label-free electrochemical dna and protein …finally, we evaluated functional pedot thin films as...

LABEL-FREE ELECTROCHEMICAL DNA AND PROTEIN

DETECTION USING RUTHENIUM COMPLEXES AND

FUNCTIONAL POLYETHYLENEDIOXYTHIOPHENES

XIE HONG

(M. Sc., NUS)

A THESIS SUBMITTED

FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

DEPARTMENT OF CHEMISTRY

NATIONAL UNIVERSITY OF SINGAPORE

2008

LIST OF PUBLICATIONS

Tansil, N. C.; Xie, H.; Xie, F.; Gao, Z. Q., Direct detection of DNA with an electrocatalytic threading intercalator. Analytical Chemistry 2005, 77, (1), 126-134. Tansil, N. C.; Xie, F.; Xie, H.; Gao, Z. Q., An ultrasensitive nucleic acid biosensor based on the catalytic oxidation of guanine by a novel redox threading intercalator. Chemical Communications 2005, (8), 1064-1066. Xie, H.; Tansil, N. C.; Gao, Z. Q., A redox active and electrochemiluminescent threading bis-intercalator and its applications in DNA assays. Frontiers in Bioscience 2006, 11, 1147-1157. Xie, H.; Yang, D. W.; Heller, A.; Gao, Z. Q., Electrocatalytic oxidation of guanine, guanosine, and guanosine monophosphate. Biophysical Journal 2007, 92, (8), L70-L72. Luo, S-C; Xie, H.; Chen, N. Y.;Yu, H-h; Ying, J. Y., Functional PEDOT thin film for electrochemical DNA biosensing and controlled cell adhesion. To be submitted. Xie, H.; Luo, S-C; Yu, H-h; Ying, J. Y., Functional PEDOT nanowires for label-free protein detection. To be submitted. Patents and Technology Disclosures: Xie, H., Gao. Z.Q., Xie, F., Determination of nucleic acid using electrocatalytic intercalators, WO 2006/025796, US 2006/0046254, Mar 2006. Yu, H-H, Ying, J. Y-R., Luo, S-C, Xie, H., Chen, N.Y., polyethylenedioxythiophene (PEDOT) biointerfaces for DNA detection, IBN Technology Disclosure, Nov 2006. Yu, H-H., Ying, J. Y-R, Xie, H., Kantchev, E. A. B, Luo, S-C., Non-fouling polyethylenedioxythiophene (PEDOT) biointerfaces for controlled adhesion of cells and proteins, IBN Technology Disclosure, Jun 2007. Conference Presentations: Xie, H.; Luo, S-C; Yu, H-h; Ying, J. Y., Non-fouling PEDOT for controled cell adhesion, Oral presentation, NanoBioEurope 2008, Barcelona, 9-13 Jun 2008. Xie, H.; Luo, S-C; Chen, N. Y.; Yu, H-h; Ying, J. Y., Functional PEDOTs for electrochemical biosensors, Oral presentation, Regional Electrochemical Meeting of South-East Aisa 2008, Singapore, 5-7 Aug 2008.

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ACKNOWLEDGEMENTS

It is a pleasure to thank many people who made this thesis possible.

I would like to start by thanking my advisor, Dr Hsiao-hua (Bruce) Yu, for his

enthusiastic supervision; and my co-advisor, Dr Choon Hong Tan, for many valuable

advices. I would also like to thank my ex-advisors, Dr Zhiqiang Gao and Dr Daiwen

Yang. Although they are unable to guide me throughout my whole PhD work, I am

grateful to their guidance and mentorship during my first year.

I am very grateful to Prof Jackie Y. Ying and Ms Noreena AbuBarka for

allowing me to pursue my dreams in Institute of Bioengineering and Nanotechnology

(IBN). I truly appreciate their constant support over the years. Without them, IBN

would not be so successful today and I would not be able to finish my projects so

smoothly. My gratitude also extends to all IBN administrative staffs for their general

support.

Many wonderful friends have kept me balanced and lighthearted through my

graduate study. They have contributed to this thesis along the way. I would like to

especially thank Dr. Shyh-Chyang Luo, Zaoli Zhang, Natalia Tansil, Emril Ali,

Naiyan Chen, Dr. Eric Kantchev, Dr Shujun Gao, Dr Han Yu, Dr. Hongwei Gu, Dr

Alex Lin, Shawn Tan, Dr Jiang Jiang, Dr Majad Khan, James Hsieh, Guangrong Peh,

Huilin Shao, Dr Peggy Chan and Lishan Wang. I am thankful for their valuable

discussions, assistance, friendship, and for making my stay in IBN enjoyable. I would

like to express my deepest gratitude to those who helped me get through the difficult

times. I thank you for all the emotional support, entertainment and caring you

provided.

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Finally I am forever indebted to my family for their love and understanding. I

would like to thank my parents for their endless support when it was most needed.

This thesis is dedicated to you.

Last but not least, I would like to thank IBN, BMRC and A*Star for the

funding.

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TABLE OF CONTENTS

List of Publications

Acknowledgements

Table of Contents

Summary

List of Abbreviations

List of Figures, Schemes and Tables

1 Introduction ................................................................................................................1

1.1 Background...........................................................................................................1

1.1.1 Electrochemical Biosensors......................................................................2

1.1.2 Label-Free Electrochemical/Electrical Assays........................................7

1.1.3 Electroactive Conducting Polymers for Biosensing .............................10

1.2 Motivation and Objectives.................................................................................11

1.3 Scope...................................................................................................................12

1.4 Thesis Outline.....................................................................................................12

2 Ruthenium-Complexed Electroactive Intercalators for Label-Free DNA

Detection.......................................................................................................................... 14

2.1 Introduction.........................................................................................................14

2.2 Experimental.......................................................................................................18

2.2.1 Materials and Reagents...........................................................................18

2.2.2 Synthesis of Electroactive DNA Intercalators ......................................19

2.2.3 Apparatus.................................................................................................22

2.2.4 Sensor Construction................................................................................23

2.3 Results and Discussion.......................................................................................25

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2.3.1 Synthesis and Characterization of Electroactive DNA Intercalators...25

2.3.2 Intercalation with DNA ..........................................................................29

2.3.3 Application for Label-free DNA Detection...........................................33

2.4 Conclusions.........................................................................................................43

3 Ruthenium-Based Polymer Complexes for Electrocatalytic Guanine

Oxidation......................................................................................................................... 44

3.1 Introduction.........................................................................................................44

3.2 Experimental.......................................................................................................46


3.2.2 Synthesis of Ruthenium-complexed Redox Polymers .........................46

3.2.3 Preparation of Redox Polymer Modified Electrodes............................49

3.2.4 Apparatus.................................................................................................49


3.3.1 Synthesis and Characterization of Redox Polymers.............................50

3.3.2 Redox Polymer Modified Electrode ......................................................53

3.3.3 Electrocatalytic Oxidation of Guanine on Modified Electrode............54

3.3.4 Redox Titration .......................................................................................56

3.3.5 Oxidation of Guanosine and Guanosine Monophosphate (GMP) .......58

3.4 Conclusions.........................................................................................................60

4 Nanostructured Functional Polyethylenedioxythiophenes (PEDOTs)........... 61

4.1 Introduction.........................................................................................................61

4.1.1 Conducting Polymers..............................................................................61

4.1.2 Nanostructured Conducting Polymers...................................................63

4.1.3 Synthesis of 1-D Conducting Polymer Nanostructures........................64

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4.1.4 1-D Polyethylenedioxythiophene (PEDOT) Nanostructures ...............65

4.2 Experimental.......................................................................................................66


4.2.2 Chemical Polymerization .......................................................................67

4.2.3 Electrochemical Polymerization ............................................................68

4.2.4 Characterization ......................................................................................68


4.3.1 Surfactant Template-Guided Nanofiber Synthesis ...............................68

4.3.2 Stepwise Electropolymerization.............................................................73

4.3.3 Electrical Field-Assisted Nanowire Growth..........................................77

4.4 Conclusions.........................................................................................................79

5 PEDOT Nanowires for Label-Free Protein Detection ...................................... 81

5.1 Introduction.........................................................................................................81

5.2 Experimental Section .........................................................................................83


5.2.2 Device Fabrication and Nanowire Synthesis.........................................83

5.2.3 Aptamer Immobilization and Protein Binding......................................84

5.2.4 Electrical Measurement ..........................................................................84


5.3.1 Device Characteristics ............................................................................85

5.3.2 Biomolecule Conjugation.......................................................................86

5.3.3 Protein Detection.....................................................................................88

5.3.4 1-D Nanostructure vs 2-D Film..............................................................91

5.4 Conclusions.........................................................................................................93

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6 Functional PEDOT Nanobiointerface: Toward in vivo Applications ............. 95

6.1 Introduction.........................................................................................................95

6.2 Experimental.......................................................................................................97


6.2.2 Electropolymerization and Film Synthesis............................................98

6.2.3 Electrochemical Characterization ..........................................................99

6.2.4 Polymer Film Analysis ...........................................................................99

6.2.5 Protein Adsorption................................................................................100

6.2.6 Cell Culture ...........................................................................................100

6.3 Results and Discussions...................................................................................102

6.3.1 Synthesis and Characterization of Functional PEDOT Thin Films ...102

6.3.2 Biocompatibility of Functional PEDOT Thin Films ..........................106

6.3.3 Adhesive and Non-adhesive PEDOT Nanobiointerfaces...................107

6.3.4 Controlled Cell Patterning....................................................................110

6.3.5 Biotin-functionalized PEDOT Nanobiointerface................................111

6.3.6 Peptide-functionalized PEDOT Nanobiointerface..............................115

6.4 Conclusions.......................................................................................................120

7 Conclusions and Outlook ..................................................................................... 121

References

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SUMMARY

This thesis presents our studies on the development of label-free

electrochemical biosensors for DNA/protein detection. The urgent need for the

development of point-of-care devices for the detection of infectious agents and

cancer-related biomarkers motivate us to keep searching for simple, fast, sensitive yet

affordable analytical tools. We have demonstrated two very different approaches for

label-free DNA/protein detection with electrochemical transduction. Ruthenium-

complexed electroactive DNA threading intercalators and aptamer-modified

polyethylenedioxythiophene (PEDOT) nanowires were used as signal reporters for the

corresponding binding events.

In part I, we studied label-free electrochemical DNA detection using

ruthenium-complexed intercalators. Two ruthenium-complexed electroactive DNA

intercalators were synthesized, characterized, and their application for label-free DNA

detection were investigated. One based on electrochemiluminescence, and the other

one based on electrocatalytic oxidation of guanine bases in the DNA sequences. The

electroactive intercalators are dual functional: selective binding of double-stranded

DNA (ds-DNA) and generation of catalytic electrochemical signals. This feature

allows simple and sensitive detection. Moreover, the oxidation potential of guanine

base and its corresponding nucleoside and nucleotide under physiological buffer

condition were determined experimentally first time by electrocatalytic oxidation

titration using ruthenium-complexed redox polymer modified electrode.

In part II, we explored the use of a conducting polymer, functionalized

polyethylenedioxythiophene (PEDOT), as an intrinsic transducer for label-free protein

sensing. Various approaches for the synthesis of 1-D PEDOT nanostructures were

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studied. Functional PEDOT nanowires were directly synthesized across the electrode

junction under the assistance of an external electric field. Such PEDOT nanowires

devices can be applied immediately after synthesis for field effect transistor (FET)

based sensing, eliminating complicated post-synthesis alignment and assembly.

Label-free detection of a blood-clogging factor, thrombin, was demonstrated using

aptamer-modified PEDOT nanowires. In comparison with 2-D thin films, 1-D

nanostructures are crucial for field effect transistor (FET) based sensing. The PEDOT

nanowire based sensing platform is applicable for label-free detection of DNA as well

as proteins which their DNA aptamers are available.

Finally, we evaluated functional PEDOT thin films as tunable

nanobiointerfaces for effective biomolecule immobilization and controlled cell

adhesion, for future cell-based sensing and other in vivo applications. Particularly,

biotin-functionalized PEDOT surface and peptide-functionalized PEDOT surface

were achieved through direct polymerization from mixed monomer solution and facile

post-polymerization functionalization. Specific protein adsorption and controlled cell

attachment were demonstrated on these biologically-relevant functionalized PEDOT

surfaces. Similar modification is also feasible on nanostructured PEDOT surfaces, and

we expect to see more exciting in vivo applications in the future.

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LIST OF ABBREVIATIONS

AFM Atomic force microscope

APS Ammonium persulfate

BSA Bovine serum albumin

Bpy 2,2′-bipyridine

CV Cyclic voltammetry

CMC Critical micelle concentration

CP Capture probe

CPNWs Conducting polymer nanowires

CTAB Cetyltrimethylammonium bromide

DNA Deoxyribonucleic acid

ds-DNA Double-stranded DNA

ss-DNA Single-stranded DNA

2/1E Half wave potential

EB Ethidium bromide

ECL Electrochemiluminescence

EDC 1-ethyl-3-[3-(dimethylamino)-propyl]carbodiimide

EDOT 3,4-ethyelendioxythiophene

EIS Electrochemical impedance spectroscopy

ELISA Enzyme-linked immunosorbent assay

FBS Fetal bovine serum

FET Field effect transistor

GMP Guanosine monophosphate

HR-MS High resolution mass spectrometry

IME Interdigitated microelectrode

ITO Indium tin oxide

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LSV Linear scan voltammograms

MLCT Metal-to-ligand charge-transfer

mRNA Messenger ribonucleic acid

ND Naphthalene diimide

NHE Normal hydrogen electrode

NHS N-hydroxysulfosuccinimde

PAA Polyacrylamide

PIND N,N′-bis[(3-propyl)imidazole]-1,4,5,8-naphthalene diimide

PBS Phosphate buffered saline

PCR Polymerase chain reaction

PDMS Polydimethylsiloxane

PEDOT Polyethylenedioxythiophene

PPy Polypyrrole

PVI Poly(4-vinylpyridine)

PVP Poly(N-vinylimidazole)

QCM Quartz crystal microbalance

SDS Sodium dodecyl sulfate

SEM Scanning electron microscope

SWV Square wave voltammetry

TE Tris-EDTA buffer

TEM Transmission electron microscope

TMEDA N,N,N′,N′-tetramethylethylenediamine

TPA Tri-n-propylamine

UV-Vis Ultra violet-visible

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LIST OF FIGURES

Figure 1-1: Schematic presentation of a biosensor .........................................................1

Figure 1-2: Schematic of nanowire based FET sensor (adapted from Ref 62)...............9

Figure 2-1: Cyclic voltammograms of the starting materials Ru(bpy)2Cl2 (····), reaction mixture after refluxing with PIND for 30 min in ethylene glycol (---), and the purified final product (―). Supporting electrolyte: PBS; Scan rate: 100 mV/s.....26

Figure 2-2: Cyclic voltammograms of the purified Ru-PIND-Ru in PBS at scan rate of (a) 100, (b) 200, (c) 300, (d) 400, and (e) 500 mV/s. ................................................26

Figure 2-3: Cyclic voltammograms of Ru(dmbpy)2Cl2 after refluxing with PIND for (a) 0, (b) 10, and (c) 30 min. Supporting electrolyte: PBS; Scan rate: 100 mV/s. .......27

Figure 2-4: UV-Vis absorption spectra of (a) PIND-Ru-PIND, (b) Ru(dmbpy)2(Im)2, (c) Ru(dmbpy)2Cl2, and (d) PIND in ethanol. ................................................................29

Figure 2-5: UV-Vis spectra of 25 µM Ru-PIND-Ru (resolution 0.10 nm) as a function of increasing concentration of salmon sperm DNA (in base pair) of (a) 0, (b) 25, (c) 50 and (d) 100 µM. Insert: Enlarged UV-Vis adsorption spectra of the intercalative binding region. ............................................................................................30

Figure 2-6: (A) Fluorescent displacement titration curve of Ru-PIND-Ru against a 5 μM hairpin oligonucleotide with EB. (B) Scatchard plot for the titration of hairpin oligonucleotide/EB with Ru-PIND-Ru. ..........................................................................32

Figure 2-7: UV-Vis absorption spectra of 20 μM PIND-Ru-PIND in 0.10 M pH 7.0 phosphate buffer with increasing concentration of salmon sperm DNA (from top, 0, 20, 40, 60, 80 and 100 μM in base pair). ........................................................................33

Figure 2-8: Cyclic voltammograms of 200 nM of (a) poly(T)40 hybridized to a non-complementary capture probe coated electrode, and (b) poly(AT)20, (c) poly(AG)20, and (d) poly(G)40 hybridized to their complementary CP coated electrode, respectively. Scan rate: 100 mV/s. ..................................................................................34

Figure 2-9: Cyclic voltammograms of TP 53 hybridized to (a) perfectly-matched and (b) one-base-mismatched biosensors. Hybridization was carried out in TE buffer containing 1.0 mg of mRNA. Scan rate: 100 mV/s........................................................36

Figure 2-10: (a) ECL intensity at 610 nm versus potential profiles, cyclic voltammograms of (b) 5.0 μM PIND-Ru-PIND in 0.10 M phosphate buffer (pH 7.0), and (c) 5.0 μM PIND-Ru-PIND in TPA saturated phosphate buffer. Scan rate: 20 mV/s. For clarity, the voltammogram of PIND-Ru-PIND was scaled up 50 times. ....38

Figure 2-11: (a) Photoluminescence spectrum of PIND-Ru-PIND (430 nm illumination) in 0.10 M phosphate buffer and (b) ECL spectrum of PIND-Ru-PIND in a TPA saturated 0.10 M phosphate buffer. .....................................................................39

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Figure 2-12: Linear scan voltammograms (LSV) of PIND-Ru-PIND bound to (a) 200 nM of complementary DNA, and (b) 1.0 μM non-complementary DNA hybridized biosensors. Supporting electrolyte: 0.10 M phosphate buffer (pH 7.0), potential scan rate 100 mV/s....................................................................................................................40

Figure 2-13: ECL responses at 610 nm of PIND-Ru-PIND bound to biosensors hybridized with (a) 1 nM non-complementary target, (b) 50 pM one-base-mismatched target, and (c) 50 pM complementary target. Poise potential: 1.0 V, ECL measurement was done in TPA saturated phosphate buffer. ................................................................42

Figure 2-14: Effect of TPA (•) and applied potential () on the ECL responses at 610 nm of 50 pM complementary DNA after incubation in 10 μM PIND-Ru-PIND.........43

Figure 3-1: Cyclic voltammograms of the reaction mixtures at different reaction time during the synthesis of PVIPAA-Ru(bpy)2Cl: (a) 0, (b) 2 h and (c) 20 h.....................51

Figure 3-2: UV-Vis spectra of Ru(bpy)2Cl2 before (―) and after (---) grafting to the polymer backbone. ...........................................................................................................52

Figure 3-3: (A) Sweep-rate dependency of the CV of a PVIPAA-Ru(bpy)2Cl2 coated ITO electrode in PBS, scan rate=20, 50, 100, 200, 500, 1000 mV/s, following arrow direction. (B) The plot of anodic peak current with scan rate. ......................................53

Figure 3-4: Cyclic voltammograms of redox polymer thin film coated ITO electrodes in PBS. From left to right: (a) PVPPAA-Ru(OCH3), (b) PVPPAA-Ru(CH3), (c) PVPPAA-Ru, (d) PVIPAA-Ru(COOCH3), (e) PVPPAA-Ru(COOCH3). ...................54

Figure 3-5: Cyclic voltammograms of blank ITO in (A) PBS and (B) PBS with 0.5 mM guanine at different pH: (―–) 10.5, (·····) 9.5, (– – –) 8.5, (· − · −) 7.5. ...............55

Figure 3-6: Cyclic voltammograms of a PVIPAA-Ru(bpy)2Cl thin film coated ITO electrode in (a) PBS and (b) PBS with 20 mM guanine. (c) Cyclic voltammogram of a bare ITO electrode with 50 mM guanine in PBS. Scan rate: 100 mV/s. ...................56

Figure 3-7: Titration curves showing the increase in the electrocatalytic guanine oxidation current when the pH is raised at redox polymer film coated ITO electrodes. From left to right, (a) PVIPAA-Ru(COOCH3), (b) PVPPAA-Ru, (c) PVPPAA-Ru(CH3), (d) PVIPAA-Ru, (e) PVPPAA-Ru(OCH3), (f) PVIPAA-Ru(OCH3)...........57

Figure 3-8: pH dependency of the threshold-potentials of (a) guanine (•), (b) guanosine (), and (c) GMP () electrooxidation, catalyzed by different polymers.....58

Figure 3-9: Chemical structures of guanine, guanosine and GMP. .............................59

Figure 4-1: Chemical structures of common conductive polymers. ............................62

Figure 4-2: Chemical structure of EDOT-OH and EDOT-COOH ..............................67

Figure 4-3: (a) SEM image and (b) TEM image of poly(EDOT-COOH) nanofibers. (c) HRTEM image of individual nanofiber. ...................................................................69

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Figure 4-4: SEM images of poly(EDOT-COOH) obtained at different surfactant concentration (SDS used as a surfactant and FeCl3 as oxidizing agent, monomer concentration was kept constant at 15 mM): (a) 30 mM, (b) 50 mM, (c) 60 mM, (d) 80 mM, and (e) 100 mM. ......................................................................................................70

Figure 4-5: SEM images of poly(EDOT-COOH) obtained at different monomer concentration: (a) 2, and (b) 20 mM, in the presence of 100 mM SDS and 40 mM FeCl3. SEM images of poly(EDOT-COOH) obtained from (c) 10x, (d) 5x, and (e) 2x dilution from the original mixture of 20 mM monomer, 100 mM SDS and 40 mM FeCl3..................................................................................................................................70

Figure 4-6: Schematic of the salt-assisted surfactant micelle transformation and formation of poly(EDOT-COOH) nanofibers. ...............................................................71

Figure 4-7: SEM image of Poly(EDOT-COOH) nanostructures obtained in the presence of 30 mM EDOT-COOH monomer and 10.5 mM CTAB with different oxidizing agent (A) 30 mM of APS (B) 60 mM FeCl3. .................................................73

Figure 4-8: FESEM image of the PEDOT nanotubes deposited on the microelectrodes consisting of a pair of gold interdigitate electrodes with 40 fingers (dimensions: 10 μm width, 4000 μm length, 50 nm thickness, 10 μm inter-electrode gap). Adapted from Ref 73 ...............................................................................................73

Figure 4-9: SEM images of (a) poly(EDOT-COOH) and (b) poly(EDOT-OH). Polymerization was done under constant current, 0.5 mA/cm2, in TBAPF6/CH3CN containing 10 mM monomer. Images at bottom panel are taken at higher magnification. ...................................................................................................................75

Figure 4-10: SEM images of poly(EDOT-COOH) polymerized under constant current, 0.25 mA/cm2, in (a) 0.1 M TBAPF6/CH3CN and (b) 0.1 M LiClO4 aqueous solution (9:1 H2O: CH3CN) containing 10 mM EDOT-COOH monomer. Images at bottom panel are taken at higher magnification. ............................................................76

Figure 4-11: SEM images of poly(EDOT-OH) polymerized under constant current, 0.1 mA/cm2, from 10 mM EDOT-OH aqueous solution with different surfactants (a) 50 mM SDS, (b) 5 mM Brij 35, and (c) 2% P123. Supporting electrolyte: 0.1 M LiClO4. ..............................................................................................................................77

Figure 4-12: (a) Optical and (b) scanning electron micrograph of poly(EDOT-COOH) nanowires grown between two Au electrodes under alternating electric field. ............78

Figure 4-13: Scanning electron micrograph of poly(EDOT-COOH) nanowires grown between two Au electrodes under DC field. ...................................................................78

Figure 5-1: Experimental setup of CPNW FET devices for protein detection. Au 1 and Au 2 represent two working electrodes (WE1 and WE2), served as source and drain respectively. Counter electrode is a Pt wire where the electrochemical gate potential is applied via the reference electrode (Ag/AgCl). A bias voltage (Vbias) applied between WE1 and WE2 (Vbias=VWE1-VWE2) is equivalent to source-drain current (Vsd). .....................................................................................................................84

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Figure 5-2: Electrical characteristics of poly(EDOT-COOH) nanowire device. (a) I-V curve, (b) Isd-Vsd characteristics at varying Vg (Vg = -0.4 to + 0.4 V, step = 0.1 V, scan rate = 1 mV/s) in 0.1 M LiClO4/buffer (pH=5), and (c) |Isd|-Vg plot at constant Vsd= -0.2 V, derived from the above Isd-Vsd characteristics.....................................................86

Figure 5-3: Isd-Vsd characteristics of the poly(EDOT-COOH) nanowire device at gate voltage of 0 V (a) before and (b) after aptamer immobilization. ..................................87

Figure 5-4: Normalized current change before and after immobilization of different biomolecules on the poly(EDOT-COOH) nanowires (Vg= 0, Vsd= 0.4 V). .................88

Figure 5-5: Typical Isd-Vsd characteristics of aptamer-modified PEDOT nanowire devices (a) before and (b) after incubation with thrombin (Vg= 0 V)...........................89

Figure 5-6: Normalized current change of aptamer-modified PEDOT nanowire devices after incubation with 100 nM thrombin: (a) 49-mer non-complementary probe, (b) 28-mer non-complementary probe, (c) thrombin-binding aptamer.........................90

Figure 5-7: (A) Overlay of Isd-Vsd curves after 1 h incubation with thrombin at concentration of 0, 1, 10, 100, 1000 nM, follow arrow direction. (B) Calibration curve of aptamer-modified PEDOT nanowire device: normalized current change (-∆I/I0) as a function of thrombin concentration. The source-drain current was measured at Vsd= 0.4 V and Vg=0 V. ............................................................................................................90

Figure 5-8: Electrical characteristics of poly(EDOT-COOH) thin film device. (A) Isd-Vsd characteristics at varying Vg (Vg = -0.4 to + 0.4 V, step = 0.1 V, scan rate = 1 mV/s) in 0.1 M LiClO4/buffer (pH=5), (B) |Isd|-Vg plot at constant Vsd (a) 0.2 V and (b) -0.2 V, derived from the Isd-Vsd characteristics. .............................................................91

Figure 5-9: Isd-Vsd characteristics of poly(EDOT-COOH) thin film device (a) before, (b) after aptamer immobilization, and (c) after thrombin binding (Vg= 0 V)...............92

Figure 5-10: Fluorescence microscope image of poly(EDOT-COOH) modified with (a) random sequence probe and (b) thrombin-binding aptamer after binding with thrombin and a Cy3-labeled 2nd aptamer. .......................................................................92

Figure 5-11: The major advantage of 1-D nanostructures (B) over 2-D thin film (A) for FET based biosensing. Adapted from Ref 211 ..........................................................93

Figure 6-1: Chemical structures of functionalized EDOT monomers .........................98

Figure 6-2: Chemical structures of (A) EDOT-C1-biotin, (B) EDOT-C8-biotin, and (C) EDOT-C15-biotin. .............................................................................................................98

Figure 6-3: (a) Electropolymerization of 10 mM of EDOT-OH monomers in CH3CN (---) and in aqueous microemulsion containing 0.05 M of SDS and 1 mM of HCl (––) with 0.1 mM of LiClO4 as supporting electrolyte at a scan rate of 100 mV/s. (b) In situ QCM measured weight gain during the electropolymerization shown in (a)......103

Figure 6-4: Electropolymerization of (A) 10 mM of EDOT-C8-biotin monomer, (B) 10 mM of EDOT-OH monomer with different ratio of EDOT-C8-biotin, and (C) 10 mM of EDOT-OH monomer with 10% of EDOT-C8 -biot in , in aqueous

xv

microemulsion containing 0.05 M of SDS and 1 mM of HCl with 0.1 mM of LiClO4 as supporting electrolyte at a scan rate of 100 mV/s....................................................104

Figure 6-5: Molecular modeling result of EDOT-C1-biotin and EDOT-C12.............105

Figure 6-6: SEM (a) and AFM (b) image of poly(EDOT-COOH) film prepared from aqueous microemulsion..................................................................................................106

Figure 6-7: Viability of NIH3T3 (gray) and HepG2 (black) cells in the presence of different PEDOT film coated ITO glass substrate. ......................................................107

Figure 6-8: Adhesion of NIH3T3 (above) and KB (below) cells on PEDOT nanobiointerfaces of different monomer compositions: (a) EDOT, (b) EDOT-OH, (c) EDOT-COOH, and (d) EDOT-EG3-OH/EDOT-OH, molar ratio=9:1. ......................108

Figure 6-9: Attachment and proliferation of seeded NIH3T3 cells on poly(EDOT-OH) biointerface after (a) 2 h, (b) 15 h, and (c) 39 h of incubation in full medium...........108

Figure 6-10: (a) Legend of monomer composition of layered PEDOT nanobiointerfaces. (b)–(g) Controlled cell adhesion from alternating layer-by-layer PEDOT nanobiointerface deposition with adhesive and non-adhesive properties. ...109

Figure 6-11: Contact angles of layer-by-layer PEDOT nanobiointerfaces deposition. The color legends for the composition of PEDOT nanobiointerfaces are as shown in Figure 6-10......................................................................................................................110

Figure 6-12: Controlled cell adhesion on patterned poly(EDOT-OH) on poly(EDOT-EG3-OH)-co-poly(EDOT-OH) surfaces. (a) Top and side views of the device patterned by selective electropolymerization using PDMS mask. Magnified microscopic images of selective cell adhesion on the patterned surface were shown in (b) and (c)........................................................................................................................111

Figure 6-13: XPS analysis of poly(EDOT-OH) before (dashed line) and after biotinylation reaction (solid line). The inset shows the amplified region corresponding to N1s emission...............................................................................................................113

Figure 6-14: QCM studies of binding of BSA and streptavidin on functional PEDOT surfaces () non-biotin functionalized PEDOT () biotin-functionalized PEDOT. .115

Figure 6-15: Water contact angle of poly(EDOT-EG3-OH)-co-(EDOT-COOH) film before and after peptide conjugation. Monomer molar ratio of EDOT-EG3-OH: EDOT-COOH = 8:2 .......................................................................................................117

Figure 6-16: Controlled cell attachment (NIH3T3) on (a) RGD-functionalized, (b) carboxylic acid-functionalized, and (c) RDG-functionalized PEDOT surfaces. Functional group density was controlled at 10% while the remaining 90% are poly(EDOT-EG3-OH). ...................................................................................................118

Figure 6-17: Cell adhesion on RGD-modified PEDOT surfaces with different RGD density (a) 50%, (b) 20%, (c) 10%, (d) 5%, and (e) 1%. The effect of RGD density on cell adhesion was plotted on the right bottom. The y-axis is the number of attached cells per cm2....................................................................................................................119

xvi

LIST OF SCHEMES

Scheme 3-1: Synthetic scheme of (A) PVP-co-PAA and (B) PVI-co-PAA. ...............47

Scheme 3-2: Synthetic scheme of Ru complexes as electroactive pendant unit..........48

Scheme 3-3: Synthesis of redox polymer (A) Ru-complexed PVP-co-PVI and (B) Ru-complexed PVI-co-PAA. .................................................................................................49

Scheme 6-1: (a) Post-polymerization biotinylation of poly(EDOT-OH) films, (b) Post-polymerization biotinylation of poly(EDOT-COOH) films................................112

LIST OF TABLES

Table 2-1: Oligonucleotide sequences for DNA hybridization assay ..........................19

Table 2-2: Oligonucleotide sequences for tumour protein gene TP53 detections.......19

Table 2-3: Hairpin oligonucleotide sequences for intercalation study.........................19

Table 2-4: QCM data of CP coated quartz crystal after hybridization and intercalation...........................................................................................................................................37

Table 3-1: Oxidation potential of polymers complexed with different substituted ruthenium redox units.......................................................................................................53

1

1 Introduction

1.1 Background

Tremendous advances have been achieved in the area of biosensors over the

past three decades. Biosensors are compact analytical devices that employ the

biochemical molecular recognition event for the detection or identification of target

analytes. They have been widely applied in various areas including clinical

diagnostics, environmental monitoring, homeland security, food and pharmaceutical

analysis.1-8 All biosensors have the basic configuration that comprises an analyte

recognition layer and a signal conversion unit (transducer).

Figure 1-1: Schematic presentation of a biosensor

Nearly all types of biointeraction can be implemented into analyte recognition

schemes, from small biomolecules, nucleic acids, enzymes and antibodies to viruses,

whole cells and microorganisms. The measurable signal can be in the form of light

(optical), frequency (acoustic) or current (electrical), depending on the transducer

used. Biosensors can be classified either according to the target analyte or the signal

generated from the transducer. A good biosensor should be sensitive, specific, fast,

easy to use, reliable and cheap.

2

Advances in molecular biology have led to a better understanding of

DNA/proteins and their specific functions. The occurrence of various cancers and

diseases usually involves altered gene/protein expression. Potential biomarkers

associated with cancer or other diseases have been identified throughout many years

research. The accurate detection of these biomarkers would be useful for the early

diagnosis of specific diseases in clinical research.

Early diagnosis of cancer is crucial for the successful disease treatment.

However, cancer markers are generally presented at an ultra-low level during early

stages of the disease. Existing diagnostic tests (e.g. ELISA) are not sensitive enough

and only detect proteins at levels corresponding to advanced stages of the disease.

Therefore, highly sensitive detection techniques are urgently needed for effective

cancer treatment and increased survival rates. Moreover, smaller, faster and cheaper

biosensor devices are highly desired for decentralized clinical test such as emergency-

room screening, bedside monitoring and home self-testing.

1.1.1 Electrochemical Biosensors

Electrochemical biosensors are sensing devices that the biological recognition

element is intimately coupled to an electrode transducer. The transducer is able to

convert the biological recognition event into a useful electrical signal, either in the

form of potential (potentiometric), current (amperometric) or impedance

(impedimetric). Considering that electrochemical reactions directly generate an

electronic signal, biosensors based on this approach greatly simplified signal

transduction, avoiding expensive equipment requirement. Over the years,

electrochemical biosensors have been demonstrated as a simple, inexpensive and yet

accurate and sensitive platform for disease diagnosis. The flagship example of

3

commercial success amperometric biosensor is for blood glucose measurement. The

first generation glucose biosensor was demonstrated by Clark and Lyons in 1962.9 To

date, easy-to-use self-testing glucose strips, coupled to pocket-size amperometers,

have dominated the $5 billion/year diabetes monitoring market.10 The continuous

growing market for the need of home monitoring devices is the key to success. Beside

blood glucose, hand-held battery operated electrochemical clinical analyzers have

been shown extremely useful for rapid point-of-care measurement of multiple

electrolytes, metabolites11 as well as bedside blood gas monitoring.12

Despite the commercial success of electrochemical biosensors for blood sugar

monitoring, cancer-related assays are far more complex than home self-testing of

glucose. Tremendous efforts have been put into the development of biosensors for

DNA/protein detection over the past two decades. Modern electrochemical

DNA/immunosensors have recently demonstrated great potential for monitoring

cancer-related protein markers and DNA mutations.13

Electrochemical nucleic acid assays

Nucleic acid assays are often involved in clinical analysis for the detection of

specific nucleotide sequences, either for the identification of a particular

microorganism that is infectious, or DNA mutations that is associated with certain

genetic diseases. For sequence specific assays, single-stranded nucleic acid sequences

are immobilized on an electrode surface as the recognizing elements. In the presence

of the target analyte, complementary sequence in the case, the hybridization event is

detected electrochemically directly or indirectly. Nucleic acid hybridization is a

thermodynamic favored process, triggered by highly specific base-pairing interactions,

where each nucleotide base strongly binds to its complementary base through

hydrogen bonds. Vast amount of literature has been published in DNA hybridization

4

detection, and many comprehensive reviews on DNA-based biosensors are

available.14-22 Therefore, we will not review the literature again here, but only

highlight different transduction strategies used in electrochemical DNA biosensors.

The transduction strategies for DNA hybridization detection can be broadly

divided into two main categories, label-free and labeled approaches. Various labels

including redox active molecules,23 enzymes,24-28 and nanoparticles29 have been used

to tag target DNA sequence for hybridization event monitoring. In label-free approach,

cationic metal complexes22, 30-33 (e.g. Ru(NH3)63+, Fe(CN)6

3-, Co(Phen)33+, Co(bpy)3

2+)

or organic compounds34-37 (e.g. methylene blue, daunomycin, AQMS:

anthranquinone-2-sulfonic acid), have been reported for the use as hybridization

indicators, based on their preferential binding to either ss-DNA or ds-DNA. Other

label-free methods for the detection of DNA hybridization rely on changes to the

electrical properties of an interface,21 the change in flexibility from ss-DNA to the

rigid ds-DNA38-40 and the electrochemical oxidation of guanine bases.41, 42 General

electrochemical techniques such as cyclic voltammetry (CV), square wave

voltammetry (SWV), AC voltammetry, pulsed amperometry, and electrochemical

impedance spectroscopy (EIS) are employed to decode the hybridization event.

Despite enormous progress made in the development of electrochemical DNA

biosensors, key issues leading to the final commercialization are still around the

sensitivity, selectivity, and simplicity.

Electrochemical immunoassays and protein assays

Moving beyond DNA, electrochemical biosensors were also employed to

detect proteins. Abnormal expression of certain proteins can indicate the presence of

various cancers. Quantitative determination of these tumor markers plays an

important role in disease screening, diagnosis and treatment. Several authors gave

5

excellent reviews on the development of electrochemical immunoassays.43-46

Electrochemical immunosensors, combining the inherent specificity of

immunoreactions with the high sensitivity and convenience of electrochemical

transducers, are becoming an important analytical tool for the detection of antibody-

antigen interactions.

In electrochemical immunoassays, changes of potential, current, conductance,

capacitance or impedance caused by the immunoreactions can be directly detected

and correlated to the level of analyte. However, the binding of an antigen to their

specific antibody is accompanied by only small physical-chemical changes, and their

sensitivity is limited for clinical applications. Therefore, different labels such as

enzymes, nanoparticles and carbon nanotubes have been used for amplifying the

response from immunoreactions.

Enzymes are the most frequently used labels due to their inherent amplification.

Although homogeneous assays, which is based on the change of the activity of

enzyme labels before and after forming immunocomplex, do not require the

separation the free enzyme labels, heterogeneous assays, with more complicated

procedures, offers better limit of detection. The sensitivity of enzyme-based

immunoassays could be further enhanced when combined with other ways of

electrochemical signal amplifying. However, the inherent drawback of this approach

is the labor-intensive processes involving long incubation periods and multiple

incubation and washing steps.44

Gold nanoparticles have recently been used for ultrasensitive electrochemical

protein detection.47 A capture antibody was immobilized on the ferrocenyl-tethered

dendrimer modified indium tin oxide electrode. The detection antibody was labeled

with 10 nm gold nanoparticles. The gold nanoparticles catalyze the reduction of p-

6

nitrophenol to p-aminophenol (AP), the catalytically-generated AP was further

electrochemically oxidized to p-quinone imine (QI) by the electron mediation of

ferrocene on the ITO surface, and QI was then chemically reduced back to AP by

NaBH4 in solution. A detection limit of 1 fg/mL for mouse immunoglobulin (IgG) and

prostate-specific antigen (PSA) was achieved. Dequaire et al also demonstrated a

sensitive immunoassay for IgG using gold nanoparticles to label the antibody.48 The

nanogold label was measured by stripping volammetry after dissolution with acid.

The large number of gold ions released from each gold nanoparticle contribute to a

substantial improvement in sensitivity, as low as 3 pM IgG was detected. Similarly,

wang’s group used different quantum dots (ZnS, PbS, Cds, CuS) to label antibodies

for each specific protein. Multiple proteins were measured simultaneously based on

stripping amperometric signal of different metal ions released from those inorganic

nanocrystals.49, 50

Beside nanoparticles, carbon nanotubes (CNTs) were also used to amplify

detection signal in electrochemical protein assays. CNTs served as a carrier for

enzyme molecules. Using alkaline phosphate (ALP)-loaded CNTs to label detection

antibody, as low as 500 fg/mL of IgG was detected in a sandwich assay.51 Similarly,

sensitive detection of PSA was demonstrated using CNTs modified with horse radish

peroxidase (HRP) labeled secondary antibody. Due to the large surface area of CNTs,

hundreds of HRP labels per binding event were achieved, and as low as 4 pg/mL PSA

was detected in 10 uL of undiluted calf serum.52

The tremendous progress in nanotechnology offers excellent prospects for

developing highly sensitive protein biosensors. The use of nanomaterials in

elelctrochemical protein assays for signal enhancement lies in two aspects. One relies

on the unique material properties of nanomaterials for sensitive signal transduction.

7

The other is based on the use of nanomaterials as carriers for the amplification of

binding events. A drawback of using nanomaterials as labels in electrochemical

protein assays is that the preparation of the labels is not very reproducible. In addition,

fouling of the electrode surface can lead to poor reproducibility.

1.1.2 Label-Free Electrochemical/Electrical Assays

As discussed in earlier sections, most of the current detection technologies

require the labeling of target analytes for signal generation or amplification. The main

disadvantage of the label-based bioassays is the long procedures involving multi-steps

of incubation and washing. Labeling process usually involves complex chemical

reaction with the biological target, which is time consuming and costly. Furthermore,

the target biomolecules may lose its biological function after labeling due to

degradation. This becomes more particular in the case of immunoassay or other

protein assays. To bypass these drawbacks, label-free bioaffinity sensors are

intensively investigated. Label-free approach is becoming a more favored choice due

to its simple and rapid analysis.

Electrochemical detection

Label-free detection of DNA hybridization can be monitored using

electrochemical techniques, relying on either the changes of electrical or physical

properties on the interface. Hybridization indicators, based on their preferential

binding to either ss-DNA or ds-DNA, were commonly used as signal reporters. For

protein sensing, various recognition strategies based on biomolecules interactions,

such as antibody/antigen, aptamer/protein and carbohydrate/protein have been

exploited.53, 54 Interactions between the immobilized antibody and the target antigen

has been directly monitored using a variety of electrochemical techniques such as

8

electrochemical impedance spectroscopy (EIS) 55-57, capacitance measurement58,

amperometry59, 60, and square wave voltammetry (SWV).61 Examples of proteins

being detected include IgG, bovine serum albumin (BSA), human serum albumin

(HSA), and hepatitis B surface antigen. Limit of detection usually in the range of

ng/mL. The label-free approach avoids the use of competitive antigens labeled with a

fluorophore or with an enzyme, such as HRP or glucose oxidase (GOX), whose

enzymatic products are electroactive. The removal of labeling step reduces the risk of

contamination and accelerates the analytical process.

Electrical detection

Nanowires and nanotubes based field effect transistor (FET) devices have

been used for the direct detection of small molecules, DNA, proteins and viruses.62-65

Binding of charged molecules on the nanowire surface caused a change in

conductance due to the field effect. For example, negative charges caused an increase

in conductance, and positive charges caused a decrease of conductance of p-doped

SiNWs. When the charged proteins were specifically captured on the antibody

modified nanowires, the corresponding conductance change of the nanowire was a

measure of the target protein (Figure 1-2). Si nanowire (SiNW) based FET sensor has

shown DNA/protein determination at the low picamolar to femtomolar level.66-68

Semiconductive carbon nanotubes (CNTs) has also been demonstrated for protein

sensing with nanomolar sensitivity.69 Direct real-time detection of glucose was

achieved using glucose oxidase (GOx) modified CNT.70

9

Figure 1-2: Schematic of nanowire based FET sensor (adapted from Ref 62)

Beside SiNWs and CNTs, conducting polymer nanowires (CPNWs) are

emerging as a promising candidate for nanowired based biosensing. Tao et al reported

glucose detection using a polyaniline (PANi) nanojunction sensor.71 Ramanathan

recently demonstrated label-free detection of biotin-DNA using avidin-functionalized

polypyrrole nanowires at 1 nM.72 The one-step incorporation of functional biological

molecules into the CPNWs during its synthesis within built-in electrical contacts is

the major advantages over those SiNWs and CNTs biosensors that require post-

synthesis functionalization, alignment and positioning. Compared to SiNWs and

CNTs, the application of CPNWs for DNA/protein biosensing is still in an early stage

of development.73, 74 Several issues including their chemical/thermal/mechanical

stability need to be addressed before they can be utilized to their full potential.

The electrical detection based on nanowire FET devices provides real-time

label-free measurement, with ease of integration in addressable arrays for

multiplexing. A large nanowire arrays could be fabricated on one chip, with hundreds

of electrically and individually addressable sensing units. Even though there has been

tremendous advancement in nanowire biosensors, there are still difficulties associated

with the fabrication and assembly for practical use. A possible limitation of nanowire

biosensors is the relatively high cost of the equipment and preparation. However, in

10

spite of the numerous challenges, nanowire biosensors offer unlimited research

opportunities.

1.1.3 Electroactive Conducting Polymers for Biosensing

π-conjugated polymers (conducting polymers) have emerged as potential

candidates for electrochemical sensors. The unique property of conducting polymers,

along with their compatibility with biological molecules in aqueous solution, has been

exploited for the fabrication of accurate, fast, and inexpensive biosensor devices. It is

believed that conducting polymers improve the sensitivity and selectivity of

electrochemical biosensors due to their electrical conductivity or charge transport

properties.

Conducting polymers have demonstrated several advantages for bio-receptor

capturing. Biomolecules, such as enzyme, antibody, DNA, aptamer etc. can be

immobilized onto conducting polymers without loss of activity. Ahuja reviewed the

biomolecular immobilization on the conducting polymers for biosensor applications,

and compared different mode of immobilization techniques such as physical

adsorption, covalent conjugation, and electrochemical immobilization.75 Conducting

polymers provide good matrix support to the biological-active molecules either by

electrostatic, covalent or non-specific interactions. Earlier studies on conducting

polymer-based biosensors mainly use conducting polymer as an immobilization

matrix, and electrochemical signals are generated from either enzymatic reactions or

other electroactive labels.76

Conducting polymers are also intrinsic electronic transducers. The

perturbations in polymer chain conformation and/or electronic structure from the

presence of the probe/target conjugates lead to a change in macroscopic material

11

properties, which can be measured by cyclic voltammetry (CV) and electrochemical

impedance spectroscopy (EIS). Numerous papers have demonstrated the possibility of

using conducting polymers as both immobilization matrix and intrinsic electronic

transducer for the label-free detection of biological molecules.77-82 Moreover,

conducting polymers can be electrochemically grown on very small sized electrode

precisely, which allows for in vivo monitoring of biomolecules.83

In summary, electrochemical/electrical biosensors promise low-cost, rapid and

simple-to-operate analytical tools and represent a broad area of emerging technologies

ideally suited for point-of-care analysis. The high sensitivity, specificity, simplicity

and miniaturization of modern electrochemical biosensors permit them to rival the

most advanced optical ones. With the development of new materials and novel

detection schemes, label-free electrochemical biosensors would find more practical

application in various fields including clinical diagnostics, environmental monitoring,

drug screening, and homeland security etc.

1.2 Motivation and Objectives

The urgent need for the development of point-of-care devices for the detection

of infectious agents and cancer-related biomarkers motivate us to keep searching for

simple, fast, sensitive yet affordable analytical tools. This project aims to design and

develop label-free electrochemical or electrical nucleic acid/protein biosensors, with

the focus on simple yet sensitive detection schemes, toward point-of-care disease

diagnosis for clinical use and future in vivo testing.

12

1.3 Scope

This thesis focuses on the study of label-free electrochemical/electrical

biosensors for sensitive detection of DNA and proteins. Design and development of

new detection schemes are the main focus. Polymer chemistry and physics, sample

preparation, microfluidics and miniaturization, multiplexing, and the incorporation of

the detection platform into a working diagnostic device are beyond the scope of this

thesis.

1.4 Thesis Outline

This thesis presents our studies on the development of label-free

electrochemical biosensor for DNA/protein detection. Chapter one provides an

overview of the background and motivation of the project. In this chapter, we review

the state-of-art detection technology, especially through electrochemical/electrical

transduction. Special attention is given to label-free electrochemical/electrical

approaches, which is also the main interest of our study. Chapter two and three

discuss ruthenium-based redox active compounds and their application for label-free

DNA sensing. Chapter two presents two ruthenium-based electroactive intercalators

for label-free DNA detection, with one based on electrochemiluminescence, and the

other based on electrocatalytic oxidation of guanine in the DNA sequences. In chapter

three, we study the catalytic guanine oxidation on ruthenium-containing polymer

complex modified electrodes, and report the determination of the apparent oxidation

potential of guanine and its family compounds under physiological buffer condition

first time by electrocatalytic oxidation titration. In chapter four and five, we present

our study of electroactive conducting polymers, functional poly(3,4-

ethylenedioxythiophene)s (PEDOTs) and their use as transducers for label-free

13

protein sensing. Chapter four discusses and compares various approaches for the

synthesis of one dimensional functional PEDOT nanostructures. Chapter five

highlights the application of PEDOT nanostructures for label-free protein biosensing.

In chapter six, we evaluate functional PEDOTs thin films as tunable nanobiointerfaces

for specific protein adsorption and controlled cell attachment, for future in vivo

applications. Finally we conclude the thesis with future direction in chapter seven.

14

2 Ruthenium-Complexed Electroactive

Intercalators for Label-Free DNA Detection

2.1 Introduction

Nucleic acid-based biosensors have a wide variety of potential applications

that range from genotyping to molecular diagnostics.41, 84, 85 The use of

electrochemical techniques instead of fluorescence allows for simpler and smaller

detectors.86, 87 The simplest of such systems would be through direct electrochemistry

of nucleic acids: a solid electrode modified with an oligonucleotide probe produces a

measurable electrochemical signal upon hybridization to a specific target gene.

However, it is generally believed that direct redox reaction of nucleic acids is

irreversible and often suffers from a pronounced fouling effect, resulting in rather

poor selectivity and reproducibility.88 Moreover, direct oxidation of water takes place

at potentials close to that of nucleic acid oxidation and significantly lifts the

background signal. The ability to directly detect nucleic acid selectively and

sensitively has been a major goal of electrochemical research.

A number of approaches have been proposed for direct electrochemical

detection of nucleic acid.42, 89-92 Substantial improvements were achieved using

baseline-corrected adsorptive stripping square-wave voltammetry. As little as 15.4

fmol of nucleic acid was detected on a carbon paste electrode.91 The poor electron

transfer kinetics of nucleic acid was also addressed using electrocatalysts. Thorp’s

group first reported the detection of attomole quantity of immobilized DNA using a

transition redox active metal complex, Ru(bpy)32+, as a homogenous catalyst.

However, since the compound is unable to interact selectively with ds-DNA, the assay

suffered from high background signal and lack of sensitivity.42 The analytical signal is

15

superimposed onto an intrinsically large background current due to the direct

oxidation of the catalyst itself and the catalytic oxidation of oligonucleotide capture

probes. Improvement was made by replacing guanine in CP with its electrochemical

inactive analogues, which eliminates most of the catalytic oxidation current from CP,

but little can be done to minimize the direct oxidation of the catalyst.89 Earlier work

from our laboratory showed that low redox potential electrocatalysts are beneficial in

enhancing the sensitivity owing to a minimized background current.26, 27

The use of electroactive DNA binding compounds as hybridization indicators

negates the need for labelling the target DNA, as commonly required in conventional

DNA detection techniques. Milan and Mikkelsen first proposed the idea of using a

electroactive indicator, tris(1,l0-phenanthroline)cobalt(III) perchlorate or Co(phen)33+,

to signify hybridization.30 Upon hybridization of the target, the modified electrode

was immersed in a solution tris(1,l0-phenanthroline) cobalt(III) perchlorate

(Co(phen)33+) to allow binding. The voltammetric analysis was subsequently carried

out in the same solution. The concentration of target DNA was correlated to the

characteristic redox signal of the cobalt complex. Since then, nucleic acid biosensors

based on voltammetric detection of electroactive organic35, 36, 93, 94 or inorganic31, 34, 95-

98 indicators interacting preferentially with double stranded DNA (ds-DNA) have

been reported. More details can be found in Erdem’s review paper.20 The organic

compounds used as reporters in electrochemical DNA detection bind to ds-DNA

either through groove binding or intercalation while the inorganic compounds are

mainly binds through electrostatic interaction. The background signal arising from the

nonspecific binding of these compounds to single stranded DNA (ss-DNA) was a big

problem. New intercalators, offering better discrimination between ss-DNA and ds-

DNA are being developed for better signal/noise ratio.

16

In recent years, metallointercalators, metal complexes that bind through

intercalation to ds-DNA, have gained attention due to their more selective binding. In

addition, the catalytic nature of metallointercalators makes them ideal candidates as

electrochemical reporters for label-free DNA detection. Takenaka and coworkers

reported an electrochemical detection scheme using a redox reporter that comprises a

ferrocene-labeled naphthalene diimide intercalating unit.99 The high binding constant

of the intercalating unit allows the reporter to form a more stable complex with ds-

DNA, while the electrocatalytic nature of ferrocene enabled signal amplification for

sensitive detection.

Transition metal complexes have been extensively studied for their

electrochemistry and photochemistry.100-103 Electrochemiluminescence (ECL) have

been demonstrated as one of the most sensitive techniques and therefore been

proposed for the ultrasensitive detection of DNA hybridization events.104-107 ECL is

the process of generating excited states in a photoactive molecule at an electrode

surface, leading to luminescence upon return to the ground state. The key to its

ultrahigh sensitivity lies in the ultralow background noise, which is a direct

consequence of having two different forms of energy for analytical signal generation

and detection. Unlike fluorescence-based techniques, ECL does not involve an

excitation light source and it theoretically produces a “zero” background. Ru(bpy)32+

has been extensively studied for its ECL, which was first reported by Bard some thirty

years ago.108 Because of its low-lying metal-to-ligand charge-transfer (MLCT) excited

states,109 high emission quantum yields (~4.2% in H2O)110 and long excited-state

lifetimes (~600 ns), the well known Ru(bpy)32+/tri-n-propylamine (TPA) system is

usually adopted in analytical applications. As demonstrated by Bard et al., as little as

1.0 fM of DNA is detected when a Ru(bpy)3 2+ doped polystyrene microbead (Ru-

17

PMB) is used as an ECL tag.104 Recently, Rusling and co-workers have reported that

ECL signals can be generated in a DNA-[Ru(bpy)2PVP] bilayer.111, 112

The marriage of a highly selective intercalator and electrocatalysis or ECL

provides novel platforms for ultrasensitive label-free detection of DNA. A promising

approach toward the enhancement of the amperometric or ECL signal is to build up

multiple electroactive tags on a single ds-DNA chain. This strategy has the advantage

of providing multiple redox sites, greatly increasing the number of charge

recombination events per target DNA molecule, and thereby enhancing the intensity

of analytical signal and lowering the detection limit. A much better selectivity and

higher stability are expected with properly designed electroactive intercalators.

Our group has been interested in using electroactive threading intercalators to

tag DNA and develop ultrasensitive DNA detection systems. In a previous report, we

described the synthesis and analytical application of an osmium-complexed

electroactive threading intercalator, which allows the detection of 50-mer target DNA

in the range of 1 – 300 pM with a detection limit of 600 fM, based on the

amperometric signal from catalytic oxidation of ascorbic acid.113 In this chapter, we

report the synthesis, characterization and analytical application of two ruthenium-

complexed electroactive DNA intercalators as signal reporters for ultrasensitive DNA

detection. In the first example, the feasibility of using a novel electroactive mono-

intercalator, N,N′-bis[(3-propyl)imidazole]-1,4,5,8-naphthalene diimide (PIND)

imidazole complexed with Ru(bpy)2Cl (Ru-PIND-Ru, bpy=2,2′-bipyridine) as a low

redox potential electrocalalytic reporter for sensitive label-free electrochemical

detection of nucleic acid was studied. A remarkable improvement in the voltammetric

response of nucleic acids were observed due to the combined catalytic function of the

imidazole-complexed [Ru(bpy)2Cl] redox moieties toward the guanine base and the

18

high selectivity of Ru-PIND-Ru towards ds-DNA. In another example, a bis-

intercalator PIND-Ru-PIND, where the electroactive unit Ru(dmbpy)2 (dmbpy=4,4'-

dimethyl-2,2'-bipyridine) was sandwiched between two intercalating units through

coordinative bonds with the two imidazole groups at the termini of PIND, was

explored as an electroactive ECL reporter for sensitive label-free DNA detection. The

intercalated PIND-Ru-PIND exhibited reversible electron-transfer and strong ECL in

the presence of TPA. A 2000-fold sensitivity enhancement over direct voltammetry

was obtained. The proposed ECL procedure demonstrates several advantages in terms

of sensitivity, selectivity and simplicity.

2.2 Experimental

2.2.1 Materials and Reagents

1-(3-aminopropyl)-imidazole (AI, 98%,) and 1,4,5,8-naphthalene

tetracarboxylic dianhydride (NTD, >95%), 4,4'-dimethyl-2,2'-dipyridyl (dmbpy,

99.5%) and ruthenium trichloride were purchased from Sigma-Aldrich (St Louis, MO,

USA). Ru(bpy)2Cl2 (99%) was from Avocado Research Chemicals Ltd (Leysham,

Lancester, UK). All chemicals were of reagent grade and used as received. Capture

probes used in this work were custom-made by Alpha-DNA (Montreal, Canada) and

all other oligonucleotides were custom-made by 1st Base Pte Ltd (Singapore).

Oligonucleotide sequences used in the work were listed in the tables below.

A 10 mM Tris-HCl/1 mM EDTA/0.1 M NaCl buffer solution (TE) was used

as hybridization buffer. A phosphate-buffer saline (PBS, pH 7.4), consisted of 0.15 M

NaCl and 20 mM phosphate buffer, was used as supporting electrolyte. To minimize

the effect of RNases on the stability of mRNA, all solutions were treated with diethyl

19

pyrocarbonate and surfaces were decontaminated with RNaseZap (Ambion, TX) for

RNA related work.

Table 2-1: Oligonucleotide sequences for DNA hybridization assay

Table 2-2: Oligonucleotide sequences for tumour protein gene TP53 detections

Table 2-3: Hairpin oligonucleotide sequences for intercalation study

2.2.2 Synthesis of Electroactive DNA Intercalators

Intercalating Unit PIND

N,N'-bis[1-(3-propyl)-imidazole]-1,4,5,8-naphthalene diimide (PIND) was

prepared following a general procedure for the synthesis of diimide.114, 115 Briefly, 0.3

20

g of 1,4,5,8-naphthalene tetracarboxylic dianhydride (NTD) was slowly added into a

magnetically stirred mixture of 3.0 mL of 1-(3-aminopropyl)-imidazole and 3.0 mL of

tetrahydrofuran (THF). The rate of addition was controlled to minimize clogging. The

reaction mixture was left refluxing for overnight and then cooled to room temperature.

Next, it was dispersed in 10 mL of acetone/water (3:1) mixture and poured into 500

mL of rapidly stirred anhydrous ether to precipitate the compound. The precipitate

was collected by suction filtration through a fine fritted funnel and washed briefly

with ethanol. The product was purified by running it through a silica gel column using

ethanol:chloroform (1:1) as the eluent and dried under vacuum at 40 ºC overnight to

give 0.46 g of yellow crystals (yield 85%). 1H NMR (300 MHz CDCl3) δ 8.76 (4H),

7.54 (2H), 7.26 (2H), 4.27 (4H), 4.12(4H), 2.31 (4H) and 1.83 10(2H). [PIND+H+] =

483.3 and [M+2H+]/2 = 242.3. HR-MS (FAB): calcd. for C6H22N6O4+H+ 483.1781

[M+H+]; found 483.1770.

Redox Active Pedant Ru(dmbpy)2Cl2

(R=−CH3)

Cis-bis(4,4'–dimethyl-2,2'-bipyridine)dichlororuthenium (Ru(dmbpy)2Cl2) was

synthesized following a literature reported procedure.116 A mixture of Ru(III)

trichloride hydrate (1 g, 3.8 mmol, 20% excess), 4,4'-dimethyl-2,2'-bipyridine (1.2 g,

6.5 mmol), and lithium chloride (1.1 g, 26 mmol) in 60 mL of DMF was stirred under

reflux for 8 hours. The solvent was removed by vigorous stirring in diethyl ether. The

21

crude product was then dissolved in chloroform and washed with water. Upon drying,

the product was obtained in the form of black powder (1.72 g, 70% yield).

Ruthenium Complexed DNA Intercalators

(a) Mono-intercalator Ru-PIND-Ru

Ru-PIND-Ru was synthesized in a single-step ligand-exchange reaction.

PIND (0.12 g, 0.25 mmol) was slowly added to a solution of Ru(bpy)2Cl2 (0.34 g,

0.55 mmol, 10% excess) in 8.0 mL of fresh-distilled ethylene glycol in small portion

over 10 min and the mixture was left refluxing for 30−40 minutes. The completion

of the ligand-exchange reaction, indicated by the disappearance of redox peak of

starting materials and formation of those of the products, was monitored by cyclic

voltammetry. The purple reaction mixture was then poured slowly into 100 mL of

rapid stirred ethanol saturated with KCl. The precipitate was collected by suction

filtration through a fine fritted funnel. The crude product was washed with PBS,

dissolved in 3.0 - 5.0 mL of ethanol and precipitated again from KCl saturated

ethanol. The precipitate was further purified by crystallization from ethanol giving

the pure product in 78% yield. The product showed a single pair of reversible redox

peaks at the gold electrode with an E1/2 of 0.63 V in PBS. To ensure a complete

double ligand-exchange at the two imidazole termini of PIND, slight excess of

Ru(bpy)2 (10–25%) is required. HR-MS: calcd. for C66H54N14O4Ru2Cl22+ 690.0953

[M2+]; found 690.0976

(b) Bis-intercalator PIND-Ru-PIND

22

(R=4,4'-dimethyl-2,2'-bipyridine)

PIND-Ru-PIND was synthesized similarly with a slight excess of PIND

instead of Ru complex in previous case. To a solution of Ru(dmbpy)2Cl2 (0.20 mmol)

in 8.0 mL fresh-distilled ethylene glycol was added 0.50 mmol PIND and the

resulting mixture was stirred for 10 min before refluxing. The course of the ligand-

exchange reaction was followed by cyclic voltammetry. The orange reaction mixture

was then poured slowly into 500 mL of rapidly stirred anhydrous ether. The

precipitate was collected by suction filtration through a fine fritted funnel. The crude

product was dissolved in 8−10 mL of water and was extracted twice with chloroform.

The precipitate was further purified by crystallization from ethanol giving the pure

product in 80% yield. A slight excess of PIND (20−25%) is required to ensure a

complete double ligand-exchange.

2.2.3 Apparatus

Electrochemical experiments were carried out using a CH Instruments model

660A electrochemical workstation coupled with a low current module (CH

Instruments, Austin, TX). A conventional three-electrode system, consisting of a 3.0-

mm-diameter gold working electrode, a non-leak Ag/AgCl (3.0 M NaCl) reference

electrode (Cypress Systems, Lawrence, KS), and a platinum wire counter electrode,

was used in all electrochemical measurements. To avoid the spreading of the sample

droplet beyond the 3.0-mm diameter working area, a patterned hydrophobic film was

applied to the gold electrode after the immobilization of the CP. All potentials

reported in this work were referred to the Ag/AgCl electrode. UV-visible spectra were

23

recorded on a V-570 UV/VIS/NIR spectrophotometer (JASCO Corp., Japan). NMR

spectroscopic study was done on a Bruker 400 MHz system from Bruker Biospin

GmbH (Karlsruhe, Germany). Mass spectrometric experiments were performed with a

Finnigan/MAT LCQ Mass Spectrometer (ThermoFinnigan, San Jose, CA).

Measurements of ECL were performed with a Fluorolog-3 spectrofluorometer

(Jobin Yvon Inc, Edison, NJ) in conjunction with a 660A electrochemical workstation.

The three-electrode system consisted of a gold working electrode, a non-leak

Ag/AgCl reference electrode, and a platinum foil counter electrode. The three

electrodes were hosted in a standard 1.0-cm fluorescence cuvette and arranged in such

a way that the working electrode faces the emission window and the other two

electrodes are behind the working electrode. All potentials reported in this work were

referred to the Ag/AgCl electrode. All experiments were carried out at room

temperature, unless otherwise stated.

2.2.4 Sensor Construction

Immobilization of capture probe

The preparation and pre-treatment of gold electrodes follow standard

procedures.113 Briefly, prior to capture probe adsorption, a gold electrode was exposed

to oxygen plasma for 5-10 min and then immediately immersed in absolute ethanol

for 20 min to reduce the oxide layer. A CP monolayer was adsorbed by immersing the

gold electrode in a 20 mM phosphate buffer (pH 7.4) solution of 100 μg/mL CP for

16−24 h. After adsorption, the electrode was copiously rinsed with and soaked in the

phosphate buffer for 20 min, rinsed again, and blown dry with a stream of air. The

surface density of CP, assessed electrochemically by the use of cationic redox probe

according to the procedure proposed by Steel,117 was found to be in the range of

1.15−1.35 x 10-11 mol/cm2. To minimize non-DNA related reporter molecules uptake

24

and improve the quality and stability of the CP monolayer, the CP-coated gold

electrode was further immersed in an ethanolic solution of 2.0 mg/mL 1-

mercaptododecane (MD) for 4−6 h. Loosely absorbed MD molecules were rinsed off

and the electrode was washed by immersion in a stirred ethanol for 10 min followed

by thorough rinsing with ethanol and water. The electrode was ready after air-dry.

Hybridization with target DNA

The hybridization of a target DNA was carried out in droplet form. First, the

CP coated electrode was placed in a moisture saturated environmental chamber

maintained at 60 ºC. A 5.0 μL droplet of hybridization solution containing the target

DNA was uniformly spread onto the electrode (low stringency, 27 ºC below the salt-

adjusted melting temperature). After 1 h of hybridization at 60 ºC, the electrode was

rinsed thoroughly with a blank hybridization solution.

Binding of electroactive reporter

After washing away non-specifically bound target DNA, the electrode was

further incubated with a 5.0 μL droplet of 100 μg/mL of electroactive reporter (Ru-

PIND-Ru or PIND-Ru-PIND) for 10 min. Reporter molecules were attached to the

hybridized DNA duplex via threading intercalation. It was then thoroughly rinsed

with NaCl-saturated 0.1 M phosphate buffer (pH 7.0) containing 10% ethanol to

remove those non-intercalation related reporter molecules, e.g. bound through

electrostatic interaction or hydrophobic interaction. The amount of reporter molecules

remaining on the electrode surface after washing should thus be proportional to that of

duplex DNA and represents the amount of complementary target.

25

2.3 Results and Discussion

2.3.1 Synthesis and Characterization of Electroactive DNA Intercalators

2.3.1.1 Cyclic Voltammetry

The redox potential of ruthenium complexes changes upon ligand exchange,118

therefore, the formation of the redox active ruthenium complexed mono-intercalator

and bis-intercalator can be conveniently monitored by cyclic voltammetry. During

reflux in ethylene glycol, cyclic voltammetric tests were conducted every 5 min.

Formation of Ru-PIND-Ru

The formation of the electroacticve Ru-PIND-Ru intercalator was monitored

by cyclic voltammetry. Figure 2-1 shows two typical voltammograms obtained in the

first 30 min. As shown in trace a, before adding PIND to Ru(bpy)2Cl2, one pair of

reversible voltammetric peaks centered at 0.40 V were obtained, corresponding to the

well-known redox process of Ru(bpy)2Cl2. Upon adding PIND, a new pair of

voltammetric peaks appeared at 0.63 V, indicating the formation of Ru-PIND-Ru

(Figure 2-1 trace b). Both electron transfer processes are clearly resolved and exhibit

all the characteristics of reversible processes, except for the slightly larger peak-to-

peak potential separation that is mainly due to a higher iR drop of the reaction

medium. The intensities of the voltammetric peaks at 0.63 V increased gradually with

reaction time, while those at 0.40 V diminished gradually. Both of the redox pairs

reached a steady-state after 30-40 min of refluxing. The minute voltammetric peaks at

0.40 V are indicative of the excess amount of Ru(bpy)2Cl2. After separation and

purification, voltammetric tests of the purified Ru-PIND-Ru showed only one pair of

voltammetric peaks implying that the purification process is very effective (Figure 2-1

trace c).

26

Figure 2-1: Cyclic voltammograms of the starting materials Ru(bpy)2Cl2 (····), reaction mixture after refluxing with PIND for 30 min in ethylene glycol (---), and the purified final product (―). Supporting electrolyte: PBS; Scan rate: 100 mV/s

Figure 2-2: Cyclic voltammograms of the purified Ru-PIND-Ru in PBS at scan rate of (a) 100, (b) 200, (c) 300, (d) 400, and (e) 500 mV/s.

Purified Ru-PIND-Ru exhibits electrochemical characteristics exactly as

expected for a highly reversible redox couple in solution. Little change was observed

after numerous repetitive potential cycling between 0.0 and +0.90 V, revealing good

stability of Ru-PIND-Ru in solution. Figure 2-2 shows the sweep rate dependency of

the voltammograms of Ru-PIND-Ru. At slow scan rates of <500 mV/s, a typical

diffusion controlled voltammogram was recorded as expected for a one-electron

27

exchange system exhibiting an ideal Nernstian behaviour: the peak current is

proportional to the square root of the potential scan rate; the peak-to-peak potential

separation is very close to the theoretical value of 59 mV and independent of potential

scan rate. Such results ascertain that all the ruthenium redox centers are involved in

reversible heterogeneous electron transfer.

Formation of PIND-Ru-PIND

Figure 2-3: Cyclic voltammograms of Ru(dmbpy)2Cl2 after refluxing with PIND for (a) 0, (b) 10, and (c) 30 min. Supporting electrolyte: PBS; Scan rate: 100 mV/s.

Similarly, the formation of PIND-Ru-PIND bis-intercalator was monitored by

cyclic voltammometry. Figure 2-3 shows the typical voltammograms obtained in the

first 30 min. Before adding PIND to Ru(dmpy)2Cl2, one pair of reversible

voltammetric peaks centered at 0.29 V were obtained, corresponding to the well-

known redox process of Ru(dmbpy)2Cl2 (Figure 2-3 trace a). After only 10 min of

refluxing, the voltammetric peaks of Ru(dmbpy)2Cl2 disappeared completely and two

new pair of voltammetric peaks appeared at 0.49 and 0.68 V, indicating the formation

of PIND-Ru and PIND-Ru-PIND, respectively (Figure 2-3 trace b). The intensities of

the voltammetric peaks at 0.68 V increased gradually with reaction time.

Simultaneously, those at 0.49 V diminished gradually. Voltammetric tests of the

28

reaction mixture after 30 min refluxing showed only one pair of voltammetric peaks

(Figure 2-3 trace c), indicating the completion of the double ligand exchange process.

2.3.1.2 UV-Vis Spectrophotometry

UV-Vis spectrum of electroactive ruthenium-based intercalators were acquired

and compared with those of the starting materials. As shown in Figure 2-4, adsorption

spectra of PIND-Ru-PIND, PIND, Ru(dmbpy)2Cl2 and a model compound

Ru(dmbpy)2(Im)2 were overlaid together. UV-Vis spectrum of PIND-Ru-PIND

(Figure 2-4 trace a) is similar to that of Ru(dmbpy)3−naphthalene diimide

compound.118-120 It exhibits intense absorption band in the UV region due to

intraligand (IL) π→π* (dmbpy) transitions and followed by a broad absorption band

in the visible region (400−600 nm) due to spin allowed Ru (dπ) → dmbpy (π*) MLCT

transition. The absorption peaks at 380 and 361 nm are mainly due to π→π* transition

in PIND with some contribution from underlying MLCT absorbance. The absorption

maximum of PIND-Ru-PIND is red-shifted from 415 to 495 nm with respect to

Ru(dmbpy)2Cl2 (Figure 2-4 trace c). The same changes were also observed in the

spectrum of the model compound Ru(dmbpy)2(Im)2 (Figure 2-4 trace b). This is likely

a direct consequence of the ligand exchange which results in two types of MLCT

transitions within the ruthenium complex: Ru (dπ) → dmbpy (π*), and Ru (dπ) →

imidazole (π*). The imidazole groups of PIND are conjugated, resulting in a lower π*

level for this ligand relative to the chloride of the complex. Moreover, the spectrum of

PIND-Ru-PIND is a composite of the absorption spectra from both the PIND moiety

(Figure 2-4 trace d) and the Ru(dmbpy)2(Im)2 complex. A simple overlay of

Ru(dmbpy)2(Im)2 and PIND generated a spectrum which is almost identical to that of

PIND-Ru-PIND, confirming the formation of PIND-Ru-PIND.

29

Figure 2-4: UV-Vis absorption spectra of (a) PIND-Ru-PIND, (b) Ru(dmbpy)2(Im)2, (c) Ru(dmbpy)2Cl2, and (d) PIND in ethanol.

2.3.1.3 Mass Spectrometry

Mass spectrometric analysis of PIND-Ru-PIND was carried out using

electrospray ionization. Predominant peaks were found at m/z 717, 483, 478, and 242,

corresponding to (PIND-Ru-PIND)2+/2, (PIND+H+), (PIND-Ru-PIND+H+)3+/3, and

(PIND+2H+)/2 respectively, which are in good agreement with the mass of the desired

compounds. No mono-grafted PIND-Ru(dmbpy)2Cl was observed in the ESI−MS

spectrum, ruling out any incomplete grafting of Ru(dmbpy)2.

2.3.2 Intercalation with DNA

UV-Vis spectrophotometry was used to study the binding of Ru-complexed

PIND with ds-DNA. As an example, Figure 2-5 is the overlay of UV-Vis spectra of

Ru-PIND-Ru at increasing amount of salmon sperm DNA. As shown in the figure,

addition of DNA to Ru-PIND-Ru at a DNA base pair/Ru-PIND-Ru ratio of 4.0

resulted in ~40% decrease and a 2 nm red-shift of the naphthalene diimide (ND)

absorbance band at 366 and 387 nm. In the UV-Vis spectrophotometry,

hypochromism (decrease in adsorption intensity) and bathochromism (red shifts) are

30

the signatures of intercalative binding, where the fused planar aromatic ring system of

a threading intercalator inserts itself between the base pairs of ds-DNA.121, 122 Similar

phenomena were previously observed with ND having aliphatic tertiary amine side

chains.119, 120 The ND absorbance band hypochromism reached a plateau at the DNA

base pair/Ru-PIND-Ru ratio > 4.0, indicating that binding of Ru-PIND-Ru to ds-DNA

takes place by preferential intercalation of the ND.

Figure 2-5: UV-Vis spectra of 25 µM Ru-PIND-Ru (resolution 0.10 nm) as a function of increasing concentration of salmon sperm DNA (in base pair) of (a) 0, (b) 25, (c) 50 and (d) 100 µM. Insert: Enlarged UV-Vis adsorption spectra of the intercalative binding region.

To have a better estimation of the intercalating property, a competition

experiment, similar to that proposed by Boger,123 was designed using AT-rich short

hairpin oligonucleotides to establish the binding constant. It has been demonstrated

that these hairpin oligonucleotides form a 1:1 complex with threading intercalators.

The basis of this methodology involves the use of two intercalators, one fluorescent

and the other non-fluorescent. The fluorescent intercalator first saturates the ds-DNA.

Then a second intercalator, here ruthenium complexed PIND, is introduced into the

system with gradual increase in concentration. Under the assumption that the two

31

intercalator would bind to similar sites in the ds-DNA, introduction of 2nd non-

fluorescent intercalator will displace the 1st fluorescent intercalator, and cause a

decrease in fluorescence intensity. A well-known threading intercalator, ethidium

bromide (EB), was chosen as fluorescent indicator. EB has been widely studied as an

efficient DNA intercalator and is one of the most popular fluorescent intercalator used

in DNA assay. It possesses relatively little sequence preference and displays a 25-fold

fluorescence enhancement upon binding to ds-DNA, which provides sufficient

sensitivity and good discrimination against free EB molecules in fluorescence

measurement. In addition, the kinetics of EB intercalation is quite fast124, which

significantly shortens the time needed to reach equilibrium. A fluorescence titration

curve was first generated to determine the binding stoichiometries. As shown in

Figure 2-6A, the change in fluorescence was plotted against molar equivalents of Ru-

PIND-Ru, from which the binding stoichiometry of 1:1 was determined. The

intersection of the pre- and postsaturation portions of the curve provides ∆Fsat and

allows for the determination of [free intercalator] based on the following equation.123

[free intercalator] =

∆∆

−sat

xT F

FXDNA][

In which, [free intercalator] = concentration of free intercalator, [DNA]T = total

concentration of DNA, X = molar equivalents of intercalator vs DNA, ∆Fx = change in

fluorescence, and ∆Fsat = change in fluorescence at the point where DNA is saturated

with the intercalator.

For cases in which the binding stoichiometriy is 1:1, binding constant can be

established by Scatchard analysis of the fluorescence titration curve.125, 126 As shown

in Figure 2-6B, plotting of ∆F/[free intercalator] vs ∆F yields a linear portion of the

Scatchard plot and binding constant was estimated from the slope of the linear curve.

32

The stability constant was found to be 3 x 107 M-1, corresponding to a ~75-fold

enhancement over ND. A plausible explanation for the stability enhancement would

be that after the ND group has intercalated with ds-DNA, the two cationic

[Ru(bpy)2Cl]+ units in Ru-PIND-Ru form ion-pairs with phosphates on each side of

the ds-DNA, making ND more tightly fixed in between the base pairs of ds-DNA.

Figure 2-6: (A) Fluorescent displacement titration curve of Ru-PIND-Ru against a 5

μM hairpin oligonucleotide with EB. (B) Scatchard plot for the titration of hairpin

oligonucleotide/EB with Ru-PIND-Ru.

Similar results were obtained for bis-intercalator PIND-Ru-PIND. As shown

in Figure 2-7, the addition of DNA to PIND-Ru-PIND at a DNA base pair/PIND-Ru-

PIND ratio of 5.0 resulted in a 45% decrease and a 3-nm-red-shift of the ND

absorption band at 364 and 385 nm. The hypochromism of the PIND absorption band

reached a plateau at the base pair/PIND-Ru-PIND ratio ~7.0, and a constant

hypochromism was observed for the ratio above this value. A single clean isosbestic

point was observed at all DNA base pair/PIND-Ru-PIND ratios, suggesting that only

one spectrally distinct PIND-Ru-PIND/DNA complex is present. Both observations

are qualitatively consistent with those observed for intercalating compounds,

33

indicating that binding of PIND-Ru-PIND to ds-DNA takes place by preferential

intercalation. In addition, after the PIND groups have intercalated with ds-DNA, the

dicationic [Ru(dmbpy)2]2+ group in PIND-Ru-PIND forms an ion-pair with a

phosphate of ds-DNA, making the two intercalated PIND groups more tightly fixed in

between the base pairs of ds-DNA.

Figure 2-7: UV-Vis absorption spectra of 20 μM PIND-Ru-PIND in 0.10 M pH 7.0 phosphate buffer with increasing concentration of salmon sperm DNA (from top, 0, 20, 40, 60, 80 and 100 μM in base pair).

2.3.3 Application for Label-free DNA Detection

2.3.3.1 Ru-PIND-Ru and DNA Detection

Ru-PIND-Ru and electrocatalytic oxidation of guanine

In section 2.3.2, we showed that Ru-PIND-Ru intercalates very strongly to ds-

DNA. Here we demonstrate its application for label-free DNA sensing. Under

optimized conditions, the target DNA was selectively bound to their complementary

capture probes and became fixed on the biosensor surface upon hybridization.

Thorough rinsing with the hybridization buffer washed off most of the non-

hybridization related binding. Ru-PIND-Ru was brought to the biosensor surface

during a subsequent incubation with hybridized duplex. Non-intercalation related Ru-

34

PIND-Ru uptake was mostly removed by extensive washing with NaCl saturated 10

mM phosphate buffer. For proof-of-concept, synthetic oligonucleotides with simple

polyT, polyG, or repeated AT and AG units were designed and tested. Cyclic

voltammograms for the complete biosensors are shown in Figure 2-8. For the non-

complementary poly(T)40, one pair of minute voltammetric peaks were observed at

the redox potential of Ru-PIND-Ru (0.62 V) after hybridization (Figure 2-8 trace a),

largely due to pure electrostatic interaction between Ru-PIND-Ru and CP on the

biosensor surface. For the complementary poly(AT)20, poly(AG)20 and poly(G)40,

slight positive shifts (8.0 ± 2.0 mV) in the redox potential were observed and the peak

currents increased substantially (Figure 2-8 traces b, c and d).

Figure 2-8: Cyclic voltammograms of 200 nM of (a) poly(T)40 hybridized to a non-complementary capture probe coated electrode, and (b) poly(AT)20, (c) poly(AG)20, and (d) poly(G)40 hybridized to their complementary CP coated electrode, respectively. Scan rate: 100 mV/s.

The number of Ru-PIND-Ru molecules producing the observed current can be

estimated from the charge under the oxidation current peak. Since two electrons are

transferred per Ru-PIND-Ru molecule, the observed current, 0.30 µA after

hybridization to 200 nM of poly(AT)20, resulted therefore from 1.3 pmol of active and

35

intercalated Ru-PIND-Ru. Assuming the Ru-PIND-Ru/DNA base pair ratio of 1/4,

0.13 pmol of target DNA is hybridized. The hybridization efficiency was evaluated

electrochemically using Tarlov’s method,117 taking 1.26 x 10-11 mol/cm2 (midrange of

the estimated values) as the surface CP coverage on a 3.0-mm-diamter gold electrode,

it was found that 13% of the target DNA and 15% of the surface bound capture probes

were actually hybridized, comparable to the values found in literature.127, 128

Interestingly, when poly(AG)20 and poly(G)40 were hybridized with their

corresponding complementary capture probe coated biosensors, noticeable increments

in anodic current and slight decreases in cathodic current were observed (Figure 2-8

traces c and d). The increment increased almost linearly with increasing guanine

content, indicating that guanine bases in the oligonucleotides are catalytically

oxidized at 0.62 V by the intercalated Ru-PIND-Ru.129 Electrocatalytic oxidation of

guanine by ruthenium complexes will be discussed in detail later in chapter 3. These

results clearly demonstrated that Ru-PIND-Ru selectively interacts with ds-DNA and

the Ru-PIND-Ru–ds-DNA adduct has a very slow dissociation rate.

As guanine bases in the target probe can be catalytically oxidized by Ru-

PIND-Ru, better sensitivity is expected when working with genomic DNA/RNA

samples due to much higher content of guanine bases. We further evaluate the

feasibility of detection of cancer susceptibility genes using Ru-PIND-Ru as the

electroactive indicator. A full length TP53 gene in mRNA was selected as our target

gene. mRNA was extracted from rat liver tissues according to standard protocols.

Prior to hybridization, the mRNA mixture was denatured at 70 °C for 10 min. 20-mer

oligonucleotides were immobilized on the surface of each individual sensor and

served as CPs. The sequences of the CPs are complementary to the sequence of the

target gene at specific region where no mutation is reported. Upon hybridization at 53

36

ºC for 30 min, TP53 from the mRNA pool was selectively bound to the biosensor

surface. Thorough rinsing with the hybridization buffer washed off all of the non-

hybridization-related mRNA.

A typical cyclic voltammogram of the biosensor after applying Ru-PIND-Ru

is shown in Figure 2-9. As seen in trace a in Figure 2-9, a considerably higher peak

current was observed for the anodic process, indicating that a larger amount of

electrons is involved in the oxidation process, most probably due to the captured long

TP53 mRNA molecules that bring many more guanine bases to the biosensor surface.

The selectivity of the biosensor was evaluated in 1.0 mg mRNA by using one-base-

mismatched capture probe under hybridization conditions set for the perfectly

matched sequence. The current increment for the one-base-mismatched sequence was

only ~40% of that for the perfectly matched sequence (Figure 2-9 trace b), readily

allowing discrimination between the perfectly matched and mismatched sequences.

Figure 2-9: Cyclic voltammograms of TP 53 hybridized to (a) perfectly-matched and (b) one-base-mismatched biosensors. Hybridization was carried out in TE buffer containing 1.0 mg of mRNA. Scan rate: 100 mV/s.

37

Integration of oxidation or reduction current peak at a low scan rate 10 mV/s

yields a surface coverage of 3.8 pmol in terms of electroactive Ru2+/Ru3+ sites. The

total amount of Ru-PIND-Ru, 1.9 x 10-11 mol/cm2, is equivalent to 32% of the CP

being hybridized and fully intercalated. To have a better understanding of the

hybridization efficiency and Ru-PIND-Ru loading level, a series of QCM

measurements were carried out on TP53 after hybridization, and after Ru-PIND-Ru

intercalation. The results are summarized in Table 2-4. As shown in Table 2-4, ~40

fmole of TP53 was hybridized. This number represents ~1.6% of the surface-bound

CP was actually hybridized, a much lower value than that of short oligonucleotides

(20–50-mers) reported in the literature.127, 128 It is not surprising that the hybridization

efficiency decreases drastically with increasing the size of the target nucleic acids. In

addition, the QCM experiments showed that one Ru-PIND-Ru molecule intercalated

per 11–14 bases of TP53, suggesting that some of the Ru-PIND-Ru molecules

intercalated into the secondary structure of TP53,130, 131 further enhancing the

sensitivity of the method. Ru-PIND-Ru loading density was found to be in the range

of 1.5–2.2 × 10-11 mol/cm2, which is in good agreement with that obtained in

voltammetric tests.

Table 2-4: QCM data of CP coated quartz crystal after hybridization and intercalation

38

2.3.3.2 PIND-Ru-PIND and DNA Detection

In this section, we report the ECL properties of a bis-intercalator PIND-Ru-

PIND and its application for label-free DNA detection.

ECL behavior of PIND-Ru-PIND in TPA solution

The cyclic voltammetric and ECL responses of 5.0 μM PIND-Ru-PIND was

measured in pH 7 phosphate buffer saturated with TPA at a gold electrode. As shown

in Figure 2-10, the peak potential for the oxidation of TPA occurred at 1.0 V (Figure

2-10 trace c) while that of PIND-Ru-PIND occurred at 0.68 V (Figure 2-10 trace b),

implying that the oxidation of PIND-Ru-PIND at the electrode surface occurs before

that of TPA required for the production of ECL. The maximum ECL intensity for this

system was observed at 0.94 V (Figure 2-10 trace a).

Figure 2-10: (a) ECL intensity at 610 nm versus potential profiles, cyclic voltammograms of (b) 5.0 μM PIND-Ru-PIND in 0.10 M phosphate buffer (pH 7.0), and (c) 5.0 μM PIND-Ru-PIND in TPA saturated phosphate buffer. Scan rate: 20 mV/s. For clarity, the voltammogram of PIND-Ru-PIND was scaled up 50 times.

39

Figure 2-11 shows the ECL spectrum of PIND-Ru-PIND compared with its

photoluminescence spectrum. As expected, within experimental error, the ECL

spectra obtained for PIND-Ru-PIND at different positive potential biases have the

same features as its photoluminescence spectrum in phosphate buffer. This is because

the emission arises from the decay of the same MLCT excited state Ru(dmbpy)22+*,

generated by either illumination or electrochemical excitation.

Figure 2-11: (a) Photoluminescence spectrum of PIND-Ru-PIND (430 nm illumination) in 0.10 M phosphate buffer and (b) ECL spectrum of PIND-Ru-PIND in a TPA saturated 0.10 M phosphate buffer.

PIND-Ru-PIND for electrochemiluminescent DNA detection

DNA biosensors with redox active moieties grafted ND as electrochemical

indicators have previously been reported.99, 113 When hybridization occurs, ND

selectively interacts with the ds-DNA and gave a greatly enhanced analytical signal

compared to non-hybridized ss-DNA. The difference in voltammetric peak current is

used for quantification purpose. However, to our knowledge, no study has been done

on the ECL detection of DNA using a threading intercalator. Similar to the other

redox active ND intercalator, PIND-Ru-PIND was firstly evaluated as an electroactive

tag for possible applications in ultrasensitive DNA sensing.

40

In the first hybridization test, a complementary and a non-complementary CP

(control) coated biosensor were hybridized to the target DNAs. Upon hybridization,

the complementary target DNAs were selectively bound to the complementary CP

and became fixed on the biosensor surface. On the other hand, hybridization with the

non-complementary CP failed to capture any of the target DNAs, and therefore little

change of the biosensor was expected. Thorough rinsing with the hybridization buffer

washed off most of the non-hybridization related DNA. PIND-Ru-PIND was brought

to the biosensor surface during a subsequent incubation with a PIND-Ru-PIND

solution.

Figure 2-12: Linear scan voltammograms (LSV) of PIND-Ru-PIND bound to (a) 200 nM of complementary DNA, and (b) 1.0 μM non-complementary DNA hybridized biosensors. Supporting electrolyte: 0.10 M phosphate buffer (pH 7.0), potential scan rate 100 mV/s.

Linear scan voltammograms (LSV) for the biosensors after hybridization are

shown in Figure 2-12. For the non-complementary CP coated biosensor, after

hybridization a minute voltammetric peak was observed at the redox potential of

PIND-Ru-PIND (Figure 2-12 trace b), largely due to pure electrostatic interaction of

residual PIND-Ru-PIND and CP on the biosensor surface. As shown in trace a, after

hybridization with the complementary CP coated biosensor, a slight positive shift in

41

the redox potential was observed and the peak current increased by as much as 100-

fold. It was found that extensive washing with NaCl-saturated pH 7.0 0.10 M

phosphate buffer removed most of the nonspecific PIND-Ru-PIND uptake.

The ECL behaviour of the hybridized biosensors before and after incubation

with PIND-Ru-PIND was examined with a positive potential bias of 1.0 V applied to

the biosensors. Figure 2-13 shows ECL responses of various biosensors after

hybridization with target DNAs and incubation with PIND-Ru-PIND. Very little ECL

response was observed for the biosensor hybridized with the non-complementary

target DNA (trace a), largely due to the presence of a very small amount of

electrostatically bound PIND-Ru-PIND to the DNA. It can be seen that the presence

of intercalated PIND-Ru-PIND in the complementary target DNA hybridized

biosensor greatly increase the ECL signal of the system with an enhancement of ~50-

fold (trace c). In contrast, no ECL signal was observed at the complementary DNA

hybridized biosensor before PIND-Ru-PIND incubation. Therefore, we concluded that

the much improved ECL response after PIND-Ru-PIND intercalation is indeed due to

a genuine ECL process of the Ru(dmpy)2 moieties. The selectivity was evaluated at

50 pM by analyzing one-base-mismatched DNA under hybridization conditions set

for the perfectly matched sequence. A ∼65% drop in ECL intensity was observed

(trace b), readily allowing discrimination between the perfectly matched and

mismatched oligonucleotides. These results clearly demonstrated that PIND-Ru-PIND

selectively interacts with ds-DNA and the PIND-Ru-PIND−ds-DNA adduct has a

very slow dissociation rate, which paves the way for developing ultrasensitive DNA

biosensors.

42

Figure 2-13: ECL responses at 610 nm of PIND-Ru-PIND bound to biosensors hybridized with (a) 1 nM non-complementary target, (b) 50 pM one-base-mismatched target, and (c) 50 pM complementary target. Poise potential: 1.0 V, ECL measurement was done in TPA saturated phosphate buffer.

We further evaluate the effect of TPA and the applied potential on the ECL

signal. Figure 2-14 shows the dependence of ECL signal on TPA concentration as

well as applied potential. The optimum concentration of TPA is found at around 0.20

M (saturated). At lower TPA concentration, the electron-transfer reaction between

activated PIND-Ru-PIND and the intermediate formed during the oxidation of TPA is

less effective. Therefore, to maximize ECL sensitivity, ECL measurements were

always conducted in the saturated TPA solution. Variation of the applied potential had

a profound effect on the ECL intensity (Figure 2-14 solid trace). The threshold

potential of ECL was found to be 0.65 V and the ECL signal increased rapidly beyond

the threshold potential until a maximum intensity was reached in the range of

0.93−1.05 V. Under the optimized conditions, the ECL signal was proportional to the

target DNA concentration in the range of 0.70−400 pM with a detection limit of 400

fM, 2000-fold higher that of the direct voltammetric detection of DNA. The ECL data

agreed well with the voltammetric results obtained earlier in solution and confirmed

43

again that PIND-Ru-PIND is electrochemiluminescent and it can be used to detect

DNA with high specificity and sensitivity.

Figure 2-14: Effect of TPA (•) and applied potential () on the ECL responses at 610 nm of 50 pM complementary DNA after incubation in 10 μM PIND-Ru-PIND.

2.4 Conclusions

Ruthenium-complexed electroactive DNA threading intercalators, Ru-PIND-

Ru and PIND-Ru-PIND, were synthesized and characterized. Spectrometric and

electrochemical characterization confirmed the formation of desired compounds.

Successful attempts were made in utilizing Ru-PIND-Ru and PIND-Ru-PIND as

effective electroactive reporters in label-free electrochemical DNA assays. The

combination of selective intercalation to ds-DNA with electrocatalytic property of

PIND-Ru-PIND or electrochemiluminescent function of Ru-PIND-Ru, provides

simple, direct, and highly sensitive non-labeling methods for DNA quantification. We

believe that methods demonstrated in this chapter could have a wide applicability to

ultrasensitive DNA assays.

44

3 Ruthenium-Based Polymer Complexes for

Electrocatalytic Guanine Oxidation

3.1 Introduction

We have seen considerable efforts in the development of electrochemical

techniques for the detection of nucleic acids in biological samples. Due to their

compatibility with advanced semiconductor technologies, electrochemical biosensors

promise to provide a simple, accurate and inexpensive platform for DNA assays. Our

interest in sensitive and selective electrochemical nucleic acids biosensors led us to

search for new electrocatalysts, lowering the potential at which DNA is electroxidized:

the lower the potential, the better the sensitivity.

Guanine has been identified as the first to oxidize DNA base, oxidized either

directly or through hole transfer along the DNA π stack.132 Its oxidation has been

studied extensively in the context of DNA damage, associated with mutation and

aging.133, 134 The oxidation potentials of guanine and guanosine were measured by

pulse radiolysis and cyclic voltammetry.135, 136 Due to differing pH and other

conditions, the estimates of the one-electron oxidation potentials vary widely. Pulsed

radiolysis, the measurement of choice when the redox reaction involves unstable

radicals in the presence of an internal reference,137 registered values of the one-

electron oxidation potentials of guanine, which varied between 0.63 and 0.83 V

versus normal hydrogen electrode (NHE) i at pH 13.135, 138 The electrochemically

measured direct oxidation potentials were ~ 0.9 V versus NHS at physiological pH.139

i For a better comparison, all literature reported values were converted to those with reference to

standard hydrogen electrode (NHE).

45

High over-potentials make difficult the accurate direct determination of the oxidation

potentials.

Guanine bases in DNA were also catalytically oxidized by [Ru(bpy)3]2+ and

polyvinylpyridine (PVP)-bound [Ru(bpy)2]2+.111, 140 The Rusling group observed

voltammetric responses to the catalytic guanine oxidation in DNA on pyrolytic

graphite electrode covered with PVP-Ru(bpy)22+ film at 0.99 V versus NHE.111 Thorp

et al measured the oxidation potential of guanine in double helical DNA indirectly, by

using trans-[Re(O)2(4-Ome-py)4]+ and related dioxorhenium(V) complexes as

mediators, reporting a potential between 1.1 V and 1.2 V versus NHE at pH 7.140

Because of the uncertainty about the redox potential for the one-electron oxidation of

guanine, it is necessary to obtain an independent and reliable value for this important

DNA base. This is an important measure of the susceptibility of cells to damage by

endogenous oxidizing radicals and exogenous oxidants.

In chapter 2, we reported a threading intercalator, N,N′-bis[3-

propylimidazole]-1,4,5,8-naphthalene diimide (PIND) complexed with Ru(bpy)2Cl

(Ru-PIND-Ru), that catalyzes the oxidation of guanine.141 In this chapter, we continue

to focus on guanine oxidation, and report the systematic determination of the apparent

oxidation potentials of guanine, guanosine and guanosine monophosphate (GMP) in

aqueous silane solution, by monitoring their electrocatalytic oxidation current, with

ruthenium-based polymer complexes as electrocatalysts. It is established that in a pH

7.4 aqueous saline solution, guanine and guanosine are catalytically oxidized on

ruthenium-complexed redox polymer modified indium tin oxide electrodes at

potentials much more reducing than previously reported.93

46

3.2 Experimental


4-vinylpyridine (99%, Fluka), N-vinylimidazole (99%, Fluka), acrylamide

(98%, Fluka), Ammonium persulfate ((NH4)2S2O8, 98%, Alfa Aesar), N,N,N′,N′-

tetramethylethylenediamine (TMEDA, 99%, Alfa Aesar), 4,4'-dimethyl-2,2'-dipyridyl

(99.5%, Aldrich), 4,4'-dimethoxyl-2,2'-dipyridyl (99.5%, Aldrich), 4,4'-dicarboxyl-

2,2'-dipyridyl (99.5%, Aldrich), ruthenium(III) trichloride (99%, Aldrich) and 2,2′-

bipyridyl ruthenium dichloride (Ru(bpy)2Cl2, 99%, Avocado Research Chemicals)

were of reagent grade and used as received. Phosphate buffered saline (PBS, pH 7.4,

1st base) was used as test buffer.

3.2.2 Synthesis of Ruthenium-complexed Redox Polymers

Synthesis of Polymer Backbone

Two types of polymer backbone were used in this study, they are

poly(vinypyridine-co-acryamide), PVP-co-PAA, and poly(vinylimidazole-co-

acrylamide), PVI-co-PAA. As shown in scheme 3-1, PVP-co-PAA was prepared by

copolymerization of 4-vinylpyridine and acrylamide.142, 143 Briefly, 2.0 mL of 4-

vinylpyridine and 2.0 g of acryamide were dissolved in 10 mL acetone-water mixture

(1:1) in a round-bottom flask. To this solution, 100 mg of ammonium persulfate

(NH4)2S2O8 and 100 µL of N,N,N',N'-tetramethylethylenediamine (TMEDA) was

added and the mixture was kept refluxing for 3-5 hours. The copolymer was

precipitated with 30-35 mL of acetone. After centrifuging, the copolymer was

redissolved in 5-10 mL of ethanol-water mixture (1:1) and precipitated again in

acetone. The final product was dried in vacuum for overnight at 50° C. PVI-co-PVP

was prepared by copolymerization of N-vinylimidazole and acrylamide in a similar

way.

47

Scheme 3-1: Synthetic scheme of (A) PVP-co-PAA and (B) PVI-co-PAA.

Synthesis of Electroactive Pendent Unit

Ruthenium(Ru) complexes containing different R substituted bis(2,2′-

bipyridine) (R= CH3, OCH3, COOCH3) were prepared following a literature reported

procedure.116 As an example, a mixture of Ru(III) trichloride hydrate (1 g, 3.8 mmol,

20% excess) of 4,4'- methyl substituted-2,2'-bipyridine (1.2 g, 6.5 mmol), and lithium

chloride (1.1 g, 26 mmol) in 60 mL of DMF was stirred under reflux for 8 hours. The

solvent was removed by vigorous stirring in diethyl ether. The crude product was then

dissolved in chloroform and extracted with water. Upon drying, the product [Ru(bpy-

CH3)2Cl2]Cl was obtained in the form of black powder (1.72 g, 70% yield).

We found that the ruthenium complex with carboxylic acid-substituted

bis(2,2'-bipyridine) (R=COOH) is difficult to purify. Therefore, carboxylic acid-

substituted bis(2,2'-bipyridine) (R=COOH) was esterified before complexing with

ruthenium. Briefly, 20 mg of 4,4'- carboxylic acid substituted-2,2′-bipyridine was

dissolved in 5 mL of methanol and 200 μL of concentrated H2SO4, and the

reaction mixture was left refluxing for 3 hrs. Excess methanol was evaporated and

5 mL of water was added into the same flask. Adjust solution pH to 7 with sodium

48

bicarbonate. Extraction with chloroform and re-crystallize in acetone/chloroform

yield white crystal of methyl ester substituted-2,2′-bipyridine.

Scheme 3-2: Synthetic scheme of ruthenium complexes as electroactive pendant unit.

Synthesis of Ruthenium-complexed Redox Polymers

Ruthenium complexes containing different substituted bis(2,2'-bipyridine)

(R=H, CH3, OCH3, COOCH3) were grafted to the polymer backbone (PVP-co-PAA

or PVI-co-PAA) according to a literature procedure with a slight modification

(Scheme 3-3).143, 144 Briefly, in a 10 mL round-bottom flask, 150 mg of polymer was

dissolved in 4 mL fresh-distilled ethylene glycol (EG). 50 mg of Ru(bpy-R)2Cl2 (R=H,

CH3, OCH3, COOCH3) was added into this solution and the mixture was stirring for

10 min before refluxing. The course of the ligand-exchange reaction was followed by

cyclic voltammetry. Samples were taken from reaction mixture at different time

points and diluted with EtOH/PBS mixture before running CV. The reaction was

stopped when stable CV curves obtained. The reaction mixture was slowly poured

into 400 mL of rapidly stirred anhydrous ether. The precipitate was collected by

suction filtration through a fine fritted funnel. The crude product was dissolved in

water and extracted twice with chloroform. The sample was rotary evaporated and

vacuum dried.

49

Scheme 3-3: Synthesis of redox polymer (A) Ru-complexed PVP-co-PVI and (B) Ru-complexed PVI-co-PAA.

3.2.3 Preparation of Redox Polymer Modified Electrodes

Ruthenium-complexed redox polymers can be easily attached to many

electrode surfaces. In our study, we choose ITO as our substrate electrode. The ITO

electrode was cut into 1 cm × 2.5 cm rectangles and cleaned using standard

procedures. The redox polymer coated electrodes can be prepared by

electrochemically cycling the clean ITO electrode in polymer solutions. Bare ITO

electrodes were electrochemically cycled in different redox polymer solution (5

mg/mL) from 0 V to 1 V for 3 cycles. Rinsed the electrodes with DI water thoroughly

and air dried. These redox polymer modified electrodes are stable for weeks.

3.2.4 Apparatus

Electrochemical experiments were carried out using a CH Instruments model

660A electrochemical workstation coupled with a low current module (CH

Instruments, Austin, TX). A conventional three-electrode system, consisting of a 3.0-

mm-diameter glass carbon working electrode or an ITO electrode (1 cm × 2.5 cm), a

50

non-leak Ag/AgCl (3.0 M NaCl) reference electrode (Cypress Systems, Lawrence,

KS), and a platinum wire counter electrode, was used in all electrochemical

measurements. All potentials reported in this work were referred to the Ag/AgCl

electrode. UV-Visible spectra were recorded on a V-570 UV/VIS/NIR

spectrophotometer (JASCO, Japan). NMR spectroscopic study was done on a Bruker

400 MHz system from Bruker Biospin GmbH (Karlsruhe, Germany). Mass

spectrometric experiments were performed with a Finnigan/MAT LCQ Mass

Spectrometer (ThermoFinnigan, San Jose, CA).


3.3.1 Synthesis and Characterization of Redox Polymers

Ruthenium-based polymer complexes have been studied in detail in late 1980s

due to their long-lived excited states and facile electron-transfer dynamics. The

application of these materials as redox catalysts, photosensitizers and molecular

diodes have been proposed.145-147 In this work, we will use ruthenium-complexed

redox polymers as electrocatalysts for the study of guanine oxidation.

We have successfully synthesized a series of redox polymers, poly(4-

vinylpyridine) (PVP) and poly(N-vinylimidazole) (PVI) copolymerized with

polyacrylamide (PAA) containing different substituted ruthenium bis(2,2'-bipyridine)

redox units. The synthesis process was monitored by checking the potential shift of

the Ru2+/3+ redox peaks using cyclic voltammetry. Figure 3-1 shows typical cyclic

voltammograms taken at different time point during a ligand-exchange reaction. It is

known that the redox potential of Ru complexes changes upon ligand exchange.

Therefore, the potential shift provides a convenient way to monitor the formation of

Ru-complexed polymer. As shown in Figure 3-1, only one pair of redox peaks around

51

0.25 V was observed at the initial stage of the reaction (curve a), corresponding the

redox reaction of starting material Ru(bpy)2Cl2. Upon refluxing with copolymer for

some period, a new pair of peaks appeared at 0.6 V (curve b), indicating the ligand

exchange on the Ru metal center and the formation of complexes with the polymer.

Both electron transfer processes are clearly resolved with redox waves. The slightly

larger peak-to-peak potential separations are mainly due to the lower conductivity of

the reaction medium. The intensity of the new peaks increased gradually with a

concurrent decrease of the peaks at 0.25 V. Diminish of peaks at 0.25 V suggests the

completion of ligand exchange reaction (curve c).

Figure 3-1: Cyclic voltammograms of the reaction mixtures at different reaction time during the synthesis of PVIPAA-Ru(bpy)2Cl: (a) 0, (b) 2 h and (c) 20 h.

The formation of the polymer complex was also followed using UV-Vis

spectrophotometry. The adsorption spectra of the starting material Ru(bpy)2Cl2 before

and after the complexion reaction were shown in Figure 3-2. Two adsorption peaks at

351 nm and 510 nm were observed for Ru(bpy)2Cl2 (spectrum a), which has been

assigned to metal-to-ligand charge-transfer (MLCT) transition of the type dπ(Ru)→

π*(bpy).148 The peak at 510 nm underwent a blue shift after the reaction, indicating

the formation of the polymer complex (spectrum b).

a

b

c

52

Figure 3-2: UV-Vis spectra of Ru(bpy)2Cl2 before (―) and after (---) grafting to the polymer backbone.

The redox potentials of the polymers complexed with different R-substituted

Ru(bpy-R)2Cl2 are summarized in Table 3-1. It has been reported that ligand

modifications alter the electron density at the metal center of the osmium (Os)

complex, resulting in shifts in the redox potential.101 Similarly, substitutions on the

bipyridine ligands change the ligands’ stability and influence the overall redox

potential of the Ru complexes. Methyl and methoxyl groups are both electron

donating groups that increase electron density and facilitate oxidation of the redox

species they are attached to, resulting in lower oxidation potentials. The methoxyl

group, with a free lone pair electron adjacent to the π system, exerts a stronger

resonance effect than the inductive effect of the methyl group. As a result, complexes

with methoxyl substitution exhibit lower redox potential compared to those with

methyl groups. On the contrary, methyl ester of carboxyl group, an electron accepting

unit, decreases the electron density near the metal center and lead to a higher

oxidation potential of the Ru complexes.

b

a

53

Table 3-1: Oxidation potentials of polymers complexed with different substituted ruthenium redox units.

Redox Polymer E1/2 (V)PVIPAA-Ru(OCH3) 0.42PVIPAA-Ru(CH3) 0.53PVPPAA-Ru(OCH3) 0.55PVIPAA-Ru 0.62PVPPAA-Ru(CH3) 0.66PVPPAA-Ru 0.74PVIPAA-Ru(COOCH3) 0.79PVPPAA-Ru(COOCH3) 0.93

3.3.2 Redox Polymer Modified Electrode

Figure 3-3 shows the cyclic voltammograms (CVs) of a PVIPAA-Ru(bpy)2Cl2

coated ITO electrode in PBS buffer solution recorded at different scan rates. The CVs

show electrochemical behavior expected for an adsorbed fast electron exchanging

species, as demonstrated by the symmetrical shape of the voltammograms, the 90-mV

width of half-peak current, the linear dependency of the peak current on the scan rate

and the close-to-zero peak separation, suggesting the formation of redox polymer thin

film on the electrode surface.

Figure 3-3: (A) Sweep-rate dependency of the CV of a PVIPAA-Ru(bpy)2Cl2 coated ITO electrode in PBS, scan rate=20, 50, 100, 200, 500, 1000 mV/s, following arrow direction. (B) The plot of anodic peak current with scan rate.

54

Figure 3-4 shows the CVs of the ITO electrodes modified with different redox

polymers in PBS buffer. By varying the substituted group on 2,2′-bipyridine, we are

able to manipulate the redox potentials of the ruthenium complexes and hence those

of the redox polymers. The wide range of tunable potential is particularly important

for their application as redox catalysts. A good example is when the polymer is used

to modify electrodes and catalyze another redox couple. Only complexes with redox

potentials greater than that of the redox couple in question are able to oxidize it and

yield catalytic current.149 To efficiently catalyze the reaction, the redox potential of

the polymer coating must be matched to the redox couple in question.

Figure 3-4: Cyclic voltammograms of redox polymer thin film coated ITO electrodes in PBS. From left to right: (a) PVPPAA-Ru(OCH3), (b) PVPPAA-Ru(CH3), (c) PVPPAA-Ru, (d) PVIPAA-Ru(COOCH3), (e) PVPPAA-Ru(COOCH3).

3.3.3 Electrocatalytic Oxidation of Guanine on Modified Electrode

ITO electrode has very wide electrochemical window in PBS at positive

potential range (Figure 3-5A), which makes it an excellent substrate for this study. It

was found that guanine oxidation occurs at around 1.0 V on blank ITO electrode

under neutral pH. This is due to the high over-potential of guanine oxidation on ITO

55

surfaces. We also found that guanine oxidation is pH dependent and becomes

relatively easier at higher pH (Figure 3-5B).

Figure 3-5: Cyclic voltammograms of blank ITO in (A) PBS and (B) PBS with 0.5 mM guanine at different pH: (―–) 10.5, (·····) 9.5, (– – –) 8.5, (· − · −) 7.5.

However, guanine oxidation occurs at much lower potential on redox polymer

modified electrode. As shown in Figure 3-6, PVIPAA-Ru(bpy)2Cl modified electrode

shows a pair of symmetrical redox peaks in PBS (trace a). When guanine was added

to the PBS solution, its reversible voltammogram changed to the one showing

characteristic of irreversible electrocatalytic oxidations, a rise in anodic current and

decrease in cathodic current (trace b). This catalytic guanine electrooxidation was not

observed on the bare ITO electrode (trace c), suggesting that guanine was catalytic

oxidized by PVIPAA-Ru(bpy)2Cl on modified electrode.89

We also observed that the rate of catalytic oxidation of guanine is pH

dependent. Because guanine oxidation is proton-releasing, the Gibbs free energy

release driving reaction, the rate increases at higher pH. Mechanistically, the electron

transfer in the guanine-Ru(III) complex is proton-coupled: the abstraction of the first

guanine electron is concomitant with the deprotonation of guanine.150

56

Figure 3-6: Cyclic voltammograms of a PVIPAA-Ru(bpy)2Cl thin film coated ITO electrode in (a) PBS and (b) PBS with 20 mM guanine. (c) Cyclic voltammogram of a bare ITO electrode with 50 mM guanine in PBS. Scan rate: 100 mV/s.

3.3.4 Redox Titration

We measured the catalytic oxidation currents of guanine across the pH values

ranging from 2 to 12 in a stirred four-electrode cell, containing a pH electrode, the

redox polymer coated working electrode, a reference and a counter electrode. With

the working electrode poised at the formal potential of its redox polymer, we

increased the pH step-wise, while monitoring the electrooxidation current. The

guanine within the redox polymer films was promptly consumed upon applying the

formal potential. Figure 3-7 shows acid-base titration curves for the electrooxidation

of the guanine in the films. Each titration curve reveals a pH threshold and an upper

limit, where the electrooxidation rate no longer changes with pH. The intersection of

the lower and middle portions of the curve provides the onset pH, at which the

electrocatalytic oxidations started. The classical S-curves establish that near the

formal potentials of the polymers, guanine electrooxidation involves an OH-

consuming step. The slopes for polymers 2, 3, 4 and 5, the closest to neutral pH

domain (pH 5- pH 10), are similar. We found that the current increases 10-fold for a

57

2-pH unit increase. With each pH unit translating to 59 mV, the behavior is Tafel-like,

i.e. a 10-fold current increase is observed upon increasing the reaction driving

potential by 2×59 mV=118 mV. 129

Figure 3-7: Titration curves showing the increase in the electrocatalytic guanine oxidation current when the pH is raised at redox polymer film coated ITO electrodes. From left to right, (a) PVIPAA-Ru(COOCH3), (b) PVPPAA-Ru, (c) PVPPAA-Ru(CH3), (d) PVIPAA-Ru, (e) PVPPAA-Ru(OCH3), (f) PVIPAA-Ru(OCH3).

The rates of the five steps (Reactions 1-5) of the one-electron electrocatalytic

oxidation of guanine denoted as GH are, by definition, equal at steady state.

Ionization: GH + H2O → G- + H3O+ (1)

Ion-pairing: G- + Ruc3+ → [G-.Ruc

3+] (2)

Electron transfer: [G-.Ruc3+] → [•G .Ruc

2+] (3)

Dissociation: [•G.Ruc2+] → •G + Ruc

2+ (4)

Bulk electrooxidation of the Ru complex: Ruc2+→ Ruc

3+ + e (5)

Reaction 1 explains the pH dependence of the guanine electrooxidation current

in Figure 3-7, for the redox polymer electrocatalysts of Figures 3-4. The rate, i.e. the

current, reaches a plateau at the pH where the rate of formation of the ion pair [G-

.Ruc3+] no longer depends on the G- concentration, because all the Ruc

3+ is exhausted.

The concentration of Ruc3+ in the film is a function of the rate of electrooxidation of

Ruc2+ (reaction 5) in the bulk of the film, determined by the redox potential of the

a b c d e f

58

Ruc2+/3+ redox couple and by the electron diffusion coefficient in the film, which in

turn depends on the rate of collisional electron exchange between the redox centers.

The potential at which catalytic oxidation of guanine by a particular redox

polymer is plotted again the onset pH as shown in Figure 3-8 (trace a).ii The slopes

are found of 60 mV per pH unit. Thus, when normalized for pH, all potentials at

which the electrooxidations observed are the same. For example, the 0.81 ± 0.01 V (vs

NHE) value at pH 7.4 for guanine is also obtained when the measured threshold

potential is adjusted by 0.059 × [threshold pH - 7.4] V. These threshold potentials are

neither reversible potentials nor thermodynamic values, but are practical values

(apparent oxidation potential).

Figure 3-8: pH dependency of the threshold-potentials of (a) guanine (•), (b) guanosine (), and (c) GMP () electrooxidation, catalyzed by different polymers.

3.3.5 Oxidation of Guanosine and Guanosine Monophosphate (GMP)

We extended our study to guanosine and GMP to explore the influence of the

sugar unit and the phosphate group on the oxidation potential (refer to Figure 3-9 for

their chemical structures). Similar to guanine, linear pH dependency with slopes of 60

ii All potentials reported in the text from this point onwards were converted with reference to NHE

59

mV were obtained for both guanosine and GMP (Figure 3-8, traces b and c), except

that the curves shifted to a more positive value.

Figure 3-9: Chemical structures of guanine, guanosine and GMP.

For biological considerations, the electrooxidation potentials of guanosine are

significantly higher than those of guanine. The threshold value for the catalytic

electrooxidation of guanine at pH 7.4 is 0.81 ± 0.01 V (vs NHE), while that of

guanosine is 1.02 ± 0.01 V (vs NHE). The difference reflects, at least in part, the

difference in the energetics of forming the G– anion in Reaction 1 by deprotonation of

the imidazole of guanine, versus by deprotonation of guanosine, which does not have

an imidazole proton. In the case of GMP, at physiological pH and low ionic strength,

where the negative charge of the phosphate group is not screened by Na+ cations, the

anionic proximal phosphate could make the forming of G– energetically unfavorable

and raise the threshold potential of oxidation of G of GMP to above that of guanosine.

However, we found that the pH dependent catalytic electrooxidation currents of GMP

are indistinguishable from those of the guanosine. The value of 0.81 V and 1.02 V (vs

NHE) oxidation potential measured for guanine and guanosine/GMP in physiological

buffer solution, is considerably lower than the earlier reported 1.17 V and 1.29 V (vs

NHE) respectively.88, 151

60

3.4 Conclusions

The electrochemical behaviour of guanine, guanosine, and guanosine

monophosphate (GMP) at ruthenium-complexed redox polymer film modified ITO

electrodes was examined by voltammetry and redox titration. Using the redox

polymer coated ITO electrodes as indicator electrodes, a new method for measuring

the oxidation potentials, based on monitoring their catalytic oxidation by different

redox polymer coated electrodes at different pH, was proposed in this work. To our

knowledge, the oxidation potentials of 0.81 V and 1.02 V versus NHE determined for

guanine and guanosine/GMP under physiological conditions were the lowest

oxidation potentials ever reported. This finding may provide useful information for

the study of complex charge migration involved in biological systems.

61

4 Nanostructured Functional

Polyethylenedioxythiophenes (PEDOTs)

4.1 Introduction

Electroactive polymers represent an emerging class of important conductive

materials, which are viewed as pioneer structures that may lead to useful

macromolecular electronic devices.145 Electroactive polymers fall into three board

categories: π-conjugated conducting polymers, polymers with covalently linked redox

groups (redox polymers), and ion-exchange polymers. The common features are semi-

rigid mechanical properties, the ability to pass electrical current, and the ability to be

oxidized or reduced by electrolysis. In the previous chapter, we discussed about

ruthenium-complexed redox polymers, their synthesis and electrochemical properties.

From this chapter onwards, we will focus on π-conjugated conducting polymers,

specifically, polyethylenedioxythiophenes (PEDOTs), their nanostructure synthesis,

properties and applications.

4.1.1 Conducting Polymers

Conducting polymers have found applications in many areas due to their

flexibility and metal-like electric conductivity. The discovery of electrically

conducting polymer (CP) dates back to 1970s. In 1977, Alan Macdiarmid, Hikedi

Shirakawa and Alan Heeger reported a 10-million-fold increase in the conductivity of

polyacetylene doped with iodine.152 Consequently, other aromatic CPs with better

stabilities and conductivities were developed (Figure 4-1).

62

Figure 4-1: Chemical structures of common conductive polymers.

Conducting polymers can be synthesized either chemically or

electrochemically, each with its own advantages and disadvantages. Chemical

synthesis allows large scale production of the materials and subsequent modification

in bulk, which is currently difficult with electrochemical synthesis. However,

electrochemical technique allows the formation of ultrathin films in the nanometer

range.

Conducting polymers are highly conjugated systems. The unique chain

structure of alternating double- and single-bonds allows the formation of delocalized

electronic states, which maximizes the sideway overlap among the π molecular

orbitals and endows the polymer with metal-like semi-conductive properties. This

delocalization allows charge carrier move along the polymer backbone and between

adjacent chains, but limited by disorder and coulombic interactions. Prior to doping,

conducting polymers are insulators (~10-10 S/cm); however, the conductivity can be

boosted up to 12 orders of magnitude upon doping. The conductivity depends on the

individual polymer system as well as the type and extent of doping.153 Doping can be

performed chemically or electrochemically. The chemical nature of the dopant not

63

only affects electrical conductivity, but also affects surface and bulk structural

properties.

Conducting polymers have been successfully applied in the area of energy

storage,154 antistatic coating,155 electrochromic devices,156 light emitting diodes,157, 158

and sensors.159, 160 More details can be found in Gurunathan’s review paper.157

Research on conducting polymers for biomedical applications expanded greatly with

the discovery that these materials are compatible with many biological molecules in

1990s.161 Furthermore, the ability to entrap and controllably release biological

molecules and the ability to transfer charges from a biochemical reaction make them

an emerging class of advanced materials for many specific applications such as

biosensor, tissue engineering scaffolds, drug delivery devices and bio-actuators.

4.1.2 Nanostructured Conducting Polymers

Conducting polymers (CPs) have been intensively studied both in the

fundamental and applied researches. Earlier studies on conducting polymers have

been focused mainly on bulk-type sample properties, such as thin films or powder

pellets. They have been used to construct organic light emitting diodes (OLED),

organic field effect transistors (OFET), electromagnetic interference shielding, anti-

static coatings and capacitors. With the growing interests of the use of nanomaterials

in biosensors, microelectronics and photonics, people have shifted their focuses

towards the synthesis and applications of nanostructured CPs. In particular, one-

dimensional (1-D) polymer nanostructures have attracted a lot of attention due to their

intriguing properties derived from small dimensions, high aspect ratio, enlarged

surface area, and flexibility. One dimensional (1-D) nanostructures are the smallest

dimension structures that can be used for efficient electron transport and thus are

critical to the function and integration of nanoelectronic devices. Because of their

64

high surface-to-volume ratio and tunable electron transport properties resulting from

the quantum confinement effect, their electrical properties are strongly influenced by

small perturbations. As opposed to their thin-film counterpart, signal loss due to

lateral current shunting can be avoided. Therefore, enhanced performances such as

greater sensitivity and faster response can be expected.

Although 1-D CP nanomaterials have demonstrated improved performances

over the conventional bulk materials in a wide applications such as sensors, controlled

drug release, bio-scaffold, wound dressing, and filtrations,71-73, 162-165 the main

challenge remains to find efficient, scalable, and site specific approaches for precisely

incorporating these 1-D nanomaterials into devices. Moreover, the difficulty in

modifying the polymer surfaces with desired functionalities hinders the realization of

their full potential.

4.1.3 Synthesis of 1-D Conducting Polymer Nanostructures

There are many different ways to synthesize 1-D conducting polymer

nanostructures, both chemically and electrochemically. Template-assisted approaches

have been widely employed to achieve well defined 1-D CP nanostructures. Hard-

templates166, 167 include anodized aluminum oxide membranes (AAO), track-etched

polycarbonate membranes and zeolite channels, and soft-templates168-172 refer to

surfactants, polyelectrolytes and liquid crystals. In addition, other template-less

methods like interfacial polymerization,173 seeded polymerization,174

electrospinning,163 and stepwise electrochemical deposition175 have also been reported.

The key to successful 1-D nanostructure synthesis is promoting conditions that favor

homogenous nucleation of the polymer chains and suppressing subsequent growth of

new polymer chains on top of one another (secondary growth).176 However, it is

65

always a challenge to precisely control the size, shape and surface functionality of

organic conducting polymer nanostructures.

4.1.4 1-D Polyethylenedioxythiophene (PEDOT) Nanostructures

PEDOT represents a technologically important class of conducting polymers

displaying high stability, moderate band gap, low redox potential and high optical

transparency in its electrically conductive state.177 Although PEDOT has been

extensively investigated for use in antistatic coatings, flexible electronic devices, and

transparent electronics,178 its 1-D electron transport properties are remain largely

unexplored due to the difficulty of producing PEDOT with well-controlled 1-D

morphology.

PEDOTs are particularly recalcitrant to fibril or tubular polymer growth due to

their enhanced hydrophobicity and slow kinetics of oxidation. Techniques such as

nanofiber seeding,179 activated seeding,180 and interfacial polymerization181 that have

been used to synthesize nanofibers of polyaniline (PAni) or polypyrrole (PPy) yield

only granular powders when extended to PEDOTs.168 Only a few studies on the

nanostructured PEDOT have been presented recently. Martin’s “template synthesis”

entails synthesizing the desired materials within the pores of a nanoporous membrane

such as track-etched membranes and porous alumina membranes. By controlling the

polymerization time, hollow tubules or solid nanofibris of PEDOT were

synthesized.182-184 Although hard-template-assisted synthesis can produce well-

defined conducting polymers with 1-D nanostructured features, this technique is

unsatisfactory for fabricating cost-effective nanostructures in large volumes for

possible applications. In addition, etching away the template might cause the polymer

nanoarrays to collapse into disoriented structures. Alternatively, Manohar’s group

described a one-step, room temperature method to chemically synthesize bulk

66

quantity of PEDOT nanofibers using a V2O5 seeding approach. The resulting PEDOT

nanofibers are 3-10 micron long and 100-180 nm in diameter and in the form of a

non-woven mesh.174 The same group has also successfully synthesized 50-100 nm

PEDOT nanotubes using a reverse microemulsion polymerization method.168 Using

similar reverse microemulsion system, Yoon et al obtained PEDOT nanorods with

diameters of 40±20 nm and length of 200±30 nm.164 Foulger et al reported the

synthesis of PEDOT nanofibers with diameters down to the 10-nm scale from an

aqueous anionic surfactant solution. According to the paper, this self-assembled

micellar soft-template approach is able to generate PEDOT fibers with aspect ratio

greater than 100 with very high yield.169

However, to the best of our knowledge, there has been no report on the

synthesis of nanostructured PEDOTs bearing side-chain functional groups. In addition,

the effect of side-chain functional groups on the polymer morphology has not been

well-studied. Driven by potential biosensor applications, we are particularly interested

in functionalized PEDOTs, their nanostructures, properties and applications. In this

chapter, we explore the synthesis of 1-D nanostructured functional PEDOTs and their

integration into functional devices. Various synthesis approaches such as surfactant

template guided chemical polymerization, stepwise electropolymerization, and

electric field-assisted polymerization were studied.

4.2 Experimental


Ethylenedioxythiophene (EDOT, Sigma-Aldrich), lithium perchlorate (LiClO4,

Fluka), sodium dodecyl sulfate (SDS, Alfa Aesar), cetyltrimethylammonium bromide

(CTAB, Sigma-Aldrich), iron tricloride (FeCl3, Sigma-Aldrich), ammonium

67

persulfate ((NH4)2S2O8, Alfa Aesar) were used as received. Hydroxymethyl-

functionalized EDOT (EDOT-OH) and carboxylic acid-functionalized EDOT

(EDOT-COOH) were synthesized in our laboratory as described elsewhere (Figure 4-

2).185 A phosphate-buffered saline (PBS) consisting of 137 mM of NaCl, 2.7 mM of

KCl, and 10 mM of phosphate buffer was used as supporting electrolyte solution.

Indium tin oxide coated glass (Delta Technologies, Ltd.) were cut into 1cm × 1 cm

square pieces and cleaned by standard procedure prior to use.

Figure 4-2: Chemical structure of EDOT-OH and EDOT-COOH

4.2.2 Chemical Polymerization

The PEDOT nanostructures were synthesized by chemical polymerization in a

mixture solution of a monomer (EDOT, EDOT-OH, or EDOT-COOH), a surfactant

(CTAB or SDS) and an oxidizing agent (APS or FeCl3). The general synthetic process

was as follows: EDOT monomers were dissolved in an aqueous surfactant solution,

and then an oxidizing agent aqueous solution was quickly added into the above

mixture to initiate the polymerization. All the reactions were left at room temperature

overnight. The reaction mixture containing final polymer products were centrifuged

and washed with DI water and methanol alternatively for at least 3 times. The molar

ratio between monomer and oxidizing agent was kept at 1:1 for APS and 1:2 for FeCl3.

68

4.2.3 Electrochemical Polymerization

Electrochemical polymerization was carried out with an Autolab PGSTAT 30

potentiostat (Metrohm) using either cyclic voltammetry or chronoamperometry

techniques. Sputter coated Au electrode with or without pattern and ITO electrodes

were used as working electrodes, and a Pt wire was as the counter electrode. A non-

leak Ag/AgCl electrode was used as the reference electrode. The

electropolymerization was done either in an organic solvent system (0.1 M

TBAPF6/CH3CN) or an aqueous system (0.1 M LiClO4/H2O, with additional 10% of

CH3CN or 0.05 M SDS).

4.2.4 Characterization

The morphology of the resulting PEDOT nanostructures was examined using

field-emission scanning electron microscope (FE-SEM, JEOL) and high-resolution

transmission electron microscope (HRTEM, JEOL). Samples for both SEM and TEM

were prepared by placing a drop of aqueous suspension of PEDOTs on silicon wafer

and carbon-coated copper grid.


4.3.1 Surfactant Template-Guided Nanofiber Synthesis

Carboxylic acid-functionalized PEDOT (poly(EDOT-COOH) nanofibers were

synthesized from an aqueous surfactant aqueous solution via a soft-template approach.

Figure 4-3 shows the SEM and TEM images of the poly(EDOT-COOH) nanofibers

synthesized in the presence of SDS as a surfactant and FeCl3 as an oxidizing agent.

69

Figure 4-3: (a) SEM image and (b) TEM image of poly(EDOT-COOH) nanofibers. (c) HRTEM image of individual nanofiber.

In a typical procedure, 17.25 mg of EDOT-COOH was dissolved in 5 mL of

100 mM SDS aqueous solution, and 24.38 mg of FeCl3 was added into the above

solution, vigorously stirring for 1 min for a homogenous mixing. FeCl3 addition

resulted in some white precipitation which might due to the formation of complexes

with surfactant molecules.169 After overnight reaction, the blue color reaction mixture

was centrifuged and washed with DI water and methanol alternatively for at least 3

times to get rid of the excess surfactant and oxidizing agent. The synthesized

nanofibers have diameter around 5-10 nm with length up to several micrometers.

Effect of monomer and Surfactant concentration

Figure 4-4 shows the effect of surfactant concentration on the formation of

nanofiber structures. A polymer film without any nanostructures was obtained when

SDS concentration was lower than 30 mM. Fibril structures were starting to form

when SDS concentration was above 50 mM, and became clearer with the increase of

SDS concentration. Higher SDS concentration (100 mM) leads to longer and more

separated nanofibers. We also found that the ratio between monomer and surfactant is

important for the formation of nanofiber structures (Figure 4-5). When the reaction

mixture was diluted and SDS concentration dropped below 50 mM, no nanofiber

structures were formed, instead more spherical nanostructures were obtained.

70

Figure 4-4: SEM images of poly(EDOT-COOH) obtained at different surfactant concentration (SDS used as a surfactant and FeCl3 as oxidizing agent, monomer concentration was kept constant at 15 mM): (a) 30 mM, (b) 50 mM, (c) 60 mM, (d) 80 mM, and (e) 100 mM.

Figure 4-5: SEM images of poly(EDOT-COOH) obtained at different monomer concentration: (a) 2, and (b) 20 mM, in the presence of 100 mM SDS and 40 mM FeCl3. SEM images of poly(EDOT-COOH) obtained from (c) 10x, (d) 5x, and (e) 2x dilution from the original mixture of 20 mM monomer, 100 mM SDS and 40 mM FeCl3.

As illustrated in Figure 4-6, the formation of PEDOT nanofibers can be

explained as follows: When SDS is dissolved in water, spherical micelles formed

when its concentration is between critical micelle concentration (CMC) and sphere-to-

a b

200 nm 200 nm

c d e

200 nm 200 nm 200 nm

a b c

d e

500 nm 500 nm

500 nm 500 nm

500 nm

71

cylinder transformation concentration. The CMC for SDS is 8.2 mM in deionized

water at room temperature186 and the second CMC for SDS to undergo spherical to

cylindrical micelle transformation is 70 mM.187 However, the concentration required

for the transformation from sphere to cylinder drops significantly with the addition of

salt.188 In addition, FeCl3 is known to be a good oxidant for the polymerization of

EDOT.169 Therefore, in this case, spherical micelles that contain EDOT-COOH

monomers transform into cylindrical micelles after the addition of FeCl3, and

polymerization of the EDOT-COOH monomer inside the cylindrical micelles

produced the PEDOT nanofibers.

Figure 4-6: Schematic of the salt-assisted surfactant micelle transformation and formation of poly(EDOT-COOH) nanofibers.

Oxidizing agents and morphology

It has been reported that oxidizing agents have great impact on the

morphology of the final conducting polymer products. When APS was used as

oxidizing agent instead of FeCl3, no precipitation was formed, and the clear solution

changed to light blue in colour after overnight polymerization. However, no polymer

products were isolated after removing the excess SDS and APS. This might due to the

formation of oligomers in stead of polymers, which are difficult to be isolated from

the reaction mixtures.

Surfactant and morphology

EDOT-COOH FeCl3

MeOH/H2O

Wash

SDS

EDOT-COOH

Poly(EDOT-COOH)

72

The surfactant effects on the morphology of PEDOT nanostructures were also

investigated using another cationic surfactant cetyltrimethylammonium bromide

(CTAB). The cmc of CTAB is 0.87 mM in deionized water and a transition from

spherical to cylindrical micelles occurs at a concentration of more than 12 CMC at

room temperature.189 The concentration of CTAB in this study was controlled in the

ranges of micelle aggregations according to the literatures.190 Surprisingly,

poly(EDOT-COOH) nanospheres with diameter around 80-90 nm were formed

instead of nanofibers when CTAB was used as surfactant (Figure 4-7), regardless of

the oxidizing agents used. It was previously reported that PPy formed ribbon- or wire-

like nanostructures in the presence of CTAB and APS, but sphere-like structures

when CTAB and FeCl3 were used.171 The authors suggested that a lamellar

mesostructure was formed due to self-assembly between the cations of CTAB and

anions of APS, leading to the growth of wire- and ribbon-like nanostructures.191

However, no fibril nanostructure was obtained when the same system was adapted to

EDOT. We did observed the formation of a white precipitate of (CTA)2S2O8

immediately after adding APS to both CTAB and CTAB/EDOT mixture in our study,

but no fibril structure was observed in final polymer product. No precipitate was

formed between CTAB and FeCl3 when FeCl3 was used as oxidant. We attributed the

different morphologies of nanostructured poly(EDOT-COOH) and PPy to their very

different nature of monomer materials. The detailed mechanism of nanospheres

formation is not clear at this moment. We believe that solution microsctructures

(micelle aggregates) could play an important role in controlling the growth of polymer

in both oxidant systems. More studies need to be done to understand this phenomenon.

73

Figure 4-7: SEM image of Poly(EDOT-COOH) nanostructures obtained in the presence of 30 mM EDOT-COOH monomer and 10.5 mM CTAB with different oxidizing agent (A) 30 mM of APS (B) 60 mM FeCl3.

4.3.2 Stepwise Electropolymerization

Surfactant-assisted chemical polymerization provides a facile and inexpensive

way of producing PEDOT nanofibers in bulk quantities; however, integration of these

nanofibers to devices with the desired alignment remains a challenge. Solution casting

has been used to incorporate nanowires/nanotubes to devices,164 but there is little

control over the distribution, position, and alignment of the nanowires/nanotubes

(Figure 4-8), which will eventually affect the device performance and limit their

applications. There is a definite need for more convenient strategies that allow direct

fabrication of conducting polymer nanowires precisely between electrode junctions.

Figure 4-8: FESEM image of the PEDOT nanotubes deposited on the microelectrodes consisting of a pair of gold interdigitate electrodes with 40 fingers (dimensions: 10 μm width, 4000 μm length, 50 nm thickness, 10 μm inter-electrode gap). Adapted from Ref 73 .

a b

200 nm 200 nm

74

Electrochemical deposition has been demonstrated as a powerful tool for

growing metal, oxide, and polymer nanostructures directly on the electrode surface.192

Oriented arrays of nanowires provide an ideal platform for a range of sensing

applications. Most of the reported oriented nanostructure arrays were obtained

through hard-template approaches.193 Removal of the template after synthesis

rendered desired nanowire arrays that replicate the size and ordering of the template.

For nanowires/nanotubes with a diameter smaller than 100 nm, etching the template

would cause the collapse of nanowire arrays into disoriented structures.192 Liang et al

developed a stepwise electrochemical deposition process to grow large arrays of

oriented conducting polymer nanowires on a variety of smooth and textured substrates

without using a supporting porous template.175 Through controlled nucleation and

growth using stepwise electrochemical deposition process, aniline or pyrrole

molecules spontaneously assembled into desired uniform nanowires. Tseng’s group

further improved this step-wise electrochemical deposition approach and reported a

template-free, site-specific elelctrochemical method to grow conducting polymer

nanowire arrays at electrode junction in an integrated microfluidic system.165, 194 The

synthesized conducting polymer nanowires (CPNWs) with diameter ranging from 50

to 200 nm formed numerous intercrossing networks between two electrodes with a

gap of 2 μm.

In this section, we report our study of growing functional PEDOT

nanostructures directly on the electrode surfaces or between electrode junctions using

electrochemical polymerization. Different electrochemical techniques, polymerization

conditions as well as surfactant effects on the polymer morphology were investigated.

Polymerization parameters and morphology

75

We applied stepwise electrochemical deposition approach on functionalized

EDOT and studied their morphology dependency on the polymerization parameters.

No PEDOT nanowire arrays were observed under all conditions we studied. Figure 4-

9 and 4-10 display some representative results of polyEDOT, poly(EDOT-OH) and

poly(EDOT-COOH) electropolymerized under different parameters. The polymer

morphology was found greatly dependent on the side-chain functional group, solvent

and current density. The formation of granular clusters might due to the poor

solubility of EDOT oligomers and the inherent non-planar molecular structure of

PEDOTs, especially those with side-chain functional group.

Figure 4-9: SEM images of (a) poly(EDOT-COOH) and (b) poly(EDOT-OH). Polymerization was done under constant current, 0.5 mA/cm2, in TBAPF6/CH3CN containing 10 mM monomer. Images at bottom panel are taken at higher magnification.

5 μm

2 μm

a

2 μm

5 μm

b

76

Figure 4-10: SEM images of poly(EDOT-COOH) polymerized under constant current, 0.25 mA/cm2, in (a) 0.1 M TBAPF6/CH3CN and (b) 0.1 M LiClO4 aqueous solution (9:1 H2O: CH3CN) containing 10 mM EDOT-COOH monomer. Images at bottom panel are taken at higher magnification.

Effect of additives

We also investigated the effect of additives on the polymer morphology.

Anionic surfactant SDS, non-ionic surfactants Brij 35 (polyoxyethyleneglycol

dodecyl ether) and pluronic P123 (triblock copolymer EO20PO70EO20) were tested.

Unlike pyrrole or aniline, thiophene and its derivatives have limited solubility in

water and their polymerization occurs usually in organic solvents. Addition of

surfactants allows their polymerization from aqueous solutions.195 In the case of

EDOT, it has been shown that the addition of anionic surfactants improves the

solubility of the monomer and lowers the anodic potential at which the

electropolymerization takes place.196 Figure 4-11 are some representative SEM

images of poly(EDOT-OH) electropolymerized from aqueous electrolyte with

different surfactants. It is obvious that surfactant has great influence on the resulting

polymer morphology. Branched porous polymer network was obtained when SDS and

2 μm

5 μm

a b

5 μm

2 μm

77

P123 were used as surfactant. Smooth and dense film structure was observed when

Brij 35 was employed as surfactant. The results suggested that the influence of

solubilising additives on the polymer morphology depends very much on the different

chemical nature of the surfactants, micelle stability and their affinity to the surface. 196

Figure 4-11: SEM images of poly(EDOT-OH) polymerized under constant current, 0.1 mA/cm2, from 10 mM EDOT-OH aqueous solution with different surfactants (a) 50 mM SDS, (b) 5 mM Brij 35, and (c) 2% P123. Supporting electrolyte: 0.1 M LiClO4.

4.3.3 Electrical Field-Assisted Nanowire Growth

Assembly of metallic and semiconductor nanowires directly between

electrodes through electrochemical methods is highly attractive for nanoelectronic

devices because it reduces lengthy, complex and expensive lithographic fabrication

procedures. The application of an electric field to fabricate palladium nanowires

between electrodes has been previously reported.197 We applied similar approach to

grow 1-D carboxylic acid-functionalized poly(3,4-ethylenedioxythiophene)

(poly(EDOT-COOH)) nanowires between two gold electrodes directly from EDOT-

COOH monomer solution.

Patterned Au electrodes with 2 µm gap were fabricated following standard

photolithography method. A drop of 0.1 M EDOT-COOH/CH3CN monomer solution

(2 µL) was placed on the substrate to fill the electrode gap. A cover slip was used to

prevent evaporation. Application of external electrical field between two parallel Au

electrodes resulted in the growth of polymer nanowires across the gap. Figure 4-12

200 nm 200 nm 200 nm

a b c

78

shows the optical and scanning electron micrograph of poly(EDOT-COOH)

nanowires grown between two Au electrodes with a 2 μm gap under alternating

electric field. Multiple nanowires with regular distance were formed between two

electrodes and bridged the gap.

Figure 4-12: (a) Optical and (b) scanning electron micrograph of poly(EDOT-COOH) nanowires grown between two Au electrodes under alternating electric field.

Figure 4-13: Scanning electron micrograph of poly(EDOT-COOH) nanowires grown between two Au electrodes under DC field.

We found that the monomer concentration, the applied voltage (field intensity)

and frequency had marked influences on the nanowire growth. Under the applied

alternating electric field, the EDOT monomer is first oxidized to its radical cation at

the anode. In the absence of other mobile ions, the ionized EDOT cations then migrate

a

b

79

to the opposite side of the electrode driven by the external electric field across the

electrode gap. High concentration of monomer radicals is maintained near the

electrode surface due to the much faster electron-transfer reaction than diffusion.

Competition between the formation of a high concentration of ions at the anode and

the migration of the ions away from the electrode controls the wire thickness near the

electrode.198 Formation of conductive nanowires between electrodes decreases the

local electric field, inhibiting the growth of new nanowires nearby. 197, 199 This

explains the observation of regular arrays of nanowires along the electrode gap. We

also noticed that wires grown under fix DC field are a few μm in diameters and have

tapered shape towards cathode (Figure 4-13), while nanowires formed under

alternating electric field is much thinner, although not as straight as those formed

under fix DC field. This can be explained that under alternating electric field,

nanowires begin to grow from both sides of the electrode and get joined in the gap.

The nanowires are curved, possibly due to the slight mismatch of their starting

position.

The adhesion between these poly(EDOT-COOH) nanowires and electrodes are

found strong and stable. The as-grown nanowires are conductive with a total parallel

resistance in the range of several kΩ. Since the polymerization and nanowire

assembly are occurred concurrently, study of their applications as functional devices

can be carried out immediately after synthesis. We will discuss the application of

these functional PEDOT nanowires for label-free protein detection later in chapter 5.

4.4 Conclusions

In this chapter, we have studied the synthesis of 1-D nanostructured functional

PEDOTs and their integration into functional devices. Various approaches such as

80

surfactant-assisted chemical polymerization, stepwise electropolymerization as well

as electric field-assisted polymerization were explored. Functional PEDOT nanofibers

with diameters in the range of 100 nm were obtained using surfactant-assisted

chemical polymerization. Electrochemical polymerization generated PEDOTs with

very different morphology, which is largely dependent on the polymerization

parameters, side-chain functional groups as well as the additives. Porous PEDOT

networks were obtained on electrode surfaces with SDS as a surfactant. Finally, we

demonstrated that electric field-assisted polymerization produces functional PEDOT

nanowires that can bridge an electrode gap. This approach eliminates post-synthesis

alignment and assembly, allowing a direct study of their applications as functional

devices immediately after synthesis.

81

5 PEDOT Nanowires for Label-Free Protein

Detection

5.1 Introduction

Simple yet sensitive detection of biological and chemical species of interest

has always been desired for various applications in the area of healthcare and life

sciences. From disease diagnostics to drug screening, nanowire-based devices have

emerged as a powerful platform for direct and sensitive electrical detection of

biological and chemical species.63, 200-202 Significant progress has been achieved in the

use of silicon nanowires (SiNWs)64, 66, 203 and carbon nanotubes (CNTs)70, 204-206 for

the detection of proteins and nucleic acids. In recent years, one-dimensional (1-D)

nanostructures of conducting polymers have attracted great attention due to their

tunable conductivity, chemical diversity and ease of fabrication. There has been

considerable progress in the synthesis of 1-D conducting polymer nanostructures.166-

176, 181-183, 191-194, 207 Sensor applications of conducting polymer nanowires/nanofibers

have also been demonstrated recently.73, 208, 209 Among all known conducting

polymers, poly(3,4-ethylenedioxythiophene) (PEDOT) has been recognized as one of

the most promising candidate for practical applications due to its remarkable

conductivity and air stability.177 Despite the success commercial application of

PEDOT-PSS as an antistatic coating material and a hole-transporting material, there is

currently limited knowledge on synthesis and application of 1-D PEDOT

nanostructures.164, 165, 168, 170, 174, 182, 184, 210, 211 In particular, functionalized PEDOTs

and their application for biosensing has not been reported.

In previous chapter, we reviewed various approaches for growing 1-D PEDOT

nanostructures. For example, well-defined nanowires/nanotubes with diameter around

82

200-300 nm were produced from hard-template assisted synthesis using anodized

alumina membrane,182, 184 nanofibers and nanotubes with diameter less than 100 nm

were obtained using surfactant as soft-template.168, 211 Despite the large scale

synthesis, integration of loose PEDOT nanowires/nanofibers onto a functioning

device is a big challenge. There is a definite need for the development of much more

convenient strategies that allow direct fabrication of conducting polymer nanowires

precisely between electrode junctions.

Polyaniline/poly(PAni/PEO) nanowires with diameter around 100 nm were

formed on gold electrodes using a scanned-tip electrospinning method, which showed

resistance change upon exposure to NH3 gas at concentration as low as 0.5 ppm.163

Polyaniline (PAni) and polypyrrole (PPy) nanowires (in the order of 100 nm) were

generated across an electrode junction through electrodeposition within e-beam-

patterned electrolyte channels.212 Tseng’s group reported a template-free, site-specific

elelctrochemical method to grow conducting polymer nanowire arrays at the electrode

junction in an integrated microfluidic system.165, 194 The synthesized conducting

polymer nanowires with diameter ranging from 50 to 200 nm formed numerous

intercrossing networks between two electrodes with a gap of 2 μm. The use of such a

device for the detection of HCl and NH3 gases was demonstrated.194

In chapter 4, we reported a simple electrochemical approach of growing 1-D

carboxylic acid-functionalized poly(3,4-ethylenedioxythiophene) (poly(EDOT-

COOH)) nanowires between two gold electrodes directly from EDOT-COOH

monomer solution by applying an alternating electric field. In this chapter, we will

investigate the performance of these functionalized PEDOT nanowire devices and

their application for label-free protein detection.

83

5.2 Experimental Section


Hydroxymethyl-functionalized EDOT (EDOT-OH) was synthesized following

previously reported procedures.213, 214 Carboxylic acid-functionalized EDOT was

subsequently synthesized from EDOT-OH through an ether linkage (EDOT-

COOH).215 Aptamers with 5′ amine modification were custom-synthesized (1st base,

Singapore). The sequences used in this work are as follows:

5′-T6GGTTGGTGTGGTTGG-3′ (thrombin-binding aptamer)

5′-T6TATCTACGAATTCATCAGGGCTAAAGAGTGCAGAGTTACTTAG-3′ (49-

mer non-complementary probe)

5′-T6TTGAGTCTGTTGCTTGGTCAGC-3′ (28-mer non-complementary probe)

Bovine serum albumin (BSA) and thrombin (from bovine plasma) were purchased

from Sigma-Aldrich and diluted with buffer to desire concentration before use. The

buffer used in this work is 50 mM Tris buffer with 140 mM NaCl and 1 mM MgCl2,

with pH adjusted to 5 for the maximum stability of polymer nanowires.216 All other

reagents were used as received.

5.2.2 Device Fabrication and Nanowire Synthesis

Device fabrication and nanowire formation across the electrode gap were

described in detail in Chapter 4. Briefly, standard photolithography method was used

to create patterned Au electrodes with 2 μm gap. A drop of 0.1 M EDOT-COOH

/acetonitrile monomer solution (2 μL) was placed on the substrate to fill the electrode

gap. A cover slip was applied on top of the electrode to prevent evaporation.

Application of an alternating electrical potential for 2 ~ 4 s (4 V, 1 kHz) resulted in

the growth of polymer nanowires across the gap. The device was then rinsed with

acetonitrile followed by deionized water and air dried. A heat treatment (70 °C in

84

vacuum for 2 h) was carried out to improve the adhesion between nanowires and the

electrodes. All the experiments were carried out with an Autolab PGSTAT 30

potentiostat (Metrohm) under ambient conditions.

5.2.3 Aptamer Immobilization and Protein Binding

The Poly(EDOT-COOH) nanowires were incubated in 100 mM EDC/40 mM

NHS for 20 min, rinsed with deionized water, further incubated in 1 μM amine-

modified aptamer solution for overnight. After washing and drying, aptamer was

covalently immobilized onto the poly(EDOT-COOH) nanowires through formation of

amide bond with surface carboxylic acid group. The aptamer-modified PEDOT

nanowires were pre-incubated in blank binding buffer for 30 min before exposed to

protein solution.

5.2.4 Electrical Measurement

Figure 5-1: Experimental setup of CPNW FET devices for protein detection. Au 1 and Au 2 represent two working electrodes (WE1 and WE2), served as source and drain respectively. Counter electrode is a Pt wire where the electrochemical gate potential is applied via the reference electrode (Ag/AgCl). A bias voltage (Vbias) applied between WE1 and WE2 (Vbias=VWE1-VWE2) is equivalent to source-drain current (Vsd).

85

After nanowire formation, the device was converted into FET devices, as

shown in Figure 5-1. A polydimethylsiloxane (PDMS) well was applied on top of the

electrode as a solution chamber. Two gold electrodes were configured as source and

drain respectively, a Pt wire counter electrode was used as gate, and a Ag/AgCl

reference electrode was used to accurately control the solution potential.74, 217 All

electrical measurements were conducted in buffer solution.


5.3.1 Device Characteristics

The functional PEDOT nanowires were formed between two Au electrodes

with a 2 μm gap as described in chapter 4 section 4.3.3. The as-grown nanowires are

conductive with a total parallel resistance in the range of several kΩ. Figure 5-2a

shows the representative I-V characteristic of a poly(EDOT-COOH) nanowire device.

A linear ohmic dependence of the current on the voltage was observed in the range of

-0.1 to 0.1 V, confirming that the PEDOT nanowires are conductive and have good

ohmic contact with both electrodes. The measurement was performed under ambient

condition in dry state and the conductivity of the PEDOT nanowires was found to be

stable for a few days, in contrast to a few hours reported for PAni and PPy

nanowires.212 The electrical properties of the PEDOT nanowire devices were also

investigated in the liquid phase. Figure 5-2b displays the dependence of the source-

drain current (Isd) versus the source-drain voltage (Vsd) on gate voltage (Vg). At

negative Vsd, the Isd value decreases upon increasing positive Vg, as illustrated in

Figure 5-2c, indicating a typical depletion mode p-type transistor characteristic of

poly(EDOT-COOH) nanowires.165 This phenomenon was however, not observed on

thin film-based devices. We noted that the Isd-Vsd curve is not stable enough when

86

cycled in solution with pH greater than 5, and the source-drain current was much

lower in pH 7 solution as compared to pH 5. We believe that the low conductivity and

slow degradation is related to the deprotonation of carboxylic acid group on the

polymer side chain. The charge effect of side chain functional group on the

conductivity of PEDOT thin film was previously reported by our group.216 Therefore,

the FET measurement and subsequent protein binding experiments were done in pH 5

buffer.

Figure 5-2: Electrical characteristics of poly(EDOT-COOH) nanowire device. (a) I-V curve, (b) Isd-Vsd characteristics at varying Vg (Vg = -0.4 to + 0.4 V, step = 0.1 V, scan rate = 1 mV/s) in 0.1 M LiClO4/buffer (pH=5), and (c) |Isd|-Vg plot at constant Vsd= -0.2 V, derived from the above Isd-Vsd characteristics.

5.3.2 Biomolecule Conjugation

For specific detection of proteins, a biorecognition layer on the sensor surface

is needed. Aptamer has been identified as a superior substitute for antibodies in

immunoassay due to higher stability, affinity and specificity.218 In addition, aptamer is

much smaller in size than its antibody counterpart, which makes it more preferable in

FET based biosensing. It has been reported that all biological binding events occurred

outside Debye length are very likely to be “screened” by the electrical double layer

and their effect on the equilibrium charge carrier distribution will not be detected.219

Debye length is simply defined as the typical distance required for screening the

surplus charge by the mobile carrier present in a material, which varies as the inverse

square root of ionic strength. In this work, the aptamer-protein interaction was chosen

b a c - 0.4 V

+ 0.4 V

87

to demonstrate the applicability of using poly(EDOT-COOH) nanowires for label-free

electrochemical protein detection.

An amine-modified thrombin-binding aptamer was covalently immobilized

onto the poly(EDOT-COOH) nanowire surface via EDC/NHS coupling. Isd-Vsd

characteristics of the poly(EDOT-COOH) nanowire devices at gate voltage of 0 V

were recorded before and after aptamer conjugation. As illustrated in Figure 5-3, the

source-drain current increased dramatically after aptamer immobilization. We

attribute this current/conductance increase to the increase in negative charge density

on the nanowire surface, as aptamers are negatively charged oligonucleotides.

Immobilization of aptamers leads to a substantial accumulation of negative charge on

the nanowire surfaces. The negative charges on the nanowire surface will change the

mobile charge carrier distribution within the nanowires and hence its conductance.

Figure 5-3: Isd-Vsd characteristics of the poly(EDOT-COOH) nanowire device at gate voltage of 0 V (a) before and (b) after aptamer immobilization.

To reaffirm this charge effect, a positively charged molecule, thrombin, was

covalently conjugated to the poly(EDOT-COOH) surface in the similar manner. Isd-

Vsd characteristics were monitored before and after thrombin immobilization. We

observed a decrease of the source-drain current after thrombin immobilization.

Because the isoelectric point of thrombin is 7.05, thrombin is positively charged in pH

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5 buffer. As a result, binding of thrombin leads to an accumulation of positive charge

on the nanowire surfaces, causing reduced conductance. No significant change in the

source-drain current was observed for nanowires incubated only in PBS buffer,

confirming that the decrease was not due to nanowire detachment or polymer

degradation (Figure 5-4). This further identified the effect of surface charge on the

nanowire conductance.

Figure 5-4: Normalized current change before and after immobilization of different biomolecules on the poly(EDOT-COOH) nanowires (Vg= 0, Vsd= 0.4 V).

5.3.3 Protein Detection

The feasibility of poly(EDOT-COOH) nanowires for label-free

electrochemical protein detection was evaluated by aptamer-modified PEDOT

nanowires. Thrombin was chosen as our model protein because of its well studied

aptamer sequence. The aptamer modified poly(EDOT-COOH) nanowire devices was

incubated with 100 nM thrombin solution in pH 5 Tris buffer (50 mM Tris, 140 mM

NaCl and 1 mM MgCl2) for 1 h. Figure 5-5 displays typical Isd-Vsd characteristics

change before and after thrombin binding. An average decrease of 50% in

conductance with variation less than 10% was observed across devices.

89

Figure 5-5: Typical Isd-Vsd characteristics of aptamer-modified PEDOT nanowire devices (a) before and (b) after incubation with thrombin (Vg= 0 V).

To further confirm that the change in source-drain current was attributed to the

specific binding of thrombin to the aptamer-modified nanowires, control experiments

were carried out by using devices modified with non-complementary sequence probes.

Two non-thrombin binding probes with different length were tested. As shown in

Figure 5-6, much smaller responses were obtained for both devices compared to the

device modified with thrombin-binding aptamer. A relatively larger signal was

observed for the device with the longer non-complementary probe, indicating greater

amount of non-specific binding on longer non-complementary probe. This is most

likely due to larger electrostatic interaction.

From our experiments, it is evident that aptamer-modified poly(EDOT-COOH)

nanowires is capable to detect the presence of thrombin specifically. We further

evaluate the device sensitivity by monitoring the change of source-drain current as a

function of thrombin concentration. Isd-Vsd curves were recorded after 1 h incubation

with thrombin at different concentrations as shown in Figure 5-7A. The source-drain

currents at fixed source-drain voltage and fixed gate voltage (Vsd = 0.4 V, Vg= 0 V)

were extracted and plotted against thrombin concentration. As shown in Figure 5-7B,

over a wide range of concentration from 1 nM to 1 μM, the normalized current

90

changes (-∆I/I0) are linear with the logarithm of thrombin concentration. The dynamic

range of our device is wide enough to cover physiologically relevant concentrations,

which ranging from a few nanomolar (resting blood) to several hundred nanomolar

(when the clotting cascade is activated).220 Upon further optimization, PEDOT

nanowire-based sensor platform could be used for in vitro diagnostics.

Figure 5-6: Normalized current change of aptamer-modified PEDOT nanowire devices after incubation with 100 nM thrombin: (a) 49-mer non-complementary probe, (b) 28-mer non-complementary probe, (c) thrombin-binding aptamer.

Figure 5-7: (A) Overlay of Isd-Vsd curves after 1 h incubation with thrombin at concentration of 0, 1, 10, 100, 1000 nM, follow arrow direction. (B) Calibration curve of aptamer-modified PEDOT nanowire device: normalized current change (-∆I/I0) as a function of thrombin concentration. The source-drain current was measured at Vsd= 0.4 V and Vg=0 V.

A B

91

5.3.4 1-D Nanostructure vs 2-D Film

It is generally believed that the 1-D nanostructures are critical to the function

of the FET based sensing. Because of their high surface-to volume ratio and tunable

electron transport properties from the quantum confinement effect, their electrical

properties are strongly influenced by minor perturbations.211 To better understand the

effect of 1-D nanostructures in conducting polymer-based FET biosensor, we

performed similar experiments on poly(EDOT-COOH) thin film device. First, a

poly(EDOT-COOH) thin film was electropolymerized onto a 5-μm interdigitated

microelectrode (IME, Abtech Scientific Inc) from an acetonitrile solution containing

10 mM monomer with 0.1 M TBAPF6 as a supporting electrolyte. After rinsing with

acetonitrile and deionized water, the polymer coated electrode was tested in 0.1 M

LiClO4 solution buffered to pH 5 with acetic acid. Aptamer immobilization and

thrombin assay were carried out similarly as describe in earlier section. Figure 5-8 and

Figure 5-9 show that no FET property was observed for thin film based devices and

binding of thrombin induced negligible change in the electrical properties of polymer

film.

Figure 5-8: Electrical characteristics of poly(EDOT-COOH) thin film device. (A) Isd-Vsd characteristics at varying Vg (Vg = -0.4 to + 0.4 V, step = 0.1 V, scan rate = 1 mV/s) in 0.1 M LiClO4/buffer (pH=5), (B) |Isd|-Vg plot at constant Vsd (a) 0.2 V and (b) -0.2 V, derived from the Isd-Vsd characteristics.

A B

- 0.4 V

+ 0.4 V

92

Figure 5-9: Isd-Vsd characteristics of poly(EDOT-COOH) thin film device (a) before, (b) after aptamer immobilization, and (c) after thrombin binding (Vg= 0 V).

To confirm the binding of thrombin to PEDOT thin film, Cy3-labeled

thrombin-binding 2nd aptamer was further incubated with the thin film device to label

binded thrombin. Fluorescence microscope image of the PEDOT thin film device

modified with thrombin-binding aptamer presented uniform fluorescence over the

surface while as shown in Figure 5-10b. Only background reflection for the device

modified with random sequence probe.

Figure 5-10: Fluorescence microscope image of poly(EDOT-COOH) modified with (a) random sequence probe and (b) thrombin-binding aptamer after binding with thrombin and a Cy3-labeled 2nd aptamer.

The different result obtained for thin film device suggested that simply binding of

protein to the film surface was not able to perturb the electronic properties of polymer

film. PEDOT thin film is not sensitive enough to be used as a signal transducer for the

93

label-free protein detection. The above findings did not surprise us. As illustrated in

Figure 5-11, binding of biomolecules to the surface leads to the accumulation or

depletion of charge carriers in the “bulk” of the nanowire in 1-D structures, as

opposed to only the surface in 2-D thin film. This gives rise to large changes in the

electrical properties that potentially enables the detection of biomolecules. 1-D

nanostructures avoid the reduction in signal intensities from lateral current shunting.

This property provides the sensing modality for label-free and direct electric readout

in nanowire based FET biosensors.

Figure 5-11: The major advantage of 1-D nanostructures (B) over 2-D thin film (A) for FET based biosensing. Adapted from Ref 211 .

5.4 Conclusions

1-D carboxylic acid-functionalized poly(3,4-ethylenedioxythiophene)

(poly(EDOT-COOH)) nanowires have been synthesized directly across the electrode

junction using electric field assisted growth. This fabrication method is simple and

easy to control because polymerization and self-alignment is achieved simultaneously.

The poly(EDOT-COOH) nanowire devices show typical depletion mode p-type field-

effect transistor (FET) properties. More importantly, the free carboxylic acid

functional group on the polymer backbone makes the bioconjugation much

straightforward. The protein-binding aptamer was conjugated to the PEDOT

nanowires as the molecular recognition element. We have demonstrated label-free

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electronic detection of blood-clotting factor thrombin using aptamer-modified

PEDOT nanowire FET devices. Upon exposure to thrombin solution, there is a

substantial decrease in current flow, attributed to the specific interaction between

thrombin and aptamer-conjugated polymer chains. The dynamic range covers

physiological relevant concentrations, suggesting the potential usefulness in practical

in vitro diagnostics, although real biological samples detection might be more difficult

than pure protein samples due to the presence of other interferences. The PEDOT

nanowire based sensing platform can be extended to other protein targets which their

DNA aptamers are available and is also applicable for label-free DNA detection.

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6 Functional PEDOT Nanobiointerface: Toward

in vivo Applications

6.1 Introduction

Among all the different research areas and healthcare applications in

bioengineering, the most critical aspect is the interface between materials and

biological systems.221 Regardless of the material is designed for use in drug delivery,

tissue engineering, implanted sensor devices or artificial organs, the biointerface

determines the biocompatibility of foreign objects and the efficiency of target

function. For synthetic polymeric materials to be used for in vivo medical applications,

one important problem is inadequate interaction between polymer and cells, leading to

foreign body reactions in vivo.222 Approaches to improve biocompatibility of these

materials include reduction of unspecific protein adsorption, enhancement of

adsorption of specific proteins and immobilizing of cell recognition moieties to obtain

controlled interaction between cells and synthetic substrates.223, 224

Conducting polymers have been widely used in the design of electrochemical

biosensors. Discussions in earlier chapters show that conductive polymers improve

the sensitivity and selectivity of electrochemical biosensors due to their electrical

conductivity or charge transport properties. It has also been demonstrated that

biomolecules, such as enzyme, antibody, DNA, and aptamer, can be immobilized onto

conducting polymers without any loss of activity. Moreover, conducting polymers can

be electrochemically grown on very small size electrode over defined areas. The

unique material properties of conducting polymers and compatibility with biological

molecules in aqueous buffer solutions open new opportunities for the use of

conducting polymer for a number of other biological applications,225 such as tissue

96

engineering scaffolds, neural probes, drug delivery devices and bio-actuators. It was

found that conducting polymers, via electrical stimulation, were able to modulate

cellular activities, including cell adhesion, migration, DNA synthesis and protein

secretion.161, 226 Therefore, many conducting polymer-based biomaterial studies are

centered around those which respond to electrical impulses, e.g. neuron,226-228 bone,229

muscle,230 and cardiac cells231 etc. The use of conducting polymers as an electrode

coating material for neural probes83, 232, 233 and as scaffolds234-236 to induce tissue

regeneration is under extensive exploration. Controlled surface morphology and

surface modification of these materials with biological moieties is desired to enhance

the biomaterials-tissue interface and to promote desired tissue responses. Although

PPy is extensively studied as a conducting polymer for biological applications, the

inherent weaknesses from backbone defects, as well as the instability to strong

biologically relevant reducing agents, such as dithiothreitol (DTT) and glutathione,

limit their commercial applications. PEDOTs are much more superior than PPy in

terms of stability, rendering them resistant to strong biological reducing agents.237

In this chapter, I will present some initial efforts on the development of

functional PEDOT nanobiointerfaces toward in vivo applications. The PEDOT

nanobiointerfaces have following characteristics: thin and smooth, composition-

tunable, capable of grafting with a variety of functional groups, fast deposition onto

selected electrode surfaces, amenable to large-scale manufacturing, conductive

enough to provide electrical stimuli when needed, and show low intrinsic cytotoxicity

and no inflammatory response upon implantation. Through proper design of side-

chain functional groups, adhesive (fouling) or non-adhesive (non-fouling) surfaces for

controlled adhesion of proteins and cells were achieved. Preliminary results show that

these PEDOT nanobiointerfaces is promising for applications such as cell patterning.

97

6.2 Experimental


Ethylenedioxythiophene (EDOT, Sigma-Aldrich), lithium perchlorate (LiClO4,

Fluka), sodium dodecyl sulfate (SDS, Alfa Aesar), 1-ethyl-3-[3-(dimethylamino)-

propyl]carbodiimide (EDC, Sigma-Aldrich), N-hydroxysulfosuccinimde (NHSS,

Sigma-alddrich), biotin (Sigma-Aldrich), Biotin N-hydroxysuccimide ester (biotin-

NHS, Pierce chemical), biotinamidohexanoyl-6-aminohexanoic acid N-

hydroxysuccimide ester (Biotin-LC-NHS, pierce chemical), bovine serum albumin

(BSA, Sigma-Aldrich) and streptavidin (Pierce chemical) were used as received.

Peptides GRGDSP and GRDGSP were ordered from GL Biochem (Shanghai).

Hydroxymethyl-functionalized EDOT (EDOT-OH), amino-functionalized EDOTs

(EDOT-C1-NH2 and EDOT-C6-NH2), carboxylic acid-functionalized EDOT (EDOT-

COOH), and triethylene glycol-functionalized EDOT (EDOT-EG3-OH) were

synthesized in our laboratory as described elsewhere in detail (refer to Figure 6-1 for

their corresponding chemical structures).185 Biotin-functionalized EDOTs were

synthesized from EDOT-C1-NH2 or EDOT-C6-NH2, through reaction with biotin-

NHS or biotin-LC-NHS. Three different biotin-functionalized EDOTs with different

linker chain (EDOT-C1-biotin, EDOT-C8-biotin, EDOT-C15-biotin) were synthesized

(Figure 6-2). A phosphate-buffered saline (PBS) consisting of 137 mM of NaCl, 2.7

mM of KCl, and 10 mM of phosphate buffer was used as supporting electrolyte

solution. Indium tin oxide 93 coated glass (Delta Technologies, Ltd.) were cut into

1cm × 1 cm square pieces and cleaned by standard procedure prior to use.

98

EDOT-C1-NH2 EDOT-C6-NH2 EDOT-EG3-OH

Figure 6-1: Chemical structures of functionalized EDOT monomers

Figure 6-2: Chemical structures of (A) EDOT-C1-biotin, (B) EDOT-C8-biotin, and (C) EDOT-C15-biotin.

6.2.2 Electropolymerization and Film Synthesis

PEDOT films from different EDOT monomers were electropolymerized on

ITO electrodes from 10 mM of EDOT aqueous solutions containing 0.1 M of LiClO4

99

as supporting electrolyte in the presence of 1 mM of HCl and 0.05 M of SDS by

cycling potentials from -0.6 to 1.1 V at a scan rate of 100 mV/s for 2-3 cycles. The

electropolymerization was also done under 1.0 V constant potential with a cut-off

charge density of 5 mC/cm2 for certain experiments. All the potential reported in this

chapter refer to Ag/AgCl. Multilayered PEDOT structures were made in a similar

manner using layer-by-layer growth from different monomer solutions.

6.2.3 Electrochemical Characterization

Cyclic voltammetric and potentiostatic experiments were performed with a

CHI electrochemical workstation in a three-electrode electrochemical cell with

platinum as the counter electrode and Ag/AgCl as the reference. Electrochemical

quartz crystal microbalance (EQCM) measurement was recorded with CHI400 time-

resolved EQCM using a crystal with a 7.995-MHz fundamental frequency, and an

evaporated gold electrode of a surface area of 0.196 cm2. In situ conductivity scan

was monitored with a PGSTAT30 Eco Chemie Autolab electrochemical instrument at

a scan rate of 1 mV/s in 0.1 M of LiClO4 solution. PEDOT was deposited on an

interdigitated microelectrode (IME) with gaps of 5 μm.

6.2.4 Polymer Film Analysis

The surface morphology of polymer films was observed with field emission

scanning electron microscopy (FESEM) and atomic force microscopy (AFM).

FESEM was carried out with JEOL JSM-7400 at a vacuum of 10-8 torr and an

accelerating voltage of 10 kV. AFM was performed in the tapping mode at room

temperature in air with BioScopeTM Digital Instruments. Rrms was obtained for a scan

range of 20 μm × 20 μm. Contact angle measurement was conducted with Contact

Angle Systems OCA (Dataphysics Instrument). 1 μL of water was introduced on the

surface of polymer films to form a droplet. Contact angle was measured using

100

software SCA20 for the OCA system. The measurement was repeated three times for

each polymer film. The chemical composition of the polymer surface was analyzed

using a VG ESCA 220i-XL imaging x-ray photoelectron spectroscopy (XPS)

equipped with an aluminum Kα source.

6.2.5 Protein Adsorption

Protein adsorption on PEDOT thin films was carried out using Q-sense 4 (Q-

sense AB, Sweden). The QCM-D, described in detail elsewhere, iii measures the

frequency change (Δf) and the energy dissipation change (ΔD) by periodically

switching off the driving power over the crystal and recording the decay of the

damped oscillation. The time constant of the decay is proportional to D and the period

of the decaying signal gives f. 5 mHz, AT-cut gold coated quartz crystals (Q-sense

AB, Sweden) were used as substrate. Measurements up to four harmonics

(fundamental frequency, 15, 25 and 35 MHz, corresponding to the overtones n=3, 5,

and 7, respectively) of the 5 MHz crystal were recorded. In this work we only present

the Δf and ΔD corresponding to the 3rd overtone. During the measurement, the

polymer coated crystal was mounted in a liquid chamber which was internally

maintained at 25 ± 0.1 ºC. Samples are introduced by a peristaltic pump at a flow rate

of 50 µL/min.

6.2.6 Cell Culture

NIH3T3 Cells, HepG2 cells and KB cells were cultured in complete culture

medium (DMEM for NIH3T3 and HepG2, RPMI for KB, standard culture medium

was supplemented with 10% Fetal Bovine Serum (FBS), 200 units/mL penicillin and

200 μg/mL streptomysin). Cells were maintained at 37 ºC in a humidified atmosphere

iii http//www.q-sense.com

http://www.q-sense.com

101

containing 5% CO2 for continuous proliferation. At confluence, cells were trypsinized,

washed and split for subsequent passage. All the cell lines were maintained below 20

passages. For cell adhesion experiments, harvested cells were resuspended in serum-

free medium at a concentration of 50,000 cells/mL.

Cell attachment experiment

The PEDOT film coated ITO pieces (substrates) were loaded into a 24-well

plate and sterilized for 1 hr in 70% ethanol before cell experiment. The substrates

were then washed with PBS buffer twice and serum-free medium once. 500 μL of cell

suspension was added into each well. After seeding, samples were kept in incubator.

Microscopic examination was done at different time point (1 h, 2 h, 3 h and

overnight). Non-adherent cells were removed by gently washing the wells three times

with PBS. Samples were visualized under Olympus inverted optical microscope IX71

using phase contrast mode. Images were taken using a CCD camera. After imaging,

complete medium was added into the sample well and the samples were put back in

incubator for proliferation.

Cell viability by MTT assay

NIH3T3 and HepG2 cells were seeded on different PEDOT film coated ITO

glass substrates and left for proliferation in the complete medium in incubator for 24 h,

48 h and 72 h. Same size uncoated microscope coverslip was used as blank control.

Change medium after 48 h. After incubation, the supernatant was discarded, and

replaced with 0.5 mL of serum-free medium for each well. 50 μL of MTT (5 mg/mL)

were added to each well and incubated for 4 h. The medium was discarded by

aspiration, and the formazon was dissolved with 500 μL of dimethyl sulfoxide

(DMSO). The absorbance was measured using plate reader at 550 nm. The cell

viability was calculated by normalizing with the blank control group.

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6.3 Results and Discussions

6.3.1 Synthesis and Characterization of Functional PEDOT Thin Films

The electropolymerization of PEDOT films in aqueous microemulsion was

monitored by EQCM in situ. Figure 6-3 shows a typical electropolymerization cyclic

voltammogram and the corresponding mass change. For microemulsion

polymerization of EDOT-OH, the weight gain in the first cycle (1.97 μg) was much

greater than that in the second cycle (1.33 μg) and the third cycle (1.27 μg). This was

most likely due to the difference in surface properties. In the initial cycle,

poly(EDOT-OH) was directly deposited onto Au surface, which provided a better

electron transfer interface. In subsequent cycles, poly(EDOT-OH) was deposited onto

the poly(EDOT-OH)-coated electrode, which has lower conductivity and hence

reduced efficiency in electron transfer, leading to a lower deposition rate. Similar

results were obtained for EDOT-COOH, mass increase during the first cycle was also

much higher than that during the subsequent cycles. However, compared to EDOT-

OH, smaller mass increase in the initial cycle for poly(EDOT-COOH) (1.84 μg) was

observed . The slower polymerization might due to the steric hindrance from the

carboxylic acid functional group. Took reported PEDOT density of 1.5 g/cm and

neglected the side-chain functional group, 1 µg of polymer would correspond to a

film of ~ 30 nm in thickness. PEDOT film with different thickness can be easily

obtained by simply controlling the number of cycles.

103

Figure 6-3: (a) Electropolymerization of 10 mM of EDOT-OH monomers in CH3CN (---) and in aqueous microemulsion containing 0.05 M of SDS and 1 mM of HCl (––) with 0.1 mM of LiClO4 as supporting electrolyte at a scan rate of 100 mV/s. (b) In situ QCM measured weight gain during the electropolymerization shown in (a).

It was noted that for EDOT-EG3-OH, we observed the dissolution of deposited

blue polymer. This is most likely due to the increased water solubility of both the

monomer and the polymer with the tethering hydrophilic ethylene glycol groups. The

solubility of the polymer was reduced when EDOT-EG3-OH was copolymerized with

another less soluble monomer. The existence of a minimum of 5% EDOT-OH in the

monomer mixture would greatly enhance polymer deposition. We kept the EDOT-

EG3-OH ratio at 90% for subsequent cell adhesion study.

Synthesis of PEDOT nanobiointerfaces with –OH, -COOH and -EG3-OH

surface functional groups have been demonstrated by direct electropolymerization

from the monomer precursors containing specific functional groups. However, when

1st cycle

2nd cycle

3rd cycle

1st cycle

First 3 cycles

1st cycle

104

we tried the same approach on biotin-functionalized EDOT (EDOT-Cx-biotin), we

found that the EDOT-Cx-biotin precursors can not be efficiently electropolymerized

into electroactive polymer alone (Figure 6-4). More studies show that EDOT-Cx-

biotin is able to copolymerize with EDOT-OH with the molar ratio less than 10%.

Figure 6-4: Electropolymerization of (A) 10 mM of EDOT-C8-biotin monomer, (B) 10 mM of EDOT-OH monomer with different ratio of EDOT-C8-biotin, and (C) 10 mM of EDOT-OH monomer with 10% of EDOT-C8-biotin, in aqueous microemulsion containing 0.05 M of SDS and 1 mM of HCl with 0.1 mM of LiClO4 as supporting electrolyte at a scan rate of 100 mV/s.

It has been reported that the attached functional groups might have direct or

indirect electronic and steric effects on the electropolymerization process and the

electronic properties of the resulting polymer,238 despite the apparent simplicity of

direct synthesis of functional conjugated polymers from electropolymerization of its

precursor derivatives with covalently attached functional groups. Initially we

suspected it is due to the steric effect of the biotin. However, the problem remains for

EDOT-Cx-biotin even when the length of the spacer is over 30 Å. For steric effects,

inserting a linker of appropriate length or reducing the number of active sites usually

helped to solve the problem by preventing possible distortion of the π-conjugated

backbone.239, 240 Because of our findings, we speculated that the biotin group on the

EDOT side chain directly interacts with the π-conjugated system. Inductive and/or

mesomeric effects modify the reactivity of the cation radical and thus affect the

105

efficiency of the electropolymerization process. This was supported by a simple

molecular modeling. Simulation results of EDOT-C1-biotin and another model

molecule EDOT-C12 was shown in Figure 6-5. We found that the biotin unit prefers to

fold back and maximize its electron overlay with that of the thiophene ring. In this

configuration, lower minimum energy is achieved by π-π stacking. This is opposed to

the case of EDOT with long alkane side chain, where extended structure has lower

minimum energy. In view of the difficulty to get biotin-functionalized PEDOT thin

film from direct electropolymerization of EDOT-Cx-biotin monomer, post-

polymerization biotinylation was adopted in our study. We will describe the post-

polymerization biotinylation in detail later in section 6.3.5.

SEM image showed that PEDOT film prepared from aqueous microemulsion

was homogeneous and smooth, not like those prepared from the organic solution

(Figure 6-6a). AFM image further confirmed the nanoscale uniformity of the polymer

film, showing an average Rrms of < 5 nm (Figure 6-6b).

Figure 6-5: Molecular modeling result of EDOT-C1-biotin and EDOT-C12.

106

Figure 6-6: SEM (a) and AFM (b) image of poly(EDOT-COOH) film prepared from aqueous microemulsion.

Hydrophobic/hydrophilic property of electropolymerized PEDOT films was

also studied by measuring their water contact angle. The water contact angle

decreased from 80.1° for EDOT to 60.8° and 40.2° for EDOT-OH and EDOT-COOH,

respectively. This indicated that the side-chain functional groups on EDOT monomers

affect the surface properties of the polymer film. These results also suggest that the

functional groups were available on the surface instead of being embedded in the

PEDOT backbone. Therefore, by changing the species and compositions of monomers

in the microemulsion, the surface properties of the resulting PEDOT films could be

tailored precisely. The functional groups on the film surface could be further applied

towards probe conjugation and biomolecule immobilization.

6.3.2 Biocompatibility of Functional PEDOT Thin Films

Figure 6-7 shows the biocompatibility of PEDOT, poly(EDOT-OH) and

poly(EDOT-COOH) films prepared from microemulsion electropolymerization (50

mM of SDS, 1 mM of HCl and 10 mM of monomers). Cell viability after 3 days of

incubation in the presence of these films was > 90%, which was higher than that of

the control (ITO glass substrate without PEDOT coating). These results clearly

indicated the low intrinsic cytotoxicity of the PEDOT films, which open the way for

further cell-related studies.

a b

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Figure 6-7: Viability of NIH3T3 (gray) and HepG2 (black) cells in the presence of different PEDOT film coated ITO glass substrate.

6.3.3 Adhesive and Non-adhesive PEDOT Nanobiointerfaces

Controlling the interface between cells and solid substrates is important for

cell-based biosensors and biochips225. For a better understanding of the interaction

between cells and functionalized PEDOT substrates, we have studied the cell

adhesion on different functional PEDOT thin films. Cells were allowed to attach to

PEDOT surfaces for 2 h under serum-free conditions. After 2 h of incubation,

unattached and loosely attached cells were gently removed by washing three times

with PBS. Substrates were then further incubated in full medium with 10% FBS

overnight. As shown in Figure 6-8, we found that both NIH3T3 and KB cells

selectively attached to polyEDOT and poly(EDOT-OH) surfaces. PEDOT surface

electropolymerized from a mixture of 10% EDOT-OH and 90% EDOT-EG3-OH were

cell-resistant as-expected. However, for PEDOT-COOH surface, cell-adhesive or cell-

resistant is depending on the cell type. Figure 6-9 further shows that cells attached

well on poly(EDOT-OH) surface, and started to spread and proliferate after further

incubation in full culture medium.

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Figure 6-8: Adhesion of NIH3T3 (above) and KB (below) cells on PEDOT nanobiointerfaces of different monomer compositions: (a) EDOT, (b) EDOT-OH, (c) EDOT-COOH, and (d) EDOT-EG3-OH/EDOT-OH, molar ratio=9:1.

Figure 6-9: Attachment and proliferation of seeded NIH3T3 cells on poly(EDOT-OH) biointerface after (a) 2 h, (b) 15 h, and (c) 39 h of incubation in full medium.

Based on these results, we further investigated the possibility of engineering

the surface “cell-adhesiveness” or “cell-resistance” through layer-by-layer deposition.

Multilayer PEDOT structures were constructed by electropolymerization of

alternative layers of poly(EDOT-EG3-OH)-co-(EDOT-OH) and pure poly(EDOT-OH)

from their respective monomer solutions (Figure 6-10). Similarly, cell adhesion

experiments showed that cells were selectively attached onto the poly(EDOT-OH)

surface, and would resist attachment onto the poly(EDOT-EG3-OH)-co-poly(EDOT-

OH) surface. In the case of multilayer structures, the cell-adhesion properties of the

device were controlled only by the top surface layer.

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Figure 6-10: (a) Legend of monomer composition of layered PEDOT nanobiointerfaces. (b)–(g) Controlled cell adhesion from alternating layer-by-layer PEDOT nanobiointerface deposition with adhesive and non-adhesive properties.

The formation of alternate layers was confirmed by the change in water

contact angles. The advancing water contact angle decreased from 85º (ITO surface)

to 48º when the first layer of poly(EDOT-EG3-OH)-co-poly(EDOT-OH) was

deposited due to the hydrophilic nature of the triethylene glycol side chains (Figure 6-

11). It then increased to 62º when a second layer of poly(EDOT-OH) was formed on

top of the first layer. The water contact angle decreased to 50º due to the formation of

the more hydrophilic poly(EDOT-EG3-OH)-co-poly(EDOT-OH) third layer. This

confirmed that the surface property of PEDOT surfaces could be tuned through layer-

by-layer electropolymerization.

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Figure 6-11: Contact angles of layer-by-layer PEDOT nanobiointerfaces deposition. The color legends for the composition of PEDOT nanobiointerfaces are as shown in Figure 6-10.

6.3.4 Controlled Cell Patterning

The non-adhesive PEDOT biointerfaces were also applied towards cell

patterning. As shown in Figure 6-12, we first covered the ITO-coated glass slides with

a non-adhesive poly(EDOT-EG3-OH)-co-poly(EDOT-OH) film. The polymer surface

was then patterned with a PDMS mask, and a 2nd layer of adhesive poly(EDOT-OH)

film was electropolymerized only at the exposed area. Cells were seeded on the

substrate surface as previously described. After washing away non-adherent cells,

cells were only observed on the poly(EDOT-OH)-covered regions, and not on

surrounding poly(EDOT-EG3-OH)-co-poly(EDOT-OH) surface, reinforcing previous

observation in section 6.3.3. This work, as a proof-of-concept, demonstrated the

successful engineering and patterning of the PEDOT nanobiointerface with cell-

adhesive and cell-resistant properties through simple layer-by-layer

electropolymerization. It would be very useful for cell patterning and chip-based

biosensor applications where controlled adhesion is critical.

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Figure 6-12: Controlled cell adhesion on patterned poly(EDOT-OH) on poly(EDOT-EG3-OH)-co-poly(EDOT-OH) surfaces. (a) Top and side views of the device patterned by selective electropolymerization using PDMS mask. Magnified microscopic images of selective cell adhesion on the patterned surface were shown in (b) and (c).

6.3.5 Biotin-functionalized PEDOT Nanobiointerface

As mentioned earlier, functionalized PEDOT film can be easily formed on

various electrode surfaces by direct electrochemical polymerization, yielding polymer

films with comparable conductivity and stability as non-functionalized PEDOT. As

demonstrated in previous sections, we have successfully synthesized PEDOT films

with various surface functional groups including –OH, -COOH, -EG3-OH by

polymerization direct from specific monomer solution. The functional groups attached

to the polymer backbone played an important role for further biomolecule

binding/repelling.

Post-polymerization biotinylation

Biotin grafted conducting polymer represents one of the most common

platforms for CP-based biological application. Biotin/avidin assembly is considered as

the important building block for CP-based biosensors.240-242 In view of the difficulty

of obtaining biotin-functionalized PEDOT through direct electropolymerization from

EDOT-Cx-biotin monomer, we adopted post-polymerization biotinylation as an

alternative solution to introduce biotin group to the PEDOT nanobiointerface. For

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poly(EDOT-OH) film, biotin was grafted to the polymer using 4-

(dimethylamino)pyridine (DMAP)-catalyzed coupling in CH3CN.243 For poly(EDOT-

COOH) film, biotin was grafted to the polymer using well-known activated amine

coupling reaction (scheme 6-1). Biotin-functionalized PEDOTs show no change in

their electroactivity and intrinsic conductivity.

Scheme 6-1: (a) Post-polymerization biotinylation of poly(EDOT-OH) films, (b) Post-polymerization biotinylation of poly(EDOT-COOH) films.

The biotinylation reactions were confirmed by X-ray photoelectron

spectroscopy (XPS) analysis. We observe an obvious peak around 400 eV,

corresponding to the emission of 1s level of nitrogen atoms for biotinylated film

(Figure 6-13). In comparison, no peak was seen at similar position for original

poly(EDOT-OH) film.

(a)

(b)

(i) EDC, sulfo-NHS, biotinamindopetylamine

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Figure 6-13: XPS analysis of poly(EDOT-OH) before (dashed line) and after biotinylation reaction (solid line). The inset shows the amplified region corresponding to N1s emission.

Protein adsorption on biotin-functionalized PEDOT nanobiointerfaces

Protein adsorption on biotin-functionalized PEDOT nanobiointerfaces was

studied by quartz crystal microbalance (QCM). Bovine serum albumin was used as a

representative protein for non-specific binding study and streptavidin for specific

protein binding study. Figure 6-14 shows the binding processes on poly(EDOT-OH)-

co-(EDOT-COOH) thin film before and after biotin functionalization. For QCM

measurement, the decrease in frequency (ΔF) of a crystal is proportion to the mass

increase on surface based on the Sauerbrey relation. As shown in Figure 6-14A,

immediately after exposing to BSA solution, there is a rapid and drastic decrease in

frequency for both films, indicating strong non-specific BSA adsorption. A steady-

state value was reached within a few minutes. After stabilizing for about 15 minutes,

streptavidin solution (0.1 mg/mL) was introduced to the flow system. For biotin-

functionalized PEDOT film, we observe another rapid decrease in frequency. Since

the polymer surface was already passivated by BSA, this decrease in frequency was

therefore attributed to the specific binding of streptavidin molecules to the polymer

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surface due to biotin-avidin interaction. It is interesting to note that streptavidin layer

shows more compact and rigid structure than the flexible, water-rich nature of non-

specific adsorbed BSA layer, as indicated by the decrease of dissipation signal

recorded at the same time (Figure 6-14B). It has been previously reported that

streptavidin film assembled on the biotin-containing surface is highly rigid with a

well-ordered structure while the streptavidin film formed through amine coupling is

highly dissipative and less structured.244 Therefore, we concluded that the post-

polymerization biotinylation leads to a biotin-functionalized conducting surface

suitable for further streptavidin binding. In a similar experiment, non-biotin

functionalized PEDOT film was tested as a control. Similar frequency decrease for

BSA adsorption and negligible frequency decrease for streptavidin suggest that the

film was unable to bind streptavidin due to the absence of the biotin moiety on its

surface. This further confirms the preceding frequency decrease is due to specific

interaction between streptavidin and surface biotin group. Similar phenomena were

observed for poly(EDOT-OH) film before and after biotinylation.

Upon further analysing the QCM results, for biotin-functionalized PEDOT

film, the 18 Hz frequency decrease due to specific streptavidin binding corresponds to

an increase in mass of 332 ng/cm2 (3.8×1012 molecules/cm2). Considering the

molecular footprint of streptavidin as a 5.2×3.65 nm rectangle,245 the maximum

coverage corresponding to a close-packed monolayer of streptavidin was estimated at

5.3×1012 molecules/cm2. Therefore a submonolayer of streptavidin was immobilized

on the electrode surface through specific biotin binding.

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Figure 6-14: QCM studies of binding of BSA and streptavidin on functional PEDOT surfaces () non-biotin functionalized PEDOT () biotin-functionalized PEDOT.

6.3.6 Peptide-functionalized PEDOT Nanobiointerface

Most of the mammalian cells are anchorage-dependent and thus must attach

and extend on a surface in order to proliferate. Interaction between cells and their

culture substrates are critical for cell growth and functions. RGD peptides have been

known to promote cell adhesion through the interaction with ECM proteins

(fibronectin, integrin, etc) in the serum or secreted by cells on the cell membrane.246,

247 In the mid 1980s, Pierschbacher and Ruoslahti found that many extracelluar matrix

(ECM) proteins contains the tripeptide RGD and this short peptide is an important

ligand for cell adhesion.247 It was subsequently found that cells employ a large family

of transmembrane proteins called integrins to recognize this and other ligands of the

ECM.248, 249 These findings were followed by many reports that demonstrated

substrates modified with the RGD peptide alone supported the adhesion and spreading

of mammalian cells.

Numerous materials have been RGD functionalized for academic study or

medical applications.250-259 Since many polymers do not have functional groups on

their surface, blending, co-polymerization, chemical and physical treatment of the

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polymers have been used to introduce functional groups for further RGD peptides

attachment. Stable linking of RGD peptides to a surface is essential to promote strong

cell adhesion.256 The formation of focal adhesions only occurs if the ligands withstand

the cells contractile forces. In order to provide stable linking, RGD peptides should be

covalently attached to the polymer via functional groups. Human umbilical vascular

endothelial cells (HUVECs) cultured on poly(1-(2-carboxyethyl) pyrrole) (PPyCOOH)

films modified with the RGD motif demonstrated improved attachment and

spreading.260 In addition, to achieve specific ligand-cell interaction, the substrate itself

must be able to prevent cells from remodeling the interface by resisting the deposition

of additional adhesive proteins expressed by the attached cells. Roberts and

Whitesides demonstrated a well-controlled surface that promote cell attachment by

the specific interaction of RGD with cell surface integrin receptors and resist

significant deposition of cell-derived matrix components, using a mixed self-

assembled monolayer (SAM) containing both EG6OGRGD and EG3OH terminated

alkanethiolates.261 Since it is relatively easier for us to generate functional PEDOT

thin film with different functional groups, we would like to explore cell attachment on

dual functionalized PEDOT surface containing both RGD peptide and EG3OH group.

In this part of work, a cell adhesive peptide GRGDSP was covalently

immobilized onto the PEDOT surface containing dual side chain functional groups, a

carboxylic acid group and an EG3OH group. The RGD-grafted PEDOT surface was

characterized by contact angle measurement, and specific cell attachment on RGD-

modified PEDOT surface was then studied.

Post-polymerization peptide conjugation

A cell adhesive peptide GRGDSP and a control peptide GRDGSP were

covalently immobilized onto a dual functionalized PEDOT surface. First,

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poly(EDOT-EG3OH)-co-poly(EDOT-COOH) was electropolymerized on ITO coated

glass substrate as described earlier in section 6.2.2, from a mixed monomer solution

containing EDOT-EG3OH and EDOT-COOH. The percentage of the side chain

carboxylic acid group in the final copolymer was controlled by varying the monomer

molar ratio between EDOT-EG3OH and EDOT-COOH. After electropolymerization,

a 100 µL of activation solution containing 100 mM EDC and 40 mM sulfo-NHS was

added onto the PEDOT film. After 20-min reaction, the carboxylic acid groups on the

PEDOT surface were converted into a stable intermediate form of sulfosuccinimide

ester, which will then readily react with the primary amine on the N terminus of the

peptide. After overnight reaction, peptides were chemically grafted onto the PEDOT

backbone. The peptide-grafted PEDOT surface was characterized by contact angle

measurement.

Figure 6-15: Water contact angle of poly(EDOT-EG3-OH)-co-(EDOT-COOH) film before and after peptide conjugation. Monomer molar ratio of EDOT-EG3-OH: EDOT-COOH = 8:2

As illustrated in Figure 6-15, water contact angle was around 50° for

unmodified copolymer film, and it dropped to around 40° after peptide conjugation.

Due to the hydrophilic nature of peptide, conjugation of peptide to the surface will

118

render a more hydrophilic surface, and thus a decrease in water contact angle.

Therefore, the decrease of water contact angle indicates a successful peptide

conjugation.

Cell attachment study

NIH3T3 mouse fibroblast cells were used in this cell attachment study. Cells

were cultured in Dulbecco’s Modified Eagle’s medium (DMEM, GIBCO)

supplemented with 10% fetal bovine serum (FBS), 200 units/mL penicillin, and 200

µg/mL streptomysin. Cultures were maintained at 37ºC in a humidified atmosphere

containing 5% CO2. Near confluence, cells were dissociated with trypsin-EDTA

solution, washed and resuspended in serum-free DMEM medium. A fixed number of

cells (15000 cells/cm2) were plated onto the PEDOT/ITO substrates with or without

peptide modification. PEDOT/ITO substrates were pre-sterilized by immersing in

70% ethanol for 1 hr. Cell attachment is assessed after an incubation of 3 h in a

humidified atmosphere containing 5% CO2. Non-adhered cells are removed by gently

washing three times with 1x PBS. Figure 6-16 shows the typical microscope image of

PEDOT surface with or without RGD peptide.

Figure 6-16: Controlled cell attachment (NIH3T3) on (a) RGD-functionalized, (b) carboxylic acid-functionalized, and (c) RDG-functionalized PEDOT surfaces. Functional group density was controlled at 10% while the remaining 90% are poly(EDOT-EG3-OH).

Cells were found to preferably attach to PEDOT surfaces modified with RGD

peptide, while little cells were attached to the surfaces with the control peptide as well

119

as surfaces without any peptide. This result indicated that RGD peptide is critical for

cell attachment through specific ligand-cell interaction. We demonstrated earlier on in

section 6.3.3, poly(EDOT-EG3OH) is considering an “inert”, non-adsorbing interface

for cell attachment. Therefore, in the absence of the RGD peptide, there is no

anchoring point for cells attached to since the background EG3OH groups resist the

deposition of extracellular matrix by the cells. A few cells attached to poly(EDOT-

EG3OH)-co-poly(EDOT-COOH) was through non-specific adsorption to the EDOT-

COOH fraction.

Effect of RGD surface density

Figure 6-17: Cell adhesion on RGD-modified PEDOT surfaces with different RGD density (a) 50%, (b) 20%, (c) 10%, (d) 5%, and (e) 1%. The effect of RGD density on cell adhesion was plotted on the right bottom. The y-axis is the number of attached cells per cm2.

The effect of RGD surface density on cell attachment was evaluated. The

RGD surface density can be easily controlled by varying the percentage of EDOT-

COOH in the copolymer. It has been reported that the number of attached cells, cell

shape and cell proliferation are related to RGD surface density.256 Figure 6-17 shows

that cells were bound to all the surfaces across the percentage range we studied, with

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maximum cell adhesion achieved at 10 ~ 20% fraction of RGD functional group.

More studies on cell spreading and proliferation are necessary to fully understand the

impact of RGD density on cell behavior.

6.4 Conclusions

We have demonstrated the synthesis of various functionalized PEDOT thin

films that is ultrasmooth and composition tunable. Through proper design of side-

chain functional group, adhesive (fouling) or non-adhesive (non-fouling) surfaces for

controlled adhesion of proteins and cells were achieved. Particularly, biotin-

functionalized PEDOT surface and peptide-functionalized PEDOT surface were

achieved through direct polymerization from mixed monomer solution and facile

post-polymerization functionalization. Biotin-functionalized PEDOT surface that

shows specific recognition towards streptavidin represents key building block for the

development of new classes of functional conjugated polymers and should lead to

interesting developments in the field of electrochemical biosensors. Whereas, RGD

peptide-functionalized PEDOT surface that promotes specific cell adhesion with

minimum non-specific adsorption will be useful for cell-based applications.

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7 Conclusions and Outlook

There is an increasing need for portable, integrated biosensors that are easy-to-

use and can even be packaged as home-test kits. These bioanalytical devices that can

accurately detect target analytes are important to the field of medical diagnostics, food

testing and drug screening. However, many existing technologies, such as ELISA and

microarray platforms, are costly, complicated and usually requires trained personnel.

Electrochemical-based label-free biosensors on the other hand, offer fast and sensitive

detection at a much lower cost due to its simple protocols and compact size. In this

thesis, we have presented the results from studies that will bring us closer to the

realization of such a device.

We have demonstrated two different approaches for the development of label-

free electrochemical biosensors for DNA/protein detection. Ruthenium-complexed

electroactive DNA threading intercalators and aptamer-modified PEDOT nanowires

were used as signal reporters for the detection of specific analytes. The ruthenium

intercalators can selectively bind ds-DNA and generate easily distinguishable

electrochemical signals. The second approach utilizes conductive polymer PEDOT

nanowires directly synthesized across an electrode junction. As a specific analyte

binds to the capture probe on the PEDOT nanowires, a detectable change in electrical

signal can again be easily detected.

In addition to our studies on biosensing, we further examined cell-based

applications of PEDOT thin films. In these studies, we showed that PEDOT surfaces

can be specifically functionalized for specific protein adsorption and also controlled

cell attachment. We envision that PEDOT surfaces with nano-features (e.g. fibers,

tubes, particles) can be used on similar applications and may give rise to new and

122

exciting observations not seen before. This study may also provide useful information

for future in vivo sensing.

With the development of new materials, detection strategies, and advances in

microfabrication and instrumentation, a highly integrated device that incorporates

sample preparation, liquid handling, detection and signal processing modules, and can

perform highly efficient, simultaneous analysis of biologically important molecules is

just around the corner. This may eventually become the mainstay of future

bioanalytical systems and will have a groundbreaking impact on the future of personal

healthcare, food, and pharmaceutical industries. We hope that these studies will

contribute to the concept of a micro-total-analysis system.

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label-free electrochemical dna and protein …finally, we evaluated functional pedot thin films as...

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