LABEL-FREE ELECTROCHEMICAL DNA AND PROTEIN …Finally, we evaluated functional PEDOT thin films as...
Transcript of 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
3.2.1 Materials and Reagents...........................................................................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 Results and Discussion.......................................................................................50
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
4.2.1 Materials and Reagents...........................................................................66
4.2.2 Chemical Polymerization .......................................................................67
4.2.3 Electrochemical Polymerization ............................................................68
4.2.4 Characterization ......................................................................................68
4.3 Results and Discussion.......................................................................................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
5.2.1 Materials and Reagents...........................................................................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 Results and Discussion.......................................................................................85
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
6.2.1 Materials and Reagents...........................................................................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
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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
3.2.1 Materials and Reagents
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 Results and Discussion
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
4.2.1 Materials and Reagents
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 Results and Discussion
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
5.2.1 Materials and Reagents
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 Results and Discussion
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
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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
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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.
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6.2 Experimental
6.2.1 Materials and Reagents
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
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.
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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.
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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
120
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.
123
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