell samples in the absence of

signaling compounds.(A) Baseline HEK293 (B) DRD2-HEK293, (C) DRD1-DRD2 expressed

HEK293, and (D) DRD1 expressed HEK293 cells (black) in the absence signaling compounds.

165

5.3.5. Raman peaks assignment of DRD1-HEK293 cells

The dopamine receptor can be controlled genetically in cells by overexpression,

knockdown, or knockout approaches. The overexpression of dopamine receptor can be

confirmed by fluorescence microscopy. We have overexpressed the tango-DRD1 and GFP-

DRD2 in HEK293 and HT22 cells; the overexpression of these genetically tagged proteins is

easily confirmed by fluorescence microscopy. After overexpression of D1 and D2 dopamine

receptor, the SERS spectra of HEK293 cells and HT22 cells are changed. Nevertheless, the

assignment of all Raman peaks from cells is difficult, here we have briefly assigned the SERS

peaks of DRD1 expressed HEK293 cells (Figure 5.14). The SERS peaks of DRD1 expressed

HEK293 cells in the low-frequency region (< 1000 cm-1) are related to 790 cm-1: 𝜔(C𝛿-H) (his);

863 cm-1: 𝜔(C𝛿-H) of hist and 𝜏(C𝜍-N𝜂2) of Arg; 916 cm-1: C-C stretch, C𝛿 wag, and C𝛼-C𝛽 of

Lys, and 𝜐(C-C) of phe. The SERS peaks in higher frequency region (>1000) are related to 1002

cm-1: ring deformation of Phe; 1136 cm-1: NH3+ asym. rocking, and -OOC-C𝛼-H𝛼 of Lys; 1241

cm-1: CH2 twisting of Met; 1275 cm-1: N𝛿-C𝛿-H; N𝛾-C𝛿-H of His.; 1330 cm-1: CH2 wag of

Met.; 1392 cm-1: NH3+-C-H and C𝛽-C𝛼-H𝛼 of Ile.; 1454 cm-1: H𝛼-C𝛼-H, N-C𝛼-H𝛼, and CH3-

asym. deformation mode of Met.; 1492 cm-1: N𝛿-C𝛿, N𝛿-C𝛿-H of His.; 1554 cm-1: amide II

bands; and 1587 cm-1: -OOC asym stretch of Ala., C𝛾-C𝛿, C𝛽-C𝛾, and N𝛿-C𝛾-C𝛿 stretch of His.

The assignment of SERS peaks obtained from DRD1-HEK293 cells are based on literature

values related to Raman frequency of amino acid residue which is summarized in table 5.2.149-158

166

Table 5.2. The assignment of SERS peaks obtained from DRD1-HEK293 cells.

5.4. Summary

The cAMP is formed in cells, when signaling compounds such as DA, AMP, MAMP, or

MDPV interact effectively with D1 dopamine receptors. We used internalized silica-coated silver

nanoparticles (AgNP@SiO2) to adsorb intracellular cAMP and enhance Raman intensity. Our

experimental approach helps us to collect SERS spectra from intracellular cAMP using low-

power laser excitation with short exposure time which helps to minimize the chance of

significant cell damage. Our experimental results and DFT calculations show that 780 cm-1 and

1503 cm-1 are signature Raman peaks to probe the cAMP formation in living cells. We found that

the SERS peak at 780 cm-1 is associated to C-O, C-C, and C-N stretching; and symmetric and

asymmetric bending of two O-H bonds of cAMP, and the SERS peak at 1503 cm-1 is found to be

contributed by O9-H3 bending mode of cAMP. Our experimental approach is successful to probe

167

the expression of D1 and D2 dopamine receptors and distinguish their interactions with signaling

compounds. This approach could be improved to study the selectivity of psychoactive drugs

towards different types of dopamine receptors.

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CHAPTER 6. RAMAN SPECTROSCOPY PROBING OF REDOX STATES AND

MECHANISM OF FLAVIN COENZYME

Redox states of Flavin mononucleotide (FMN) play important and regulating roles in living

systems. To understand the involvement and contribution of FMN coenzyme in different

biological processes, probing and characterizing of the associated FMN redox states using

powerful experimental approaches are fundamental and crucial. In our study, we have generated

several typical FMN redox states in Britton-Robinson (B-R) buffer at different pH environments

by applying electric potentials. The electric potential and pH-dependent events of protonation,

deprotonation, and electron transfer process of FMN are probed and characterized by surface-

enhanced Raman spectroscopy (SERS) or Resonance SERS (SERRS) using silica-coated silver

nanoparticles (AgNP@SiO2) as SERS substrate. In addition to experimental SERS analysis, we,

using Density Functional Theory (DFT), computationally calculated Raman spectra to identify

the spectral signatures of the FMN redox-state sensitive Raman modes. Here, we have

specifically probed, analyzed, and characterized signature Raman modes of different redox states

of FMN coenzyme including FMNH2•+ (1508-1510 cm-1), FMNH2 (1512-1514 cm-1), FMN2-•

(1498 cm-1), and FMN3- (1492 cm-1) and proposed the redox reaction schemes of FMN in

different experimental conditions.

6.1. Introduction

The family of flavoproteins commonly contains flavin coenzymes like flavin adenine

dinucleotide (FAD) and flavin mononucleotide (FMN) as the prosthetic group. Flavoproteins

catalyze many redox reactions in biological systems where FMN and FAD have key roles for

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Figure 6.1. The DFT Raman spectra from different redox states of Flavin mononucleotide(FMN)

with geometry optimization on the B3LYP level with basis set of 6-31G(d), and Gaussian 09

package with scaling factor 0.9614.

the functions of flavoproteins. Flavoproteins are involved in a broad range of biological

processes including bioluminescence, photosynthesis, DNA repair, apoptosis, and elimination of

reactive oxygen species (ROS).1-4 ROS has been associated with the induction and complications

of diabetes mellitus, age-related eye disease, and neurodegenerative diseases such as Parkinson's

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disease. Malfunctioning of flavoproteins is also related to oxidative stress and the damage of

extensive range of molecular species like lipids, proteins, and nucleic acids, initiation, and

development of cancer, as well as the side-effects of radiation and chemotherapy. Thus,

flavoproteins and flavin coenzymes are extensively studied and highly significant in protein

science. In addition, flavin coenzymes catalyze electron-transfer reactions in diverse ways on

flavoproteins which involve in catalysis of both one and two electrons transfer process.5,6

Sometimes, they are involved in the catalysis of electron transfer reactions between two-electron

donor and one-electron acceptor.7,8 The electron transfer mechanism of flavin coenzymes

changes as their interactions with the flavoenzymes and the surrounding environment changes.

The X-ray crystallographic study shows that the electroactive site of flavin coenzymes like a

FAD in aqueous environments is accessible to water molecules 9 and does not bind covalently to

enzymes. Under such conditions, flavin coenzymes undergo a two-electrons/two-proton (2e-

/2H+ ) reduction process and generate the fully reduced form of flavin coenzymes (FADH2) 7.

However, in a non-aqueous environment, coenzymes bind covalently to flavoenzyme and form

stable flavin semiquinone forms (FADH•). The difference is also reported in proton-coupled

electron-transfer (PCET) mechanism of flavin coenzymes in an aprotic organic solvent, and the

aqueous solution.10,11The electrochemical studies show that the redox pathways of the FMN

molecules in aqueous solutions are pH dependent.12-24 The redox states of FMN has an

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Figure 6.2. Zoom in view of integrated SERS spectra of Flavin mononucleotide in B-R buffer at

different pH environments.(A) pH 11, (B) pH 9, (C) pH 7.6, (D) pH 7.4, (E) pH 7, (F) pH 5, and

(G) pH 3. SERS spectra of FMN collected in each pH condition at different states are

represented with different colors; Red = dried, blue = wet in KCl, green = oxidized, and black =

reduced states.

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important role in blue-light photoreceptor, and electron transport process during ATP

synthesis. During ATP synthesis, FMN couples with the series of iron-sulfur (Fe/S) clusters in

the NADH dehydrogenase (complex I) to transport the electron from NADH to ubiquinone (Q)

and pump the protons from the mitochondrial matrix to its intermembrane space, this is one of

the important example where different redox states of FMN involve in a biological process

(Figure. 6.3). As earlier studies show, FMN is involved in both one and two-electron transfer

redox processes taking part in reversible interconversion to generate its oxidized (FMN),

semiquinone (FMNH•) and reduced form (FMNH2). 5-8 Many studies have been carried out on

flavin coenzymes to understand biological properties and the importance of their redox states.

Webb and coworker applied the redox fluorometry technique based on the inherent fluorescence

from NAD(P)H and/or flavoprotein to monitor cellular energy metabolism as a function of

substrate availability.25

Figure 6.3. Schematic representation of the inner membrane of mitochondria showing the

involvement of FMN cofactor in electron transport process during ATP synthesis.The redox

states of FMN couples with series of iron-sulfur clusters to transport electron from NADH to

ubiquinone.

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Figure 6.4. The experimental SERS spectrum of FMN at the redox-sensitive region in pH 11

(A), pH 9 (B), pH 7.6 (C), pH 7.4 (D), pH 7 (E), pH 5 (F), and pH 3 (G).SERS spectra of FMN

collected in each pH condition at different states are represented with different colors; dried state

(red), the wet state with KCl solution (blue), oxidized state (green), and reduced state (black).

Each integrated SERS spectrum is the average of 40 typical SERS spectra.

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The kinetics of reduction of the flavocytochrome have been investigated by using laser

flash photolysis.26 Webster and coworker studied the electrochemical reduction mechanisms of

FMN in buffered aqueous solutions using variable-scan-rate cyclic voltammetry, controlled-

potential bulk electrolysis, UV−Vis spectroscopy, and rotating-disk-electrode voltammetry.11

Similarly, Hazra and coworker used the steady-state and time-resolved fluorescence quenching

of flavin, circular dichroism and thermal melting techniques to predict the structural reformation

on aptamer due to flavin-aptamer binding.27 Begum and coworker studied the translocation of

Flavo enzyme in the nucleus of Dictyostelium discoideum by utilizing immunofluorescence.28

Despite the excellent performances of the aforementioned techniques, in many circumstances,

they are not always useful to characterize complex mixtures of different redox states of flavin

coenzymes because of their intrinsic limitations. For instance, the fluorescence-based study is

restricted to observe the “on” and “off” states of redox states and is not effective for their fine

structural characterization. In addition, the photobleaching effect, phototoxicity of the shorter

wavelength used for excitation, and byproducts formed during photochemical reaction makes

this technique less applicable in living systems.29-34 Similarly, UV−Vis spectroscopy and

voltammetry and electrolysis based techniques are also not useful to study minor structural

changes of flavin coenzymes in living environments. Here, we found that surface-enhanced

Raman spectroscopy (SERS) is a powerful experimental approach for the fine characterization of

redox states of flavin coenzymes and redox reaction schemes in different pH environments.

Although, the original form of Raman scattering cross-section of most of the molecules are very

smaller, the technical advancement in Raman spectroscopy has greatly increases significance of

this techniques to study wide varieties of samples.35-80 For instance, SERS technique is

established as a powerful analytical approach for the label-free characterization and analysis of

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biomolecules like heme protein, porphyrins, membrane protein, and highly organized systems

such as membrane preparation and photosynthetic bacteria, and photosynthetic reaction

centers.81-88 In this work, we have utilized the SERS technique for the analysis and

characterization of electrochemically generated redox states of flavin mononucleotide (FMN),

which helps to determine the electrochemical reaction pathway of FMN in different pH

environments.

The SERS technique commonly utilized electromagnetic field enhancement and chemical

enhancement mechanisms provided by roughened metallic surface or nanoparticles, especially

gold or silver to overcome the limitation of conventional Raman spectroscopy.89 However, many

biological molecules like proteins and enzymes are denatured during SERS experiment due to

direct interaction of these molecules with Ag+ ions or AgNPs. It is reported that Ag+ ions or

AgNPs interact with sulfhydryl group of proteins or enzymes and induce the denaturation of

these molecules.90,91 Therefore, the inventions of effective functionalization approach of SERS

substrates are highly desirable and recommended to prevent the direct interaction between

metallic surface and biological molecules. For this purpose, we have coated silver nanoparticles

with the ultra-thin (4-6 nm) layer of silica, which is highly effective to prevent the possible

chemical interactions of Ag+ ions or AgNPs with biological molecules without reducing

electromagnetic field enhancement.92,93 The fluorescence free and high-quality Raman spectra

are obtained when silica-coated silver nanoparticles (AgNP@SiO2) are used as SERS substrate,

which indicates that metal nanostructures have a higher fluorescence quenching efficiency in

contrast to the quenching efficiency of protein or potassium iodide.94-96 The metal

nanostructures used in the SERS technique help to quench the fluorescence background

produced from FMN molecules, and obtain higher signal-to-noise ratios in the Raman spectra,97

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This advantage of the SERS technique is very appealing because the initial efforts to obtain

Raman spectra of flavins using conventional Resonance Raman (RR) were useless due to

interference of highly intense free flavin emission at around 530 nm.98

Figure 6.5. (A) The UV-Vis absorption spectrum, and (B) emission spectrum of FMN (10 µM)

in B-R buffer solution.The wavelengths of absorption and emission maximum are found 445 nm

and 526 nm respectively for all pH environment.

Based on our control experiment and reports in the literature, the single photon resonance

Raman (RR) measurement for flavins could encounter the interference from high fluorescence

background around 530 nm (Figure. 6.5B). Therefore, we did not prefer conventional resonance

Raman (RR) for this study. Another importance of the SERS technique is its effectiveness under

physiological conditions. Since the Raman cross section of water is significantly lower than that

of targeted molecules, this technique is less suffered from water background effect.99 In our

experiment, we have observed the absorption band of FMN at 445 nm (Figure. 6.5A). This band

represents the transition from ground electronic states (ν0) to the first excited electronic state of

FMN (ν1). Moreover, a shoulder at around 470 nm in the absorption spectra of FMN for some

flavoproteins is also reported.100 This shoulder could be originated due to the transition from

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ground electronic state (ν0) to first excited electronic state (ν0) of FMN. In our Raman

measurement, we used the CW argon ion laser at 488 nm for the sample excitation, which is

closer to the 470 nm wavelength required for the ν0-ν0 transition. Therefore, the enhancement in

our Raman spectra could be mainly contributed from SERS, but we cannot rule out a possible

contribution from the resonance SERS (SERRS) to the enhancement of Raman signal intensity.

The SERS technique has been extensively used to characterize the flavins after Copeland

and co-worker first recorded the high-quality spectra of free flavins101 and flavoproteins102 using

colloidal silver as SERS substrate. Using the roughened silver electrodes, and controlling the pH

values of the buffer solutions, Xu and co-workers studied the in situ electrochemical SERS for

the semiquinone radical of FMN.103 Similarly, Zheng and co-workers first utilized the resonance

Raman experiment, and quantum calculations approach to characterize the fully reduced

flavin.104 The numbers of studies including DFT calculation, and isotopic substitution and semi-

empirical calculation have provided important information on the assignment of fully oxidized

and reduced isoalloxazine ring of flavins.105-110 However, the Raman modes of FMN redox

states and mechanism in the broad range of experimental environments are not still clearly

demonstrated. In our work, SERS has proved its importance to probe and characterize complex

mixtures of FMN redox states in the broad range of experimental conditions. We applied the

different electric potential (varied from + 0.3 to -1.0 V) in the FMN solution prepared in the

different pH value of Britton-Robinson (B-R) buffer (varied from 3 to 11 pH) to generate its

different redox states. Here, FMN coenzymes leads to different electrochemical reaction

mechanisms like protonation, deprotonation, or electron transfer process depending on pH

environments. The information obtained from several SERS experiments in different-different

pH environments and DFT Raman calculations are analyzed precisely to determine the presence

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of FMN redox states and electrochemical mechanisms of the entire redox processes for the

provided experimental conditions.

6.2. Experimental section

6.2.1. Synthesis of silver nanoparticles and sample preparation

Silver nitrate, sodium citrate, potassium chloride, sodium silicate, acetic acid, phosphoric

acid, and boric acid were purchased from Sigma Aldrich and used without further purification.

Silver nanoparticles (AgNPs) were synthesized by a standard sodium citrate reduction method,111

followed by the addition of active sodium silicate to generate ultrathin silica shells over

AgNPs.112 The synthesized nanoparticles were characterized by Transmission electron

microscopy (TEM) and Varian UV-Vis spectrophotometer (EL07013173). According to our TEM

measurement, the size of AgNP@SiO2 was found 58 ± 5 nm with SiO2 shell thickness 5 ± 1 nm.

The UV-Vis spectroscopy shows that the maxima (λmax) of the plasmon resonance band position

related to silver nanoparticles is found at 420 nm which was shifted to 407 nm after generating

the ultra-thin layer of silica on its surface which is shown in figure 2.13 in chapter 2. The

Britton-Robinson (B-R) buffer was prepared by adding equal volumes of an equimolar

concentration of acetic acid, phosphoric acid, and boric acid (0.04 M). Then the pH was adjusted

by careful addition of NaOH solution (0.2 M). The B-R buffer was then added to FMN to form

its 100×10-9 M solution.


Geometry optimization and Raman wavenumber calculations were performed using the

density functional theory (DFT) method on a B3LYP level with a basis set of 6-31G (d) and

Gaussian 09 package to observe Raman wavenumber of the redox-sensitive mode of FMN

different redox species of FMN molecules. The literature show that the scaling factors of 0.9804

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could be more applicable for the DFT calculated vibrational wavenumber at the lower

wavenumber region (<1000 cm-1), and 0.9625 for the wavenumber at higher wavenumber region

(>1000 cm-1).113,114 Based on our control experiment for the calibration of the scaling factor

(Figure 2.12, chapter 2), and a comprehensive evaluation report115, we have scaled the DFT

calculated Raman frequencies by a factor of 0.9614, which is reasonable to our region of interest

(higher wavenumber region). The molecular orbitals were calculated with the same basis set and

visualized with Avogadro software (Avogadro: an open-source molecular builder and

visualization tool. Version1.XX.). All calculations were carried out on a vector processor (Ohio

Supercomputer Center, Columbus, Ohio).

6.2.3. Surface-enhanced Raman measurements and electrochemical control

All SERS spectra were recorded by using a home-modified confocal Raman

microscope116 (Figure. 2.9, chapter 2) with a 30 second integration time. A continuous-wave

(CW) of 488 nm argon ion laser was used to excite the sample at approximately 10-15 µW for

SERS. The setups were carefully calibrated using a mercury lamp and cyclohexane before

Raman measurements with a spectral resolution of 2 cm-1. A CHI 600C electrochemical

workstation was used for performing electrochemical control equipped with a home-made

electro spectroscopic cell (Figure 2.9B in chapter 2)117 (working electrode: Indium tin oxide

(ITO)/glass cover glass; counter electrode: platinum wire; reference electrode: silver wire). The

~40 µl solution of FMN (prepared in B-R buffer and 0.4 M KCl) and silver nanoparticles are

incubated overnight on the ITO surface and dried, then a solution of 0.1 M KCl was put on the

top of the ITO as a supporting electrolyte in the homemade electroscopic cell (Figure 2.9B in

chapter 2). This solution was deoxygenated by purging with high-purity nitrogen gas before the

voltammetry analysis; all voltammetry experiments were conducted at room temperature. The

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cyclic voltammetry (CV) was first performed before each Raman measurements to determine the

supply of appropriate values of electric potential for the complete oxidation and reduction of

FMN coenzyme in respective experimental conditions. We have tested different scan rates and

identified that the scan rate in 0.1 V/s is optimal to obtain CV signals at an adequate signal-to-

noise ratio (Figure. 6.6). A more negative or positive electrochemical potential than the formal

redox potential was applied to keep the sample at a fully reduced or oxidized state, respectively.

The cyclic voltammetry provides the driving force which may induce certain transitions and

produces different redox states of FMN as shown in scheme 1 and 2, otherwise these FMN redox

states would not have occurred physiologically in the provided pH range.

6.2.4. Two-Dimensional distribution SERS Plot vs conventional SERS spectrum

Two-dimensional SERS spectra of FMN molecules in different experimental

environments are generated by combining 40 high-quality SERS spectra (Figure. 6.7, 6.8, 6.9,

6.10, and 6.11). The inevitable complications due to temporal fluctuation of peak position and

intensity in conventional SERS spectrum are removed by generating two-dimensional (2-D)

distribution SERS spectra. 2-D SERS spectrum of a sample is obtained by interpolating Raman

wavenumber (cm-1) and intensity (arbitrary unit) of 40 conventional SERS spectra. In 2-D SERS

spectra, Raman shifts are characterized by colorful strips instead of signal peaks of conventional

SERS spectrum. The color scale of the strips shows the intensity, the broadening of a strip along

x-axis shows the trend of temporal fluctuation in Raman wavenumber, and lengthening of the

strip along the y-axis shows the recurrence of a Raman mode in the numbers of SERS spectra. In

our analysis, the center of the strip along the x-axis is taken as the mean position of the Raman

frequencies which helps to adjust the impact of temporal fluctuation of the Raman wavenumber.

The mean position of the colored strips and peak position of integrated Raman spectrum are

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highly consistence (Figure. 6.8, 6.9, 6.10, and 6.11). However, it is not convenient to use the

mean position of the colored strip to resolve the small shifting of peaks. Therefore, we have used

the peak position of integrated SERS spectra to resolve the small shifting of peaks positions

during redox processes. The Raman analysis using Integrated SERS spectra and 2-D SERS

spectra are helpful for the selection of wavenumber of certain Raman modes and help to get rid

of the inevitable temporal signal fluctuation problem. Besides these benefits, the problems of

unusual spike and noise in conventional SERS spectra are removed using 2-D SERS spectra. 2-D

SERS spectra are mostly consistent to conventional signal-peak intensity-based SERS spectra

and helpful for identifying SERS signals in addition to the traditional Raman spectra for the

Raman analytical study.


Table 6.1. DFT calculated and experimental Raman wavenumber of the redox-sensitive

mode of different redox states of FMN with the energy of its HOMO and LUMO.

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Table 6.2. Redox potential of FMN (50nM) in different pH of Britton-Robinson buffer at

optimal scan rate 0.1 V/S.

The physiologically important redox states of FMN coenzyme are produced by applying

different electric potential and pH environments to the FMN solutions prepared in B-R buffer

medium. The redox potentials of FMN at different pH value are given in table 6.2. At most of the

pH environments, the CV curves (Figure 6.6) of FMN have the single redox peaks, which is

likely because of the formation of the complex mixture from its neutral and ionic redox species

with average redox potential.103 The initial species of FMN at the acidic buffer medium (pH 3-5)

is likely to be FMN only, it is because the pKa of FMN is closer to 10, and it does not lose amine

proton in the acidic buffer solutions. In the environment of pH 3-5, the proton concentration is

sufficient for the protonation of the FMN, so it is possible that the FMN undergoes a two-

electron/two-proton reduction in a voltammetry induced electric potential to form the fully

reduced flavin FMNH2.10,11,13-15,21,118 At the neutral, and moderately alkaline buffer solution (pH

7-9), the initial species of FMN is still FMN, however the proton concentration in this pH range

is much lower than the FMN concentration, therefore the reduced forms of FMN may not

undergo instantaneous protonation reaction. But, at strong alkaline medium, when the pH of

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buffer medium is greater than the pKa of FMN (pKa 10), the FMN more possibly loses its amine

proton and generates the FMN- as its initial redox species. In this condition, the FMN undergoes

a two-electron reduction process and produces the redox species like FMN2-•, and FMN3-.

Table 6.3. The Raman assignment of different redox states of FMN coenzymes.

The application of the SERS technique is found remarkably effective to probe and

characterize the electric potential and pH dependent redox states of FMN. Based on the

literature, we have characterized the SERS spectra of FMN in to two regions: (1) the spectral

region above 1000 cm-1, which is called the high-wavenumber or high-intensity region, and (2)

the region below 1000 cm-1, which is called the low-wavenumber or low-intensity region.119 The

low wavenumber region contains in- and out-plane vibrations of the ribityl chain or the flavin

ring system, which are not our focus in this study. The conventional visual speculation of redox

states of FMN also validates the formation of different redox states during electrochemical

processes; FMN is yellow in the oxidized state, red or purple in the semi-reduced anionic state,

blue in the neutral semiquinone state and colorless in the completely reduced state.120 The DFT

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Raman calculation of thermodynamically feasible redox states of FMN provides useful

information about its redox-sensitive Raman modes. The consistency is found between DFT

calculated and experimental Raman spectra which is worthwhile to determine the redox states of

FMN in specific experimental conditions (Figure. 6.13). The assignment of redox-sensitive

Raman peak of those FMN redox states are achievable in comparison of the experimental results

(Figure. 6.2) and the DFT calculated Raman mode signals (Figure. 6.1). The presence of the

redox state of FMN in an experimental condition is characterized by examining the consistency

found between experimental SERS spectra and DFT Raman spectra. The visualization of DFT

calculated molecular vibration, and consideration over electrochemical redox processes of FMN

in the B-R buffer solution show that the redox-sensitive mode of FMN is related to N1=C10a-

C4a=N5 bond, this mode is indicated by the dotted ellipsoid in scheme 1. This Raman mode is

varied in the range of 1490 to 1520 cm-1 based on the electric potential and pH value supplied

into FMN solutions.

Besides the above-mentioned redox-sensitive region, there are prominent SERS peaks in

a range of 1520-1538 cm-1 which are related to five redox species of FMN commonly found in

all experimental conditions irrespective of acidic, neutral, or alkaline buffer mediums and

applied electric potentials. According to DFT Raman calculation, these redox species are related

to N5=C4a symmetric stretching, C10a=N1, C9a-N10-C10a twisting, and N10-C10a-C4a twisting of the

redox species FMN (1523 cm-1), FMN•-(1524 cm-1), FMN2- (1526 cm-1), FMNH• (1524 cm-1),

and FMN•-(1525 cm-1). The pictures of corresponding Raman mode assignment of these redox

states are shown by the red arrow lines in scheme 1, and 2. In experimental spectra, these peaks

are observed in the range of 1520-1538 cm-1 and seated around the dotted black line (Figure.

6.6). In the alkaline B-R buffer medium (pH 7.4-11), the SERS spectrum of oxidized state of

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FMN has a Raman peak at 1498 cm-1; this mode is primarily contributed by N5-H bending, and

partially related to N1=C10a stretching, and C4a-N5-C5a stretching.121 This mode is downshifted to

1492 cm-1 when FMN is reduced by supplying negative potential (1V) (Figure. 6.2A-D); the

downshifting is induced by the weakening of C4a-N5, and N1=C10a stretching due to the decrease

of oxidation states of N5 and N1 respectively.122,123 The experiment shows that the process is

reversible, i.e. the supply of the positive electric potential to the reduced state leads to the

disappearance of Raman peak at 1492 cm-1 and the reappearance of the peak at 1498 cm-1.

Figure 6.6. The Cyclic voltammetry curve obtained from the mixture of FMN (50nM) and

AgNP@SiO2 in B-R bufferin different pH using a CHI 600C electrochemical workstation

equipped with a home-modified electroscopic cell (Figure 2.9 in chapter 2). Indium tin oxide

(ITO) is used as a working electrode, silver chloride (AgCl) as a reference electrode, and

197

Platinum (Pt) as a counter electrode. The optimal scan rate for the adequate signal-noise ratio is

found 0.1 V/s. The redox potentials of FMN in the optimal scan rate are given in table 6.2.

Figure 6.7. Two-dimensional (2-D) distribution Raman spectrum of FMN in acidic and alkaline

medium.(A) 2-D distribution SERS spectrum of oxidized and (B) reduced states of FMN at pH

5. (C) The 2-D distribution SERS spectrum of oxidized and (D) reduced state of FMN at pH 11.

The blue ellipses are used to emphasize the regions of redox-sensitive Raman peak.

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Figure 6.8. Two-dimensional (2-D) distribution Raman spectrum of FMN at pH 3 and 5.2-D

distribution Raman spectrum of FMN from (A) oxidized and (B) reduced state at pH 3. The 2-D

distribution Raman spectrum of FMN from (C) oxidized and (D) reduced states at pH 5

respectively. The integrated SERS spectrum (red) has been placed over 2-D distribution Raman

spectrum to test the consistency between conventional SERS spectrum and 2-D distribution

Raman spectrum.

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Figure 6.9. Two-dimensional (2-D) distribution Raman spectrum of FMN at pH 7 and 7.4.2-D

distribution Raman spectrum of FMN from (A) oxidized and (B) reduced state at pH 7. The 2-D

distribution Raman spectrum of FMN from (C) oxidized and (D) reduced states at pH 7.4



Raman spectrum.

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Figure 6.10. Two-dimensional (2-D) distribution Raman spectrum of FMN at pH 7.6 and 9.2-D

distribution Raman spectrum of FMN from (A) oxidized and (B) reduced state at pH 7.6. The 2-

D distribution Raman spectrum of FMN from (C) oxidized and (D) reduced states at pH 9



Raman spectrum.

201

Figure 6.11. Two-dimensional (2-D) distribution Raman spectrum of FMN at pH 11.The 2-D

distribution Raman spectrum from (A) oxidized, and (B) reduced states of FMN at pH 11. The

SERS spectra from FMN at (C) dried states and (D) wet states (KCl solution) without use of

applied potential. The integrated SERS spectra (red) have been placed over 2-D distribution

Raman spectrum to test the consistency between conventional SERS spectrum and 2-D

distribution Raman spectrum.

Above mentioned experimental evidence proves that these SERS peaks are related to

redox-sensitive modes. The Raman peaks for the oxidized states are seated on the dotted red line

located at 1498 cm-1 while The Raman peaks for reduced states are slightly off from the red

dotted line and positioned at 1492 cm-1 on the blue dotted line (Figure 6.6). On the basis of our

DFT calculations these two Raman signals are related to 1502, and 1489 cm-1 respectively which

are originated from N5-H bending, N1-C10a stretching and the asymmetric C4a-N5-C5a

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stretching.12 These modes are the characteristic redox-sensitive modes of FMN and correlated to

protonation, deprotonation, and electron transfer processes.

Similarly, our experiment shows that the Raman peaks of FMN in the acidic medium (pH

3-5) are downshifted from 1512-1514 cm-1 to 1508-1510 cm-1 when it is brought into the reduced

state by supplying the negative potential (Figure 6.2F-G). The SERS peaks of oxidized state of

FMN are seated on a solid red line at the region of 1512-1514 cm-1, and that of the reduced state

located at 1508-1510 cm-1 are seated on the solid blue line. The downshift of Raman

wavenumber is explained as a waning of the C10a=N1 and N5a=C4a bonds and the formation of the

newer bonds N5-H, N1-H, and N5-C4a=C10a-N1. When a positive potential is supplied into a newly

formed reduced state, the Raman peak of FMN at 1508-1510 cm-1 gradually disappears with the

reappearance of a peak at 1512-1514 cm-1, which means these reversibly reproducible peaks are

characteristic signatures for the redox process of the FMN in the acidic medium. The

experimental results are consistent to DFT calculation, where the DFT Raman peak position of

FMN located at 1523 cm-1 is shifted to 1511 cm-1 for FMNH2•+, and to 1514 cm-1 for FMNH2

redox states (Figure 6.1). After analysis of experimental data and DFT calculation, we could tell

that the reappearance and disappearance of experimental Raman peak at 1508-1510 cm-1 and

1512-1514 cm-1 are contributed by the formation of FMNH2•+ and FMNH2 redox states

respectively.

The stretching modes: C10a=N1 and N5a=C4a bonds are stronger for the redox species

FMNH•, FMNH• -, and FMN, which are responsible for producing Raman peaks at around 1523-

1525 cm-1. Alternatively, the stretching of N5-H, N1-H, and N5-C4a=C10a-N1 observed in the redox

species FMNH2•+

and FMNH2 are weaker in the redox species FMNH•, FMNH•-, and FMN

resulting the disappearance of Raman modes at 1508-1510 cm-1 (FMNH2•+) and 1512-1514 cm-1

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(FMNH2), and appearance of Raman peaks at around 1524 cm-1. The redox-sensitive peaks of

FMN in the buffer medium of pH 7 is broadened at the region of 1495-1520 cm-1 making it

harder to distinguish the oxidized state and reduced state of FMN (Figure 6.2E). The broadening

of Raman peak could be explained as the ambiguous pH environment, which neither allows to

follow the acidic pathways merely nor the alkaline pathways for the redox processes.

Furthermore, the redox species FMNH2•+ and FMNH2 have DFT calculated Raman peaks

at 1511 cm-1 and 1514 cm-1 respectively. These DFT calculated Raman peaks are consistent to

experimentally determined characteristic reduction and oxidation region of FMN in the acidic

medium located at 1508-1510 cm-1 and 1512-1514 cm-1 respectively. The HOMO-LUMO energy

of FMNH2•+ and FMNH2 shows that these redox states are energetically more favorable (Figure

6.12), and literature shows that they are only formed in the acidic aqueous/buffer medium.11 The

Raman wavenumber of FMNH2•+ at 1508-1510 cm-1 is summarized as the collective contribution

of N5-C4a-C10a=N1 stretching and stretching of N5-C4a in rings. In addition, the assignment of

Raman wavenumber of FMNH2 at 1512-1514 cm-1 is summarized as the combined contribution

of wagging of two hydrogen atoms bound to N5 and N1 atoms, stretching of C10a-N1 and N5-C4a,

and twisting of C4a= C10a, the brief picture of vibrational mode of FMNH2 related to 1512-1514

cm-1 has been shown in scheme 1. The experimental SERS peaks at 1508-1510 and 1512-1514

cm-1 observed in the FMN molecule in the acidic buffer medium are also found in DFT Raman

calculations for FMNH2•+ and FMNH2 redox species. In DFT calculated Raman spectra,

FMNH2•+ redox species has a Raman peak at 1511 cm-1; this Raman shift is in the range of

experimental value 1508-1512 cm-1.

204

Figure 6.12. The calculated HOMO-LUMO energy state of different redox species of

FMNusing density functional theory (DFT) method on a B3LYP level with a basis set of 6-31G

(d). The HOMO-LUMO energy calculation shows that the redox states FMN, FMNH•, FMNH2•,

and FMNH2•, and FMNH2 are energetically more stable redox states.

Similarly, the DFT Raman spectra of FMNH2 has a peak at 1514 cm-1 which is consistent

with the experimentally found value at 1512-1514 cm-1. The consistencies are found between

experimental results and theoretical calculations, which are important to predict the formation of

specific redox states of FMN under the experimental conditions. The FMN molecule in alkaline

buffer does not produce FMNH2•+ and FMNH2 species and does not show peaks at the region of

1508-1510 cm-1 and 1512-1514 cm-1, instead it shows downshifting of redox-sensitive Raman

peak from the region of 1498 cm-1 to 1492 cm-1 when it is supplied with negative electric

potentials which is due to the formation of redox species FMN3- (1492 cm-1) from FMN2-• (1498

cm-1). In the alkaline medium, FMN2-• is formed by the reduction of FMN- or oxidation of FMN3-

205

by one electron, and FMN3- is either formed by the reduction of FMN- with two electrons in the

two-step mechanism or reduction of FMN2-• by one electron (scheme 2). Beside Peaks at 1498

cm-1 and 1492 cm-1, FMN in the alkaline buffer medium also gives characteristic Raman peaks in

each applied electric potential at around 1555-1559 cm-1 (seated on the black solid line in Figure

6.6 A-D). After analysis over the redox mechanism of FMN in the acidic and alkaline buffer

medium, it is found that FMN- is the species that is only formed in the alkaline buffer medium

and gives the Raman peak at the range of 1555-1559 cm-1. The experimental and theoretical facts

support that the experimental SERS peak of FMN at 1555-1559 cm-1 in the alkaline buffer

medium is contributed by a C4-N3 symmetric stretch of redox species FMN-. The DFT Raman

spectrum of FMNH- redox state has a peak at 1557 cm-1; which is contributed by C10a=N1

stretching. This redox state is present in both acidic and alkaline mediums of FMN, but

experimental Raman peak at 1555-1559 cm-1 is only observed in the alkaline buffer medium. The

Raman peaks of FMNH- in the acidic medium could be coupled to the peaks at the region of

1565-1573 cm-1, which are seated on the solid green line (Figure 6.6D-G). It is not a surprise for

shifting or disappearance of experimental SERS peak for FMNH- in acidic medium because its

concentration can be decreased by other kinetically favorable redox species during the reversible

redox mechanism. After careful analysis of our observation from experimental SERS spectra,

DFT calculated Raman spectra of many redox states of FMN coenzymes, and electrochemical

properties of different-different types of chemical bond available in the redox states of FMN, we

can assemble this information to determine the pathways for the pH-dependent redox process of

FMN which are summarized in Scheme 1, and scheme 2 respectively. This experimental and

analytical approach is successful to probe the varieties of redox species and the redox pathway of

FMN in non-biological environments.

206

Figure 6.13. The consistency of experimentally collected SERS peak position and DFT

calculated Raman peak position of different redox states of FMNcoenzyme for their redox-

sensitive mode. The Raman wavenumber of redox species of FMN are written inside parenthesis

in red color for experimental SERS spectra, and in black for DFT Raman spectra.

In addition of purified FMN, we have also observed the consistent behaviors from the

FMN cofactors present in wild-type neural nitric oxide synthase (nNOS) (Figure 6.14). In the B-

R buffer solution at pH 5, the SERS signal of FMN cofactor present in the nNOS enzyme shows

the significant redox-sensitivity of oxidized state (1513 cm-1) and reduced state (1507 cm-1). This

result is consistent with the redox-sensitivity of the free FMN at pH 5 at its oxidized state (1508-

1510 cm-1) or reduced state (1512-1514 cm-1). This result suggests that the FMN cofactor in both

non-biological and biological systems likely follows the similar redox mechanism. These types

207

of studies could be helpful to understand the biochemical mechanisms of biologically important

molecules in cells and living systems.

Figure 6.14. The experimental results from the study of FMN in nNOS of rat (Rattus

norvegicus) using combined approach of cyclic voltammetry and SERS technique.(A) The cyclic

voltammetry curve obtained from the 50 nM solution of nNOS complex in the B-R buffer

solution at pH 5 (B) The crystal structure (PDB 1tll) of the neuronal nitric-oxide synthase

(nNOS) of rat (Rattus norvegicus), where the FMN domain is magenta, and the FAD domain is

grey. (C) The comparison of SERS spectra obtained from the fully reduced (-0.5 V) and oxidized

(+0.3 V) states of neuronal nitric oxide synthase (nNOS) in the B-R buffer solution at pH 5. The

reduction of the FMN molecules present in nNOS causes the downshifting of Raman mode from

1513 cm-1 to 1507 cm-1.

208

Scheme 1. The Redox mechanism of FMN coenzyme in the B-R buffer at pH 3-5

determined based on the DFT calculated Raman spectra of its redox species, experimental SERS

spectra in the different electrochemical conditions, and electrochemical properties of various

chemical bonds. The red arrow lines represent the Raman vibrational modes related to Raman

wavenumbers of given redox species.

In our experiment, we have observed the shifting of redox-sensitive Raman peak of FMN

from 1513 cm-1 (FMNH2) to 1498 (FMN2-•) and 1508 (FMNH2•+) during redox processes. These

results from our FMN SERS study are comparable to the results in the published literature. The

correspondence is observed between our SERS measurements on FMN and the Raman

measurements on Lumiflavin by Carey and coworker, where the Raman peak position of

209

Lumiflavin at 1514 cm-1 has been shifted to 1496, 1504 and 1506 cm-1 in different experimental

conditions.110 It is also found that our experimental data has good agreement to the results from

FAD Raman measurements, where FAD Raman peaks are shifted from 1500 cm-1 to 1499, 1502

and 1503 cm-1 due to different experimental environments and redox processes.104 It is also

found that the wavenumber of the redox-sensitive Raman mode of the oxidized or natural form

of flavoproteins or flavin cofactors are downshifted when they are fully reduced which is also

consistent with our current results. We observed a redox-sensitive SERS peak at around 1500 cm-

1 in our current results from FMN. In literature, similar results are consistently found for many

flavin cofactors regardless of their source flavoproteins.124 The appearance of this redox-

sensitive Raman peak at around 1500 cm-1 in all flavoproteins and flavin cofactors could be

explained as the presence of shared redox-sensitive modes N1=C10a-C4a=N5 as represented by

blue dotted ellipsoid in the scheme 1. Nevertheless, there is some variation for the Raman

wavenumber of redox-sensitive Raman modes between our results and results in the previously

published literature. This discrepancy of wavenumber could be explained as the result of

environmental effects.

Scheme 2. The Redox mechanism of FMN coenzyme in B-R buffer at pH 7-11

determined based on DFT calculated Raman spectra of its redox species, experimental SERS

spectra in different electrochemical conditions, and electrochemical properties of various

chemical bonds. The red arrow lines represent the Raman vibrational modes related to the Raman

wavenumber of given redox species.

210

6.4. Summary

We have utilized the SERS or resonance SERS (SERRS) and DFT Raman calculation

technique to probe and characterize pH dependents redox states of FMN and entire redox

pathways. In our experiments, the varieties of redox states of FMN are generated using electric

potentials in the acidic and alkaline (3-11 pH) Britton-Robinson (B-R) buffer. The comparative

analytical approach of the DFT calculated Raman spectra of various redox states of FMN with

results from SERS experiments are found useful to characterize pH and electric potential

dependent redox states of FMN and determine the mechanism of redox processes. We have

characterized the redox-sensitive Raman peaks observed at 1498 cm-1 and 1492 cm-1 in the

alkaline buffer as the result of the formation of energetically less stable redox species: FMN3- and

211

FMN2-• respectively. Similarly, we have also characterized the energetically more favorable

redox-sensitive redox species in the acidic medium: FMNH2 (1512-1514 cm-1), and FMNH2•+

(1508-1510 cm-1). Likewise, we have also characterized the pH specific redox species FMN-

(1555-1559 cm-1) in the alkaline medium. Our experimental and analytical approach can probe

the redox species and redox pathways of FMN in both non-biological and biological

applications.

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